1 Department of Molecular Cell Biology, Weizmann Institute of Science Rehovot
76100 Israel
2 Sigma Aldrich Israel, Park Rabin, Rehovot 76100, Israel
* Present address: The Department of Ornamental Horticulture, The Volcani
Center, Beit-Dagan, P.O.B. 6, 50250 Israel
Author for correspondence (e-mail:
benny.geiger{at}weizmann.ac.il
)
Accepted 23 April 2002
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Summary |
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Key words: ß-catenin, GSK-3ß, APC
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Introduction |
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Wnt signaling is essential for proper embryonal development. In
Xenopus embryos, ectopic expression of Wnt can induce secondary axis
formation (Sokol, 1999). In
the fruit fly, the homologous wingless pathway is involved in the
establishment of segment polarity, wing formation and differentiation of the
endoderm (Cadigan and Nusse,
1997
). In mice, the targeting of different wnt isoforms leads to
different phenotypes; Lack of Wnt 1 results in the deletion of part of the
midbrain (McMahon and Bradley,
1990
), the ablation of Wnt-4 affects the kidney
(Stark et al., 1994
), Wnt 7a
affects limb development (Parr and
McMahon, 1995
) and wnt3 knockout mice are deficient in
the formation of the anterior-posterior axis
(Liu et al., 1999
).
In adult tissues, components of the Wnt signaling pathway, such as
ß-catenin and APC, regulate cell proliferation in epithelial cells lining
the colon (Polakis, 1999;
Polakis, 2000
). Mutations that
perturb the function of the ß-catenin degradation complex, such as
truncation of APC or mutations in the GSK-3ß phosphorylation sites of
ß-catenin, are present in 90% of colon cancers and in other types of
tumors (Polakis, 2000
).
Although phosphorylation of ß-catenin by GSK-3ß plays a pivotal role
in regulating the fate and activity of ß-catenin, the properties of the
phosphorylated intermediate of ß-catenin have not been directly
characterized yet. In this study we used a novel antibody that specifically
recognizes S33/S37 phosphorylated ß-catenin to study the kinetics of its
phosphorylation, dephosphorylation and ubiquitination in normal and colon
cancer cells, and we determined its subcellular distribution. We show that
normal cells contain very low levels of pß-catenin (<1-2% of the
protein), but the levels of pß-catenin increase upon overexpression of
the protein or after blocking proteasomal degradation. pß-catenin can be
dephosphorylated by an as yet unknown phosphatase at a rate comparable to its
GSK-3ß-mediated phosphorylation in cells overexpressing ß-catenin.
pß-catenin accumulates mainly in the nucleus but fails to form a ternary
complex with LEF-1 and DNA and is not associated with AJ, except shortly after
junction formation. We further show that colon cancer cells expressing
different APC mutants differ in their phosphorylated ß-catenin levels,
suggesting that APC, truncated at position 1338 (as in SW480 cells), can still
support phosphorylation but fails to promote degradation whereas truncation of
APC at position 1555 (as in HT29 cells) allows limited phosphorylation and
degradation of ß-catenin.
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Materials and Methods |
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Cells and transfections
Bovine endothelial cells (BCAP), MDCK, 293T, Rat1 and Rat1 ras and the
human colon carcinoma cell lines SW480, HCT116 and HT29 were cultured in
Dulbecco's modified Eagle's medium (DMEM) supplemented with 10% fetal calf
serum (FCS). For transfection, cells were plated to form a 50%-70% confluent
culture in 30 mm dishes. MDCK cells were transfected using lipofectamine
(GIBCO BRL, Rockville USA), and 293T cells were transfected by the calcium
phosphate method. Treatments with 30 mM LiCl or 25 µM MG132 were for the
time periods indicated.
Plasmids
Plasmids expressing HA-ß-catenin, HA-S33Y ß-catenin,
VSV-ß-catenin and GFP-ß-catenin were as described previously
(Shtutman et al., 1999;
Simcha et al., 1998
;
Zhurinsky et al., 2000b
).
HA-ß-catenin S37A was a kind gift from S. Byers
(Orford et al., 1999
). The
chicken N-cadherin and
F-ß-TrCP expressing plasmids were
previously described (Sadot et al.,
1998
; Sadot et al.,
2000
).
Protein analysis
Protein levels were monitored by western blotting. The following antibodies
were used: monoclonal anti-phosphorylated-ß-catenin and polyclonal
anti-ß-catenin were from Sigma (Rehovot, Israel); monoclonal
anti-ß-catenin (clone 14c19220) was from Transduction Laboratories
(Lexington, KY). A monoclonal anti-HA antibody (clone 12CA5) was from Roche
(Germany) and polyclonal anti-HA (Y11 sc-805) was from Santa Cruz
Biotechnology (Santa Cruz, CA). Western blots were developed using the ECL
method (Amersham UK). Autoradiograms were scanned by a GS-700 imaging
densitometer (Bio Rad Laboratotries, Hercules, CA) using the FotoLook PS
2.07.2 software. The intensity of the bands was quantified using the NIH image
1.61 software.
Immunofluorescence microscopy
Cells were cultured on glass coverslips, fixed with 3% paraformaldehyde in
phosphate-buffered saline (PBS) and permeabilized with 0.5% Triton X-100. The
coverslips were incubated with the primary antibodies as described above. The
secondary antibodies were Alexa-488-conjugated goat anti-mouse or anti-rabbit
IgG (Molecular Probes, Eugene, OR) and Cy3-conjugated goat anti-mouse or
anti-rabbit IgG (Jackson ImmunoResearch Laboratories West Grove PA). Images
were acquired using the DeltaVision system (Applied Precision, Issaqua, WA)
equipped with a Zeiss Axiovert 100 microscope (Oberkochen Germany) and
Photometrics 300 series scientific-grade cooled CCD camera (Tucson, AZ),
reading 12 bit images, and using a x100/1.3 NA plan-Neofluar objective.
For quantitative image processing, the Priism software was employed
(Kam et al., 1993). Ratio
imaging analysis was done as described previously
(Zamir et al., 1999
).
Nuclear extracts and DNA mobility gel shift analysis
293T cells grown in 90 mm diameter dishes were transfected with 2 µg of
LEF-1 and 8 µg of ß-catenin expression plasmids or with the control
pCIneo vector. 36 hours after transfection, nuclear extracts were prepared as
previously described (Shtutman et al.,
1999). Briefly, cells were incubated for 15 minutes in low-salt
buffer, then NP-40 was added, nuclei were pelleted by centrifugation, and
nuclear proteins were extracted with high-salt buffer at 4°C. Protein
concentrations were determined using the bicinchoninic acid protein assay
reagent (Pierce) and bovine serum albumin was used as a standard. For DNA
binding assays, 6 µg of nuclear extracts were used. 1 µg of antibody was
added to the binding reactions for analyzing the DNA mobility supershift.
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Results |
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To determine the specific interaction of the antibody with full-length
ß-catenin, 293T cells were transfected with wild-type ß-catenin or
with the two corresponding ß-catenin mutants, S33Y
(Simcha et al., 1998) and S37A
(Orford et al., 1999
), which
were tagged with HA. Although the expression of the wild-type and mutant
ß-catenins were comparable (Fig.
2A, bottom panel), wild-type ß-catenin was recognized by the
pß-catenin antibody, whereas the S33Y mutant reacted only weakly,
and the S37A mutant was not recognized at all
(Fig. 2A, upper panel). To test
what fraction of ß-catenin is phosphorylated, 293T cells were transfected
with ß-catenin together with HA-tagged
F-ß-TrCP that can bind
to phosphorylated ß-catenin (Hart et
al., 1999
; Sadot et al.,
2000
) but is unable to promote its ubiquitination and degradation.
pß-catenin was then immunoprecipitated from the transfected cells as a
complex with
F-ß-TrCP using anti-HA antibodies and immunoblotted
with either the
pß-catenin antibody or with a general antibody
against ß-catenin. Comparison of the labeling intensities with the two
antibodies was used for calibration (Fig.
2B). These experiments indicated that in
ß-catenin-overexpressing 293T cells, 35-55% of the transfected
ß-catenin molecules were phosphorylated
(Fig. 2B).
|
The endogenous ß-catenin of 293T cells was not recognized by the
pß-catenin antibody, but treatment with the proteasomal inhibitor
MG132 resulted in a dramatic increase in pß-catenin levels
(Fig. 2C, lane 3 upper panel)
and in only a two-fold increase in the levels of total ß-catenin
(Fig. 2C, lane 3 lower panel).
Treatment with LiCl (which inhibits GSK-3ß activity) induced an
accumulation of total ß-catenin (comparable to that found in
MG132-treated cells), yet the accumulated protein was not phosphorylated
(Fig. 2C, lane 2 upper panel).
Taken together these results suggest that the
pß-catenin antibody
is specific to the S33/S37 phosphorylated ß-catenin and that transfected
ß-catenin accumulates as a phosphorylated molecule
(Fig. 2A, lane 1). In
non-transfected cells, pß-catenin is a short lived molecule
(Fig. 2C, lane 1, compare with
Fig. 2A, lane 1 upper panel and
see below).
Interestingly, the pß-catenin antibody did not recognize
transfected plakoglobin in 293T cells (Fig.
2D, compare lane 3 with 5) despite the high sequence homology with
ß-catenin in the GSK-3ß phosphorylation domain
(Fig. 1A). To rule out the
possibility that
pß-catenin antibody does not recognize
transfected plakoglobin because it is not phosphorylated, we
co-immunoprecipitated the phosphorylated form of plakoglobin with
F-ß-TrCP. As shown in Fig.
2D (lanes 7 and 8),
F-ß-TrCP-bound plakoglobin (which
is most probably phosphorylated on the corresponding serine residues) is not
recognized by the
pß-catenin antibody. It is worth noting that
there is an increase in endogenous pß-catenin in the presence of excess
plakoglobin (Fig. 2D, compare
lane 3 with lane 1). This is in agreement with previous reports showing that
overexpression of plakoglobin attenuates the degradation of ß-catenin
(Miller and Moon, 1997
;
Simcha et al., 1998
;
Zhurinsky et al., 2000a
), but,
as shown here, not its phosphorylation.
The dynamics and fate of phosphorylated ß-catenin
In the presence of Wnt signaling, ß-catenin phosphorylation is
inhibited, thus leading to a reduction in its poly-ubiquitination and
proteasomal degradation (Polakis,
2000). By an alternative process, ß-catenin could be
dephosphorylated by a phosphatase. To test whether dephosphorylation of
ß-catenin takes place in live cells, GFP-ß-catenin was transfected
into 293T cells, and 24 hours later, proteasomal degradation was blocked by
MG132 for 1, 2 and 4 hours in the presence or absence of LiCl to block
GSK-3ß activity. After 2 hours with MG132 and LiCl, the medium was
replaced with fresh medium containing only MG132. After incubation for
different periods of time (from 10 to 120 minutes), the level of
pß-catenin was determined. As shown in
Fig. 3A after 10 minutes of
treatment with LiCl, a dramatic drop (<50%) in the content of
pß-catenin was observed (Fig.
3A, lane 5 upper panel). Since proteasomal activity was blocked by
MG132 during this time and no reduction in total ß-catenin was observed
(Fig. 3A, lower panel), we
propose that the reduction in pß-catenin results from its
dephosphorylation rather than its degradation. Removal of LiCl was followed by
a rapid increase in pß-catenin levels
(Fig. 3A, lanes 9-12, upper
panel). To determine the phosphorylation-dephosphorylation kinetics in
non-transfected cells, 293T cells were treated with MG132 for different time
periods to induce accumulation of pß-catenin
(Fig. 3B, lanes 1-5). The
levels of pß-catenin increased linearly up to four-fold after 5 hours of
MG132 treatment (Fig. 3B,C).
Western blotting for total ß-catenin
(Fig. 3B lanel 1-5 lower panel)
showed an increase in the poly-ubiquitinated protein. To determine whether
dephosphorylation of pß-catenin takes place, 293T cells were first
treated with MG132 for 2 hours, and then LiCl was added for different periods
ranging from 10 minutes to 2 hours. The addition of LiCl resulted in a rapid
decline in pß-catenin level (a decrease of
60% within 10 minutes,
Fig. 3C) and after 30-60
minutes with LiCl essentially no pß-catenin was detected
(Fig. 3B, lanes 6-9). It is
noteworthy that only a limited decrease in total ß-catenin was detected
following this treatment (Fig.
3B, lower panel). To compare the rate of ß-catenin
dephosphorylation (in the presence of LiCl) to its rate of phosphorylation,
cells treated with MG132 and LiCl (Fig.
3B lane 9) were washed and further incubated, in the absence of
LiCl, for 10-60 minutes. As shown in Fig.
3B (lanes 10-12), removal of LiCl resulted in the fast recovery of
pß-catenin levels at a rate similar to the decline observed after
addition of LiCl (note that MG132 was present throughout this treatment to
avoid proteasomal degradation of the protein).
|
The molecular partners of phosphorylated ß-catenin
Next, we asked whether pß-catenin can interact with ß-catenin's
transcriptional (LEF-1) and junctional (cadherin) partners. HA-LEF-1 and
VSV-ß-catenin were co-transfected into 293T cells, immunoprecipitated
with HA antibody and immunobloted with
pß-catenin. As shown
in Fig. 4A (upper panel),
pß-catenin could form a complex with LEF-1. We next examined whether
pß-catenin can form a ternary complex with LEF-1 and DNA. First we
verified that the
pß-catenin antibody effectively
immunoprecipitated pß-catenin from cell extracts in the buffer used for
DNA mobility gel shift assays (Fig.
4B). Then, a gel shift analysis was performed with a radiolabelled
probe corresponding to the consensus DNA sequence of the LEF/TCF-binding site
incubated with nuclear extracts from 293T cells transfected with LEF-1 and
ß-catenin. As shown in Fig.
4C, although a general antibody against ß-catenin
supershifted the ternary complex of DNALEF-1ß-catenin
(Fig. 4C, lane 5), the
pß-catenin antibody was unable to do so (lane 6). This suggests
that pß-catenin is inefficient in forming a ternary complex with LEF-1
and DNA.
|
Interestingly, pß-catenin (unlike its non-phosphorylated counterpart)
was also inefficient in associating with transfected N-cadherin
(Fig. 5 lane 5) compared with
its association with F-ß-TrCP
(Fig. 5 lane 6), whereas the
levels of both total and pß-catenin were elevated in
N-cadherin-transfected cells (Fig.
5 lane 2). This suggests that pß-catenin is normally not
associated with N-cadherin.
|
The subcellular distribution of phosphorylated ß-catenin
Immunofluorescence labeling with pß-catenin from different
cultured cells, including MDCKs (epithelial), BCAPs and PEACs (endothelial),
as well as Rat1 and Rat1 ras (fibroblasts), was largely negative. However, the
inhibition of proteasomal degradation with MG132 in BCAP cells
(Fig. 6A), or after
overexpression of ß-catenin in MDCK cells
(Fig. 7a), resulted in the
accumulation of pß-catenin in the nuclei of these cells. Examination of
the transfected MDCK cells using three dimensional ratio imaging
(Fig. 7a) indicated that the
transfected ß-catenin (visualized using regular
ß-catenin
antibody) is associated with both adherens junctions and the nucleus (blue in
the ratio image), whereas the pß-catenin is exclusively nuclear
(Fig. 7a, yellow in the ratio
image) and was absent from cell-cell junctions. To verify that the absence of
junctional labeling for pß-catenin indeed resulted from the absence of
the phosphorylated molecule we subjected the cells to different
permeabilization procedures, none of which revealed junctional labeling (data
not shown), indicating that the lack of antibody accessibility is unlikely.
Interestingly, we found that pß-catenin is transiently localized in newly
formed adherens junctions of BCAP cells
(Fig. 7b). Thus, adherens
junctions of BCAP cells, formed within 10 hours of plating
(Fig. 7bA) or after 3 hours
recovery from treatment with EGTA (Fig.
7bF) were labeled with
pß-catenin, unlike those
stained after 20-30 hours of incubation
(Fig. 7bB,C). This fluorescence
labeling could be readily inhibited by the synthetic, doubly phosphorylated
ß-catenin peptide (insert, Fig.
7bA). These results suggest that pß-catenin can be
incorporated into newly formed, but not mature, junctions and that the
junction-bound phosphorylated protein is either turning over after a longer
time or undergoing dephosphorylation.
|
|
Phosphorylation of ß-catenin in colon cancer cells
The majority of colon tumors contain APC mutations, which are characterized
by large deletions in the C-terminus of the protein
(Polakis, 2000). This region
is involved in ß-catenin and axin binding, and its deletion leads to the
accumulation of ß-catenin. Using the novel
pß-catenin
antibody we determined the levels of pß-catenin before and after blocking
proteasomal degradation in several colon cancer cell lines. HT29 (APC
truncation at position 1555), SW480 (APC truncation at position 1338) and
HCT116 (wild-type APC and
Ser45 ß-catenin) were compared with 293T
cells containing wild-type APC and ß-catenin. The levels of
ß-catenin were found to be higher in the three colon cancer cell lines
compared with 293T cells, as expected (Fig.
8, lane 1, compare lane 3 with 5 and 7). Interestingly, blocking
proteasomal degradation resulted in a five-fold increase in the content of
pß-catenin in HT29 and HCT116 (Fig.
8, compare lanes 3 to 4 and 7 to 8), whereas a more than 50-fold
increase was observed in 293T cells (Fig.
8, lanes 1 and 2). This suggests that the mutant APC in HT29 is at
least partially capable of supporting degradation of ß-catenin. In HCT116
cells the pß-catenin may be derived from the wild-type molecule generated
from the intact allele (Morin et al.,
1997
). In addition, a high content of pß-catenin (20% of the
total ß-catenin) was observed in SW480 cells. This is in agreement with a
previous observation that ß-TrCP can bind to ß-catenin from SW480
cells in the absence or presence of a proteasomal inhibitor
(Hart et al., 1999
). This
suggests that the relatively high amounts of ß-catenin accumulating in
SW480 cells might saturate the ß-catenin degradation machinery or that
the region between amino acid 1338 (SW480) and 1555 (HT29) in the APC molecule
may play an important role in the degradation or dephosphorylation of
pß-catenin.
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Discussion |
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An essential pre-requisite for addressing these questions was to obtain a
highly specific antibody (pß-catenin) for tracing the
phosphorylated molecule. Our study indicates that this antibody interacts
primarily with the doubly phosphorylated ß-catenin, either the endogenous
or when transfected, tagged (with HA, VSV or GFP) or untagged. ß-catenin
phosphorylated at position S37 showed low reactivity with
pß-catenin, whereas ß-catenin phosphorylated on S33 and the
closely homologous protein plakoglobin were negative. It was thus concluded
that
pß-catenin is highly specific for pß-catenin on both S33
and S37.
Examination of different epithelial, endothelial and mesenchymal cell lines indicated that all, except for the colon cancer cell line SW480 (see below), contained undetectable levels of phosphorylated ß-catenin. Transfection with ß-catenin cDNA or inhibition of proteasomal degradation resulted in a fast and extensive increase in pß-catenin levels. Interestingly, according to its electrophoretic mobility, the phosphorylated protein is largely non-polyubiquitinated, suggesting that either phosphorylation by GSK-3ß is considerably more efficient than the subsequent step in ß-catenin turnover, namely ubiquitination, or that excess ß-catenin specifically attenuates the ubiquitination process.
As pointed out above, among the cells tested for pß-catenin, the colon
cancer SW480 cell line was exceptional as it accumulated pß-catenin. This
result shed new light on the differential regulation by APC of the
phosphorylation of ß-catenin and its consequent degradation or
dephosphorylation. It is well established that the tumor suppressor protein
APC and the associated protein axin/conductin provide the scaffold for the
phosphorylation of ß-catenin
(Polakis, 2000) and that
ß-TrCP is present in a complex with axin and APC
(Kitagawa et al., 1999
). It is
not clear, however, whether APC is playing a role in the actual ubiquitination
and degradation of ß-catenin. Mutations in APC, which occur in over 80%
of human colon cancers, are believed to lead to the accumulation of
ß-catenin and presumably to the activation of specific genes that confer
malignant properties on these cells, for example, Myc
(He et al., 1998
) and cyclin D1
(Shtutman et al., 1999
;
Tetsu and McCormick, 1999
). It
was previously demonstrated that deletion mutants of APC lacking amino acids
1941 to 2644 could not inhibit ß-catenin-mediated transactivation in
SW480 cells (Morin et al.,
1997
), suggesting that this domain of APC is necessary for
ß-catenin downregulation (Morin et
al., 1997
). The results presented here indicate that
phosphorylation and ubiquitination of ß-catenin may be differentially
regulated by distinct regions of the APC molecule. Thus, truncation of the APC
molecule at position 1338 (like in SW480 cells) can support some
phosphorylation, but such APC lacks the region that is necessary for promoting
the degradation of ß-catenin. Alternatively, these cells may be deficient
in ß-catenin dephosphorylation, resulting in the accumulation of
pß-catenin. APC truncated at a more C-terminal position (1555 as found in
HT29 cells) can support some phosphorylation, although at a lower level,
compared with wild-type APC in 293T cells, but the subsequent ubiquitination
and degradation of the molecule are efficient, since the phosphorylated
molecule is detected in these cells only after blocking proteasomal
degradation. It thus appears that the region between amino acids 1338 and 1555
of the APC containing the second 20 amino acid repeat region is particularly
important for the turnover of pß-catenin by either promoting degradation
or affecting its dephosphorylation.
A related intriguing observation, highlighted by this study, is the
capacity of pß-catenin to undergo rapid dephosphorylation and thus avoid
proteasomal degradation. According to the common view, ß-catenin
phosphorylation is largely a unidirectional process, leading to the
ubiquitination and degradation of the protein. To test whether ß-catenin
phosphorylation is a reversible process, we blocked the phosphorylation of
ß-catenin by LiCl and its proteasomal degradation by MG132, and monitored
the changes in pß-catenin levels with the pß-catenin
antibody. Such treatment resulted in a rapid decline in pß-catenin levels
(t1/2=10 minutes), suggesting that the molecules can undergo rapid
dephosphorylation. The rate of ß-catenin dephosphorylation was comparable
to that of its phosphorylation, which was an approximately seven-fold decrease
or increase in phosphorylated ß-catenin after 1 hour of treatment. The
regulation of ß-catenin levels was commonly attributed to the activation
of Wnt signaling, which includes the inhibition of GSK-3ß activity that
leads to ß-catenin accumulation
(Zhurinsky et al., 2000b
). The
results described in this study suggest that dephosphorylation of
ß-catenin might be an alternative pathway that can protect ß-catenin
and induce its accumulation, a process equivalent to the activation of Wnt
signaling. Whether such process indeed takes place, and what might be the
exact activation mechanism, remains to be determined. Effects of
dephosphorylation by protein phosphatase 2A (PP2A) in the degradation complex
of APCAxinGSK3ßß-catenin were previously
suggested. PP2A was found to dephosphorylate axin following Wnt signaling,
thereby releasing ß-catenin from the degradation complex
(Willert et al., 1999
). On the
other hand, it has been shown that PP2A can promote the degradation of
ß-catenin and inhibit Wnt signaling
(Li et al., 2001
;
Seeling et al., 1999
;
Yamamoto et al., 2001
). A
dephosphorylation activity, directed to the ß-catenin molecule itself,
has not yet been described.
Examination of the effect of ß-catenin phosphorylation on the
interaction with its different molecular partners revealed some intriguing
features. As is evident from coimmunoprecipitation experiments, phosphorylated
ß-catenin binds to LEF1, yet it apparently fails to efficiently form a
ternary complex with DNA and is thus probably transcriptionally inactive. The
basis for these properties of pß-catenin is not clear, but it appears
likely that phosphorylation may affect its transactivation capacity. This
notion is supported by the observation that mutations in the putative
GSK-3ß phosphorylation site of ß-catenin increase its signaling
activity when compared with equal amounts of wild-type ß-catenin
(Guger and Gumbiner, 2000). In
addition a recent finding demonstrated that phosphorylated ß-catenin,
which accumulates after blocking proteasomal degradation, is unable to
activate a synthetic LEF/TCF reporter (TOPFLASH), whereas non-phosphorylated
ß-catenin accumulating after Wnt or LiCl treatments is transcriptionally
active (Staal et al., 2001
).
This may suggest that phosphorylation of the N-terminal serines of
ß-catenin have a regulatory role in ß-catenin signaling in addition
to its effect on protein stability.
Another intriguing feature of pß-catenin is its conspicuous absence
from mature cell-cell adherens junctions; yet it has a transient association
with newly formed adhesions. The latter observation suggests that
pß-catenin has the intrinsic capacity to interact with the various
junctional partners (e.g. cadherin and/or -catenin), yet these
interactions can take place only during early stages of junction formation
(first several hours after cell plating, or EGTA treatment), when presumably
new cytoplasmic ß-catenin molecules are recruited to the membrane, but
not at later stages, when the turnover of the recruited ß-catenin
molecules is rather limited. It is not surprising that GSK-3ß-mediated
phosphorylation does not occur in adherens junctions since ß-catenin
interacts with junctional cadherin via its arm repeats that are also involved
in its association with APC that is essential for ß-catenin
phosphorylation (Hulsken et al.,
1994
). The subcellular distribution of the ß-catenin
destruction complex, including APC, axin, GSK-3ß and dishevelled in
polarized epithelial cells, is mostly cytoplasmic
(Reinacher-Schick and Gumbiner,
2001
). In addition, an association of PP2A with the
cadherinß-catenin complex was reported
(Gotz et al., 2000
), suggesting
that even when pß-catenin forms a complex with cadherin it may be prone
to dephosphorylation activity. The putative role and significance of this
association of pß-catenin with newly formed junctions is still to be
determined. The transient localization of ß-catenin in junctions or its
accumulation in the nuclei may reflect a compartmentalized shelter from
degradation, whose physiological significance is also unclear at present.
Taken together the observations of this study suggest that pß-catenin is a short lived, transcriptionally inactive intermediate, with a half life determined by the finetuned balance between phosphorylation, dephosphorylation, ubiquitination, proteasomal degradation and subcellular localization.
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