Department of Cell Biology and Physiology, University of Pittsburgh School of Medicine, Pittsburgh, Pennsylvania 15261
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ABSTRACT |
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cystic fibrosis transmembrane conductance regulator; membrane traffic; chloride channel; protein secretion
The subject of this review is CFTR trafficking in the distal portion of the protein secretory pathway, which lies between the trans-Golgi network and the plasma membrane. Historically, studies of this pathway were motivated by the question of whether it would be possible to explain all of the manifestations of CF disease on the basis of a missing anion conductance at the apical membranes of epithelial cells. Accordingly, functional roles for CFTR other than its primary role as an apical membrane anion conductance pathway have been proposed to contribute to CF pathology. These include disease-associated changes in regulated protein secretion (65), the chemistry of macromolecular secretions (22, 33, 90), and the properties of the airway microenvironment that lead to bacterial colonization of CF airways (89).
Although the attention of numerous investigators has focused on CFTR trafficking through the distal protein secretory pathway as a possible contributor to the complexity of CF disease, a consensus regarding the importance of intracellular CFTR function and the process of regulated CFTR trafficking within this subcellular domain has yet to emerge. As an integral membrane protein, CFTR will reside at least transiently in all compartments of the protein secretory pathway as it migrates toward the cell surface. Thus one can expect to find CFTR in membranes of the endoplasmic reticulum (ER), Golgi, and endosomal and lysosomal compartments of epithelial cells (Fig. 1). Although there is no question that CFTR channels function as a regulated conductance pathway when they are resident in the apical membrane, numerous studies (see CFTR LOCALIZATION IN CELLS) have also indicated that a significant quantity of mature (i.e., post-Golgi) CFTR is contained within intracellular compartments, that CFTR within these compartments is functional (9, 78), and that cAMP stimulation can modulate the distribution of CFTR between these compartments and the plasma membrane (see REGULATED CFTR TRAFFIC). Therefore, the issue is not whether apical Cl conductance is due to regulation of the gating of membrane-resident CFTR channels or whether it results from the insertion of CFTR channels into the membrane, because both processes occur. We want to know to what extent these forms of CFTR regulation are linked mechanistically and how trafficking of CFTR contributes to the overall conductance response.
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This area of CFTR cell biology has been among the most contentious and unresolved subjects in CF research, and it continues to be so. In a relatively recent review of this subject, Bradbury (10) argued that disparate conclusions regarding the regulation of the distribution of CFTR between plasma membrane and intracellular compartments might be reconciled if regulated CFTR trafficking was a property specific to epithelial cells, i.e., studies showing a lack of CFTR trafficking were generally performed in fibroblasts. Nevertheless, four years after the Bradbury review, the literature has not been clarified. Accordingly, the task of bringing some synthesis to this conflicting data set is not an easy one. Yet, we feel that it is possible to do so and that the attempt should add to our understanding of the control of plasma membrane CFTR density, a critical issue in CF disease.
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CFTR LOCALIZATION IN CELLS |
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Location of mature CFTR (band C). From a biochemical standpoint,
the prevailing view is that core-glycosylated, immature (band B) CFTR is
localized in the ER, whereas complex glycosylated, mature (band C) CFTR is
present primarily in compartments distal to the Golgi, including the apical
membrane. In contrast with many heterologous expression systems, relatively
little band B is observed by immunoblot in most epithelial cells expressing
CFTR endogenously at steady state. For example, the ratio of band C to B in
Calu-3 airway cells is 10
(100). This indicates that
most of the CFTR expressed endogenously in epithelial cells is located in
distal compartments of the protein secretory pathway. The first reported
studies of CFTR localization, performed in COS-7 cells transiently expressing
CFTR, showed a weak, diffuse plasma membrane staining pattern. Most of the
signal was intracellular, probably due to protein overexpression
(23). In polarized epithelial
cells that express CFTR endogenously (T84, HT29, Caco-2, and airway
epithelia), this signal was located unambiguously at the apical membrane
domain (27,
28). No staining was detected
in the basolateral membrane, and little perinuclear staining of CFTR was
observed in epithelial monolayers, implying that little CFTR is present in the
ER at steady state (consistent with the low levels of band B). The studies of
Denning et al. (27)
highlighted the importance of localizing CFTR within epithelial cells that
express the protein endogenously. In the same study, a mostly intracellular
location of exogenously expressed CFTR was found in non-polar HeLa and 3T3
cells.
Biochemical methods. The central question concerning the issue of
regulated CFTR trafficking is whether the protein that contributes to the
plasma membrane anion conductance is present strictly in the apical plasma
membranes of epithelial cells or whether it is present also in a subapical,
post-Golgi compartment from which it is mobilized to the cell surface during
stimulation. A related issue is whether CFTR is stable in the plasma membrane
or undergoes recycling to intracellular compartments. In this regard, a rapid
rate of CFTR endocytosis from the cell surface has been observed in both
epithelial and nonepithelial systems
(13,
61,
85). These studies used
biotinylation procedures to monitor the rate of CFTR internalization from the
cell surface. In T84 cells, it was estimated that 50% of cell surface
CFTR is retrieved within a few minutes
(85); in Chinese hamster ovary
(CHO) cells the internalization rate was
5% per minute
(61). These rates of
endocytosis are not different from those of constitutively recycling
receptors, such as those for transferrin or LDL. These findings, together with
the estimated 24- to 48-h half-life of mature, plasma membrane CFTR
(7,
41), indicate that virtually
all of the CFTR that is internalized by endocytosis must be returned to the
plasma membrane by a recycling/exocytic pathway. The presence of this rapid
CFTR recycling pathway is a minimal requirement for a regulated
insertion-retrieval model that controls cellular CFTR distribution. Thus the
question should not be whether CFTR undergoes trafficking at the plasma
membrane but whether this process contributes to a change in the distribution
of CFTR between plasma membrane and intracellular compartments in response to
cAMP/PKA regulation.
In addition to these kinetic considerations, the presence of CFTR in endosomes under steady-state conditions has been established with the use of a combination of biochemical techniques and functional assays that rely on CFTR's anion channel activity. Bradbury et al. (12) resolved CFTR in the early stages of endosomal retrieval within clathrin-coated vesicles from epithelial cells. CFTR was demonstrated to be present in this compartment by immunoblot. After removal of the clathrin lattice by uncoatase and fusion of the resulting membranes with planar lipid bilayers, currents with biophysical and regulatory properties characteristic of CFTR were observed. Because clathrin removal was a necessary prerequisite for reconstitution, CFTR was definitely a component of this endocytic compartment. Additional studies have demonstrated that the rapid endocytosis of CFTR from the cell surface depends on clathrin-mediated pathways and that CFTR is not significantly internalized into caveolae (11, 85). Several transport and/or channel proteins have been found to internalize via clathrin-dependent endocytosis, including aquaporin-2 (AQP-2) (101), Na/H exchanger 3 (NHE3) (24), renal outer medulla K channels (ROMK) (114), and epithelial Na channels (ENaC) (96). Recent studies of the mechanism of CFTR endocytosis (see Functional studies of endocytosis) indicate a direct physical interaction of CFTR with the plasma membrane adapter complex, making CFTR a specific target of this endocytic machinery.
Other methods for demonstrating the presence of CFTR in endosomes have
relied on the ability of CFTR to provide a pathway for anion transport that
parallels the vacuolar H-ATPase. Lukacs et al.
(60) allowed CHO cells
expressing CFTR to take up the pH-sensitive dye FITC-dextran into endosomes,
where the rate of dissipation of a pH gradient across endosomal membranes was
stimulated by addition of a proton ionophore. In these intact cells, pH
gradient dissipation was limited by the counterion. Activation of cAMP
production elicited a twofold increase in the rate of pH change in cells
expressing CFTR but not in parental or mock-transfected cells. The results
were duplicated in microsomes isolated from these cells. Biwersi and Verkman
(9) used a similar method for
monitoring endosomal pH in CFTR-transfected 3T3 cells and in T84 cells
endogenously expressing CFTR. Treatment of the cells with forskolin before
microsome isolation produced an approximately twofold increase in the rate of
subsequently measured pH gradient dissipation but had no effect on microsomes
isolated from nontransfected or F508 CFTR-expressing 3T3 cells. These
data are also consistent with the presence of functional CFTR in endosomes
that, in principle, could contribute to CFTR recycling.
Morphological methods. Using immunofluorescence and confocal
microscopy for detection of cellular CFTR, investigators have generated data
that arrive at somewhat divergent conclusions with respect to the importance
of regulated CFTR trafficking. On one side of this issue is the study of
Lehrich et al. (55) in shark
rectal gland, a tissue that expresses CFTR at high levels endogenously. These
investigators used quantitative confocal microscopy to show that CFTR
immunofluorescence extended from the apical membrane into subapical,
supernuclear regions of the cell and that during stimulation with
secretagogues, the overall depth of this CFTR signal decreased by 50%.
Similar conclusions were reached when the histogram of CFTR fluorescence
intensity as a function of distance from the apical membrane was quantitated.
These cAMP agonist effects on CFTR distribution, imposed in perfused, intact
rectal glands before fixation, were reversible. Whether this shift in
fluorescence signal represents insertion of vesicles containing CFTR into the
apical membrane remains unknown; however, the findings are consistent with the
acute hormonal regulation of CFTR trafficking in an intact epithelial
tissue.
Similarly, Ameen et al. (1, 2) used immunofluorescence microscopy to quantify the cellular location of CFTR in rat small intestine and the influence of cAMP-dependent agonists on its distribution. Vasoactive intestinal peptide (VIP) elicited a bicarbonate-rich fluid secretory response in isolated perfused intestinal loops and elicited, within the same time frame, a redistribution of CFTR to the apical membrane domain. CFTR association with the apical membrane was quantified by colocalization of F-actin as a brush border marker. In duodenal villus cells, VIP elicited a reversible, threefold increase in brush border associated CFTR, and it redistributed CFTR within the cell apex to the brush border within 30 min. These studies were confirmed by immunoelectron microscopic methods (see Immunoelectron microscopy). The results provide support for a physiological role for cAMP-induced CFTR-containing membrane traffic in the regulation of the apical anion conductance in a native, endogenously expressing epithelium.
Other investigators have reached different conclusions with respect to regulated CFTR trafficking. Moyer et al. (70) expressed a GFP-CFTR fusion protein in MDCK type I epithelial cells and used quantitative confocal fluorescence microscopy and cell surface biotinylation to determine the effect of cAMP stimulation on apical membrane CFTR localization. Their findings showed that cAMP did not stimulate detectable GFP-CFTR translocation from intracellular compartments to the apical membrane, along an apical-to-basal GFP fluorescence gradient that extended throughout the cell. In these cells, cAMP did not regulate GFP-CFTR endocytosis, determined by apical biotinylation and subsequent immunoprecipitation. Likewise, disruption of microtubules with colchicine did not affect cAMP-stimulation of Cl secretion or the expression of GFP-CFTR in the apical membrane. Thus the authors concluded that cAMP stimulates CFTR-mediated Cl secretion in MDCK type I cells by activating only the channels that are resident in the apical membrane. A regulated CFTR trafficking event may be lacking in this cell type; however, it is also reasonable to ask whether the exogenous expression of CFTR driven by the CMV promoter leads to saturation of regulated trafficking pathways (see Functional studies of exocytosis).
A related study (58) involving measurements of cellular CFTR immunolocalization in endogenously expressing Calu-3 epithelia by fluorescence confocal microscopy detected no change in apical CFTR staining upon cAMP stimulation. Surface biotinylation experiments reported in the same study (58) also showed no stimulation-dependent increase in apical membrane CFTR. Two biotinylated proteins were detected by CFTR antibody precipitation following biotinylation; they were of equal intensity and 200220 kDa in molecular size. Yet, others have demonstrated by immunoblot that CFTR in Calu-3 cells shows the typical pattern of a predominant band C of 150180 kDa and relatively little band B; the latter is presumably intracellular (c.f., Refs. 18 and 100). Thus it is not clear that the biotin-labeled proteins identified in these experiments were CFTR.
It is possible that differences in the conclusions drawn from fluorescence
measurements in native and cultured epithelial cells stem from variations in
the distribution of CFTR in subapical compartments under nonstimulated
conditions. In the studies of native secretory tissues (shark rectal gland and
rat intestine), the apical fluorescence signal extended into the cell
sufficiently to detect a clear change in its distribution with stimulation. In
MDCK cells expressing GFP-CFTR
(70), there was an even deeper
distribution of expressed protein, but this may reflect its deposition in
nonphysiological compartments, as described earlier in this article. Given
quantitative limitations of GFP visualization, 7 molecules/pixel
(79), it may be necessary to
express CFTR at nonphysiological levels to obtain a CFTR signal. In Calu-3
cells (58), the CFTR signal
was much more densely localized at the apical membrane domain. Nevertheless,
it is reasonable to ask whether a band of apical fluorescence detected in
epithelial cross sections can resolve apical membrane CFTR from that present
in a nearby subapical recycling pool. Consideration of the point-spread
function of fluorescence intensity variation with distance suggests that
structures separated by >0.2 µm will not be resolved using light
microscopy. Thus immunofluorescence, even coupled with confocal microscopy,
may lack the ability to resolve fluorescence signals from membrane vesicles
having a diameter of 0.1 µm or less, which are components of the CFTR
recycling pathway. This vesicle size expectation is based on neurosecretory
and other regulated trafficking vesicles (see APPENDIX). For
example, intracellular GLUT-4 staining by thin-section immunoelectron
microscopy was localized to vesiculotubular structures 5070 nm in
diameter (62). Thus it is
possible that different conclusions at the light microscopic level may emerge
from different cellular distributions of intracellular CFTR trafficking
compartments and their constituents (see CONSEQUENCES OF REGULATED CFTR
TRAFFICKING).
Immunoelectron microscopy. The results of several studies using immunoelectron microscopic methods are consistent with a subapical population of post-Golgi CFTR (c.f. Refs. 26, 42, 86, and 107). Puschelle et al. (86) used immunogold labeling to localize CFTR in vesicles present beneath the plasma membrane of human airway epithelial cells. They also observed vesicles in the process of fusion or retrieval from the plasma membrane. Immunogold labeling of CFTR-expressing L cells and Sf9 cells also demonstrated the presence of CFTR beneath the plasma membrane as well as in the rough ER (26). Studies of the striated duct of rat submandibular gland cells by Webster et al. (107) showed immunogold labeling of CFTR along the apical membrane as well as in numerous subapical membrane vesicles. Their estimates of labeling density suggested that more CFTR is present in subapical intracellular compartments than in the plasma membrane under "resting" conditions. Some of the CFTR-labeled vesicles also stained with antibodies against the transferrin receptor and rab4, two endosomal markers, indicating that these vesicles are likely part of an endosomal/recycling pathway for CFTR.
The fluorescence measurements of Ameen et al. (1), performed in rat small intestine, were confirmed by immunoelectron microscopy. CFTR was identified in subapical vesicles in several intestinal cell types, including cells from the crypt, Brunner glands, and a subpopulation of villar cells where CFTR expression predominated (so-called CFTR high expressors, or CHE cells). cAMP stimulation elicited a two- to threefold increase in CFTR labeling of the apical microvilli of CHE cells in response to cholera toxin, providing evidence for regulated insertion of CFTR into the apical membranes of a native epithelium.
The study of Howard et al. (42) examined apical membrane domain CFTR localization as a function of CFTR expression level. In an earlier study (43), these investigators showed that CFTR located in the plasma membrane could be discriminated from intracellular CFTR by using nonpermeabilized MDCK cells expressing an extracellular epitope-tagged CFTR (where the flag, M2 epitope, was added to the 4th extracellular loop of CFTR, after amino acid 901). They found that cAMP stimulation increased the cell surface signal of CFTR two-to threefold in the steady state. In the more recent studies, they induced CFTR expression by using a recombinant adenovirus that encoded M2-901/CFTR. Virally expressed, FLAG-tagged CFTR was functional and could be detected on the apical surface of forskolin-stimulated, polarized MDCK (type II) cells by immunofluorescence performed on nonpermeabilized epithelial monolayers. At a low multiplicity of infection (MOI) (i.e., lower CFTR expression level), forskolin stimulated the insertion of M2-901/CFTR into the apical membrane, but at higher MOI and M2-901/CFTR expression levels, no agonist-dependent increase in surface expression could be detected. Immunoelectron microscopy confirmed the redistribution of CFTR to the apical membrane upon forskolin stimulation at the lower CFTR expression levels and demonstrated that the apically inserted CFTR originated from a population of subapical vesicles. Results similar to these have been obtained with the use of immunoelectron microscopy in Calu-3 cells endogenously expressing CFTR, where a two- to threefold increase in apical CFTR was observed in response to acute forskolin stimulation (Hug MJ and Frizzell RA, unpublished observation).
The observations of Howard et al. (42) may reconcile at least some of the prior conflicting reports regarding the effect of cAMP stimulation on CFTR trafficking. They show that a high level of CFTR expression, obtained by using strong promoters, for example (70), can saturate pathways for regulated CFTR trafficking, even in epithelial systems. Also in oocytes, the CFTR trafficking signal (cAMP-induced increase in membrane capacitance) saturated as the CFTR expression level was elevated by injection of more RNA (102). Inappropriate trafficking of AQP-2, in relation to the actions of dominant-negative AQP-2 mutants, has also been observed at high protein expression levels (51). Later in this article, we discuss candidate protein interactions that may mediate regulated CFTR trafficking at the apical membrane. If cells express different levels of the relevant traffic regulatory proteins, then it is likely that the contribution of regulated CFTR trafficking to the total anion conductance (as opposed to regulated gating of membrane resident CFTR) will vary in different epithelial or nonepithelial systems.
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REGULATED CFTR TRAFFIC |
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The first studies that implicated CFTR function in regulated endocytosis
were performed in colonic and pancreatic epithelial cell lines, with the
latter transduced to express CFTR using retrovirus vectors. Bradbury et al.
(13,
14) measured
detergent-sensitive horseradish peroxidase and FITC-dextran uptake to monitor
fluid phase endocytosis in T84 and CFPAC-1 cells. They found that cAMP
significantly inhibited fluid-phase endocytosis, and in pancreatic cells, this
depended on the expression of wild-type (wt) CFTR. No cAMP inhibition of
endocytic activity was observed in parental CFPAC-1 cells or those transduced
by empty vector. A similar 60% inhibition of endocytosis was observed
during cAMP stimulation of endogenous CFTR-expressing T84
(13) and 9HTEo-cells
(94). In T84 cells, PKC
stimulation did not affect either basal endocytosis or the inhibitory effect
elicited by cAMP stimulation.
Methods employing biotinylation of the cell surface have also demonstrated
that CFTR internalization is inhibited by cAMP/PKA. Prince et al.
(85) biotinylated carbohydrate
residues on the extracellular surface of T84 cells using NHS-biotin.
Subsequent immunoprecipitation and radioisotope labeling of CFTR with the use
of [32P]ATP and PKA was performed to distinguish between CFTR at
the cell surface and in intracellular compartments. These workers found that
apical membrane CFTR was rapidly internalized by T84 cells (half-time of
3 min), and they showed that cAMP decreased this internalization rate to
an extent similar to that found in prior studies of fluid-phase endocytosis
(
60% inhibition). They also showed that the internalization rate of G551D
CFTR was similar to that of the wild-type protein but that, interestingly, its
endocytosis was not affected by cAMP. This finding suggested that either the
channel activity of CFTR is related to its ability to be retrieved from the
cell surface or the altered structure of the G551D mutant precludes regulation
of both gating and trafficking. It was difficult to detect a change in the
total amount of CFTR at the cell surface using the method of Prince et al.
(85). Even at short times (1
min), only a 12% increase in apical membrane CFTR was detected. However, the
use of in vitro phosphorylation to detect CFTR after its biotinylation and
immunoprecipitation may be influenced by the existing phosphorylation status
of the protein, thus compromising surface CFTR detection. That is, if CFTR is
already phosphorylated by agonist, the subsequent CFTR signal would be reduced
because phosphorylation sites are already occupied. A method that avoids this
complication employs a second CFTR antibody to detect biotinylated CFTR, which
avoids the potential for back-phosphorylation to reduce the signal. This
approach also demonstrates a forskolin-induced inhibition of CFTR endocytosis,
a depletion of the protein from intracellular compartments, and a consequent
increase in plasma membrane CFTR
(61).
Studies documenting the presence of functional CFTR in endosomes have been extended to show a role for CFTR in postendocytic membrane trafficking. Biwersi et al. (8) implicated a role for CFTR in cAMP-stimulated endosome-endosome fusion. They labeled endosomes of CFTR-expressing and control 3T3 cells with fluorophores such that a fluorescence signal would be detected if endosomes from separately labeled cell populations fused with one another. These investigators asked whether the presence of CFTR in these endosomes, and its stimulation by cAMP, influenced the fusion process. Endosomes from nonstimulated CFTR-expressing cells fused with each other at a rate that did not differ from that observed in control cells not expressing CFTR. The endosomes isolated from CFTR-expressing cells that were stimulated by forskolin, however, showed a 2.6-fold increase in fusion rate. This was abolished by endosomal Cl depletion, suggesting that a CFTR-mediated Cl flux, or the conformational changes in CFTR associated with Cl conduction, somehow contribute to this process. These data are consistent with the concept that CFTR stimulation can alter membrane traffic in endosomal recycling pathways.
Recent findings have been able to establish certain structural features of
CFTR that govern its rapid removal from the cell surface. Studies by Prince et
al. (84) and Weixel and
Bradbury (109) have
demonstrated that the internalization of CFTR via clathrin-dependent
mechanisms depends on the presence of a YXX motif (where
is a bulky
hydrophobic residue) at the CFTR COOH terminus. Similar tyrosine-based
endocytic motifs are present on other proteins that are rapidly retrieved from
the cell surface (see Ref. 80
for review). In vivo cross-linking and in vitro pull-down assays showed that
CFTR binds to the plasma membrane endocytic adaptor complex AP-2. The CFTR
COOH terminus was able to bind AP-2 but did not bind the Golgi-specific
adaptor complex AP-1. Mutation of the tyrosine residue at position 1424 of
CFTR significantly reduced the ability of an isolated CFTR COOH terminus to
bind AP-2. The YDSI sequence of CFTR interacts specifically with the µ2
subunit of AP-2 (108), a site
implicated in adapter binding to similar endocytic motifs
(74). Protein binding studies
showed that the COOH terminus binds selectively to this adapter subunit. Cells
expressing either a dominant negative µ2 or a CFTR lacking the
tyrosine-based internalization motif at the COOH terminus (Y1424A) fail to
endocytose CFTR efficiently. These studies indicate that the interaction of
CFTR's COOH terminus with AP-2 guides it into the clathrin-mediated endocytic
pathway, as observed for many cell surface receptors. Although these studies
provide compelling evidence for specific protein interactions that mediate
CFTR retrieval at the cell surface, they do not yet provide an indication of
how this process may be regulated by cAMP/PKA. We have speculated on possible
mechanisms for regulated CFTR endocytosis (see MECHANISMS OF REGULATED
CFTR TRAFFICKING: PERPETRATOR OR BYSTANDER? and
Fig. 2).
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Functional studies of exocytosis. The studies of Bradbury et al. (13) on CFTR-dependent endocytosis also examined CFTR-dependent exocytic recycling to the cell surface. To monitor exocytosis, the plasma membranes were pulse-labeled with a biotinylated lectin (WGA), and after its removal from the bath, the cells were allowed to take up lectin into intracellular compartments. Remaining cell surface WGA was blocked with unlabeled avidin, and at various times the recycling of internalized WGA to the cell surface was assayed using Texas Red-labeled avidin. In cells expressing wt CFTR, recycling and exocytosis of internalized marker was increased approximately threefold by forskolin, whereas cAMP stimulation had no effect on WGA recycling in either the parental CFPAC-1 cell line or in empty vector controls. Importantly, these studies showed that cAMP-responsive recycling/exocytosis required CFTR expression.
Similar findings were obtained in human airway cell lines (94) where recycling/exocytosis was monitored as the release of previously internalized FITC-dextran (a fluid-phase marker of endocytosis) from human airway cells. Treatment of cells expressing wt CFTR with a cAMP analog elicited increased release of FITC-dextran, and this was accompanied by increases in membrane capacitance monitored during whole cell patch-clamp measurements. Again, trafficking was CFTR dependent; cAMP had no effect on membrane capacitance or recycling/exocytosis in airway cells derived from a CF patient. The investigators concluded that cAMP stimulates exocytosis and the CFTR Cl conductance of normal but not CF cells and that it does so by stimulating the delivery of CFTR channels from an intracellular pool to the plasma membrane.
Much of the effort to determine whether cAMP stimulates CFTR-dependent membrane trafficking has employed measurements of membrane capacitance in cells that express CFTR endogenously or cells in which CFTR expression was induced. These studies are summarized in Table 1, which tabulates the methods, conditions, and experimental results of these studies. As discussed below, the diversity of methods and conditions used may contribute to their variable outcome.
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Previous work in nonepithelial cell lines (e.g., CHO cells) failed to show CFTR- and cAMP-dependent membrane trafficking (29, 47), again suggesting that the machinery for regulated CFTR recycling may be expressed only in epithelial cells. However, studies of Xenopus oocytes generally have shown a good correlation between cAMP stimulation of CFTR currents and increases in membrane capacitance (81, 102, 105). In the studies of Takahashi et al. (102), the apparent changes in cell surface area with stimulation, determined from membrane capacitance measurements, were confirmed by using blinded measurements of membrane morphometry in stimulated and nonstimulated cells. In addition, recruitment of CFTR to the cell surface was demonstrated by using epitope-tagged M2-901/CFTR labeling of nonpermeabilized cells (81). Together, these findings are consistent with the exocytic insertion of CFTR into the oocyte plasma membrane during cAMP stimulation. The cell surface immunofluorescence studies of Peters et al. (81) suggested a low level of CFTR expression in the plasma membrane of nonstimulated oocytes (i.e., fluorescence levels were near background in the absence of stimulation). These data suggested that recruitment of CFTR to the cell surface is a significant component of CFTR current stimulation in this system. However, this conclusion assumes that the flag-CFTR labeling approach can detect low levels of cell surface CFTR. Although technically difficult, it would be optimal to determine a dose-response relation between cAMP-dependent cell surface CFTR labeling and some independent parameter such as the corresponding change in membrane capacitance. It is possible, for example, that some threshold level of CFTR residing in the plasma membrane cannot be detected by this method, underestimating levels of CFTR at the cell surface under basal conditions. However, this concern does not compromise the data, obtained from three different methods, consistent with stimulation-dependent recruitment of CFTR to the surface of these cells.
The electrophysiological aspects of these studies were confirmed by the
work of Weber et al. (105),
who used impedance analysis to assess membrane conductance and capacitance and
correlated the changes in these parameters induced by cAMP. In agreement with
prior reports (81,
102), cAMP stimulation of
CFTR-expressing oocytes evoked significant increases in both current and
capacitance that were not observed in noninjected or F508-expressing
cells. Injection of Rp-cAMP (a specific cAMP antagonist) abolished
the effects of the cAMP stimulation on both current and capacitance.
Interestingly, the less specific PKA inhibitors, KT5720 and H8, after
prolonged (overnight) exposure, primarily blocked the capacitance increases
evoked by cAMP while having less effect on the corresponding current
activation. However, these reagents have been shown to alter cytoskeletal
properties. For example, KT5720 induces microtubule rearrangement by
inhibition of mitogen-activated protein kinase
(76), and H8 has been shown to
alter cytoskeletal protein phosphorylation through inhibition of PKC
(63). It would be interesting
to explore these actions further, because both of these less selective kinase
inhibitors produce cytoskeletal disruption, which may account for their
inhibitory effects on membrane trafficking. Indeed, Weber et al.
(106) showed that microtubule
disruption markedly inhibits current and capacitance increases evoked by cAMP
in CFTR-expressing oocytes, and similar conclusions have been reached in
CFTR-expressing epithelial cells
(103).
Although two groups have independently demonstrated cAMP-dependent changes in membrane capacitance and other parameters indicative of regulated CFTR trafficking in Xenopus oocytes, using different analytic methods Liu et al. (57) arrived at a different conclusion. They employed a CFTR construct containing a cysteine substitution in the postulated CFTR conduction pathway to arrive at the conclusion that essentially all conductance activation by cAMP in oocytes was due to CFTR channels that are already present in the plasma membrane. This approach relied on changes in conductance induced by treatment of this CFTR mutant with cysteine-reactive methanethiosulfonate (MTS) reagents, whose effects have been attributed to altered charge shielding properties in the CFTR conduction pathway. When oocytes were treated briefly with 2-(trimethylammonium)ethyl methanethiosulfonate (MTSET; a positively charged modifier) for as little as 20 s before cAMP activation, the activated channels behaved as if their properties were already modified. Because MTSET has been demonstrated in other systems to be cell impermeant, the data suggest that the modified channels must have already been present at the cell surface. These findings are consistent with a lack of regulated CFTR redistribution in oocytes, but they need not negate the results of prior studies for two reasons. First, this study used CFTR expression levels (based on conductance) that are 510 times higher than the prior work in which capacitance measurements were made. As discussed earlier in this article, overexpression of CFTR can saturate pathways available for regulated trafficking. Consistent with this idea, Takahashi et al. (102) observed that increasing CFTR expression levels increased CFTR currents, but the corresponding capacitance changes plateaued, as if regulated trafficking pathways had saturated. Constitutive delivery of CFTR to the plasma membrane at high expression levels would limit one's ability to detect a population of trafficking channels. Second, Liu et al. (57) did not measure membrane capacitance, cell surface CFTR, or any other measure of CFTR trafficking. Nevertheless, there is not a facile explanation for differences between these carefully performed studies and those cited above. It should be determined, using MTS CFTR modification together with capacitance measurements or surface CFTR labeling, whether high CFTR expression levels would account for these results.
Two recent studies, performed in mammalian epithelial cells that endogenously express CFTR, have reported negative findings with respect to changes in membrane capacitance during cAMP stimulation. As an independent assay, both studies also employed FM 1-43 dye labeling in an attempt to monitor membrane addition to the plasma membrane that would result from exocytosis stimulated by cAMP. In the studies of Chen et al. (21), membrane capacitance was measured in Calu-3 airway cells by imposing alternating sinusoidal and square voltage waveforms and monitoring the resulting currents to assess membrane capacitance and conductance. The calculated capacitance changes induced by cAMP were insignificant. In addition, they found no increase in steady-state plasma membrane labeling by FM 1-43 when the cells were stimulated by cAMP during fluorescence measurements. Enhanced dye labeling (fluorescence intensity) would be expected if additional membrane were exposed at the cell surface as a result of exocytosis. Thus the authors concluded that cAMP does not stimulate CFTR currents by increasing CFTR trafficking to the plasma membrane but stimulates only membrane-resident CFTR channels in Calu-3 cells.
In a similar study, Chang et al. (20) used sinusoidal voltage-current phase-lag analysis to estimate cell capacitance and fluorescence measurements of FM 1-43 membrane dye labeling in colonic HT29-Cl.19A and airway 14HBEo-cells, both of which express CFTR endogenously. The outcome in the airway cell line was similar to that in study by Chen et al. (21), i.e., no detectable cAMP-induced changes in membrane capacitance or FM dye labeling. In HT29 cells, increases in membrane capacitance with cAMP stimulation occurred only when endocytosis was blocked with the use of a dynamin antibody. As the authors indicated, the absence of a change without dynamin inhibition may result from parallel increases in both exocytosis and endocytosis during stimulation, i.e., no net increase in membrane area.
The studies of Chen et al.
(21) and Chang et al.
(20), despite the
sophisticated methodologies employed, have two significant shortcomings that
preclude definitive conclusions regarding regulated CFTR trafficking in these
systems. First, both studies were performed at room temperature, a condition
that is known to impair regulated membrane traffic in a variety of systems.
For example, the release of amylase from pancreatic exocrine cells is
decreased 70% at 25°C compared with normal temperature. The release of
insulin from rat pancreatic islets is markedly inhibited at 25°C, with a
90% reduction in the membrane capacitance response to stimulation (the
Q10 was 11.6) (75,
87). In addition,
cAMP-regulated AQP-2 recycling in renal cells is blocked at 20°C
(38). Indeed, recent data
collected in this laboratory
(45) indicate that studies
performed at 37°C in Calu-3 cells using either sinusoidal waveform
analysis or current transient analysis show a 1 pF increase in membrane
capacitance (basal membrane capacitance averages 20 pF) in response to cAMP
stimulation that parallels the increase in membrane conductance. When
identical recordings were made at room temperature with 5 or 10 mM EGTA in the
pipette solution (conditions of the Chen and Chang studies), cAMP elicited no
significant increase in membrane capacitance. It is interesting that early
recordings of capacitance changes in airway cells
(94) showed cAMP-dependent
increases at 23°C (Table
1); however, these experiments employed low EGTA concentrations
(Ca buffering at normal resting levels) in the recording pipette. Thus the
recording conditions have not been consistent.
The assays of FM dye labeling (also performed at room temperature) were
conducted by using a protocol that is insensitive to small increases in
membrane area. The cells were labeled to steady-state intensity with FM dye,
and their fluorescence intensity was monitored during cAMP stimulation.
However, if the increase in plasma membrane area induced by cAMP is only
3% over the basal membrane area (suggested from capacitance measurements
at 37°C; Ref. 46), this
signal would not be detected during steady-state dye labeling. Rather,
subsequent to stimulation, it is necessary to wash out dye and agonist from
the membrane to minimize plasma membrane (background) staining. This permits
visualization of the dye retrieved by subsequent endocytosis of labeled plasma
membrane, assuming that the cell recovers to its original area following
reversal of stimulation. As recent studies in this laboratory performed at
37°C have demonstrated, measurements performed in this manner indicate
that FM 1-43 labeling is increased by cAMP stimulation in Calu-3
(45). The protocol used in the
studies of Chen et al. (21)
and Chang et al. (20) is
similar to methods employed to monitor secretory granule release in systems
that incorporate large amounts of membrane into the cell surface during
stimulation (e.g., pancreatic acinar cells), generally in response to
stimulation by a cellular calcium rise
(36). In these systems, the
addition of membrane (and increase in FM dye signal) at steady state is easily
demonstrated, as it is in HT2916E cells during stimulation of mucin
secretion by Ca-dependent agonists (Bertrand CA, Laboisse C, Hopfer U, Bridges
RJ, and Frizzell RA, unpublished observations).
Summary of functional data. Estimates of the numbers of CFTR channels in intracellular vesicles obtained by using data derived from Calu-3 cell and oocyte measurements are provided in the APPENDIX. Important assumptions in these calculations include the single-channel parameters and assumed vesicle radius (100 nm). The calculations are based on reported values of membrane current and capacitance changes from patch-clamp studies of cAMP-stimulated cells (46, 102). These calculations and assumptions yield estimates of 110 channels per vesicle in these systems; they are not likely to provide more than one order of magnitude estimate of channel density. The mean oocyte current and capacitance changes reported by Weber et al. (105) were both approximately fourfold smaller than those in the Takahashi study (102) so that the calculated outcome would be unaffected. For comparison, Wright et al. (110) estimated the number of Na-dependent glucose transporters per oocyte vesicle at 1020, based on freeze-fracture scanning electron microscopy. The size of the imaged oocyte vesicles was 100120 nm. The value of 110 CFTRs per vesicle is instructive when considering whether one should be able to visualize vesicular GFP-CFTR by light microscopic methods. As discussed earlier in this article, the sensitivity of GFP detection (79), together with these considerations, suggests that CFTR density in a single transport vesicle is near the limit of detection.
Relation to other systems. It is worth emphasizing that the source of CFTR current generation, whether membrane-resident or inserted channels, need not be a black-and-white issue. In most systems, plasma membrane CFTR currents are likely to arise from both stimulation of membrane-resident channels and acutely trafficked CFTR channels, and the proportions of these may differ among cell types or experimental conditions (see CONSEQUENCES OF REGULATED CFTR TRAFFICKING). Data from studies of the regulated trafficking of GLUT-4 or AQP-2 indicate that they undergo recycling even under basal conditions (15, 17). During stimulation, the rate coefficients of the steps involved in recycling these proteins are altered, changing their distribution to increase the number of transporters and/or channels in the plasma membrane. Presumably, regulated CFTR recycling behaves similarly. As appears to be true of CFTR, the cellular background that can support regulated GLUT-4 or AQP-2 trafficking is very important. Thus the expression of GLUT-4 in nonmuscle or nonadipose derived cells eliminates the insulin-regulated redistribution of GLUT-4 (40, 44). Similarly for AQP-2, its endogenous expression in vas deferens results in a constitutive apical membrane localization; the channel is not found in a cAMP/PKA-regulated internal compartment as it is in renal epithelia, and there is no effect of stimulation on apical channel density (99). A requirement for the proper cellular context is likely to also characterize the regulated trafficking of CFTR. The AQP-2 data suggest that such differences can exist not only between epithelial and nonepithelial cells but also between epithelial systems. This conclusion underscores the need to determine which cellular components provide the proper context.
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MECHANISMS OF REGULATED CFTR TRAFFICKING: PERPETRATOR OR BYSTANDER? |
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Endocytosis. The physical interaction between the CFTR COOH terminus and the µ2 subunit of the AP-2 adapter complex, indicated by the results of recent studies (108), suggests a model like that shown in Fig. 2 for PKA-mediated regulation of endocytosis. This scheme is based on observations concerning the inherently rapid rate of CFTR-dependent endocytosis and the inhibitory effect of cAMP on this process (13, 14, 61, 85). It assumes that CFTR is normally endocytosed at a rapid rate because of its interaction with the AP-2 adapter complex at the YDSI motif of the COOH terminus. Recent data indicate that this interaction requires also phosphorylation of the µ2 subunit, which facilitates AP-2 interactions with membrane lipids (25). The model of Fig. 2 implies that the PKA-mediated inhibition of CFTR endocytosis may result from occlusion or obstruction of the adapter protein binding site at the COOH terminus by a CFTR phosphorylation event. For example, this could result from an as yet unidentified physical interaction of the phosphorylated R domain with the COOH-terminal tail where adapters bind, or it could be mediated by another interacting protein whose association with the COOH terminus is proportional to CFTR phosphorylation. The key feature of this model relies on phosphorylation-dependent accessibility of the COOH terminus site for adapter protein binding.
A second possibility is that the YXX motif itself is altered by
phosphorylation. In species where CFTR's sequence has been described, the YDSI
motif is highly conserved
(108). Although it does not
conform to a canonical PKA consensus sequence, it is possible that the Y+2
serine is phosphorylated by PKA or another kinase, which acts to disrupt the
adapter binding motif. The possibility that AMP kinase, implicated in binding
to this region of CFTR (39),
is involved in phosphorylation of the YDSI internalization motif requires
evaluation. However, the function of this kinase is presumably to downregulate
CFTR during metabolic stress, which would be expected to promote, not inhibit,
CFTR retrieval. Nevertheless, a phosphorylation event at or near this site
could reduce the endocytic rate of CFTR in parallel with the activation of
channel gating. It remains to be determined whether such a mechanism can
account for the finding, made in both epithelial and nonepithelial systems
(13,
14,
85,
94), that PKA phosphorylation
inhibits CFTR endocytosis, leading to retention of the channel on the cell
surface.
Exocytosis. What determines the ability of CFTR to enter a
regulated (as opposed to constitutive; Fig.
1) secretory pathway during its progression to the cell surface?
In Xenopus oocytes, a redistribution of CFTR between intracellular
and plasma membrane compartments is demonstrable by immunofluorescence
measurements, as discussed above. These data can be interpreted to suggest
that CFTR is stabilized, kinetically, in an intracellular compartment under
nonstimulated conditions. Therefore, it is reasonable to ask what structural
feature(s) of CFTR provides for its entry into a regulated secretory pathway
under basal conditions. CFTR carries its own regulatory domain, and the
phosphorylation of this domain stimulates both channel gating and trafficking.
Therefore, it is logical to ask whether the R domain is responsible for the
entry of CFTR into a regulated pathway. For this purpose, CFTR lacking the R
domain has been expressed from two injected RNAs, each corresponding to a CFTR
half molecule: N-TM1-NBD1 and TM2-NBD2-C
(56). Oocytes expressing these
constructs showed spontaneous currents quantitatively similar to the
stimulated currents of cells expressing wt CFTR, and they were not further
augmented by cAMP stimulation. In addition, there was no change in membrane
capacitance during stimulation, indicating that regulated CFTR trafficking
required the R domain. The direct targeting of R-CFTR to the plasma
membrane in these cells is consistent with the concept that the
nonphosphorylated R domain may allow CFTR to access a regulated intracellular
compartment and that CFTR within this compartment can be redistributed to the
plasma membrane when the R domain is phosphorylated. Therefore, it is
reasonable to ask the following: What are the mediators of this process? Do
protein interactions involving the R domain govern the movement of CFTR
through this regulated trafficking pathway?
Although these questions cannot be answered with certainty at present, the relatively recent discovery by Kirk and colleagues (72, 73) that CFTR interacts physically and functionally with syntaxin 1A (S1A) suggests soluble N-ethylmaleimide-sensitive factor attachment protein receptor (SNARE) proteins as candidates for participation in this regulated trafficking process. The interaction of S1A with the NH2 terminus of CFTR (73, 81) and the inhibition of CFTR currents during S1A overexpression (72) have been interpreted as the consequence of a direct interaction of S1A with CFTR that reduces its open probability (20). However, in addition, Peters et al. (81) showed that S1A overexpression decreased the density of CFTR channels in the plasma membrane. They concluded that disruption of SNARE complex formation by overexpressed S1A interfered with CFTR trafficking (presumably with exocytosis). As regards the functional involvement of syntaxin with CFTR, there is no reason to discount one mechanism in favor of the other; that is, syntaxin may regulate CFTR gating and also function as a component of the machinery that brings CFTR into the plasma membrane, or it may act as a regulator of that process.
How might SNARE proteins be involved in regulated CFTR exocytosis? To pose an answer to this question, we should recount several observations. First, the Kirk laboratory has shown that the NH2 terminus of CFTR, to which S1A binds, also interacts with a proximal portion of the R domain in protein binding studies (71). A series of basic amino acid residues at the NH2 terminus is thought to mediate this association. Mutation of these residues altered the magnitude and kinetics of CFTR current stimulation in oocytes and decreased the open probability of single CFTR channels (34), perhaps by interfering with NH2 terminus-R domain interactions that influence CFTR gating. The NH2 terminus is also the site of S1A binding (73). Thus it is possible that NH2 terminus-R domain interactions influence the association of CFTR with S1A or that it affects the ability of S1A to interact with other proteins necessary for CFTR trafficking.
In relation to another possible scenario, we recently reported physical and functional interactions of another synaptic terminal protein, the cysteine string protein (Csp), with CFTR (115). Csp is a resident of synaptic vesicles, and its knockout interferes with regulated neurotransmitter release (116), as does a S1A knockout (111). In protein binding studies, Csp interacted with both the CFTR NH2 terminus and R domain. It seems likely that the interactions of S1A and Csp with the NH2 terminus and the additional interactions of the NH2 terminus and Csp with the R domain may provide a means for linking proteins involved in regulated membrane trafficking to the regulatory (phosphorylation) status of the R domain (similar in general concept to the proposal above for regulated CFTR endocytosis). According to the model shown in Fig. 3, phosphorylation of the R domain would alter these protein interactions in a manner that would lead to SNARE protein associations appropriate for fusion of CFTR-containing vesicles with the plasma membrane. It is also of interest that Csp can bind to several SNARE proteins, including S1A, syntaxin 4, and vesicle-associated membrane protein 2 (VAMP-2) and that Csp has been proposed as a modulator of SNARE protein interactions (19). According to this concept, NH2 terminus-R domain interactions within CFTR can alter single-channel properties, but they may also couple the phosphorylation-dependent activation of CFTR channel gating to proteins that regulate CFTR trafficking. It is possible that a protein such as Csp, through its ability to interact with SNARE proteins on one hand and with both the NH2 terminus and R-domain of CFTR on the other, provides a transduction mechanism for coupling CFTR channel activation to channel trafficking mediated by the SNARE machinery.
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CONSEQUENCES OF REGULATED CFTR TRAFFICKING |
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In addition, the contribution of trafficking events to anion secretion may vary with the differentiation status of the epithelium (6, 37). It has been found, for example, that CFTR in some epithelial cells does not find its way to the plasma membrane until the tight junctions have formed and the cells polarize (68). As the epithelium is developing, cells may utilize regulated trafficking as a means to fine tune the apical Cl conductance until the expression levels of other transporters, metabolic or regulatory components, on which overall secretory or absorptive ion movements depend are expressed at appropriate levels. At that stage, the mature cell may find it reasonable to better anchor CFTR in the plasma membrane by virtue of PDZ domain (69, 97) and other interactions, to shift the balance of channel distribution in favor of the plasma membrane. Yet, even in well-differentiated native epithelia, significant CFTR trafficking signals can be demonstrated (see REGULATED CFTR TRAFFIC). Not until we know, from a mechanistic viewpoint, the protein interactions that mediate CFTR trafficking events will we be able to determine how such a balance is achieved or how it may be perturbed to counter the effects of disease.
Relation to CF disease. Disease-related mutations that influence
CFTR trafficking in distal compartments of the protein secretory pathway have
been identified. It is probably true that mutations in proteins that control
general trafficking pathways would have a negative selective advantage and may
be lethal; however, CFTR mutations that influence its trafficking have now
been described. Several reports have indicated that the half-life of
F508 CFTR at the plasma membrane, and perhaps in other post-Golgi
compartments, is markedly reduced relative to that of wt CFTR. Using
functional measurements of CFTR activity in CHO cells, Lukacs et al.
(59) found that the plasma
membrane currents associated with
F508 CFTR (recruited to the cell
surface at low temperature) decayed with a half-time of <4 h, whereas the
functional half-life of wt CFTR currents was >24 h.
Heda et al. (41) measured
the half-life of F508 vs. wt CFTR in the plasma membrane of LLC-PK1
cells using cell surface biotinylation, streptavidin-mediated isolation, and
immunoblot analysis. Mutant CFTR was brought to the cell surface with the use
of a combination of low temperature and butyrate preincubation. They found
that the half-life of plasma membrane
F508 CFTR was
4 h, whereas
the corresponding half-life of wt CFTR exceeded 48 h in these cells. The
cAMP-dependent iodide efflux correlated with CFTR expression at the cell
surface. These authors concluded that
F508 CFTR is more rapidly
internalized and perhaps targeted to a degradation pathway. The parallel study
of Sharma et al. (95) reached
a similar conclusion. Mutant CFTR was encouraged to escape the ER with the use
of a combination of reduced temperature and chemical chaperone (glycerol)
treatment. They observed also a rapid degradation rate of
F508 CFTR,
which they attribute to decreased conformational stability: the protein was
more susceptible to aggregation and protease digestion. These authors favor
the concept that folding defects of this CFTR mutant persist in the mature,
post-Golgi protein (most studies agree that it lacks normal channel gating
kinetics and open probability; c.f. Ref.
92) and that this leads to its
more rapid degradation, perhaps by ubiquitin-proteasome pathways. It is also
possible, in view of the model for regulated CFTR endocytosis illustrated in
Fig. 2, that the rapid
degradation of
F508 CFTR reflects its more rapid endocytic rate
(41,
95), increasing its exposure
to degradation mechanisms. It is evident from these findings that the altered
stability of mutant CFTR in distal trafficking pathways will require attention
should rescue of
F508 CFTR from ER degradation pathways be an
achievable therapeutic target for CF disease.
Another interesting disease mutation that influences distal pathway CFTR
trafficking has been identified by Silvis et al.
(98). These investigators
searched the mutation database for disease mutations that may lead to
increased CFTR endocytosis through creation of an endocytic motif. The N287Y
mutant resides in the second intracellular loop of CFTR and results in mild
disease when expressed in combination with F508 CFTR. This mutant did
not exhibit a folding defect, because the N287Y and wt CFTRs showed similar
maturation kinetics. However, there was roughly 50% of the mutant at the
plasma membrane at steady state relative to wt CFTR. An increased
colocalization of the mutant with the endocytic marker EEA1 suggested that
this mutation alters the distribution of CFTR between plasma membrane and
intracellular compartments. Cl transport was reduced in proportion to altered
cell surface CFTR, but the single-channel properties of N287Y were similar to
those of wt CFTR. Biotinylation experiments showed that N287Y CFTR was
internalized approximately twice as fast as wt CFTR, which is expected to
alter its distribution between plasma membrane and intracellular compartments.
These findings provide evidence of disease mutations where the primary defect
lies in altered kinetics of CFTR recycling at the plasma membrane, resulting
in subnormal apical membrane CFTR levels.
Intracellular CFTR channel function. A second consequence of regulated CFTR trafficking is that intracellular CFTR distribution may vary with the secretory status of the epithelium. Despite indications that intracellular CFTR channels can be activated by cAMP/PKA (9, 77), the model of Barasch et al. (5), suggesting that vesicular CFTR activity alters internal compartment pH, cellular glycoprotein processing, and bacterial colonization, has not been supported by subsequent studies (32, 35, 60, 83). Nevertheless, there is a fundamental need for resolving the causes for increased bacterial adherence to CF cells and their glycoprotein secretory products and for the preferential binding of P. aeruginosa to undersialylated CF mucins (16, 31, 48, 53, 91, 113). These findings argue that the deletion of CFTR from the distal secretory pathway influences epithelial cell surface chemistry. These properties of CF cells and their secretions could be affected by CFTR-dependent membrane trafficking events in intracellular compartments as well as at the plasma membrane domain. For example, Biwersi et al. (8) demonstrated that CFTR promotes endosome-endosome fusion, and, as with plasma membrane CFTR redistribution, this required both CFTR expression and cAMP stimulation. If the presence of CFTR in internal membranes alters patterns of posttranslational glycoprotein modification by changing the compartmentation of enzymes or substrates involved in these processes (78), the surface chemistry of CF cells may be affected.
Protein secretion. An issue related to the above discussion
concerns the role of CFTR in epithelial tissue protein secretory activity,
which is largely suggested by the data of McPherson and colleagues
(65,
67), who examined mucin and
amylase secretion by salivary glands of control and CF patients. Metabolic
incorporation of isotopic label into the carbohydrate residues of mucins was
used to quantitate secretory rates. The results showed that the
-adrenergic stimulation of mucin and amylase secretion was reduced 60%
in submandibular salivary glands derived from CF subjects. An extensive series
of studies showed that alterations in signal transduction mechanisms were not
responsible for this difference and that the mucin content of control and CF
glands was similar. Recently, similar results have been obtained by using the
salivary glands of CFTR knockout mice
(67). Again, the
cAMP-dependent secretory component of secretion was impaired. Mergey et al.
(66) have performed
experiments of this type in several human airway cell lines. They demonstrated
independent regulation of 14C-labeled glycoconjugate secretion by
PKA and PKC pathways (i.e., their effects were additive). The PKA agonist
isoproterenol increased mucin release by 40% in control cells, but this effect
was virtually eliminated (reduced to 3% stimulation) in cells derived from CF
subjects. A difference in PKC-mediated secretion between control and CF cell
lines was not detected, similar to the observations of Bradbury et al.
(13) on cAMP-regulated
membrane trafficking in T84 cells. Moreover, the attenuated PKA response in CF
airway cells could be restored by adenovirus-mediated CFTR expression. The
results suggest that CFTR can play an important role in cAMP-mediated
glycoconjugate secretion. Although not explicitly implicating CFTR in mucin
release, the studies of Kuver et al.
(54) have demonstrated
cAMP-dependent regulation of protein or mucin secretion in CFTR-expressing
canine gallbladder epithelial cells. CFTR was detected by immunostaining on
mucin granule membranes.
In studies that may have implications for epithelial protein secretion,
Yilla et al. (112) examined
the influence of vacuolar H-ATPase inhibition on the constitutive protein
secretory pathway in HepG2 cells. Although treatment with concanamycin B did
not influence ER-to-Golgi transit, the kinetics of protein traffic between the
Golgi and plasma membrane were significantly impaired. The secretion of
albumin, 1-antitrypsin, and transferrin was delayed, and
processing of N-linked glycans by sialyltransferases was inhibited, resulting
in the secretion of less extensively modified glycoproteins. This study
implicates intracellular compartment acidification in the sialylation and
N-linked glycan modification of secreted glycoproteins and with the rate of
protein secretion. In a related study, Jilling and Kirk
(50) demonstrated that cAMP
increased the secretion of several proteins, including
1-antitrypsin, into the apical but not the basolateral
compartment of T84 cells. This effect was Cl dependent and resulted also in an
increase in protein sialylation. An involvement of CFTR in regulated
exocytosis could underlie the differences in protein secretion observed in
studies of cells and/or tissues derived from normal and CF patients or CF
knockout mice.
Common threads. Once we understand the reasons for the ability of CFTR to alter membrane traffic, it may become apparent that the underlying mechanisms involved in the progression of CFTR between different compartments have certain features in common. A critical question in this regard concerns the exit of CFTR protein from the ER. Reasons for thinking along these lines include the common features controlling vesicle transit along compartments of the secretory pathway, e.g., a combination of SNARE and coat protein associations, which often involve interactions with the cargo being conveyed (3).
Our recent findings regarding Csp interactions with CFTR (115) also may provide an example of how common trafficking events may occur in the early and later components of the protein secretory pathway. Csp regulates exocytosis in neurosecretory cells, and implications for a similar role in CFTR exocytosis were discussed above. In addition, Csp plays a role in CFTR maturation in the ER. Csp antibodies co-precipitate a large proportion of band B CFTR and localize Csp protein to the ER, in addition to its presence at the apical domain. The major influence of Csp overexpression was a disruption of the biogenesis of mature CFTR (band C). Because Csp is an Hsc70 binding protein, which also binds to CFTR, it is likely that Csp serves as a CFTR co-chaperone. Csp overexpression may decrease CFTR biogenesis by prolonging its association with Hsc70, which can lead to CFTR degradation (88). Much work needs to be done to determine the role of Csp in CFTR maturation, but it is interesting to speculate that the steps involved in the egress of nascent CFTR from the ER and from a regulated compartment of the distal secretory pathway to the plasma membrane may involve similar protein interactions. Therefore, identifying the mechanisms underlying stage-specific CFTR trafficking events may also shed light on traffic occurring at other compartments of the cell.
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SUMMARY |
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APPENDIX |
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Relationship among membrane current, capacitance increase, and the
numbers of active CFTR channels and fused vesicles. The pertinent
equations for the number of active CFTR channels (NC) and
fused vesicles (NV) are
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For a Calu-3 cell
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For a Xenopus oocyte
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ACKNOWLEDGMENTS |
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Work from the authors' laboratories was supported by National Institute of Diabetes and Digestive and Kidney Diseases Grants DK-54814 and DK-56490 and the Cystic Fibrosis Foundation.
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FOOTNOTES |
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1 The reversal potential in the oocyte experiments was not determined and is
estimated. If vesicle diameter is >100 nm, the number of channels per
vesicle increases proportionately.
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REFERENCES |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
2. Ameen NA,
Martensson B, Bourguinon L, Marino C, Isenberg J, and McLaughlin GE. CFTR
channel insertion to the apical surface in rat duodenal villus epithelial
cells is upregulated by VIP in vivo. J Cell Sci
112: 887894,
1999.
3. Aridor M,
Bannykh SI, Rowe T, and Balch WE. Cargo can modulate COPII vesicle
formation from the endoplasmic reticulum. J Biol Chem
274: 43894399,
1999.
4. Atia F, Zeiske W, and van Driessche W. Secretory apical Cl channels in A6 cells: possible control by cell Ca2+ and cAMP. Pflügers Arch 438: 344353, 1999.[ISI][Medline]
5. Barasch J, Kiss B, Prince A, Saiman L, Gruenert D, and al-Awqati Q. Defective acidification of intracellular organelles in cystic fibrosis. Nature 352: 7073, 1991.[ISI][Medline]
6. Bebok Z,
Tousson A, Schwiebert LM, and Venglarik CJ. Improved oxygenation promotes
CFTR maturation and trafficking in MDCK monolayers. Am J Physiol
Cell Physiol 280:
C135C145, 2001.
7. Benharouga M,
Haaardt M, Kartner N, and Lukacs GL. COOH-terminal truncations promote
proteasome-dependent degradation of mature cystic fibrosis transmembrane
conductance regulator from post-Golgi compartments. J Cell
Biol 153:
957970, 2001.
8. Biwersi J,
Emans N, and Verkman AS. Cystic fibrosis transmembrane conductance
regulator activation stimulates endosome fusion in vivo. Proc Natl
Acad Sci USA 93:
1248412489, 1996.
9. Biwersi J and
Verkman AS. Functional CFTR in endosomal compartment of CFTR-expressing
fibroblasts and T84 cells. Am J Physiol Cell Physiol
266: C149C156,
1994.
10. Bradbury NA. Intracellular CFTR: localization and function. Physiol Rev 79: S175S191, 1999.[Medline]
11. Bradbury NA,
Clark JA, Watkins SC, Widnell CC, Smith HS, and Bridges RJ.
Characterization of the internalization pathways for the cystic fibrosis
transmembrane conductance regulator. Am J Physiol Lung Cell Mol
Physiol 276:
L659L668, 1999.
12. Bradbury NA,
Cohn JA, Venglarik CJ, and Bridges RJ. Biochemical and biophysical
identification of cystic fibrosis transmembrane conductance regulator chloride
channels as components of endocytic clathrin-coated vesicles. J
Biol Chem 269:
82968302, 1994.
13. Bradbury NA, Jilling T, Berta G, Sorscher EJ, Bridges RJ, and Kirk KL. Regulation of plasma membrane recycling by CFTR. Science 256: 530532, 1992.[ISI][Medline]
14. Bradbury NA,
Jilling T, Kirk KL, and Bridges RJ. Regulated endocytosis in a chloride
secretory epithelial cell line. Am J Physiol Cell
Physiol 262:
C752C759, 1992.
15. Brown D,
Katsura T, and Gustafson CE. Cellular mechanisms of aquaporin trafficking.
Am J Physiol Renal Physiol 275:
F328F331, 1998.
16. Bryan R, Kube
D, Perez A, David P, and Prince A. Over-production of the CFTR R domain
leads to increased levels of asialoGM1 and increased Pseudomonas
aeruginosa binding by epithelial cells. Am J Respir Cell Mol
Biol 19:
269277, 1998.
17. Bryant NJ, Govers R, and James DE. Regulated transport of the glucose transporter GLUT4. Nature 3: 267277, 2002.
18. Bulteau L,
Derand R, Mettey Y, Metaye T, Morris MR, McNeilly CM, Folli C, Galietta LJ,
Zegarra-Moran O, Pereira MM, Jougla C, Dormer RL, Vierfond JM, Joffre M, and
Becq F. Properties of CFTR activated by the xanthine derivative X-33 in
human airway Calu-3 cells. Am J Physiol Cell Physiol
279: C1925C1937,
2000.
19. Chamberlain LH and Burgoyne RD. Cysteine-string protein: the chaperone at the synapse. J Neurochem 74: 17811789, 2000.[ISI][Medline]
20. Chang SY, Di A,
Naren AP, Palfrey HC, Kirk KL, and Nelson DJ. Mechanisms of CFTR
regulation by syntaxin 1A and PKA. J Cell Sci
115: 783791,
2002.
21. Chen P, Hwant
GC, and Gillis KD. The relationship between cAMP,
Ca2+, and transport of CFTR to the plasma membrane.
J Gen Physiol 118:
135144, 2001.
22. Cheng PW, Sherman JM, Boat TF, and Bruce M. Quantitation of radiolabeled mucous glycoproteins secreted by tracheal explants. Anal Biochem 117: 301306, 1981.[ISI][Medline]
23. Cheng SH, Gregory RJ, Marshall J, Paul S, Souza DW, White GA, O'Riordan CR, and Smith AE. Defective intracellular transport and processing of CFTR is the molecular basis of most cystic fibrosis. Cell 63: 827834, 1990.[ISI][Medline]
24. Chow CW,
Khurana S, Woodside M, Grinstein S, and Orlowski J. The epithelial
Na+/H+ exchanger, NHE3, is internalized through a
clathrin-mediated pathway. J Biol Chem
274: 3755137558,
1999.
25. Collins BM, McCoy AJ, Kent HM, Evans PR, and Owen DJ. Molecular architecture and functional model of the endocytic AP2 complex. Cell 109: 523535, 2002.[ISI][Medline]
26. Dalemans W, Hinnrasky J, Slos P, Dreyer D, Fuchey C, Pavirani A, and Puchelle E. Immunocytochemical analysis reveals differences between the subcellular localization of normal and delta Phe508 recombinant cystic fibrosis transmembrane conductance regulator. Exp Cell Res 201: 235240, 1992.[ISI][Medline]
27. Denning GM, Ostedgaard LS, Cheng SH, Smith AE, and Welsh MJ. Localization of cystic fibrosis transmembrane conductance regulator in chloride secretory epithelia. J Clin Invest 89: 339340, 1992.[ISI][Medline]
28. Denning GM, Ostedgaard LS, and Welsh MJ. Abnormal localization of cystic fibrosis transmembrane conductance regulator in primary cultures of cystic fibrosis airway epithelia. J Cell Biol 118: 551559, 1992.[Abstract]
29. Dho S, Grinstein S, and Foskett JK. Plasma membrane recycling in CFTR-expressing CHO cells. Biochim Biophys Acta 1225: 7882, 1993.[ISI][Medline]
30. Dmitrieva NI, Michea LF, Rocha GM, and Burg MB. Cell cycle delay and apoptosis in response to osmotic stress. Comp Biochem Physiol A 130: 411420, 2001.[ISI]
31. Dosanjh A,
Lencer W, Brown D, Ausiello DA, and Stow JL. Heterologous expression of
F508 CFTR results in decreased sialylation of membrane glycoconjugates.
Am J Physiol Cell Physiol 266:
C360C366, 1994.
32. Dunn KW, Park
J, Semrad CE, Gelman DL, Shevell T, and McGraw TE. Regulation of endocytic
trafficking and acidification are independent of the cystic fibrosis
transmembrane regulator. J Biol Chem
269: 53365345,
1994.
33. Frates RC Jr, Kaizu TT, and Last JA. Mucus glycoproteins secreted by respiratory epithelial tissue from cystic fibrosis patients. Pediatr Res 17: 3034, 1983.[Abstract]
34. Fu J, Ji HL,
Naren AP, and Kirk KL. A cluster of negative charges at the amino terminal
tail of CFTR regulates ATP-dependent channel gating. J
Physiol 536:
459470, 2001.
35. Gibson GA, Hill
WG, and Weisz OA. Evidence against the acidification hypothesis in cystic
fibrosis. Am J Physiol Cell Physiol
279: C1088C1099,
2000.
36. Giovannucci DR, Yule DI, and Stuenkel EL. Optical measurement of stimulus-evoked membrane dynamics in single pancreatic acinar cells. Am J Physiol Cell Physiol 275: C732C739, 1998.[Abstract]
36. Greger R, Thiele II, Warth R, and Bleich M. Does stimulation of NaCl secretion in in vitro perfused rectal gland tubules of Squalus acanthias increase membrane capacitance? Pflügers Arch 436: 538544, 1998.[ISI][Medline]
37. Guggino WB.
Focus on "Exocytosis is not involved in activation of
C1 secretion via CFTR in Calu-3 airway epithelial
cells." Am J Physiol Cell Physiol
275: C911C912,
1998.
38. Gustafson CE,
Katsura T, McKee M, Bouley R, Casanova JE, and Brown D. Recycling of AQP2
occurs through a temperature- and bafilomycin-sensitive
trans-Golgi-associated compartment. Am J Physiol Renal
Physiol 278:
F317F326, 2000.
39. Hallows KR,
Raghuram V, Kemp BE, Witters LA, and Foskett JK. Inhibition of cystic
fibrosis transmembrane conductance regulator by novel interaction with the
metabolic sensor AMP-activated protein kinase. J Clin
Invest 105:
17111721, 2000.
40. Haney PM, Slot JW, Piper RC, James DE, and Mueckler M. Intracellular targeting of the insulin-regulatable glucose transporter (GLUT4) is isoform specific and independent of cell type. J Cell Biol 114: 689699, 1991.[Abstract]
41. Heda GD,
Tanwani M, and Marino CR. The F508 mutation shortens the
biochemical half-life of plasma membrane CFTR in polarized epithelial cells.
Am J Physiol Cell Physiol 280:
C166C174, 2001.
42. Howard M, Jiang
X, Stolz DB, Hill WG, Johnson JA, Watkins SC, Frizzell RA, Bruton CM, Robbins
PD, and Weisz OA. Forskolin-induced apical membrane insertion of virally
expressed, epitope-tagged CFTR in polarized MDCK cells. Am J
Physiol Cell Physiol 279:
C375C382, 2000.
43. Howard M, Jilling T, and DuVall M. cAMP-regulated trafficking of epitope-tagged CFTR. Kidney Int 49: 16421648, 1996.[ISI][Medline]
44. Hudson AW, Ruiz M, and Birnbaum MJ. Isoform-specific subcellular targeting of glucose transporters in mouse fibroblasts. J Cell Biol 116: 785797, 1992.[Abstract]
45. Hug MJ, Bertrand CA, and Frizzell RA. cAMP increases FM 1-43 labeled membrane uptake in Calu-3 cells (Abstract). Pediatr Pulmonol 34: 240, 2002.
46. Hug MJ, Sun F, and Frizzell RA. cAMP increases membrane conductance and membrane capacitance in Calu-3 cells (Abstract). Pediatr Pulmonol 28: 183, 1999.
47. Hug MJ, Thiele IE, and Greger R. The role of exocytosis in the activation of the chloride conductance in Chinese hamster ovary cells (CHO) stably expressing CFTR. Pflügers Arch 434: 779784, 1997.[ISI][Medline]
48. Imundo L,
Barasch J, Prince A, and Al-Awqati Q. Cystic fibrosis epithelial cells
have a regulator for pathogenic bacteria on their apical surface.
Proc Natl Acad Sci USA 92:
30193023, 1995.
49. Jentsch TJ,
Stein V, Weinreich F, and Zdebik AA. Molecular structure and physiological
function of chloride channels. Physiol Rev
82: 503568,
2002.
50. Jilling T and
Kirk KL. Cyclic AMP and chloride-dependent regulation of the apical
constitutive secretory pathway in colonic epithelial cells. J Biol
Chem 271:
43814387, 1996.
51. Kamsteeg EJ and
Deen PMT. Importance of aquaporin-2 expression levels in
genotype-phenotype studies in nephrogenic diabetes insipidus. Am J
Physiol Renal Physiol 279:
F778F784, 2000.
52. Kartner N,
Augustinas O, Jensen TJ, Naismith AL, and Riordan JR. Mislocation of
F508 CFTR in cystic fibrosis sweat gland. Nat
Genet 1:
321327, 1992.[ISI][Medline]
53. Krivan HC, Ginsburg V, and Roberts DD. Pseudomonas aeruginosa and Pseudomonas cepacia isolated from cystic fibrosis patients bind specifically to gangliotetraosylceramide (asialo GM1) and gangliotriaosylceramide (asialo GM2). Arch Biochem Biophys 260: 493496, 1988.[ISI][Medline]
54. Kuver R,
Klinkspoor JH, Osborne WR, and Lee SP. Mucous granule exocytosis and CFTR
expression in gallbladder epithelium. Glycobiology
10: 149157,
2000.
55. Lehrich RW,
Aller SG, Webster P, Marino CR, and Forrest JN. Vasoactive intestinal
peptide, forskolin, and genistein increase apical CFTR trafficking in the
rectal gland of the spiny dogfish, Squalus acanthias. Acute
regulation of CFTR trafficking in an intact epithelium. J Clin
Invest 101:
737745, 1998.
56. Lewarchik CM, Peters KW, Qi J, Dudley R, and Frizzell RA. R domain regulation of CFTR trafficking (Abstract). Pediatr Pulmonol 30: 176, 2000.
57. Liu X, Smith
SS, Sun F, and Dawson DC. CFTR: covalent modification of
cysteine-substituted channels expressed in Xenopus oocytes shows that
activation is due to the opening of channels resident in the plasma membrane.
J Gen Physiol 118:
433446, 2001.
58. Loffing J,
Moyer BD, McCoy D, and Stanton BA. Exocytosis is not involved in
activation of Cl secretion via CFTR in Calu-3 airway
epithelial cells. Am J Physiol Cell Physiol
275: C913C920,
1998.
59. Lukacs GL,
Chang XB, Bear C, Kartner N, Mohamed A, Riordan JR, and Grinstein S. The
F508 mutation decreases the stability of cystic fibrosis transmembrane
conductance regulator in the plasma membrane. Determination of functional
half-lives on transfected cells. J Biol Chem
268: 592598,
1993.
60. Lukacs GL,
Chang XB, Kartner N, Rotstein OD, Riordan JR, and Grinstein S. The cystic
fibrosis transmembrane regulator is present and functional in endosomes. Role
as a determinant of endosomal pH. J Biol Chem
267: 1456814672,
1992.
61. Lukacs GL, Segal G, Kartner N, Grinstein S, and Zhang F. Constitutive internalization of cystic fibrosis transmembrane conductance regulator occurs via clathrin-dependent endocytosis and is regulated by protein phosphorylation. Biochem J 328: 353361, 1997.[ISI][Medline]
62. Malide D, Ramm
G, Cushman SW, and Slot JW. Immunoelectron microscopic evidence that GLUT4
translocation explains the stimulation of glucose transport in isolated rat
white adipose cells. J Cell Sci
113: 42034210,
2000.
63. Mangoura D, Sogos V, and Dawson G. Phorbol esters and PKC signaling regulate proliferation, vimentin cytoskeleton assembly and glutamine synthetase activity of chick embryo cerebrum astrocytes in culture. Brain Res Dev Brain Res 87: 111, 1995.[ISI][Medline]
64. McPherson MA and Dormer RL. Cystic fibrosis: a defect in stimulus-response coupling. Trends Biochem Sci 13: 1013, 1988.[ISI][Medline]
65. McPherson MA, Pereira MM, Russell D, McNeilly CM, Morris RM, Stratford FL, and Dormer RL. The CFTR-mediated protein secretion defect: Pharmacological correction. Pflügers Arch 443: S121S126, 2001.[ISI][Medline]
66. Mergey M,
Lemnaouar M, Veissiere D, Perricaudet M, Gruenert DC, Picard J, Capeau J,
Brahimi-Horn MC, and Paul A. CFTR gene transfer corrects defective
glycoconjugate secretion in human CF epithelial tracheal cells. Am
J Physiol Lung Cell Mol Physiol 269:
L855L864, 1995.
67. Mills CL, Dorin JR, Davidson DJ, Porteus DJ, Alton EW, Dormer RL, and McPherson MA. Decreased beta-adrenergic stimulation of glycoprotein secretion in CF mice submandibular glands: reversal by the methylxanthine, IBMX. Biochem Biophys Res Commun 215: 67481, 1995.[ISI][Medline]
68. Morris AP,
Cunningham SA, Tousson A, Benos DJ, and Frizzell RA.
Polarization-dependent apical membrane CFTR targeting underlies
cAMP-stimulated Cl secretion in epithelial cells.
Am J Physiol Cell Physiol 266:
C254C268, 1994.
69. Moyer BD,
Duhaime M, Shaw C, Denton J, Reynolds D, Karlson KH, Pfeiffer J, Wang S,
Mickle JE, Milewski M, Cutting GR, Guggino WB, Li M, and Stanton BA. The
PDZ-interacting domain of cystic fibrosis transmembrane conductance regulator
is required for functional expression in the apical plasma membrane.
J Biol Chem 275:
2706927074, 2000.
70. Moyer BD,
Loffing J, Schwiebert EM, Loffing-Cueni D, Halpin PA, Karlson KH, Ismailov II,
Guggino WB, Langford GM, and Stanton BA. Membrane trafficking of the
cystic fibrosis gene product, cystic fibrosis transmembrane conductance
regulator, tagged with green fluorescent protein in Madin-Darby canine kidney
cells. J Biol Chem 273:
2175921768, 1998.
71. Naren AP,
Cormet-Boyaka E, Fu J, Villain M, Blalock JE, Quick MW, and Kirk KL. CFTR
chloride channel regulation by an interdomain interaction.
Science 286:
544548, 1999.
72. Naren AP, Nelson DJ, Xie W, Jovov B, Pevsner J, Bennett MK, Benos DJ, Quick MW, and Kirk KL. Regulation of CFTR chloride channels by syntaxin and Munc18 isoforms. Nature 390: 302305, 1997.[ISI][Medline]
73. Naren AP, Quick
MW, Collawn JF, Nelson DJ, and Kirk KL. Syntaxin 1A inhibits CFTR chloride
channels by means of domain-specific protein-protein interactions.
Proc Natl Acad Sci USA 95:
1097210977, 1998.
74. Nesterov A, Carter RE, Sorkina T, Gill GN, and Sorkin A. Inhibition of the receptor-binding function of clathrin adaptor protein AP-2 by dominant-negative mutant mu2 subunit and its effects on endocytosis. EMBO J 18: 14891499, 1999.
75. Niwa K, Shibuya I, and Kanno T. Temperature dependence of processes proximal and distal to the glucose-induced [Ca2+]i rise in stimulus-secretion coupling in rat pancreatic islets. Biol Signals 5: 3043, 1996.[ISI][Medline]
76. Olsen MK, Reszka AA, and Abraham I. KT5720 and U-98017 inhibit MAPK and alter the cytoskeleton and cell morphology. J Cell Physiol 176: 525536, 1998.[ISI][Medline]
77. Pasyk EA and
Foskett JK. Cystic fibrosis transmembrane conductance regulator-associated
ATP and adenosine 3'-phosphate 5'-phosphosulfate channels in
endoplasmic reticulum and plasma membranes. J Biol
Chem 272:
77467751, 1997.
78. Pasyk EA and
Foskett JK. Mutant (F508) cystic fibrosis transmembrane conductance
regulator Cl channel is functional when retained in
endoplasmic reticulum of mammalian cells. J Biol Chem
270: 1234712350,
1995.
79. Paterson GH, Knobel SM, Sharif WD, Kain SR, and Piston DW. Use of the green fluorescent protein and its mutants in quantitative fluorescence microscopy. Biophys J 73: 27822790, 1997.[Abstract]
80. Pearse BM, Smith CJ, and Owen DJ. Clathrin coat construction in endocytosis. Curr Opin Struct Biol 10: 220228, 2000.[ISI][Medline]
81. Peters W, Qi
JJ, Watkins SC, and Frizzell RA. Syntaxin 1A inhibits regulated CFTR
trafficking in Xenopus oocytes. Am J Physiol Cell
Physiol 277:
C174C180, 1999.
82. Pilewski JM and Frizzell RA. Role of CFTR in airway disease. Physiol Rev 79: S215S255, 1999.[Medline]
83. Poschet JF,
Boucher JC, Tatterson L, Skidmore J, Van Dyke RW, and Deretic V. Molecular
basis for defective glycosylation and Pseudomonas pathogenesis in
cystic fibrosis lung. Proc Natl Acad Sci USA
98: 1397213977,
2001.
84. Prince LS,
Peter K, Hatton SR, Zaliauskiene L, Cotlin LF, Clancy JP, Marchase RB, and
Collawn JF. Efficient endocytosis of the cystic fibrosis transmembrane
conductance regulator requires a tyrosine-based signal. J Biol
Chem 274:
36023609, 1999.
85. Prince LS, Workman RB Jr, and Marchase RB. Rapid endocytosis of the cystic fibrosis transmembrane conductance regulator chloride channel. Proc Natl Acad Sci USA 91: 51925196, 1994.[Abstract]
86. Puchelle E, Gaillard D, Ploton D, Hinnrasky J, Fuchey C, Boutterin MC, Jacquot J, Dreyer D, Pavirani A, and Dalemans W. Differential localization of the cystic fibrosis transmembrane conductance regulator in normal and cystic fibrosis airway epithelium. Am J Respir Cell Mol Biol 7: 485491, 1992.[ISI][Medline]
87. Renstrom E, Eliasson L, Bokvist K, and Rorsman P. Cooling inhibits exocytosis in single mouse pancreatic B-cells by suppression of granule mobilization. J Physiol 494: 4152, 1996.[Abstract]
88. Rubenstein RC and Zeitlin PL. Sodium 4-phenylbutyrate downregulates Hsc70: implications
for intracellular trafficking of F508-CFTR. Am J Physiol
Cell Physiol 278:
C259C267, 2000.
89. Saiman L and Prince A. Pseudomonas aeruginosa pili bind to asialoGM1 which is increased on the surface of cystic fibrosis epithelial cells. J Clin Invest 92: 18751880, 1993.[ISI][Medline]
90. Scanlin TF and Glick MC. Glycosylation and the cystic fibrosis transmembrane conductance regulator. Respir Res 2: 276279, 2001.[ISI][Medline]
91. Scanlin TF and Glick MC. Terminal glycosylation in cystic fibrosis. Biochim Biophys Acta 1455: 241253, 1999.[ISI][Medline]
92. Schultz BD,
Bridges RJ, and Frizzell RA. Rescue of dysfunctional F508 CFTR
chloride activity by IBMX. J Physiol
170: 5166,
1999.
93. Schwiebert EM, Benos DJ, Egan ME, Stutts MJ, and Guggino WB. CFTR is a conductance regulator as well as a chloride channel. Physiol Rev 79: S145S166, 1999.[Medline]
94. Schwiebert EM,
Gesek F, Ercolani L, Wjasow C, Gruenert DC, Karlson K, and Stanton BA.
Heterotrimeric G proteins, vesicle trafficking, and CFTR Cl
channels. Am J Physiol Cell Physiol
267: C272C281,
1994.
95. Sharma M,
Benharouga M, Hu W, and Lukacs GL. Conformational and
temperature-sensitive stability defects of the F508 cystic fibrosis
transmembrane conductance regulator in post-endoplasmic reticulum
compartments. J Biol Chem 276:
89428950, 2001.
96. Shimkets RA,
Lifton RP, and Canessa CM. The activity of the epithelial sodium channel
is regulated by clathrin-mediated endocytosis. J Biol
Chem 272:
2553725541, 1997.
97. Short DB,
Trotter KW, Reczek D, Kreda SM, Bretscher A, Boucher RC, Stutts MJ, and
Milgram SL. An apical PDZ protein anchors the cystic fibrosis
transmembrane conductance regulator to the cytoskeleton. J Biol
Chem 273:
1979719801, 1998.
98. Silvis MR,
Picciano JA, Bertrand C, Weixel K, Bridges RJ, and Bradbury NA. A mutation
in the cystic fibrosis transmembrane conductance regulator generates a novel
internalization sequence and enhances endocytic rates. J Biol
Chem 278:
1155411560, 2003.
99. Stevens AL,
Breton S, Gustafson CE, Bouley R, Nelson RD, Kohan DE, and Brown D.
Aquaporin 2 is a vasopressin-independent, constitutive apical membrane protein
in rat vas deferens. Am J Physiol Cell Physiol
278: C791C802,
2000.
100. Sun F, Hug MJ,
Lewarchik CM, Yun C, Bradbury NA, and Frizzell RA. E3KARP mediates the
association of ezrin and protein kinase A with the cystic fibrosis
transmembrane conductance regulator in airway cells. J Biol
Chem 275:
2953929546, 2000.
101. Sun T-X, Van
Hoek A, Huang Y, Bouley R, McLaughlin M, and Brown D. Aquaporin-2
localization in clathrin-coated pits: inhibition of endocytosis by
dominant-negative dynamin. Am J Physiol Renal Physiol
282: F998F1011,
2002.
102. Takahashi A,
Watkins SC, Howard MB, and Frizzell RA. CFTR-dependent membrane insertion
is linked to stimulation of the CFTR chloride conductance. Am J
Physiol Cell Physiol 271:
C1887C1894, 1996.
103. Tousson A,
Fuller CM, and Benos DJ. Apical recruitment of CFTR in T-84 cells is
dependent on cAMP and microtubules but not Ca2+ or
microfilaments. J Cell Sci 109:
13251334, 1996.
104. Trezise AE and Buchwald M. In vivo cell-specific expression of the cystic fibrosis transmembrane conductance regulator. Nature 353: 434437, 1991.[ISI][Medline]
105. Weber WM, Cuppens H, Cassiman J-J, Clauss W, and Van Driessche W. Capacitance measurements reveal different pathways for the activation of CFTR. Pflügers Arch 438: 561569, 1999.[ISI][Medline]
106. Weber WM, Segal A, Vankeerberghen A, Cassiman JJ, and Van Driessche W. Different activation mechanisms of cystic fibrosis transmembrane conductance regulator expressed in Xenopus laevis oocytes. Comp Biochem Physiol A 139: 521531, 2001.
107. Webster P,
Vanacore L, Nairn AC, and Marino CR. Subcellular localization of CFTR to
endosomes in a ductal epithelium. Am J Physiol Cell
Physiol 267:
C340C348, 1994.
108. Weixel KM and
Bradbury NA. Mu2 binding directs the cystic fibrosis transmembrane
conductance regulator to the clathrin-mediated endocytic pathway. J
Biol Chem 276:
4625146259, 2001.
109. Weixel KM and
Bradbury NA. The carboxyl terminus of the cystic fibrosis transmembrane
conductance regulator binds to AP-2 clathrin adaptors. J Biol
Chem 275:
36553660, 2000.
110. Wright EM,
Hirsch JR, Loo DD, and Zampighi GA. Regulation of Na+/glucose
cotransporters. J Exp Biol 200:
287293, 1997.
111. Wu MN, Fergestad T, Lloyd TE, He Y, Broadie K, and Bellen HJ. Syntaxin 1A interacts with multiple exocytic proteins to regulate neurotransmitter release in vivo. Neuron 23: 593605, 1999.[ISI][Medline]
112. Yilla M, Tan A,
Ito K, Milwa K, and Ploegh HL. Involvement of the vacuolar
H+-ATPases in the secretory pathway of HepG2 cells. J
Biol Chem 268:
1909219100, 1993.
113. Zar H, Saiman L, Quittell L, and Prince A. Binding of Pseudomonas aeruginosa to respiratory epithelial cells from patients with various mutations in the cystic fibrosis transmembrane regulator. J Pediatr 126: 230233, 1995.[ISI][Medline]
114. Zeng WZ, Babich
V, Ortega B, Quigley R, White SJ, Welling PA, and Huang CL. Evidence for
endocytosis of ROMK potassium channel via clathrin-coated vesicles.
Am J Physiol Renal Physiol 283:
F630F639, 2002.
115. Zhang H, Peters
KW, Sun F, Marino CR, Lang J, Burgoyne RD, and Frizzell RA. Cysteine
string protein interacts with and modulates the maturation of CFTR.
J Biol Chem 277:
2894828958, 2002.
116. Zinsmaier KE, Hofbauer A, Heimbeck G, Pflugfelder GO, Buchner S, and Buchner E. A cysteine-string protein is expressed in retina and brain of Drosophila. J Neurogenet 7: 1529, 1990.[ISI][Medline]