1 Research and 3 Medical Services, Veterans Affairs Medical Center and Departments of 2 Medicine and 4 Physiology and Biophysics, The University of Tennessee Health Sciences Center, Memphis, Tennessee 38163
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ABSTRACT |
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Although the
biosynthetic arrest of the F508 mutant of cystic fibrosis
transmembrane conductance regulator (CFTR) can be partially reversed by
physical and chemical means, recent evidence suggests that the
functional stability of the mutant protein after reaching the cell
surface is compromised. To understand the molecular basis for this
observation, the current study directly measured the half-life of
F508 and wild-type CFTR at the cell surface of transfected
LLC-PK1 cells. Plasma membrane CFTR expression over time
was characterized biochemically and functionally in these polarized
epithelial cells. Surface biotinylation, streptavidin extraction, and
quantitative immunoblot analysis determined the biochemical half-life
of plasma membrane
F508 CFTR to be ~4 h, whereas the plasma
membrane half-life of wild-type CFTR exceeded 48 h. This
difference in biochemical stability correlated with CFTR-mediated
transport function. These findings indicate that the
F508 mutation
decreases the biochemical stability of CFTR at the cell surface. We
conclude that the
F508 mutation triggers more rapid internalization
of CFTR and/or its preferential sorting to a pathway of rapid degradation.
cystic fibrosis; regulation; membrane protein; endocytosis; chloride channel; cystic fibrosis transmembrane conductance regulator
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INTRODUCTION |
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MOST CYSTIC
FIBROSIS (CF) is caused by a gene mutation that results in the
deletion of the phenylalanine at position 508 of CFTR, the cystic
fibrosis transmembrane conductance regulator (CFTR) protein
(13). The F508 deletion results in a misfolding of the
nascent CFTR molecule in the endoplasmic reticulum (ER), leading to its
retention in the ER by chaparone proteins and its subsequent
ubiquitination and premature degradation by the cytoplasmic proteosome
complex (reviewed in Ref. 16). As a consequence, little if
any
F508 CFTR is processed to its mature, fully glycosylated form or
trafficked to the plasma membrane where CFTR normally functions as a
cAMP-activated chloride channel. Because the
F508 mutant retains
some intrinsic chloride transport function (8, 18),
interest in approaches that drive
F508 CFTR past the ER quality
control system and to the cell surface has grown.
Several in vitro studies have shown that surface expression of F508
CFTR can be upregulated by chemical or physical means. Butyrate
compounds have been shown to create cAMP-activated chloride currents in
cultured
F508 cells, presumably by overwhelming the ER quality
control system through an increase in gene transcription (4,
28). Glycerol (30) and low temperature
(8) stabilize the conformation of newly synthesized
F508, allowing some of it to bypass the ER quality control system
and reach biochemical maturity. After each of these treatments, an
increase in cAMP-activated chloride transport can be detected in the
mutant cells. These findings suggest that pharmacological agents that
downregulate the ER quality control system might be useful in
correcting the chloride transport defect in
F508 cells.
For this approach to be clinically effective, the F508 protein must
maintain some degree of biochemical stability after reaching the cell
surface. Little is known, however, of the fate of the CFTR protein
after it reaches the cell surface. Initial work suggested that
F508
and wild-type CFTR had similar plasma membrane half-lives (8), but a subsequent in vitro study has shown that
cAMP-activated chloride currents in nonpolarized
F508 cells are less
stable than those in wild-type cells (23). Because plasma
membrane expression of the CFTR proteins was not directly examined in
that study, the molecular basis for this observation is not known. The
functional instability in
F508 cells could have been due to more
rapid inactivation of
F508 channels at the cell surface or to more
rapid internalization and/or degradation of the mutant protein.
Functional inactivation is supported by data from several studies that
show the conduction properties of
F508 CFTR differ from those of
wild-type CFTR (6, 9, 11). In fact, a recent electrophysiological study has since confirmed that
F508 CFTR is
more rapidly inactivated than wild-type CFTR in excised membrane patches (31).
Documented differences in vesicle trafficking of F508 and wild-type
CFTR (2, 32) suggest a potential role for membrane trafficking in the regulation of plasma membrane CFTR function. However, technical difficulties in getting measurable quantities of the
F508 protein to the cell surface have hindered efforts to compare
plasma membrane
F508 and wild-type CFTR protein expression. The
current study overcame that obstacle by the simultaneous treatment of
cells with low temperature and sodium butyrate, which act
synergistically to markedly upregulate surface CFTR expression
(12). With readily detectable levels of
F508 CFTR at
the cell surface, it became possible to test the hypothesis that the
F508 protein is more biochemically unstable than wild-type CFTR.
Performed in a polarized epithelial cell line, the experiments
presented confirm that the biochemical half-life of plasma membrane
F508 CFTR is much shorter than that of wild-type CFTR. These
differences in biochemical half-life correlate with transport function,
suggesting that rapid internalization and/or degradation of
F508
CFTR alone can account for the rapid loss of chloride transport
function characteristic of
F508 cells.
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MATERIALS AND METHODS |
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Cell line.
Pig kidney epithelial cells (LLC-PK1), stably transfected
with human wild-type or F508-CFTR cDNA were kindly provided by Dr.
Seng Cheng (Genzyme, Boston, MA). Cells were grown in low-glucose DMEM
supplemented with 10% FBS and 400 µg/ml Genticin (Life Technologies, Grand Island, NY) at 37°C on plastic dishes coated with human placental type IV collagen (Sigma Chemical, St. Louis, MO). For 125I efflux experiments, cells were grown on clear
transwell membrane inserts coated with type IV collagen (Corning
Costar, Cambridge, MA). For immunocytochemical experiments, cells were
grown on collagen-coated glass coverslips.
Upregulation of surface CFTR expression. Transfected LLC-PK1 cells grown to ~70% confluence as described above were treated for 60 h at 27°C in the presence of 5 mM sodium butyrate to upregulate surface CFTR expression (12). Because low temperature partially inhibits cell proliferation, cell overgrowth did not occur during this 60-h treatment, and confluency was rarely achieved.
Biochemical determination of surface CFTR expression over time. After upregulation of surface CFTR expression, cells were washed with butyrate-free media and were incubated at 37°C in media containing 20 µg/ml of cycloheximide, an inhibitor of protein synthesis, for designated time intervals up to 48 h. Cells were then washed with ice-cold PBS, pH 7.4, containing 0.1 mM CaCl2 and 1 mM MgCl2 to inhibit vesicle trafficking, and surface biotinylation was performed as described previously (19). Briefly, the glycosidic moieties of surface membrane proteins were derivatized with sodium periodate and biotinylated using biotin-LC-hydrazide according to company protocol (Pierce, Rockford, IL). The efficiency of surface biotinylation was tested by examining the effect of increasing biotin-LC-hydrazide exposure times on surface CFTR expression. No time-dependent increase in surface CFTR expression occurred with prolonged reagent exposure (data not shown).
After surface biotinylation, cells were lysed with 1% SDS containing 0.2 mM phenylmethylsulfonyl fluoride and 1 mM benzamidine, sonicated to shear DNA molecules, and centrifuged at 10,000 g for 10 min at 4°C to remove cellular debris. The clear supernatants, normalized to 50 µg of total protein, were nutated for 30 min with an excess of streptavidin-coated agarose (SA) beads or uncoated control agarose (CN) beads (Sigma). After incubation, the beads were pelleted, and the supernatants were subjected to 6.5% SDS-PAGE followed by transfer to Hybond-P polyvinylidene difluoride membrane (Amersham, Sunnyvale, CA). The transfer was blocked with 5% nonfat dry milk in 137 mM NaCl, 0.1% Tween 20, and 20 mM Tris · HCl, pH 7.6, and immunoblotted with 1:100 dilution of R3194, an affinity-purified polyclonal anti-CFTR antibody. The specificity of R3194 for CFTR has been characterized previously in transgenic mice (37), in CFTR- and mock-transfected HEK-293 cells (17), and by COOH-terminal CFTR peptide competition experiments (37). CFTR bands were visualized by enhanced chemifluorescence (Amersham) and were quantified using a STORM 860 imaging system with ImageQuant software (Molecular Dynamics, Sunnyvale, CA). With the use of the above approach, biotinylated surface proteins were irreversibly extracted by the SA bead processing. The efficiency of this extraction step was examined by stripping the immunoblots and reblotting with streptavidin-conjugated horseradish peroxidase followed by Sigma Fast diaminobenzidine color development to confirm the absence of biotinylated proteins in SA bead processed samples (see Fig. 3). Thus the post-SA bead supernatants contain only intracellular (unbiotinylated) proteins, whereas CN bead supernatants contain all cellular proteins (biotinylated and unbiotinylated). Surface CFTR expression was then determined by subtracting the intensity of the CFTR signal after SA bead extraction from the intensity of the total CFTR signal from CN bead processed samples.Functional determination of surface CFTR expression over time.
Chloride secretion was determined by isotopic efflux of
125I from preloaded cells as previously described
(33). Briefly, membrane inserts containing
LLC-PK1 cells treated with 20 µg/ml of cycloheximide at
37°C were excised, and cells were washed with efflux buffer (140 mM
NaCl, 4.7 mM KCl, 1.2 mM CaCl2, 10 mM glucose, and 10 mM
HEPES, pH 7.4). The cells were then loaded with 125I by
incubation with 3 µCi of 125I-labeled Na (New England
Nuclear, Boston, MA) for 1 h at 37°C. Cell monolayers were
washed free of excess 125I, and the time-dependent efflux
of the isotope into the media was measured after stimulation with the
cAMP agonists forskolin (10 µM) and isobutyl methylxanthine (IBMX,
1.5 mM). Cells were then solubilized with 0.1 N NaOH, and
125I radioactivity in the media samples and the final cell
lysate was determined by counting gamma emissions (Packard Minaxi gamma counter, series 5000). The rate coefficient of iodide efflux
(r) was calculated using the following formula
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Immunocytochemical determination of surface CFTR expression.
Butyrate/low-temperature-treated cells, grown on collagen-coated
coverslips, were fixed for 1 h with 4% paraformaldehyde and then
permeabilized with 0.25% saponin in PBS for 10 min. Free aldehyde
residues were quenched for 30 min with 50 mM NH4Cl, and cells were blocked with 1% BSA, both prepared in PBS. Cells were then
incubated overnight at 4°C with R3194 (polyclonal anti-CFTR) and/or
monoclonal antibody 6H (monoclonal against the -subunit of rat
Na+-K+-ATPase, generously provided by Michael
J. Caplan, Yale School of Medicine, New Haven, CT). Anti-CFTR labeling
was detected with FITC-conjugated goat anti-rabbit F(ab')2
fragments, and anti-Na+-K+-ATPase labeling was
detected with tetramethylrhodamine isothiocyanate-conjugated goat
anti-mouse F(ab')2 fragments (Jackson ImmunoResearch Labs, West Grove, PA). For double-label experiments, primary antibodies were
applied simultaneously as were the fluorescent-conjugated secondary
antibodies. Fluorescent signals were visualized on an Axiophot
fluorescent microscope (Zeiss) and digitally stored using Photoshop
4.01 software (Adobe Systems, Mountain View, CA). Photoshop was not
used to modify images other than to adjust contrast for improved signal
definition. No signal was detected in the absence of primary antibody,
indicating that background labeling was low under the experimental
conditions employed (data not shown).
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RESULTS |
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Because there is no natural cell type that expresses both F508
and wild-type CFTR, transfected cell lines have served as the standard
model system for studying differences in the processing and trafficking
of CFTR. The most widely used cell lines (CHO, 3T3, C127, and HEK),
however, are either unpolarized or of undetermined polarity. Because
vesicle trafficking in polarized and unpolarized cells may differ, we
sought to examine CFTR surface expression in a transfected mammalian
cell line with evidence of cellular polarity. LLC-PK1 cells
are transformed epithelial cells from the proximal tubule of pig
kidney. Morphological evidence for polarity was obtained by electron
microscopy, which demonstrated the appearance of apical tight junctions
in monolayers of transfected cells (Fig.
1A). Functional polarity was
demonstrated by double-label immunofluorescent microscopy experiments
using apical and basolateral membrane markers. As shown in Fig.
1B, Na+-K+-ATPase expression in
LLC-PK1 cells is primarily surface and basal in location,
consistent with its known distribution along the basolateral membrane
of polarized epithelial cells. Although CFTR has both a surface and an
intracellular signal in these high-expressing cells, the surface signal
is more apically distributed and does not colocalize with that of
Na+-K+-ATPase. The prominent intracellular CFTR
signal seen in these cells is discussed in greater detail below. These
experiments indicate that CFTR-transfected LLC-PK1 cells
have both morphological and functional features characteristic of
polarized epithelial cells.
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For direct biochemical study of F508 expression at the plasma
membrane, sufficient quantities of the protein had to be driven to the
cell surface. To accomplish this, cells were treated at 27°C in the
presence of 5 mM sodium butyrate to markedly upregulate surface CFTR
expression (12). Previous time course experiments identified 60 h of treatment as optimal for the upregulation of CFTR expression (12). Although wild-type cells did not
require butyrate or low temperature treatment for the detection of
plasma membrane CFTR, we chose to control our experiments by treating all cells identically. As shown in Fig.
2, mature (band C)
F508 CFTR was not detected in LLC-PK1 cells grown at 37°C in
the presence or absence of sodium butyrate, nor was it detected by
immunoblot analysis in cells grown at 27°C alone. When sodium
butyrate and low-temperature treatments were combined, however, there
was a marked increase in total
F508 CFTR expression that was
accompanied by the appearance of the mature, fully glycosylated form
(band C). The upregulation of CFTR expression under these
experimental conditions was even more pronounced in wild-type
LLC-PK1 cells.
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Although mature (band C) CFTR has routinely been used as a
marker of plasma membrane CFTR expression (5, 8, 30),
studies in both native (26, 36) and transfected
(21) cells have shown that not all mature CFTR resides at
the cell surface. To confirm that F508 CFTR was driven to the
surface of LLC-PK1 cells by butyrate and low-temperature
treatment, plasma membrane CFTR expression was measured biochemically,
functionally, and cytochemically. Surface biotinylation and
streptavidin extraction were used to determine how much
F508 protein
actually reached the cell surface and what molecular form(s) of CFTR
were targeted there. As shown in Fig. 3,
most of the CFTR in treated
F508 and wild-type LLC-PK1 cells did not reach the cell surface. Based on replicate experiments (n = 5), ~35% of CFTR was found to reside at the
cell surface (33.9 ± 4.7% for
F508 and 35.5 ± 3.4% for
wild-type CFTR). These biotinylation experiments also demonstrated that
only the mature band C form of CFTR was expressed at the
plasma membrane. The immature band B form was unaffected by
streptavidin extraction, indicating that it was not biotinylated and
therefore not present at the cell surface. Surface expression of
F508 CFTR after butyrate and low-temperature treatment was
subsequently confirmed cytochemically and functionally (Fig.
4). The relative distribution of CFTR
between surface and intracellular compartments, including the prominent intracellular signal, correlated with the above biotinylation data.
Functional evidence for surface CFTR expression in both cell lines was
obtained by forskolin/IBMX-stimulated 125I efflux assay
(Fig. 4).
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With the use of the surface biotinylation procedure, the half-life of
plasma membrane CFTR in F508 and wild-type cells was examined next
(Fig. 5). In these experiments, surface
CFTR expression was upregulated by pretreatment with sodium butyrate
and low temperature. Cells were then transferred to physiological
temperature (37°C) and treated for up to 48 h in the presence of
cycloheximide. Cells were then surface biotinylated, solubilized, and
streptavidin-extracted, with immunoblot analysis being used to quantify
CFTR protein expression. The rapid fall in immature (band B)
CFTR expression in the
F508 cells was due to the inhibition of new
protein synthesis by cycloheximide, combined with ER degradation and
some conversion to the mature band C form. Although not
shown in Fig. 5, band B CFTR was also rapidly degraded in
wild-type cells (the signal was undetectable by 4 h, the earliest
time point we examined). These findings are consistent with the
observations of Ward and Kopito (34), who demonstrated
that the immature forms of
F508 and wild-type CFTR have similar
rates of degradation.
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In F508 cells, total band C CFTR expression was nearly
eliminated by 6 h. In contrast, total band C CFTR
expression in the wild-type cells persisted for 48 h. Although
most of the band C CFTR was intracellular, the persistence
of a streptavidin-extractable pool for as long as 48 h provides
biochemical evidence for the presence of plasma membrane CFTR
expression in wild-type cells for at least this length of time. Because
the relative distribution of CFTR between intracellular and plasma
membrane compartments did not change appreciably over time (data not
shown), the plasma membrane half-life of each protein was estimated by
quantifying the rate of degradation of total band C CFTR.
From replicate experiments (n = 7), we calculated the
biochemical half-life of plasma membrane CFTR in
F508 cells to be
~4 h, whereas the biochemical half-life of plasma membrane CFTR in
wild-type cells exceeded 48 h.
To rule out the possibility that the relatively long plasma membrane
half-life of wild-type CFTR was an artifact of overexpression, these
biochemical and functional studies were repeated under conditions of
comparable total CFTR expression in the two cells lines. Based on data
from Fig. 2, wild-type cells treated with sodium butyrate at 37°C had
CFTR expression levels that were comparable to those found in F508
cells treated with sodium butyrate at 27°C. Thus cells were
pretreated in this manner and then processed and analyzed as described
previously. Under these conditions of comparable total CFTR expression,
the biochemical half-life of band C CFTR in wild-type cells
continued to exceed 24 h. Surface biotinylation experiments also
confirmed the presence of plasma membrane CFTR for at least this length
of time (Fig. 6). This contrasts with the
near absence of any detectable
F508 protein after 6 h of chase.
Although we were able to detect some functional
F508 CFTR at these
early time points by the qualitative 125I efflux assay, no
functional CFTR expression was detected in
F508 cells after 24 h of chase, which contrasts sharply with the persistence of functional
CFTR in the wild-type cells at this time. Similar kinetic studies were
performed in transfected (unpolarized) C127 cells, where
F508
expression exceeds that of wild-type CFTR after butyrate and
low-temperature treatment, and similar half-life values for
F508 and
wild-type CFTR were obtained (data not shown). Thus several lines of
evidence indicate that the difference in plasma membrane expression of
F508 and wild-type CFTR reflects the biology of these two protein
species and not simply overexpression.
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DISCUSSION |
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Until the long-term goals of gene therapy are realized, strategies
designed to upregulate plasma membrane expression of F508 CFTR
remain an important potential therapeutic option in CF (16, 29). Much emphasis has been placed on the biogenesis of
F508 CFTR, with the goal being to overcome its biosynthetic arrest and
thereby increase the delivery of functional
F508 CFTR channels to
the cell surface. Effective correction of the CF defect, however, will
require stable expression of
F508 after reaching the plasma membrane, and one study has shown that the functional half-life of
plasma membrane
F508 in intact cells is less than that of wild-type
CFTR (23). The current study sought to test the hypothesis that the short functional half-life in
F508 cells results from more
rapid removal of CFTR channels from the cell surface.
To study the biochemical stability of plasma membrane F508 CFTR,
sufficient quantities of the mutant protein needed to be driven to the
cell surface. The current study took advantage of combination therapy
with sodium butyrate and low temperature to increase surface
F508
protein expression to levels sufficient for the direct measurement of
CFTR protein half-life by a quantitative immunoblot technique
(12). Thus, in all experiments, cells were pretreated with
sodium butyrate and low temperature to upregulate CFTR protein
expression and then were followed at 37°C to examine the plasma
membrane half-life of each CFTR species. Because cell polarity can
affect membrane trafficking, all experiments were performed in a
polarized epithelial cell line (LLC-PK1).
After butyrate and low-temperature treatment, most of the CFTR in these
cells was intracellular. Surface biotinylation experiments demonstrated
that only the mature, fully glycosylated (band C) form of
CFTR reached the cell surface. No evidence for the direct targeting of
immature (electrophoretic bands A and B) CFTR to the plasma membrane was found. Quantitative analysis of CFTR
distribution in butyrate/low-temperature-treated LLC-PK1
cells demonstrated that ~65% of band C CFTR remained
intracellular. This was true for both F508 and wild-type CFTR and is
similar to observations made in other cell systems (26,
36). Although changes in band C CFTR expression
correlated with changes in plasma membrane CFTR expression in
transfected LLC-PK1 cells, this may not be true in all cell
types. Thus surface protein labeling techniques remain the most
accurate means of measuring plasma membrane CFTR expression.
The biochemical finding of a prominent intracellular compartment of
CFTR was confirmed immunocytochemically. After butyrate/low-temperature treatment, both F508 and wild-type cells labeled similarly, with modest surface but prominent intracellular signals. The identity of the
intracellular signal in each cell type remains unknown. In the
wild-type cells, surface biotinylation demonstrated that most of
the unbiotinylated CFTR is of the band C form and therefore is presumably in a post-ER compartment (Golgi, endosomes, transport vesicles, etc.). In
F508 cells, where most of the unbiotinylated CFTR is of the band B form, a large ER component to the
intracellular signal would be expected. Identification of the sources
of intracellular CFTR labeling in these cells will, however, require
much additional study.
The kinetics of F508 and wild-type CFTR degradation over time was
examined at physiological temperature and under conditions of protein
synthesis inhibition. The rate of CFTR degradation was quantified
biochemically and confirmed by functional assay. Replicate experiments
indicate that the biochemical half-life of plasma membrane
F508 CFTR
is ~4 h, whereas the biochemical half-life of plasma membrane
wild-type CFTR exceeds 48 h. These values correlate with changes
in 125I efflux and are remarkably similar to the functional
data generated by Lukacs et al. (23) in nonpolarized C127
cells. This time-dependent correlation between the rate of plasma
membrane CFTR degradation and the loss of cAMP-stimulated
125I efflux indicates that the functional instability in
F508 cells can be theoretically attributed to more rapid degradation
of plasma membrane
F508 CFTR. The data, however, do not exclude a
contributory role for channel inactivation in this process.
These findings establish that the F508 mutation has a negative
effect on the biochemical stability of CFTR at the cell surface. This
biochemical instability must be due to more rapid internalization of
mutant protein and/or its selective targeting for rapid degradation. CFTR is endocytosed in clathrin-coated vesicles (1, 22), and the molecular signal for CFTR internalization may reside in its
cytoplasmic tail (10, 24, 25). The
F508 mutation,
however, is in the first nucleotide-binding domain and is some distance from the cytoplasmic end of the molecule. Thus CFTR must have another
internalization signal, or the
F508 mutation must have an indirect
effect on the COOH-terminal signal(s). Based on our understanding of
CFTR folding during biogenesis, it is attractive to hypothesize that
the
F508 mutation affects CFTR folding in such a manner that the
COOH-terminal internalization signal is altered. The fundamental
question is what is the signal? COOH-terminal tyrosine-based sequences
have been implicated in the positioning of membrane proteins in coated
pits (7, 20), and one recent study has implicated
phosphorylation of tyrosine-1424 in the regulation of CFTR endocytosis
(25). On the other hand, the rate of internalization of
Ste6, a yeast homolog of CFTR, and Ste3p, another yeast membrane protein, are regulated by ubiquitination (14, 27). Because ubiquitination is the major signal for the degradation of both wild-type and
F508 CFTR during biogenesis (35), it is
provocative to speculate that it is also involved in plasma membrane
CFTR degradation. During CFTR biosynthesis, ubiquitination targets the
immature forms of both
F508 and wild-type CFTR to rapid proteosome degradation with similar kinetics (34). A pool of
wild-type CFTR, however, escapes this fate, becomes fully glycosylated, and reaches the plasma membrane. If ubiquitination was also the signal
for plasma membrane CFTR degradation, one would have to postulate that
the wild-type and mutant proteins were differentially ubiquitinated at
the cell surface to account for their different rates of degradation.
Because the yeast data suggest that the proteasome complex is not the
final target of ubiquitinated plasma membrane Ste6 (15),
one must also consider the possibility that ubiquitination of plasma
membrane CFTR serves a different role (e.g., signaling for
internalization and/or sorting rather than proteosome degradation).
Although provocative, a role for ubiquitination in the downregulation
of surface CFTR expression remains highly speculative at this time.
Alternatively, mutant and wild-type CFTR may have similar rates of
internalization but are differentially sorted in the endocytic compartment, with F508 being targeted for rapid lysosomal
degradation while wild-type CFTR is recycled back to the plasma
membrane. In this case, the
F508 mutation might be affecting the
interaction of CFTR with specific GTP-binding (Rab) proteins that
regulate vesicle trafficking (reviewed in Ref. 3). Rab4
has been most closely associated with sorting endosomes, so an
understanding of its interactions with
F508 and wild-type CFTR would
be of particular interest. Last, one must consider the effect of
phosphorylation on CFTR trafficking, particularly in light of data
showing that the rate of endocytosis differs in
F508 and wild-type
cells after cAMP activation (2). Although the current
study did not examine the biochemical stability of plasma membrane CFTR
under stimulatory conditions, the basal phosphorylation state of these
two CFTR species may differ, and this, in turn, could expose different internalization signals.
In conclusion, the F508 mutation dramatically reduces the residence
time of CFTR at the cell surface. This must be due to an effect of the
mutation on the rate of CFTR internalization and/or its intracellular
targeting. Although little is known about this aspect of CFTR biology,
it has clear implications for any therapeutic strategies that rely on
delivering more
F508 protein to the cell surface.
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ACKNOWLEDGEMENTS |
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We thank Dr. Raymond A. Frizzell for helpful discussions and acknowledge the technical assistance of Virginia Jeanes.
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FOOTNOTES |
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This work was supported by the Office of Research and Development, Medical Research Service, Department of Veterans Affairs (C. R. Marino) and by the Cystic Fibrosis Foundation (C. R. Marino).
Address for reprint requests and other correspondence: C. R. Marino, Medical Service (111), VA Medical Center, 1030 Jefferson Ave., Memphis, TN 38104 (E-mail: cmarino{at}utmem.edu).
The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
Received 24 May 2000; accepted in final form 9 August 2000.
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