The inositol 5-phosphatase SHIP2 is an effector of RhoA and is involved in cell polarity and migration.
ABSTRACT Cell migration is essential for various physiological and pathological processes. Polarization in motile cells requires the coordination of several key signaling molecules, including RhoA small GTPases and phosphoinositides. Although RhoA participates in a front-rear polarization in migrating cells, little is known about the functional interaction between RhoA and lipid turnover. We find here that src-homology 2-containing inositol-5-phosphatase 2 (SHIP2) interacts with RhoA in a GTP-dependent manner. The association between SHIP2 and RhoA is observed in spreading and migrating U251 glioma cells. The depletion of SHIP2 attenuates cell polarization and migration, which is rescued by wild-type SHIP2 but not by a mutant defective in RhoA binding. In addition, the depletion of SHIP2 impairs the proper localization of phosphatidylinositol 3,4,5-trisphosphate, which is not restored by a mutant defective in RhoA binding. These results suggest that RhoA associates with SHIP2 to regulate cell polarization and migration.
- SourceAvailable from: PubMed Central[Show abstract] [Hide abstract]
ABSTRACT: The aim of this study was to investigate whether the phosphoinositide 3-kinase (PI3K)/Akt signaling pathway affects the implantation of mouse embryos by regulating the expression of RhoA. The expression of PI3K, Akt, phosphorylated (p-)Akt, phosphatase and tensin homolog (PTEN) and RhoA in the uterus of mice on day 5 of pregnancy (D5) and in pseudopregnant mice was examined by quantitative reverse transcription polymerase chain reaction (qRT-PCR), immunohistochemistry and western blot analysis. A functional analysis of these genes was also performed by the intrauterine injection with the PI3K inhibitor, LY294002, on day 2 of pregnancy (D2). The expression levels of PI3K, p-Akt, RhoA at the implantation site were higher than those at the inter-implantation site in the endometrium; however, opposite effects were observed for PTEN expression. The expression levels of the above genes in the pseudopregnant group and in the group injected with the PI3K/Akt inhibitor, LY294002, were markedly lower than those in the pregnant group. Functional experiments revealed that the number of implantation sites had been significantly decreased (P<0.05) following the intrauterine injection of the PI3K inhibitor, LY294002, on day 2 of gestation compared with the contralateral injection of phosphate-buffered saline (PBS). These results suggest that the PI3K/Akt signaling pathway affects embryo implantation by regulating the expression of RhoA.International Journal of Molecular Medicine 03/2014; · 1.96 Impact Factor
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ABSTRACT: The SH2 containing inositol 5-phosphatase SHIP2 is a member of the mammalian phosphoinositide polyphosphate 5-phosphatase family. It is a multi-domain protein comprising a central catalytic domain, an SH2 domain at its N-terminus, proline rich sequences and SAM domain at its C-terminus. It can dephosphorylate both phosphatidylinositol 3,4,5-trisphosphate (PI(3,4,5)P3) and phosphatidylinositol 4,5-bisphosphate (PI(4,5)P2) and can participate in multiple signalling events in response to growth factors such as EGF, FGF or PDGF. Human SHIP2 can be phosphorylated at two major tyrosine residues Tyr986 and Tyr1135. Here, we report two intracellular localizations of pSHIP2(Y1135): pSHIP2(Y1135)-ir localizes at focal adhesions in EGF-stimulated HeLa cells and at the mitotic spindle in HeLa, in GIST882 cells, a human model of gastrointestinal stromal tumors derived cells, and in human astrocytoma 1321N1 cells. pSHIP2(Y1135)-ir is maximal at metaphase. In N1 cells, evidence is provided that the SHIP2 pathway impacts the distribution of mitotic centrosomes, particularly ү-tubulin. Our data reinforce the concept that SHIP2 localization in intact cells is dependent on phosphorylation mechanisms on both Ser/Thr and Tyr residues, i.e. Y1135, in three cancer cell lines.Cellular Signalling 02/2014; · 4.47 Impact Factor
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ABSTRACT: The mammalian striatin family consists of three proteins, striatin, S/G2 nuclear autoantigen, and zinedin. Striatin family members have no intrinsic catalytic activity, but rather function as scaffolding proteins. Remarkably, they organize multiple diverse, large signaling complexes that participate in a variety of cellular processes. Moreover, they appear to be regulatory/targeting subunits for the major eukaryotic serine/threonine protein phosphatase 2A. In addition, striatin family members associate with germinal center kinase III kinases as well as other novel components, earning these assemblies the name striatin-interacting phosphatase and kinase (STRIPAK) complexes. Recently, there has been a great increase in functional and mechanistic studies aimed at identifying and understanding the roles of STRIPAK-like complexes in cellular processes of multiple organisms. These studies have identified novel STRIPAK or STRIPAK-like complexes and have explored their roles in specific signaling pathways. Together, the results of these studies have sparked increased interest in striatin family complexes because they have revealed roles in signaling, cell cycle control, apoptosis, vesicular trafficking, Golgi assembly, cell polarity, cell migration, neural and vascular development, and cardiac function. Moreover, STRIPAK complexes have been connected to clinical conditions, including cardiac disease, diabetes, autism, and cerebral cavernous malformation. In this review, we discuss the expression, localization, and protein domain structure of striatin family members. Then we consider the diverse complexes these proteins and their homologs form in various organisms, emphasizing what is known regarding function and regulation. Finally, we will explore possible roles of striatin family complexes in disease, especially cerebral cavernous malformation.The international journal of biochemistry & cell biology 12/2013; · 4.89 Impact Factor
Volume 23 July 1, 2012
MBoC | ARTICLE
The inositol 5-phosphatase SHIP2 is an effector
of RhoA and is involved in cell polarity
Katsuhiro Katoa,b, Tsubasa Yazawaa, Kentaro Takia, Kazutaka Moria,b, Shujie Wanga,c,
Tomoki Nishiokaa, Tomonari Hamaguchia, Toshiki Itohd, Tadaomi Takenawae, Chikako Kataokaf,
Yoshiharu Matsuuraf, Mutsuki Amanoa, Toyoaki Muroharab, and Kozo Kaibuchia
aDepartment of Cell Pharmacology and bDepartment of Cardiology, Nagoya University Graduate School of Medicine,
65 Tsurumai, Showa, Nagoya, Aichi 466-8550, Japan; cDepartment of Anatomy, School of Medicine, Mie University,
2-174 Edobashi, Tsu, Mie 514-8507, Japan; dDivision of Membrane Biology and eDivision of Lipid Biochemistry,
Department of Biochemistry and Molecular Biology, Kobe University Graduate School of Medicine, 7-5-1 Kusunoki,
Chuo, Kobe, Hyogo 650-0017, Japan; fDepartment of Molecular Virology, Research Institute for Microbial Diseases,
Osaka University, 3-1, Yamadaoka, Suita, Osaka 565-0871, Japan
ABSTRACT Cell migration is essential for various physiological and pathological processes.
Polarization in motile cells requires the coordination of several key signaling molecules, in-
cluding RhoA small GTPases and phosphoinositides. Although RhoA participates in a front–
rear polarization in migrating cells, little is known about the functional interaction between
RhoA and lipid turnover. We find here that src-homology 2–containing inositol-5-phosphatase
2 (SHIP2) interacts with RhoA in a GTP-dependent manner. The association between SHIP2
and RhoA is observed in spreading and migrating U251 glioma cells. The depletion of SHIP2
attenuates cell polarization and migration, which is rescued by wild-type SHIP2 but not by a
mutant defective in RhoA binding. In addition, the depletion of SHIP2 impairs the proper
localization of phosphatidylinositol 3,4,5-trisphosphate, which is not restored by a mutant
defective in RhoA binding. These results suggest that RhoA associates with SHIP2 to regulate
cell polarization and migration.
Cell migration plays an important role in tissue development, im-
mune function, and wound healing. Most migrating cells have a
highly polarized morphology, such as a front leading edge and rear
tail, and front–rear polarity is critical for the efficiency of cell migration
(Ridley et al., 2003; Petrie et al., 2009). One of the major regulators
of front–rear polarity is the Rho family of small GTPases, including
Rho, Rac, and Cdc42. The activities of Rho GTPases are controlled in
a temporal and spatial manner to regulate actin cytoskeleton reorga-
nization, cell polarity, and migration (Jaffe and Hall, 2005). RhoA is
mainly activated in the rear and central regions to promote the as-
sembly and activation of contractile actomyosin networks and to re-
strict protrusions. RhoA is also activated at the leading edge
(Kurokawa and Matsuda, 2005; Pertz et al., 2006). Rac1 and Cdc42
are predominantly activated at the leading edge to generate a vecto-
rial protrusion in the direction of migration (Kraynov et al., 2000; Itoh
et al., 2002). These Rho GTPases regulate various cellular functions
through downstream effectors. Rho-kinase/ROCK/ROK, which is one
of the best-characterized effectors of RhoA, is believed to be a key
regulator of front–rear polarity (Riento and Ridley, 2003; Narumiya
et al., 2009; Amano et al., 2010). Rho-kinase phosphorylates Par-3, a
component of the Par polarity complex, or FilGAP, the negative regu-
lator for Rac, resulting in Rac1 inactivation (Ohta et al., 2006;
Nakayama et al., 2008). In addition to Rho-kinase, other RhoA effec-
tors might also be involved in front–rear polarity, because the cell
Received: Nov 29, 2011
Revised: Apr 10, 2012
Accepted: May 8, 2012
This article was published online ahead of print in MBoC in Press (http://www
.molbiolcell.org/cgi/doi/10.1091/mbc.E11-11-0958) on May 16, 2012.
Address correspondence to: Kozo Kaibuchi (email@example.com).
Abbreviations used: FN, fibronectin; LC-MS/MS, liquid chromatography–tandem
mass spectrometry; PI3K, phosphatidylinositol 3-kinase; PI(3,4,5)P3, phosphati-
dylinositol 3,4,5-triphosphate; PLA, proximity ligation assay.
© 2012 Kato et al. This article is distributed by The American Society for Cell Biol-
ogy under license from the author(s). Two months after publication it is available
to the public under an Attribution–Noncommercial–Share Alike 3.0 Unported
Creative Commons License (http://creativecommons.org/licenses/by-nc-sa/3.0).
“ASCB®,” “The American Society for Cell Biology®,” and “Molecular Biology of
the Cell®” are registered trademarks of The American Society of Cell Biology.
2594 | K. Kato et al. Molecular Biology of the Cell
dominant-negative mutant) as glutathione S-transferase (GST)–fusion
proteins and used them as ligands for the affinity columns. A cytoso-
lic fraction of rat heart was applied to affinity beads coated with GST
alone or GST-RhoA mutants. Numerous proteins were detected in
the eluates from the columns coated with GST-RhoA mutants (Figure
2A). Eluates from the GST and GST-RhoA mutant affinity columns
were used for shotgun analysis using LC-MS/MS. The list of represen-
tative RhoA-interacting proteins is shown in Supplemental Table S1.
Several known effectors, including Rho-kinase, PKN, Rhotekin, and
MYPT1, were detected in the eluates from the GTPγS⋅GST-RhoA and
active GST-RhoA-L63 columns (Supplemental Table S1).
In addition to the known RhoA-interacting proteins, we identi-
fied many proteins that specifically interacted with the GTPγS-bound
active form or active mutant of RhoA but not the GDP-bound inac-
tive form or dominant-negative mutant. Some of these proteins ap-
pear to be novel effectors of RhoA. Several proteins, including
acetyl-coenzyme A acetyltransferase 1 (ACAT1), SHIP2, filamin A in-
teracting protein (FILIP), and striatin, calmodulin–binding protein 3
(STRN3), were identified as candidate effectors. To confirm the
amounts of these proteins in eluates, we performed immunoblot
analysis using specific antibodies (Figure 2B). The bands recognized
by the anti–Rho-kinase antibody were detected in the eluates from
the GTPγS⋅GST-RhoA– and GST-RhoA-L63–immobilized columns
but not in those from the GST-, GDP⋅GST-RhoA–, or GST-RhoA-
N19–immobilized columns. RhoGDI was detected in the eluates of
the GDP⋅GST-RhoA–immobilized column. Consistent with the mass
spectrometric analysis, ACAT1 and SHIP2 were detected in the elu-
ates from the GTPγS⋅GST-RhoA– and GST-RhoA-L63–immobilized
columns (Figure 2B). SHIP2 dephosphorylates PI(3,4,5)P3 into phos-
phatidylinositol 3,4-bisphosphate. SHIP2 is believed to be a nega-
tive regulator of PI3K pathways; in addition, it regulates the actin
cytoskeleton and is involved in cell migration, cell adhesion, endo-
cytosis, and metastasis of cancer (Ooms et al., 2009). We decided to
morphology phenotype of cells treated with the Rho-kinase inhibitor
was different from that of RhoA-depleted cells (Figure 1B).
Another key regulator of front–rear polarity is the cellular phos-
phoinositide signaling system. Among the phosphoinositides, phos-
phatidylinositol 3,4,5-trisphosphate (PI(3,4,5)P3) accumulates at the
front of directionally migrating cells (Ridley et al., 2003). PI(3,4,5)P3
is produced by the phosphorylation of PI(4,5)P2 by a family of en-
zymes known as phosphatidylinositol 3-kinases (PI3Ks) and regulates
the localization of specific proteins by binding to their pleckstrin
homology domains (Cantley, 2002). The levels of phospholipids are
tightly regulated by kinases, phosphatases, and phospholipases (Di
Paolo and De Camilli, 2006). Interactions between Rho GTPases
and phosphoinositide metabolism are implicated in cell polarity.
However, the details of how Rho GTPases and phosphoinositide sig-
naling are coordinated remain unclear.
In this study, we performed affinity chromatography to search for
effectors of RhoA involved in front–rear polarity. We identified sev-
eral candidates of novel RhoA effectors, including src-homology
2–containing inositol-5-phosphatase 2 (SHIP2), a PI(3,4,5)P3-5 phos-
phatase. SHIP2 directly interacted with GTP-bound but not GDP-
bound RhoA. SHIP2 appeared to restrict PI(3,4,5)P3 localization at
the leading edge in migrating cells to thereby control the cell polar-
ity downstream of RhoA. We demonstrate a novel linkage between
Rho family GTPases and lipid signaling via a SHIP2–RhoA interaction
in the establishment of cell polarity.
Front–rear polarity and the Rho pathway
When suspended cells, such as Vero fibroblasts and U251 glioma
cells, are plated on extracellular matrix proteins, the cells begin to
spread. Next the cells begin to ruffle, with the accumulation of actin
filaments around the cell periphery, the so-called lamellipodia, and
develop membrane extensions. Then the lamellipodia gradually di-
vide into several regions. Finally, the lamellipodia localize to one side,
and the cells acquire a distinct front–rear polarized morphology.
Polarized cells have a single leading edge with an accumulation of
actin filaments. In this study, we used U251 glioma cells because they
have a distinct front–rear polarized morphology and persistently mi-
grate when seeded on a fibronectin (FN)-coated surface (Figure 1A).
To investigate the roles of the Rho pathway in front–rear polarity,
we first compared RhoA knockdown with Y-27632 treatment, a spe-
cific Rho-kinase inhibitor. RhoA-depleted cells or cells treated with
Y-27632 were suspended and seeded on FN-coated glasses. Ap-
proximately 40% of U251 cells transfected with control siRNA had a
single leading edge with a polarized morphology 8 h after plating
(Figure 1B). Cells with a single leading edge were observed less in
both RhoA-depleted cells and cells treated with Y-27632. RhoA-de-
pleted cells had an elongated cell shape or an extended, flattened
morphology with multiple small protrusions, whereas cells treated
with Y-27632 had long, thin processes and flattened, elongated pro-
trusions without stress fibers (Figure 1B). Given that the cell mor-
phology phenotype of cells treated with Y-27632 did not resemble
that of RhoA knockdown, RhoA regulates cell polarity through not
only Rho-kinase, but also other effectors.
SHIP2 as a novel effector of RhoA
To explore the other RhoA effectors involved in front–rear polarity,
we attempted to identify novel RhoA effectors with RhoA affinity col-
umn chromatography followed by a liquid chromatography–tandem
mass spectrometry (LC-MS/MS) shotgun analysis. We prepared
GDP⋅RhoA, GTPγS (a nonhydrolyzable GTP analogue)⋅RhoA, and
RhoA mutants (RhoA-L63, constitutively active mutant; RhoA-N19,
FIGURE 1: Regulation of front–rear polarity downstream of RhoA.
(A) Polarization of U251 cells. U251 cells were plated on FN-coated
glass and stained with Alexa 488–phalloidin. Bar, 10 μm. (B) Effect of
RhoA depletion or Y-27632 treatment in U251 cells. U251 cells
transfected with the indicated siRNAs for 72 h were treated with or
without 20 μM Y-27632, reseeded on FN-coated glasses for 8 h, fixed,
and stained with Alexa 488–phalloidin. Bar, 20 μm. All results are
representative of at least three independent experiments.
siControl siRhoA#1 Y-27632
Volume 23 July 1, 2012 RhoA effector SHIP2 and cell polarity | 2595
found that SHIP1 did not interact with ac-
tive RhoA under the same experimental
conditions (Supplemental Figure S1A).
We also found that SHIP2 was expressed
in U251 glioma, MDA-MB-231 epithelial,
and RAW264.7 macrophage cells, whereas
SHIP1 was expressed in RAW264.7 mac-
rophage cells but was undetectable in
U251 and MDA-MB-231 cells (Supplemental
Figure S1B), suggesting that SHIP2 is a ma-
jor SHIP in U251 and MDA-MB-231 cells. To
reveal the signal linkage between SHIP2 and
Rho family GTPases, we assessed the speci-
ficity of their interactions and confirmed that
SHIP2 specifically interacted with RhoA but
not Rac1 or Cdc42 (Figure 2D).
Direct interaction of SHIP2
with active RhoA
To narrow down the RhoA-binding region in
SHIP2, we generated several SHIP2 frag-
ments (Figure 3A) and found that green flu-
orescent protein (GFP)–SHIP2-N interacted
with GTPγS⋅GST-RhoA, whereas GFP-SHIP2-
cat or C did not (Figure 3B). To further nar-
row down and confirm the direct interaction
of active RhoA with SHIP2, we performed
an in vitro binding assay. GST-, GDP⋅GST-
RhoA–, and GTPγS⋅GST-RhoA–immobilized
beads were incubated with each maltose-
binding protein (MBP)-SHIP2 fragment.
GTPγS⋅GST-RhoA interacted with MBP-
SHIP2-N and MBP-SHIP2-NΔSH2 but not
MBP-SHIP2-SH2 (Figure 3C). The SHIP2-
NΔSH2-4 fragment (124–314 amino acids)
was sufficient to bind to RhoA (Supplemen-
tal Figure S2A). These results indicate that
active RhoA directly interacts with SHIP2 at
the N-terminal region between the SH2 and
catalytic domains. The binding activity of
SHIP2-NΔSH2-4 fragment was comparable
with that of the Rho-kinase-Rho-binding
(RB) fragment containing RhoA-binding do-
main (Supplemental Figure S2B).
We next examined whether the SHIP2-
NΔSH2-4 contains a conserved binding motif to RhoA. It has been
shown that PKN/PRK1 interacts with RhoA through a leucine zipper-
like motif (Maesaki et al., 1999). Because SHIP2-NΔSH2-4 contains
this motif, it could be important for RhoA binding (Supplemental
Figure S2C). A series of SHIP2 point mutants (D193/E195A, S202/
N203A, R216/R217A, and D223/K224A) was generated based on
several reports regarding structure and mutation analysis (Peck
et al., 2002; Owen et al., 2003; Shimizu et al., 2003; Blumenstein
and Ahmadian, 2004). The binding activities of MBP-SHIP2-NΔSH2-
4-D193/E195A and D223/K224A to active RhoA were much lower
than that of MBP-SHIP2-NΔSH2-4 under these conditions (Supple-
mental Figure S2D). We then generated a full-length SHIP2-D193/
E195A mutant; the binding activity of myc-SHIP2-D193/E195A for
active RhoA was dramatically reduced compared with myc-SHIP2-
wild type (WT) (Figure 3D). These results suggest that the leucine
zipper–like motif in the N-terminus of SHIP2 is necessary for RhoA
focus on SHIP2 because it is involved in cell polarity and actin
Characterization of SHIP2–RhoA binding
To determine whether SHIP2 associates with active RhoA in COS7
cells, we performed pull-down assays using the constitutively active
RhoA mutant. Extracts of COS7 cells cotransfected with myc-SHIP2
and GST or GST-RhoA mutants were prepared and mixed with glu-
tathione–Sepharose beads. SHIP2 interacted with GST-RhoA-L63
but not GST, GST-RhoA-N19, or GST-RhoA-A37L63, which contains
an amino acid substitution in the effector domain and has no ability
to bind the effectors (Figure 2C). These results suggest that SHIP2 is
a novel effector of RhoA.
Among the 10 mammalian inositol polyphosphate 5-phos-
phatases, SHIP1 and SHIP2 are structurally similar proteins with high
sequence identity (Ooms et al., 2009). To examine whether SHIP1
also associates with active RhoA, we performed binding assays and
FIGURE 2: Identification of RhoA-binding proteins. (A) Isolation of RhoA-binding proteins by
affinity column chromatography. GST, GDP⋅GST-RhoA, GTPγS⋅GST-RhoA, or GST-RhoA mutants
were immobilized on beads and incubated with the rat heart lysate. The bound proteins were
eluted with buffer containing 1 M NaCl and subjected to SDS–PAGE, followed by silver staining.
(B) Validation of LC-MS/MS results by immunoblotting. The eluates from affinity column
chromatography were subjected to immunoblotting using anti–Rho-kinase, anti-RhoGDI,
anti-SHIP2, and anti-ACAT1 antibodies. (C) Interaction of SHIP2 with active RhoA in COS7 cells.
Glutathione–Sepharose beads were incubated with COS7 cell lysate expressing both myc-SHIP2
and GST-RhoA mutants. The bound proteins were analyzed by immunoblotting with anti-myc
and anti-GST antibodies. The dashed line indicates separate membranes. (D) Specific interaction
of SHIP2 with RhoA but not Rac1 or Cdc42. Glutathione–Sepharose beads were incubated with
COS7 cell lysate expressing both myc-SHIP2 and GST-RhoA, GST-Rac1, or GST-Cdc42 mutants.
The bound proteins were analyzed by immunoblotting with anti-myc and anti-GST antibodies.
All results are representative of at least three independent experiments.
GST-RhoAGST-Rac1 GST-RhoA GST-Cdc42GST-Cdc42
2596 | K. Kato et al. Molecular Biology of the Cell
We further addressed the function of SHIP2-RhoA interaction.
When both myc-SHIP2 and control vector were transfected in U251
cells, most of the myc-SHIP2 was detected in the cytosol fraction.
Coexpression with HA-RhoA-L63 increased the amount of myc-
SHIP2 in the membrane fractions, whereas coexpression with
To determine whether active RhoA could modulate the PI(3,4,5)P3
5-phosphatase activity of SHIP2, we performed in vitro phosphatase
assays and found that GTPγS⋅GST-RhoA did not affect the PI(3,4,5)P3
5-phosphatase activity of SHIP2 under the specific experimental con-
ditions (Supplemental Figure S3).
FIGURE 3: Interaction of SHIP2 with active RhoA. (A) Domain structure and deletion constructs of SHIP2. (B) Mapping
of the SHIP2 region required for binding to active RhoA. GST, GDP⋅GST-RhoA, or GTPγS⋅GST-RhoA, immobilized on
beads, was incubated with COS7 cell lysate expressing GFP-SHIP2-N, GFP-SHIP2-cat, or GFP-SHIP2-C. The bound
proteins were analyzed by immunoblotting with the anti-GFP antibody. (C) Direct binding of SHIP2 with active RhoA.
GST, GDP⋅GST-RhoA, or GTPγS⋅GST-RhoA, immobilized on beads, was incubated with various MBP-SHIP2 fragments.
The bound proteins were analyzed by silver staining. Black dots indicate respective intact bands, and arrowheads
indicate the bound proteins. (D) Characterization of binding-deficient mutants of SHIP2 in COS7 cells. Glutathione–
Sepharose beads were incubated with COS7 cell lysate expressing both myc-SHIP2-WT or myc-SHIP2-D193/E195A and
GST-RhoA mutants. The bound proteins were analyzed by immunoblotting with anti-myc and anti-GST antibodies. All
results are representative of at least three independent experiments.
SH2: Src Homology 2 domain
PRR: Proline Rich Region
SAM: Sterile alpha motif
catalytic: 5′-phosphatase domain
Volume 23 July 1, 2012 RhoA effector SHIP2 and cell polarity | 2597
sites (Figure 4D). In addition, the distribution of PI(3,4,5)P3 seemed
different from that of the SHIP2-RhoA interaction.
Regulation of cell polarity by SHIP2 and RhoA
To examine the roles of SHIP2 and RhoA in cell polarization, we
performed SHIP2 and RhoA siRNA knockdown in U251 cells. Cells
depleted of SHIP2 or RhoA were suspended and seeded on FN-
coated glasses. U251 cells depleted of SHIP2 were less polarized
than controls and had spreading cell shapes (Figure 5A and Supple-
mental Figure S6A). These phenotypes were not resistant to treat-
ment with Y-27632, which led to similar phenotypes. Phalloidin
staining revealed that F-actin was observed at the peripheral re-
gions of SHIP2-depleted cells, whereas RhoA-depleted cells be-
came thinner and had multiple small, irregular protrusions (Figures
1B and 5A). Multiple small protrusions were also observed in RhoA-
depleted PC3 prostate cancer cells during spreading (Vega et al.,
To examine the functional significance of the interaction of SHIP2
with RhoA, we performed rescue experiments. The expression of
siRNA-resistant (RNAiR)-SHIP2-WT rescued the polarization defects,
whereas RNAiR-SHIP2-D193/E195A or D608A, a phosphatase-defi-
cient mutant, did not fully rescue the defects (Figure 5, B and C,
and Supplemental Figure S6, B and C), suggesting that the RhoA-
binding and phosphatase activities of SHIP2 are required for cell
polarization. Similar observations were obtained in MDA-MB-231
epithelial cells (Supplemental Figure S7, A and B). We did not ob-
serve the phenotype of the overexpression of SHIP2-WT, possibly
due to toxicity. Moderate expression of SHIP2-WT did not affect cell
polarization. In addition, the overexpression of RhoA-WT and RhoA-
L63 induced cell rounding.
Next we attempted to examine the effect of SHIP2 or RhoA
knockdown on the localization of PI(3,4,5)P3, because the depletion
of SHIP1 or SHIP2 affects the levels or localization of PI(3,4,5)P3 in
several cell types (Blero et al., 2005; Mandl et al., 2007; Nishio et al.,
2007; Nakatsu et al., 2010). The local accumulation of PI(3,4,5)P3 at
the leading edge is crucial to maintain front–rear polarity (Ridley
et al., 2003). Accumulation of PI(3,4,5)P3 was observed at the lead-
ing edge in control U251 cells, whereas PI(3,4,5)P3 was observed
throughout the cell periphery in SHIP2-knockdown cells. In RhoA-
knockdown cells, PI(3,4,5)P3 did not accumulate at the leading edge
and was instead found as multiple small, irregular protrusions
(Figure 6, A and B, and Supplemental Figure S6D). PI(3,4,5)P3 stain-
ing was markedly diminished in LY294002-treated cells (Figure 6A
and Supplemental Figure S6D) or in cells overexpressing GFP-
tagged SHIP2 or PTEN (Supplemental Figure S8). Furthermore, the
expression of RNAiR-SHIP2-WT rescued the restricted accumulation,
whereas RNAiR-SHIP2-D193/E195A or D608A did not fully rescue
the accumulation (Figure 6, C and D, and Supplemental Figure S6,
E and F). Similar observations were obtained in MDA-MB-231 epi-
thelial cells (Supplemental Figure S7, C and D). These results sug-
gest that SHIP2 regulates the proper localization of PI(3,4,5)P3
downstream of RhoA, leading to the maintenance of cell polarity.
Requirement of the RhoA-binding activity of SHIP2
for cell migration
Finally, to examine the functional significance of the interaction be-
tween SHIP2 and RhoA in cell migration, we used a Boyden cham-
ber assay. U251 cells transfected with either the indicated siRNA or
siRNA along with indicated RNAiR-SHIP2 were subjected to the Boy-
den chamber assay for 2 h. The depletion of SHIP2 or RhoA signifi-
cantly impaired epidermal growth factor (EGF)-stimulated cell mi-
gration from the upper to the lower membrane. The two different
HA-RhoA-N19 or HA-RhoA-L63S190, which is an active RhoA mu-
tant with mutation at the prenylation site and thereby fails to target
the membrane, had minimal effect (Supplemental Figure S4A). We
next asked whether RhoA could recruit SHIP2 in U251 cells using a
strategy to engage RhoA on the mitochondria. Expression of a GFP
(EGFP)-RhoA-L63 fused with mitochondrial targeting sequence
(MitoEGFP-RhoA-L63) in U251 cells partly recruited SHIP2-mCherry
to the mitochondria, whereas that of MitoEGFP-RhoA-N19 or
MitoEGFP in U251 cells did not (Supplemental Figure S4B). These
results suggest that RhoA can tether SHIP2 to the juxtamembrane
region in an activity-dependent manner, where SHIP2 catalyzes the
hydrolysis of PI(3,4,5)P3.
Association of SHIP2 with RhoA in intact cells
To examine the subcellular localization of SHIP2 and RhoA, we im-
munostained U251 cells seeded on FN-coated glasses with anti-
SHIP2 and anti-RhoA antibodies. SHIP2 and RhoA were partially
colocalized at both the central region and leading edge in U251
cells (Figure 4A). To test the specificity of the anti-SHIP2 and anti-
RhoA antibodies, endogenous SHIP2 and RhoA were depleted by
two independent small interfering RNAs (siRNAs) or siRNA mixtures.
The staining of SHIP2 and RhoA was strongly reduced upon knock-
down of the proteins by siRNA (Supplemental Figure S5, A and B).
We also confirmed that depletion of SHIP2 did not affect the RhoA
activity in U251 cells (Supplemental Figure S5C). To examine whether
SHIP2 associates with RhoA in intact U251 cells, we performed prox-
imity ligation assays (PLAs). PLA allows for the detection of native
complexes with minimal cellular disruption (Fredriksson et al., 2002;
Soderberg et al., 2006). A signal is obtained only if oligonucleotides
coupled to two separate antibodies are sufficiently close to allow
enzymatic ligation; detection then relies on amplification from the
ligated template, followed by hybridization with a fluorescent probe.
This assay reveals protein interactions in intact cells. We detected
many ubiquitous signals in the presence of the SHIP2 and RhoA
antibodies, whereas few signals were obtained in the presence of
the SHIP2 or RhoA antibody alone (Figure 4B). These signals were
assumed to be the SHIP2–RhoA interaction. Furthermore, the spots
of the SHIP2–RhoA interaction were reduced in RhoA-depleted cells
(Figure 4B). We also observed many signals using Rho-kinase and
RhoA antibodies (Supplemental Figure S5D). Quantitative analysis
revealed that the signals of the SHIP2–RhoA interaction were more
pronounced than the background obtained with each SHIP2 or
RhoA antibody alone and the signals in RhoA-depleted cells (Figure
4C). These results indicate that SHIP2 associates with RhoA in intact
To further explore the interaction between SHIP2 and RhoA, we
performed PLAs during cell spreading and polarization. When U251
cells were plated on FN-coated glasses, lamellipodia extension was
promoted after 30 min to 1 h. Cells initiated polarization ∼1 h after
spreading. After polarization, cells finally acquired a distinct front–
rear polarized morphology and began to migrate. Many signals
were ubiquitously detected with PLAs, and SHIP2 and RhoA were
partially colocalized at both the central region and the cell periphery
during cell spreading and polarization, as detected by immunostain-
ing (Figure 4D). These results suggest that an association between
SHIP2 and RhoA is present during spreading and polarization.
We then tried to observe the localization of PI(3,4,5)P3 because
PI(3,4,5)P3 is a key signaling molecule that becomes rapidly and
highly polarized in cells, such as spreading cells (Weiger et al., 2009).
We observed PI(3,4,5)P3 accumulation throughout the cell periph-
ery 30 min and 1 h after plating. After initiation of polarization, the
distribution of PI(3,4,5)P3 tended to localize at actin accumulation
2598 | K. Kato et al. Molecular Biology of the Cell
FIGURE 4: Association of SHIP2 with RhoA in intact cells. (A) Localization of SHIP2 and RhoA in U251 cells. U251 cells
were double labeled with anti-SHIP2 (green) and anti-RhoA (red) antibodies. Right, the merged image. Bar, 10 μm.
(B) Association of SHIP2 with RhoA by proximity ligation assays in U251 cells. U251 cells were treated with control
siRNA or siRNA against RhoA. PLA was performed using antibodies against SHIP2, RhoA, or both. Images represent
focused views of several confocal sections that covered the entire region of cells. Cell edges are marked with a dotted
line. Nuclei were stained with 4′,6-diamidino-2-phenylindole (DAPI; blue). Bar, 10 μm. (C) Quantification of PLAs.
Quantification of PLAs conducted in B. Asterisks indicate a difference from the value of SHIP2/RhoA PLA cells at
p < 0.01. Error bars indicate ± SD. (D) Association of SHIP2 with RhoA by PLA in spreading U251 cells. U251 cells were
suspended and seeded on FN-coated glasses and fixed at 30 min and 1, 2, 4, and, 8 h after plating. PLA was performed
using antibodies against both SHIP2 and RhoA. Cells were stained with anti-PI(3,4,5)P3, anti-RhoA, and anti-SHIP2
antibodies. PLA images represent focused views of several confocal sections that covered the entire region of cells. Cell
edges are marked by a dotted line. Nuclei were stained with DAPI (blue). Bar, 10 μm. All results are representative of at
least three independent experiments. At least 50 cells were counted per experiment.
Average number of signal per cell
RhoA Merged SHIP2
RhoA MergedSHIP2 PLAs PI(3,4,5)P3
Volume 23 July 1, 2012 RhoA effector SHIP2 and cell polarity | 2599
proximity ligation assays revealed the asso-
ciation of SHIP2 with RhoA in U251 cells.
The depletion of SHIP2 impaired not only
the front–rear polarity and polarized distri-
bution of PI(3,4,5)P3 but also cell migration.
Wild-type SHIP2 rescued the SHIP2-knock-
down phenotypes, including defects in
front–rear polarity and haptotactic migra-
tion, whereas the SHIP2 mutant defective in
RhoA binding could not rescue these phe-
notypes. Thus SHIP2 appears to regulate
PI(3,4,5)P3 levels and restrict its localization
to a single site, thereby controlling cell po-
larity and cell migration downstream of
Rho family GTPases, including RhoA,
Rac1, and Cdc42, have been implicated in
establishing the front–rear polarity of mi-
grating cells. Rac1 and Cdc42 are predomi-
nantly activated at the front leading edge to
generate a vectorial protrusion. The leading
protrusion is stabilized by its adherence to
the surrounding extracellular matrix through
adhesion receptors, such as integrins, which
in turn induces PI(3,4,5)P3 production and
thereby activates Rac1 and Cdc42 through
guanine nucleotide exchange factors (GEFs)
to further induce new protrusions (Rameh
et al., 1997; Han et al., 1998). PI(3,4,5)P3
controls the localization of DOCK2, a GEF
for Rac, leading to the regulation of mem-
brane translocation and Rac activation
(Kunisaki et al., 2006). PI(3,4,5)P3 also acti-
vates Akt to maintain the actin stress fibers
and cortical actin filaments for directional
cell migration through Girdin, a downstream
substrate of Akt (Weng et al., 2010). In con-
trast, RhoA activity is mainly activated in the
central and rear regions, which may ensure
front–rear polarity (Jaffe and Hall, 2005;
Etienne-Manneville, 2008; Iden and Collard,
2008). RhoA is believed to inhibit Rac1 activ-
ity, thereby preventing ectopic protrusion in
the rear region (Worthylake and Burridge,
2003). It has been reported that RhoA inhib-
its the Rac1 activity acting via Rho-kinase to
phosphorylate FilGAP and Par-3 (Ohta et al.,
2006; Nakayama et al., 2008). Rho-kinase
also phosphorylates myosin phosphatase
and inactivates it, thereby increasing the
phosphorylation of myosin light chain and
activating myosin II (Kimura et al., 1996), which may inhibit Rac1
activity through Rac1 GEF (Lee et al., 2010). In addition to Rho-
kinase, we found that RhoA maintains the proper localization and
levels of PI(3,4,5)P3 via SHIP2 in migrating cells, thereby contribut-
ing to the establishment of front–rear polarity.
How does RhoA regulate SHIP2 functions? We examined the ef-
fect of RhoA on the phosphatase activity of SHIP2 in a cell-free sys-
tem and found that the GTP-bound RhoA did not affect phosphatase
activity toward PI(3,4,5)P3 under our assay conditions, suggesting
that RhoA does not modulate the enzymatic activity of SHIP2 (Sup-
plemental Figure S3). However, we cannot rule out the possibility
sets of siRNAs to SHIP2 and RhoA showed similar effects on cell
migration. The inhibitory effect of SHIP2 depletion was rescued by
the expression of RNAiR-SHIP2-WT but not by the expression of
RNAiR-SHIP2-D193/E195A or D608A (Figure 7), suggesting that the
interaction of RhoA and SHIP2 is required for cell migration.
In this study, we identified several novel RhoA effector candidates,
including SHIP2, by affinity chromatography of the GTP-bound form
of RhoA. The N-terminal domain of SHIP2 directly and specifically
interacted with the GTP-bound, but not GDP-bound, RhoA. The
FIGURE 5: Requirement of SHIP2 for proper polarization in U251 cells. (A) Effect of the
depletion of SHIP2 and RhoA in U251 cells. U251 cells transfected with the indicated siRNAs for
72 h were reseeded on FN-coated glasses for 8 h, fixed, and stained with tetramethylrhodamine
isothiocyanate (TRITC)–phalloidin. Bar, 10 μm. (B) Rescue experiments of SHIP2 knockdown in
U251 cells. U251 cells transfected with both the indicated siRNAs and plasmids were reseeded
on FN-coated glasses for 8 h, fixed, and stained with TRITC–phalloidin and the anti-GFP
antibody. White arrows indicate transfected cells. Bar, 20 μm. (C) Quantification of the effects of
SHIP2 and RhoA mutants on cell polarity. The percentage of the cells that had a single leading
edge is shown. Asterisks indicate a difference from the value of control cells at p < 0.01. Error
bars indicate ± SD. All results are representative of at least three independent experiments. At
least 100 cells were counted per experiment.
siSHIP2#1 + #2
siSHIP2#1 + #2
% polarized cells
2600 | K. Kato et al. Molecular Biology of the Cell
speculate that RhoA functions as a cutoff fil-
ter for PI(3,4,5)P3 to suppress ectopic accu-
mulation through local regulation of SHIP2
activity and/or localization. The SHIP2–RhoA
interaction was observed ubiquitously in
cells (Figure 4B), but we could not identify
the exact cellular compartments in which
the PLA signals were located due to the low
resolution of this analysis. The distribution of
PI(3,4,5)P3 was found not only at the plasma
membrane, but also localized to endocy-
tosed vesicles, nuclei, and intracellular
membranes (Sato et al., 2003; Horiguchi
et al., 2006; Lindsay et al., 2006). Of note,
we cannot exclude the possibility that polar-
ized PI(3,4,5)P3 localization is disturbed as
the consequence of disruption in polarized
leading edge formation or other molecules
in the regulation and localization of SHIP2
are involved. Further analysis of the site of
colocalization is necessary to understand
the link between SHIP2 and RhoA
In addition to SHIP2, we identified novel
RhoA effector candidates, including ACAT1,
FILIP1, STRN3, and BZW (Supplemental
Table S1). These candidates specifically in-
teract with GTP-bound RhoA but not the
GDP-bound form. For example, ACAT1 is
important in pathways for the synthesis and
degradation of ketone bodies and for the
degradation of 2-methylacetoacetyl-CoA
(Haapalainen et al., 2007). FILIP negatively
controls the function of filamin A through
the degradation of filamin A, leading to
the onset of neocortical migration from the
ventricular zone (Sato and Nagano, 2005).
Although we cannot exclude the possibility
that these candidates can indirectly interact
with GTP-bound RhoA, they provide us with
a useful platform with which to further un-
derstand RhoA functions. It should be noted
that there are several intriguing proteins,
such as SLK, VAT1, PLCB4, and ACSL1, that
interact with both GTP- and GDP-bound
RhoA but not the GST control and could be
novel types of RhoA regulators. Detailed
analyses of these putative effectors and
partners are underway.
MATERIALS AND METHODS
Materials and chemicals
The cDNA encoding mouse SHIP2 was
kindly provided by M. Matsuda (Kyoto
University, Kyoto, Japan). The pCAGGS vec-
tor was kindly provided by M. Nakafuku (Cincinnati Children’s Hospi-
tal, Cincinnati, OH). The antibodies used were as follows: monoclo-
nal anti-GFP (Roche Diagnostics, Mannheim, Germany); polyclonal
anti-GFP (MBL, Nagoya, Japan); monoclonal anti-SHIP2 and anti-HA
(Cell Signaling Technology, Beverly, MA); polyclonal anti-INPPL1 (Ab-
cam, Cambridge, United Kingdom); monoclonal anti-PI(3,4,5)P3
(Echelon Bioscience, Salt Lake City, UT); monoclonal anti-RhoA
that we cannot satisfactorily reconstitute the in vivo situation with
purified RhoA and SHIP2. Because RhoA-binding activity is required
for SHIP2 to control cell polarization and migration (Figures 5, B and
C, and 7) and SHIP2 is partly recruited by RhoA in a RhoA activity–
dependent manner (Supplemental Figure S4), RhoA may be needed
to properly localize SHIP2 at the membranes to prevent the over-
production of PI(3,4,5)P3, which may inhibit ectopic protrusion. We
FIGURE 6: Requirement of SHIP2 for the proper accumulation of PI(3,4,5)P3. (A) PI(3,4,5)P3
localization in SHIP2- or RhoA-depleted U251 cells. U251 cells transfected with the indicated
siRNAs for 72 h were reseeded on FN-coated glasses for 8 h, cultured for 1 h in the presence or
absence of 50 μM LY294002, fixed, and stained with Alexa 488–phalloidin and the anti-PI(3,4,5)P3
antibody. Bar, 10 μm. (B) Quantification of the effects of SHIP2 or RhoA knockdown on the
accumulation of PI(3,4,5)P3 at the leading edge. The percentage of the cells in which the
accumulation of PI(3,4,5)P3 was observed at a single leading edge is shown. Asterisks indicate a
difference from the value of control cells at p < 0.01. Error bars indicate ± SD. (C) Rescue
experiments of SHIP2 knockdown in U251 cells. U251 cells transfected with both the indicated
siRNAs and plasmids were reseeded on FN-coated glasses for 8 h, fixed, and stained with
anti-PI(3,4,5)P3 and anti-GFP antibodies. White arrows indicate transfected cells. Bar, 20 μm
(D) Quantification of the effects of SHIP2 knockdown and rescue mutants on the accumulation
of PI(3,4,5)P3 at the leading edge. The percentage of the cells in which the accumulation of
PI(3,4,5)P3 was observed at a single leading edge is shown. Asterisks indicate a difference from
the value of control cells at p < 0.01. Error bars indicate ± SD. All results are representative of at
least three independent experiments. At least 100 cells were counted per experiment.
siSHIP2#1 + #2
at leading edge (%)
siSHIP2#1 + #2
at leading edge (%)
Volume 23 July 1, 2012 RhoA effector SHIP2 and cell polarity | 2601
into pGEX (GE Biohealthcare Bioscience, Piscataway, NJ), pMAL
(New England Biolabs, Beverly, MA), pCAGGS-myc, pEGFP (Takara,
Otsu, Japan), pmCherry, pEF-BOS-GST, and pEF-BOS-HA vectors.
pMitoEGFP vector was prepared as previously described (Rojo
et al., 2002). RhoA-A37L63, RhoA-L63, RhoA-L63S190, Rac1-L61,
Cdc42-L61, SHIP2-D608A, SHIP2-D193/E195A, SHIP2-S202/
N203A, SHIP2-R216/R217A, and SHIP2-D223/K224A cDNAs were
generated by site-directed mutagenesis. siRNA-resistant SHIP2 mu-
tants were generated by site-directed mutagenesis that introduced
silent mutations within the siRNA target sequence. All fragments
were confirmed by DNA sequencing.
GST- and MBP-tagged proteins were produced in BL21 (DE3)
Escherichia coli cells and purified on glutathione–Sepharose 4B
beads (GE Biohealthcare Bioscience) and amylose resin (New
England Biolabs), respectively. GST-RhoA-WT for phosphatase
assay was produced in Sf9 cells with a baculovirus system and
purified as previously described (Mizuno et al., 1991). In brief,
GST-RhoA-WT–expressing Sf9 cells (1 × 108 cells) were suspended
with 10 ml of homogenate buffer (10 mM Tris-HCl, pH 7.5, 1 mM
dithiothreitol [DTT], 10 mM MgCl2, 100 μM (p-amidinophenyl)
methanesulfonyl fluoride, and 10 μg/ml leupeptin). This suspen-
sion was sonicated and centrifuged at 100,000 × g for 1 h at 4°C.
The posttranslationally processed form of GST-RhoA was purified
from the membrane fraction. The 100,000 × g pellet was resus-
pended in buffer (20 mM Tris-HCl, pH 8.0, 1 mM EDTA, 1 mM DTT,
5 mM MgCl2, and 0.6% 3-[(3-cholamidopropyl)dimethylammonio]-
1-propanesulfonate [CHAPS]) , stirred for 30 min, and then centri-
fuged at 100,000 × g for 1 h at 4°C. The supernatant was purified
on glutathione–Sepharose 4B beads. Recombinant Flag-tagged
SHIP2 protein was purified as previously described (Hasegawa
et al., 2011).
Preparation of the rat heart cytosolic fraction
Rat heart (30 g) was quickly frozen with liquid nitrogen and pulver-
ized with a Cryo-Press (15 s × 8; Microtec Co., Chiba, Japan). The
frozen heart powder was homogenized in 100 ml of homogenizing
buffer (25 mM Tris-HCl, pH 7.5, 5 mM EDTA, 1 mM DTT, 5 mM
MgCl2, 500 mM NaCl, 100 μM (p-amidinophenyl)methanesulfonyl
fluoride, 2 μg/ml leupeptin, and 10% sucrose) and centrifuged at
20,000 × g for 30 min at 4°C. The supernatant was centrifuged at
100,000 × g for 1 h at 4°C. The supernatant was dialyzed three times
against buffer A (20 mM Tris-HCl, pH 7.5, 1 mM EDTA, 1 mM DTT,
and 5 mM MgCl2). After dialysis, the cytosolic fraction was centri-
fuged at 20,000 × g for 1 h at 4°C. Saturated ammonium sulfate was
then added to a final concentration of 60% saturation. After centrifu-
gation at 20,000 × g for 1 h at 4°C, the precipitate was dissolved in
10 ml of buffer A, dialyzed three times against buffer A, and used as
the cytosolic fraction.
Affinity column chromatography
Affinity column chromatography was performed as previously de-
scribed (Hikita et al., 2009). Briefly, the rat heart cytosol fraction was
loaded onto a glutathione–Sepharose 4B affinity column on which
5 nmol of GST, GDP⋅GST-RhoA, GTPγS⋅GST-RhoA, GST-RhoA-N19,
or GST-RhoA-L63 was immobilized. The guanine nucleotide–bound
forms of GST-RhoA were made by incubating GST-RhoA for 20 min
at 30°C with 150 μM GDP or GTPγS in a reaction mixture (50 mM
Tris-HCl, pH 7.5, 10 mM EDTA, 1 mM DTT, and 5 mM MgCl2). After
washing of the columns three times with buffer A and three times
with buffer A containing 50 mM NaCl, the bound proteins were
and anti-SHIP1, polyclonal anti-SHIP2 and anti-RhoGDI (Santa Cruz
Biotechnology, Santa Cruz, CA); polyclonal anti-ACAT1 (GeneTex,
Irvine, CA); monoclonal anti–α-tubulin (Sigma-Aldrich, St. Louis,
MO); monoclonal anti-myc (Wako Pure Chemical Industries, Osaka,
Japan); and monoclonal anti–N-cadherin (BD Biosciences, San Jose,
CA). The polyclonal anti–Rho-kinase antibody was prepared as previ-
ously described (Katoh et al., 2001). The polyclonal anti-GST anti-
body was produced as previously described (Nishimura et al., 2004).
Fibronectin was purchased from BD Biosciences. Y-27632 was from
Enzo Life Sciences (Farmingdale, NY). Wortmannin and LY294002
were from Calbiochem (San Diego, CA). The Duolink II Detection Kit
with PLA PLUS and MINUS Probes for mouse and rabbit were
purchased from Olink Bioscience (Uppsala, Sweden). (±)-3-Hydroxy-
3-methyl-5-pentanolide was purchased from Wako Pure Chemical
Industries. Purified phosphatidylserine was purchased from Avanti
Polar Lipids, (Alabaster, AL). PI(3,4,5)P3 was obtained from CellSig-
nals (Columbus, OH). siRNA sequences were as follows: SHIP2#1,
5′-GGUGUUUGACCAGCAGAGC-3′; control, 5′-CAGUCGCGUUU-
GCGACUGG-3′. These siRNAs with dTdT overhangs at each 3′ ter-
minus were obtained from Greiner-Japan (Tokyo, Japan). siRNAs
with dTdT overhangs at each 3′ terminus to SHIP2#2 and RhoA were
purchased from Sigma-Aldrich: SHIP2#2, SASI_Hs01_00013990;
RhoA#1, 5′-AGCAGGUAGAGUUGGCUUU-3′; RhoA#2, 5′-GGAUC-
UUCGGAAUGAUGAG-3′. Other materials and chemicals were ob-
tained from commercial sources.
The cDNA encoding full-length PTEN was cloned from a human
fetal brain cDNA library (Clontech Laboratories, Palo Alto, CA).
RhoA and SHIP2 fragments were amplified by PCR and subcloned
FIGURE 7: Requirement of the RhoA-binding activity of SHIP2 for cell
migration. U251 cells transfected with siRNA along with the indicated
plasmids were subjected to the Boyden chamber assay. The cells were
allowed to migrate for 2 h. Cells were fixed and stained with the
anti-GFP antibody. All results are representative of at least three
independent experiments. Asterisks indicate a difference from the
value of control cells at p < 0.01. Error bars indicate ± SD. At least
300 EGFP-positive cells were counted per experiment.
siControl + GFP-GST
siSHIP2#1 + GFP-GST siSHIP2#2 + GFP-GST
siRhoA#1 + GFP-GST siRhoA#2 + GFP-GST
2602 | K. Kato et al. Molecular Biology of the Cell
100 μM (p-amidinophenyl)methanesulfonyl fluoride, 2 μg/ml leu-
peptin, and 2 μg/ml aprotinin), sonicated, and centrifuged at 700 ×
g for 5 min. The homogenate was centrifuged at 100,000 × g for
30 min. The supernatant was used as the cytosol fraction. The pellet
was resuspended in lysis buffer containing 1% NP-40, rocked for 1
h, and then centrifuged at 100,000 × g for 30 min. The supernatant
was used as the membrane fraction. Aliquots of the cytosol and
membrane fractions were subjected to SDS–PAGE and probed with
the anti-myc or HA antibody.
In vitro phosphoinositide phosphatase assay
Phosphoinositide phosphatase activities were examined as previ-
ously described (Maehama and Dixon, 1998). In brief, phosphatase
assay was performed at 37°C in buffer consisting of 50 mM
4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid–NaOH, pH 7.4,
5 mM MgCl2, 2 mM DTT, 50 μM PI(3,4,5)P3, 0.5 mM phosphatidyl-
serine, 0.05% CHAPS, and 0.13 μM Flag-SHIP2 in the presence or
absence of 1.2 μM GST, GDP⋅GST-RhoA, or GTPγS⋅GST-RhoA for 5
min. The reaction was terminated by the addition of 0.1 M EDTA.
Released free phosphate was detected with BIOMOL GREEN (Enzo
GTP-Rho pull-down assay
The activity of RhoA was determined by pull-down assay with the
GST-Rho–binding domain of Rhotekin (GST-Rhotekin-RBD) as previ-
ously described (Mori et al., 2009). In brief, the cells were washed
with ice-cold phosphate-buffered saline (PBS) and lysed in lysis buf-
fer (50 mM Tris-HCl, pH 7.5, 1 mM EGTA, 10 mM MgCl2, 500 mM
NaCl, 0.5% NP-40, 0.1 mM (p-amidinophenyl)methanesulfonyl fluo-
ride, 2.5 mg/ml aprotinin, and 2.5 μg/ml leupeptin) containing 20 μg
of GST-Rhotekin-RBD. The lysates were centrifuged at 20,000 × g
for 3 min at 4°C, and the supernatants were incubated with glutathi-
one–Sepharose 4B beads for 30 min at 4°C. The beads were washed
with an excess of lysis buffer and then eluted with SDS-sample buf-
fer. The eluates were subjected to SDS–PAGE, followed by immuno-
blot analysis with the anti-RhoA antibody.
Cell culture and transfection
U251 and COS7 cells were maintained in DMEM (Sigma-Aldrich)
supplemented with 10% fetal bovine serum (FBS; SAFC Biosciences,
St. Louis, MO). MDA-MB-231 cells were maintained in Leibovitz’s
L-15 Medium (Invitrogen, Carlsbad, CA) supplemented with 10%
FBS. U251 cells were transfected with plasmids or siRNA by Oligo-
fectamine or Lipofectamine LTX (Invitrogen) or by Amaxa Nucleo-
fector (Lonza, Basel, Switzerland) according to the manufacturers’
protocols. COS7 cells were transfected by Lipofectamine 2000 or
Lipofectamine (Invitrogen) according to the manufacturer’s proto-
cols. MDA-MB-231 cells were transfected with siRNA by Lipo-
fectamine RNAiMAX (Invitrogen) or plasmids by Neon Transfection
System (Invitrogen) according to the manufacturer’s protocols.
U251 cells were fixed with PBS containing 3.7% formaldehyde for
10 min at room temperature (RT). followed by permeabilization
with 0.2% Triton X-100 for 10 min at RT. After washing with PBS, the
cells were blocked with 1% BSA (Nacalai Tesque, Kyoto, Japan) for
30 min at RT and incubated with primary antibody for 1 h at RT. For
anti-RhoA staining, the cells were fixed with ice-cold 10% trichlo-
roacetic acid for 15 min at 4°C and then permeabilized with 0.1%
Triton X-100 for 10 min at RT. After washing with PBS, cells were
incubated with primary antibody for 1 h at RT. The secondary anti-
bodies were Alexa 488– and 555–conjugated antibodies against
eluted three times with buffer A containing 1 M NaCl. The first elu-
ates were subjected to SDS–PAGE, and proteins were detected by
The proteins in the eluate were digested by trypsin for 16 h at 37°C
after reduction, alkylation, demineralization, and concentration.
Nanoelectrospray tandem mass analysis was performed using an
LTQ Orbitrap XL mass spectrometry system (ThermoFisher Scien-
tific, Waltham, MA) combined with a Paradigm MS4 HPLC System
(Michrom BioResources, Auburn, CA). Samples were injected onto
the Paradigm MS4 HPLC System equipped with a Magic C18AQ
column 0.1 mm in diameter and 50 or 150 mm in length (Michrom
BioResources). Reverse-phase chromatography was performed
with a linear gradient (0 min, 5% B; 100 min, 50% B) of solvent A
(2% acetonitrile with 0.1% formic acid) and solvent B (90% ace-
tonitrile with 0.1% formic acid) at an estimated flow rate of 1 μl/min.
Ionization was performed with an ADVANCE Captive Spray Source
(Michrom BioResources) with a capillary voltage of 1.6 kV and tem-
perature of 150°C. A precursor ion scan was carried out using mass-
to-charge ratio (m/z) of 400–2000 before MS/MS analysis. Multiple
MS/MS spectra were submitted to the Mascot program, version
2.3.02 (Matrix Science, Boston, MA), for the MS/MS ion search.
Peptides and proteins were searched against the Swiss Prot data-
base (Swiss Prot_2010_09) by Mascot. Search parameters for Mas-
cot included the following: variable modifications carbamidomethyl
(C) and oxidation (M); mass values, monoisotopic; protein mass, un-
restricted; peptide mass tolerance, 10 ppm; fragment mass toler-
ance, 0.8 Da; max missed cleavages, 1; instrument type, ESI-TRAP.
Scaffold including X! Tandem, version 3.0 (Proteome Software, Port-
land, OR), was used for validation. Protein identifications were ac-
cepted if they could be established at >95.0% probability and con-
tained at least two identified peptides.
In vitro binding assays
In vitro binding assays were performed as previously described
(Nishimura et al., 2004). Briefly, deletion mutants were immobilized
separately onto glutathione–Sepharose 4B beads. The immobilized
beads were incubated with MBP-SHIP2 deletion mutants in buffer A
containing 50 mM NaCl, 0.1% NP-40, and 0.2 mg/ml bovine serum
albumin (BSA) for 1 h at 4°C. Beads were washed five times with
buffer A containing 50 mM NaCl and 0.1% NP-40 and then sus-
pended in SDS–PAGE sample buffer. The bound proteins were sub-
jected to SDS–PAGE, and proteins were detected by silver staining.
For pull-down assay, the immobilized beads or glutathione-Sephar-
ose 4B beads were incubated with COS7 cell lysate transfected with
the indicated plasmid in lysis buffer (50 mM Tris-HCl, pH 7.5, 1 mM
ethylene glycol tetraacetic acid [EGTA], 1 mM DTT, 10 mM MgCl2,
150 mM NaCl, 1% NP-40, 100 μM (p-amidinophenyl)methanesulfo-
nyl fluoride, 2 μg/ml leupeptin, and 2 μg/ml aprotinin) for 1 h at 4°C.
The beads were washed three times with lysis buffer, and then the
washed beads were suspended in SDS–PAGE sample buffer. The
bound proteins were subjected to an immunoblot analysis with the
Localization of SHIP2 regulated by RhoA in U251 cells
The subcellular localization of SHIP2 and RhoA was examined as
previously described (Hinoi et al., 1996). In brief, U251 cells trans-
fected with the indicated plasmids were suspended in lysis buffer
(50 mM Tris-HCl, pH 7.5, 1 mM EGTA, 1 mM DTT, 5 mM MgCl2,
Volume 23 July 1, 2012 RhoA effector SHIP2 and cell polarity | 2603
mouse immunoglobulin G (IgG) or rabbit IgG (Invitrogen). For anti-
PI(3,4,5)P3 staining in U251 cells, cells were fixed with 4% paraform-
aldehyde overnight at 4°C, followed by permeabilization for 45 min
with PBS containing 10% normal donkey serum and 0.5% saponin,
overnight incubation with the anti-PI(3,4,5)P3 antibody at 4°C, and
detection by overnight treatment using Alexa 555–conjugated anti-
body against mouse IgG at 4°C. For anti-PI(3,4,5)P3 staining in
MDA-MB-231 cells, we followed a previous protocol (Yip et al.,
2008). In brief, cells, fixed/permeabilized by 4% paraformalde-
hyde/0.1% glutaraldehyde in 0.15 mg/ml saponin solution for 1 h at
37°C were stained sequentially with the anti-PI(3,4,5)P3 antibody for
1 h and the Alexa 555–conjugated secondary antibody for 45 min.
Confocal images were recorded by LSM780, by LSM5 Pascal,
by LSM5 built around Axio Observer Z1, Axiovert 200M, or 100M
with Plan-Apochromat 20× (numerical aperture [NA] 0.75), Plan-
Apochromat 20× (NA 0.8), Plan-NEO Fluar 40× (NA 0.75),
C-Apochromat 40× (NA 1.2), or Plan Apochromat 63× (NA 1.40)
lenses under the control of LSM software (Carl Zeiss, Jena, Germany)
or by a Nikon A1 confocal laser scanning microscope built around
ECLIPSE Ti with CFI Apo 40×WIλS (NA 1.25), Plan Apo VC 60×WI
(NA 1.2), or Plan Apo VC 60× (NA 1.4) lenses under the control of
NIS-elements software (Nikon, Tokyo, Japan). Images were pro-
cessed using Photoshop (Adobe, San Jose, CA). Duolink images
were processed using NIS-elements software and Photoshop.
Proximity ligation in situ assay (Duolink)
The interactions between SHIP2 and RhoA in U251 cells were ana-
lyzed using the Duolink II proximity ligation in situ assay (Fredriksson
et al., 2002; Soderberg et al., 2006) according to the manufacturer’s
instructions. The anti-INPPL1 rabbit polyclonal antibody was com-
bined with the anti-RhoA mouse monoclonal antibody. Fluorescence
spots generated were automatically counted, and the average num-
ber of spots per cell was calculated using MetaMorph (Molecular
Devices, Downingtown, PA).
Quantification of polarity
Polarity was defined as cells with a single leading edge as evaluated
by phalloidin staining. Quantification of polarity was performed with
at least three independent experiments. The evaluations were per-
formed with at least 100 cells in each experiment.
Boyden chamber assay
The Boyden chamber assay was performed as previously described
(Watanabe et al., 2009). Briefly, siRNA was transfected into U251
cells with the indicated plasmid. Cell migration assays were per-
formed using Transwell plates (pore size, 8 μm; HTS FluoroBlok In-
sert; BD Biosciences). The undersurface of the membrane was
coated for 1 h at RT with 10 μg/ml fibronectin diluted in distilled
water. The cells were seeded in the upper chamber (1 × 104 per well)
in 500 μl of DMEM with 0.1% BSA. DMEM supplemented with 0.1%
BSA and 10 ng/ml EGF was added to the lower chamber. The cells
were allowed to migrate for 2 h. After fixation, both nonmigrated
and migrated EGFP-positive cells were counted by EGFP fluores-
cence, and the ratio of migrated cells to total (migrated plus nonmi-
grated) cells was calculated and determined as the migration index.
At least 300 EGFP-positive cells were counted in each group for
each experiment. The results were normalized and expressed as a
Student’s t test and one-way analysis of variance (ANOVA) were per-
formed after data were confirmed to satisfy the criteria of normal
We thank M. Matsuda (Kyoto University, Kyoto, Japan) for kindly
providing SHIP2 cDNA and helpful discussion. We thank M. Inagaki
(Aichi Cancer Center Research Institute, Nagoya, Japan) for kindly
providing U251 cells. We thank Frank B. Gertler (Massachusetts In-
stitute of Technology, Cambridge, MA) and T. Nishimura (RIKEN
Center for Developmental Biology, Kobe, Japan) for helpful discus-
sions. We thank F. Ishidate for help with acquiring and analyzing
images and A. Iwamatsu (Protein Research Network, Kanagawa,
Japan), Y. Yamakawa (Division for Medical Research Engineering,
Nagoya University, Japan), ThermoFisher Scientific, and AMR
(Tokyo, Japan) for mass spectrometric analysis. We also thank all
members of the Kaibuchi lab for discussions and technical support,
Y. Kanazawa for technical assistance, and T. Ishii for secretarial
assistance. We also thank T. Watanabe for helpful discussions,
T. Namba, S. Nakamuta, K. Kuroda, and Y. Funahashi (our labora-
tory) for help with microscopy, and H. Shohag and X. Zhang for
technical support. We acknowledge the Division of Medical Re-
search Engineering of the Nagoya University Graduate School of
Medicine for the use of ImageQuant LAS 4010 (GE Biohealthcare
Bioscience) and Nikon-A1 microscopy, and we thank the Radioiso-
tope Center Medical Branch, Nagoya University School of Medi-
cine (Technical Staff, N. Hamada and Y. Nakamura). This research
was supported in part by Nagoya University Global COE Program,
Integrated Functional Molecular Medicine for Neuronal and Neo-
plastic Disorders, Grants-in-Aid for Scientific Research (S; 20227006)
and (C; 23590357) from the Ministry of Education, Culture, Sports,
Science and Technology of Japan, and the Kyosaidan Foundation,
the Nagono Medical Foundation, and the Japan Foundation for
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