1 Abteilung Stammzellbiologie, CMPB, Georg-August-Universität
Göttingen, Justus-von-Liebig-Weg 11, 37077 Göttingen, Germany
2 Institut für Genetik, Heinrich-Heine-Universität Düsseldorf,
Universitätsstr. 1, 40225 Düsseldorf, Germany
3 Institut für Neurobiologie, Universität Münster, Badestr. 9,
48149 Münster, Germany
* Author for correspondence (e-mail: wodarz{at}uni-duesseldorf.de)
Accepted 31 January 2005
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SUMMARY |
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Key words: Cell polarity, PAR proteins, PTEN, Actin cytoskeleton, Drosophila
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Introduction |
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The architecture of these different cell types is very diverse, raising the
question of how the PAR/aPKC complex communicates with downstream effector
molecules that specifically control different aspects of cell polarity,
including cell shape, the type and position of intercellular junctions and the
asymmetric localization of cortical and transmembrane proteins. One attractive
possibility would be that the PAR/aPKC complex affects the organization of the
submembrane cytoskeleton, which could have an impact on every aspect of
polarity mentioned above. Indeed, it has been shown that, depending on the
cell type, both actin and microtubules are involved in the control of cell
polarity (Broadus and Doe,
1997; Clark et al.,
1997
; Wallenfang and Seydoux,
2000
). However, except for the fact that the small GTPase Cdc42 is
a potent regulator of the actin cytoskeleton, no direct link between
components of the PAR/aPKC complex and cytoskeletal regulators has been
uncovered so far.
In order to find molecules that bind to Baz/PAR-3 and that may provide a
link between the PAR/aPKC complex and the cortical cytoskeleton, we have
performed a yeast two-hybrid screen using the three PDZ domains of Baz as
bait. We isolated three independent clones of the lipid phosphatase PTEN that
specifically bound to Baz. PTEN catalyzes the dephosphorylation of
phosphoinositide lipids at the D3 position of the inositol ring
(Leslie and Downes, 2002). One
substrate of particular importance is the lipid phosphatidylinositol (3,4,5)
trisphosphate [PtdIns(3,4,5)P3], which is converted by the
activity of PTEN to phosphatidylinositol (4,5) bisphosphate
[PtdIns(4,5)P2]. PtdIns(3,4,5)P3 is
produced by activation of phosphatidylinositol 3-kinase (PI3-kinase) in
response to stimulation by a multitude of growth factors and cytokines.
Interestingly, PtdIns(3,4,5)P3 locally activates Cdc42 by
recruitment of guanine nucleotide exchange factors (GEFs) that promote the
exchange of GDP for GTP specifically on Cdc42
(Zheng, 2001
). Moreover, in
mammalian cells PtdIns(3,4,5)P3 recruits phosphoinositide
dependent kinase 1 (PDK1; Pk61C FlyBase) which activates aPKC by
direct phosphorylation of a conserved threonine residue in the activation loop
of the kinase (Le Good et al.,
1998
). Thus, PtdIns(3,4,5)P3 is likely to
activate two components of the PAR/aPKC complex, Cdc42 and aPKC. Because PTEN
is predicted to antagonize the activation of both Cdc42
(Liliental et al., 2000
) and
aPKC by lowering the level of PtdIns(3,4,5)P3 in the
plasma membrane, the association of PTEN with Baz may have a significant
impact on the activity of these two key components of the PAR/aPKC
complex.
Recently, PI3-kinase signaling and PTEN have been implicated in the
polarization of Dictyostelium amoebae in response to a source of
chemoattractant (Funamoto et al.,
2002; Iijima and Devreotes,
2002
). PI3-kinase and PTEN are localized to the leading edge and
uropod, respectively, in a very dynamic fashion. PI3-kinase signaling also
appears to be required for directed migration of leukocytes
(Servant et al., 2000
;
Wang et al., 2002
). In both
cases, PI3-kinase and PTEN are thought to participate in a self-sustaining
loop that intracellularly amplifies the shallow concentration gradient of the
chemoattractant. PI3-kinase and PTEN also affect the polarization of
hippocampal neurons in culture and, more specifically, the localization of
PAR-3 and aPKC to the tip of the neurite that is going to become the axon
(Shi et al., 2003
;
Jiang et al., 2005
). Thus,
there is increasing evidence that PTEN and the PAR/aPKC complex may cooperate
in the control of cell polarity.
In Drosophila, the function of PTEN has mainly been studied with
respect to its role in the regulation of growth and proliferation in larval
and adult tissues (Stocker and Hafen,
2000). Pten mutant cells have elevated
PtdIns(3,4,5)P3 levels and are larger than wild-type cells
owing to increased growth (Goberdhan et
al., 1999
; Huang et al.,
1999
; Gao et al.,
2000
; Stocker et al.,
2002
). Clones of Pten mutant cells in imaginal discs also
show subtle defects in the organization the actin cytoskeleton
(Goberdhan et al., 1999
).
Pten interacts genetically with components of the insulin signaling
pathway including the insulin receptor, the insulin receptor substrate
IRS-1/Chico, PI3-kinase and protein kinase B (PKB)
(Goberdhan et al., 1999
;
Huang et al., 1999
;
Gao et al., 2000
;
Oldham et al., 2002
;
Stocker et al., 2002
). These
findings provided solid evidence for an antagonistic relationship between PTEN
and PI3-kinase and showed that the regulation of phosphoinositide levels is
the main vital function of PTEN.
Here, we show that PTEN directly binds to Baz/PAR-3 and colocalizes with Baz in the apical cortex of epithelia and neuroblasts. Pten mutant embryos lacking maternal and zygotic Pten function show defects during early embryonic development that point to a function for Pten in the organization of the actin cytoskeleton. Removal of Pten function from the germline in ovaries causes abnormal actin organization in nurse cells and in the oocyte. We propose that recruitment of PTEN by Baz may contribute to the polarization of the actin cytoskeleton, most likely by creating local differences in the balance between PtdIns(3,4,5)P3 and PtdIns(4,5)P2 in the plasma membrane. Moreover, PTEN may affect the activity of two key components of the PAR/aPKC complex, aPKC and Cdc42. The binding of PTEN to Baz provides the first molecular link between the PAR/aPKC complex, the actin cytoskeleton and phosphoinositide signaling.
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Materials and methods |
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Fly stocks, antibodies and immunohistochemistry
We used the amorphic allele PtenDJ189
(Gao et al., 2000) and the
phosphatase dead allele Ptenc494
(Huang et al., 1999
).
Germ-line clones of both alleles were produced in females of the genotype
y w P[ry+ FLP]12; Pten
P[hs neo ry+
FRT]2L-40A/P[mini w+
ovoD1]2L13X13 P[hs neo
ry+ FRT]2L-40A. To mark germ-line clones
in ovaries by absence of GFP expression, P[w+mC
Ubi-GFP(S65T)nls}2L P[hs neo ry+
FRT]2L-40A was used. Expression of PTEN in embryos
with the UAS GAL4 system (Brand and
Perrimon, 1993
) was done using UAS PTEN2
(Huang et al., 1999
) and UAS
PTEN3 (Goberdhan et al.,
1999
). UAS PLC
-PH-GFP flies were a gift from L. Cooley.
P{w+mC=matalpha4-GAL-VP16}V67 and
P{w+mC=Act5C-GAL4}17bF01 were used as drivers. Oregon R was used as
wild type.
To generate specific antibodies against PTEN, rabbits and rats were
immunized with a GST fusion protein containing amino acids 316-511 of PTEN2.
The rabbit antibody was affinity purified against the immobilized GST fusion
protein. Additional antibodies used were rabbit and rat anti-Baz
(Wodarz et al., 1999;
Wodarz et al., 2000
), mouse
anti-Neurotactin BP106 (Developmental Studies Hybridoma Bank), rat anti-Vasa
(Tomancak et al., 1998
),
rabbit anti-Staufen (St Johnston et al.,
1991
), goat anti-PKC zeta C20 (Santa Cruz) and mouse anti-alpha
tubulin (SIGMA). Actin was visualized with AlexaFluor 568-phalloidin
(Molecular Probes) and DNA was stained with YOYO-1 (Molecular Probes). For
whole-mount immunohistochemistry of embryos with PTEN antibodies, embryos were
fixed according to the heat-methanol procedure described by Müller and
Wieschaus (Müller and Wieschaus,
1996
). For actin and tubulin staining, embryos were fixed in 37%
formaldehyde/heptane. Secondary antibodies conjugated to Cy2, Cy3 or Cy5 were
obtained from Jackson Laboratories. Images were taken on a Leica TCSNT
confocal microscope or on a Zeiss Axioplan 2 fluorescence microscope and
processed using Photoshop (Adobe) and Canvas (Deneba) software.
Western blots and immunoprecipitation
Western blotting was done according to standard procedures. Rat anti-PTEN
was used at 1:500, rat anti-Baz (Wodarz et
al., 1999) and goat anti-PKC zeta C20 (Santa Cruz) were used at
1:1000. For immunoprecipitation, 8 µl of affinity purified rabbit anti PTEN
or 2 µl of rabbit anti-Baz serum were added to embryonic or S2 cell
extracts containing 2 mg of total protein in TNT (1% Triton X-100, 150 mM
NaCl, 50 mM Tris-Cl pH 7,5) supplemented with protease inhibitors. Immune
complexes were harvested using protein A-conjugated agarose (Roche), washed
four times in TNT and boiled in 1 xSDS sample buffer before SDS-PAGE and
western blot.
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Results |
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To test whether the association of PTEN with Baz occurred only with the PTEN2 isoform, as predicted from the yeast two-hybrid assays, we performed co-immunoprecipitation experiments with extracts of S2 cells co-transfected either with PTEN2 or PTEN3 and Baz together. In extracts of untransfected S2 cells, neither the preimmune nor the anti-PTEN immune serum precipitated detectable amounts of Baz (Fig. 2C). This finding can be explained with the very low level of endogenous Baz expression in S2 cells (Fig. 2C, top panel). In S2 cells co-transfected with PTEN2 and Baz together, Baz was co-immunoprecipitated with the PTEN antibody, but not with the preimmune serum, demonstrating the association of Baz and PTEN2 in a protein complex (Fig. 2C). By contrast, Baz did not co-immunoprecipitate with PTEN3 in co-transfected cells (Fig. 2C), confirming that Baz binds specifically to the PTEN2 isoform, which contains the PDZ-binding motif at its C terminus. To test whether an additional component of the PAR/aPKC complex is present in the complex of PTEN2 with Baz, we probed the immunoprecipitates for the presence of aPKC. aPKC co-immunoprecipitated with PTEN2 and Baz, but not with PTEN3 and Baz (Fig. 2C, bottom panel), indicating that Baz may function as a scaffold that links PTEN2 with aPKC.
In conclusion, we have shown that PTEN2 binds to the region containing PDZ domains 2 and 3 of Baz and that this interaction depends on the presence of the PDZ-binding motif at the very C terminus of PTEN2. Consistent with these in vitro binding data, PTEN2 and Baz form a complex in wild-type embryos and transfected S2 cells that also contains aPKC.
Bazooka colocalizes with PTEN in epithelia and neuroblasts
We next analyzed the subcellular localization of PTEN in embryonic
epithelia and neuroblasts. PTEN mRNA is ubiquitously expressed both maternally
and zygotically, as determined by RNA in situ hybridization (data not shown).
We could not detect endogenous PTEN protein with our antibody, presumably
because of low expression levels or insufficient sensitivity of the antibody.
However, when we expressed PTEN with the UAS-GAL4 system
(Brand and Perrimon, 1993), we
could readily detect the protein in embryonic tissues
(Fig. 3). In embryos at the
extended germ band stage (stage 10), PTEN2 was strongly enriched in the apical
cortex of the neuroectodermal epithelium and in the apical cortex of
neuroblasts (Fig. 3A). In both
cell types, PTEN2 colocalized with endogenous Baz
(Fig. 3A). At later stages of
embryogenesis, the apical enrichment and colocalization of PTEN2 with Baz was
even more pronounced (Fig. 3B).
Intriguingly, apical enrichment of PTEN2 depends on the PDZ-binding motif, as
the PTEN3 protein, which lacks the PDZ binding motif but is otherwise
identical to PTEN2 (Smith et al.,
1999
), was present on the whole plasma membrane and in the
cytoplasm (Fig. 3C). In
baz mutant embryos lacking maternal and zygotic Baz, PTEN2 showed
diffuse cytoplasmic localization and was not enriched in the apical cortex
(data not shown). However, we cannot conclude with certainty that the
mislocalization of PTEN2 is a primary consequence of the absence of Baz
function, because baz mutant embryos are already undergoing massive
degeneration at the time when PTEN2 expression driven with the GAL4 system was
strong enough to be detected by immunohistochemical staining
(Müller and Wieschaus,
1996
; Schober et al.,
1999
; Wodarz et al.,
1999
; Wodarz et al.,
2000
). Together, we have shown that PTEN2 precisely colocalizes
with Baz in epithelia and neuroblasts. Colocalization depends on the presence
of the PDZ-binding motif of PTEN2, consistent with recruitment of PTEN2 by PDZ
domains 2 and 3 of Baz.
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PTEN mutant embryos show defects in early embryonic development
Embryos lacking zygotic Pten function die late in embryogenesis or
early larval stages and do not show any obvious defects in embryonic
development (Goberdhan et al.,
1999; Huang et al.,
1999
) (data not shown). By contrast, embryos derived from
Pten germ-line clones lacking maternal and zygotic Pten
function (Ptenmat,zyg) showed severe developmental
abnormalities already in freshly laid eggs. Eggs derived from Pten
germ-line clones were generally smaller and more roundish than wild-type eggs
(Fig. 6C,D), and many did not
show any development. In those Ptenmat,zyg embryos that
initiated development, we only rarely observed the formation of pole cells
(Fig. 4G,
Fig. 5F,H) (see Movie 2 in the
supplementary material). In the few cases where pole cells were formed, their
number was very low, typically two or three, when compared with an average of
35 in wild type (Campos-Ortega and
Hartenstein, 1997
). The lack of pole cells pointed to a potential
defect in the assembly or maintenance of the germ plasm
(Rongo and Lehmann, 1996
). In
Ptenmat,zyg embryos, the mRNA of the germ plasm
determinant oskar was either diffusely localized to the posterior
pole or was completely undetectable (Fig.
4B,C). In wild type, the germ cell determinant Vasa becomes
localized to the posterior of the oocyte during oogenesis and is incorporated
into the pole cells during early embryonic development
(Fig. 4D,F,H)
(Hay et al., 1988
;
Lasko and Ashburner, 1990
). In
early Ptenmat,zyg embryos, Vasa staining at the posterior
pole was strongly reduced and was undetectable at later stages, consistent
with the failure to form germ cells (Fig.
4E,G,I).
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By contrast, in Ptenmat,zyg embryos, axial expansion did not occur, resulting in a spherical rather than ellipsoid arrangement of nuclei prior to cortical migration (Fig. 4E) and an abnormally low nuclear density in the posterior region of the embryo at the syncytial blastoderm stage (Fig. 5B). In addition, the synchrony of the cell cycle during cleavage divisions was lost (Fig. 5B,D). The pattern of mitoses was not completely randomized but can be described as occurring in waves that always started at the posterior of the embryo (see Movie 2 in the supplementary material). Consequently, mitotic figures representing all stages of the cell cycle could be seen next to each other in Ptenmat,zyg embryos (Fig. 5D). Thus, at the beginning of cellularization in cycle 14, the blastoderm nuclei showed different morphology and different condensation states of the chromatin in distinct regions of the embryo (Fig. 5F). These regional differences were reflected by the behavior of the actin network at the cellularization front, which advanced much more rapidly in the anterior region of the embryo than in the posterior region (Fig. 5F,H; see Movie 2 in the supplementary material). The delay of cellularization at the posterior pole was often accompanied by severe defects in the morphogenetic movements of gastrulation, which frequently led to rapid degeneration of Ptenmat,zyg embryos shortly after the onset of gastrulation movements (data not shown).
However, a significant number of Ptenmat,zyg embryos recovered surprisingly well from these early developmental defects and completed embryogenesis without any gross morphological abnormalities. The survival beyond gastrulation was independent of whether the embryos received a wild-type allele of Pten from their father, demonstrating that only the maternal supply of PTEN is crucial for proper development of the early embryo. Immunohistochemical staining of embryos that continued development beyond gastrulation with antibodies against proteins that localize specifically to the apical or basal pole of epithelial cells and neuroblasts, including Baz, Neurotactin, Inscuteable and Miranda, did not reveal any obvious defect in polarization of both cell types (data not shown).
PTEN controls the organization of the actin cytoskeleton in the female germline
The small size and aberrant shape of eggs derived from Pten
germ-line clones (Fig. 6A,C,D)
and the failure to localize the posterior determinants oskar mRNA and
Vasa protein in early Ptenmat,zyg embryos
(Fig. 4) point to a function
for Pten during oogenesis. Small egg size and roundish egg shape have
been reported for mutants in which the actin cytoskeleton is disorganized,
leading to inefficient transport of material from the nurse cells into the
oocyte (Robinson and Cooley,
1997). In wild-type egg chambers at stage 10
(Spradling, 1993
), actin
localizes along the cell borders of the nurse cells and is enriched in ring
canals, which stabilize the cytoplasmic bridges between nurse cells and the
oocyte (Fig. 6E'). In
addition, actin is prominently enriched in the cortex underlying the plasma
membrane of the oocyte (Fig.
6E').
By contrast, in egg chambers at stage 10 in which the germline was mutant for Pten, the actin cytoskeleton had a very disorganized structure and filled the nurse cell cytoplasm instead of localizing to cell borders (Fig. 6F'). The enrichment of actin in the oocyte cortex was strongly reduced in Pten germ-line clones (Fig. 6F'). In addition, we frequently observed fusion of nurse cells and mispositioned nurse cell nuclei that appeared to have moved into the oocyte in Pten germline clones (Fig. 6H'). We suppose that these severely abnormal oocytes give rise to the very small roundish eggs, such as the one shown in Fig. 6D. The vast majority of these very small eggs did not initiate embryonic development. Despite of the severe misorganization of the actin cytoskeleton, most Pten mutant oocytes showed normal localization of the polarity determinants Staufen (Fig. 6F''), oskar mRNA and Vasa protein (data not shown). Thus, we conclude that the failure to localize oskar mRNA and Vasa protein properly in early Ptenmat,zyg embryos points to a function for Pten in maintenance rather than establishment of posterior determinant localization. The subcellular localization of Baz was also unaffected in germ-line clones and somatic clones mutant for Pten (data not shown), demonstrating again that Pten is not required for establishment of polarity in oocytes and follicle cells. We also attempted to determine the subcellular localization of endogenous PTEN in wild-type and baz mutant ovaries but could not detect any specific staining above background, as in embryos. Thus, the question of whether Baz is required for the correct subcellular localization of PTEN in ovaries cannot be answered at this point.
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Discussion |
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Polarized localization of PTEN to the rear of the cell has been reported
for Dictyostelium cells migrating towards a source of chemoattractant
(Funamoto et al., 2002;
Iijima and Devreotes, 2002
).
This localization was complementary to the localization of
PtdIns(3,4,5)P3 at the leading edge, consistent with a
function for PTEN in lowering the local concentration of
PtdIns(3,4,5)P3 in the plasma membrane. Moreover, both
loss of Pten function and PTEN overexpression led to the loss of cell
polarity in Dictyostelium and strongly impaired the movement of the
cells towards the source of the chemoattractant
(Funamoto et al., 2002
;
Iijima and Devreotes, 2002
).
How PTEN is targeted to the rear of the cell in migrating cells is
unknown.
Mammalian PTEN contains a canonical PDZ-binding motif at its C terminus,
and this motif has been reported to interact with the multi-PDZ proteins
MAGI-2 and MAGI-3 (Wu et al.,
2000a; Wu et al.,
2000b
). Both PDZ proteins localize to tight junctions in mammalian
epithelia and cooperate with PTEN to control the activity of the downstream
kinase PKB/Akt (Wu et al.,
2000a
; Wu et al.,
2000b
; Kotelevets et al.,
2005
), indicating that subcellular targeting of PTEN may be
important for its biological activity. This hypothesis is supported by studies
of a deletion mutant of PTEN lacking the PDZ-binding motif. Although this
mutant retained lipid phosphatase activity, its activity differed from the
full-length wild-type form of PTEN in several biological assays
(Leslie et al., 2000
;
Leslie et al., 2001
).
Together, these observations demonstrate that targeting of PTEN to a specific
subcellular location may be essential for its proper function in the control
of cell polarity. Our data show for the first time that PTEN is specifically
recruited to the apical plasma membrane of epithelia and neuroblasts by direct
association with Baz/PAR-3, a key regulator of cell polarity.
PTEN activity is required for the control of several actin dependent processes in Drosophila
In order to address the issue of whether Pten activity is required
for the control of cell polarity in Drosophila, we analyzed the
phenotype of mutant ovaries and embryos lacking maternal and zygotic
Pten activity. The organization of the actin cytoskeleton in nurse
cells and in the oocyte of Pten germ-line clones becomes abnormal
from stage 9 onwards, resulting in the production of smaller, misshapen eggs.
Ptenmat,zyg embryos show defects in the axial expansion of
nuclei during nuclear division cycles 4-7 and fail to synchronize the cell
cycle in syncytial blastoderm nuclei. In addition, pole cells are strongly
reduced in number or are missing altogether, which is accompanied by the
failure to maintain oskar mRNA and Vasa protein localization at the
posterior pole. Very similar phenotypes have been reported for embryos treated
with the actin depolymerizing drug cytochalasin D and for mutants in genes
that are required for the organization of the actin cytoskeleton
(Hatanaka and Okada, 1991;
Erdelyi et al., 1995
;
Wheatley et al., 1995
;
Tetzlaff et al., 1996
;
Lantz et al., 1999
). Mutations
in the gene shackleton also show defects in axial expansion and lack
pole cells, but the posterior localization of oskar mRNA is normal,
indicating that defects in axial expansion alone are sufficient to cause the
lack of pole cells (Yohn et al.,
2003
). Interestingly, although germ plasm determinants were
mislocalized or absent in early Ptenmat,zyg embryos, they
were still localized normally during oogenesis, pointing to a function for
Pten in maintenance, rather than establishment, of germ plasm
determinant localization. Studies on ovaries and embryos mutant for the
actin-binding protein tropomyosin II gave essentially the same results
(Erdelyi et al., 1995
;
Tetzlaff et al., 1996
). Thus,
all of the phenotypes of Ptenmat,zyg mutant ovaries and
embryos described here can be related to a function for PTEN in
actin-dependent processes.
The links between PTEN and actin are obviously the substrate and the
product of the enzymatic activity of PTEN, PtdIns(3,4,5)P3
and PtdIns(4,5)P2. Both phosphoinositide lipids are
important regulators of the actin cytoskeleton.
PtdIns(4,5)P2 acts mostly by direct binding to
actin-associated proteins that link the actin cytoskeleton to the plasma
membrane or by binding to proteins that are involved in the initiation of de
novo actin polymerization, e.g. profilin and WASP
(Fig. 7)
(Yin and Janmey, 2003).
PtdIns(3,4,5)P3 in turn acts on the actin cytoskeleton via
recruitment of guanine nucleotide exchange factors (GEFs) for the small
GTPases Rac1, Rho and Cdc42 (Zheng,
2001
), which can activate WASP proteins and the Arp2/3 complex
(Fig. 7). Because we do not
know the subcellular localization of endogenous PTEN, we cannot predict at
present how exactly PTEN may affect the organization of the actin cytoskeleton
during early embryonic development. However, the fact that overexpressed PTEN2
colocalizes with PtdIns(4,5)P2 to the junctional region of
epithelial cells indicates that PTEN may locally alter the balance between
PtdIns(4,5)P2 and PtdIns(3,4,5)P3 in
the plasma membrane, leading to a modification of the actin cytoskeleton in
defined regions of the cytocortex. Studies of PTEN knockout cells and
Pten mutants in Drosophila have indeed shown that loss of
Pten leads to a significant increase in the amount of
PtdIns(3,4,5)P3 in the plasma membrane
(Stambolic et al., 1998
;
Oldham et al., 2002
).
|
A potential function for PTEN in regulation of aPKC and Cdc42
Besides its function in the regulation of actin, PTEN may regulate the
catalytic activity of aPKC, a core component of the PAR/aPKC complex that
directly binds to Baz (Wodarz et al.,
2000) and associates in a protein complex with PTEN2
(Fig. 2C;
Fig. 7). The mammalian homologs
of aPKC, the atypical PKC isoforms
and
, require
phosphorylation by the upstream kinase PDK1 in order to become fully active
(Chou et al., 1998
;
Le Good et al., 1998
;
Standaert et al., 2001
). PDK1
is recruited to the plasma membrane by direct binding of its pleckstrin
homology (PH) domain to PtdIns(3,4,5)P3. PDK1, PTEN and
several downstream effectors of the PI3-kinase signaling pathway in
Drosophila show strong genetic interactions and are crucial for the
regulation of cell growth and proliferation
(Goberdhan et al., 1999
;
Huang et al., 1999
;
Gao et al., 2000
;
Scanga et al., 2000
;
Oldham et al., 2002
;
Radimerski et al., 2002
;
Stocker et al., 2002
). We have
obtained biochemical evidence that aPKC is a substrate for PDK1 (A.G. and
A.W., unpublished) and propose that aPKC is phosphorylated in response to
elevated PtdIns(3,4,5)P3 levels. According to this
hypothesis, PTEN would be a negative regulator of the kinase activity of
aPKC.
In addition to PDK1, PtdIns(3,4,5)P3 recruits GEFs that
activate the small GTPases Cdc42 and Rac1
(Fig. 7)
(Zheng, 2001). Intriguingly,
active GTP-bound Cdc42 is also a component of the PAR/aPKC complex in
mammalian cells and in Drosophila
(Joberty et al., 2000
;
Johansson et al., 2000
;
Lin et al., 2000
;
Qiu et al., 2000
;
Hutterer et al., 2004
) (D.
Egger and A.W., unpublished). GTP-bound Cdc42 binds directly to the CRIB
domain of PAR-6 and this interaction could elevate the kinase activity of
aPKC, as has been shown in mammalian cells
(Yamanaka et al., 2001
). Thus,
PtdIns(3,4,5)P3 could activate aPKC both by recruitment of
PDK1, which directly phosphorylates aPKC, and by recruitment of GEFs, which
activate aPKC via Cdc42 and PAR6. Studies on PTEN knockout cells have indeed
shown that PTEN inhibits Rac1 and Cdc42
(Liliental et al., 2000
). The
presence of PTEN in one complex together with aPKC, Cdc42 and PAR-6 should
therefore lead to inhibition of both pathways that activate aPKCs, revealing a
novel way to control the activity of a key component of the PAR/aPKC
complex.
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ACKNOWLEDGMENTS |
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![]() |
Footnotes |
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Supplementary material for this article is available at http://dev.biologists.org/cgi/content/full/132/7/1675/DC1
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