|
|
|
|
|
|
|
||||
| Home Help Feedback Subscriptions Archive Search Table of Contents | ||||
First published online 13 June 2007
doi: 10.1242/dev.002824
| |||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||
1 Children's Hospital Research Foundation, Division of Developmental Biology,
Cincinnati, OH 45229, USA.
2 Molecular and Developmental Biology Graduate Program, University of Cincinnati
College of Medicine, OH 45219, USA.
3 Department of Biochemistry and Molecular Biology, Program in Genes and
Development, University of Texas Graduate School of Biomedical Sciences,
Houston, TX 77030, USA.
Author for correspondence (e-mail:
Christopher.Wylie{at}cchmc.org)
Accepted 8 May 2007
 |
SUMMARY |
|---|
 TOP SUMMARY INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
|---|
Key words: Cortical actin, Actin assembly, Xenopus, GPCR, Cadherins, C-cadherin, p120 catenin, Xflop, LPA
|
INTRODUCTION |
|---|
|
TOPSUMMARYINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
|---|
;
Tao et al., 2005). The
mechanism by which the cortical actin skeleton is generated and maintained is
therefore an important problem in early development.
We have shown previously that the pattern and density of cortical actin
changes through the cell cycle. During interphase, each blastomere possesses a
dense cortical actin network, which is replaced during mitosis by a much
sparser network of actin filament bundles
(Lloyd et al., 2005).
Dissociation of the blastomeres, so that cell contact and intercellular
signals are prevented, causes all blastomeres to adopt the sparse
configuration of actin, which is reversible by reassociation
(Lloyd et al., 2005). Previous
expression screening experiments identified two proteins that are required for
assembly of cortical actin. Receptors for the signaling phospholipid
lysophosphatidic acid (LPA) and an orphan G-protein-coupled receptor (GPCR)
named Xflop, were each found, by gain- and loss-of-function experiments, to be
both necessary and sufficient for assembly of the dense cortical actin network
(Lloyd et al., 2005;
Tao et al., 2005). These
experiments indicated the novel fact that intercellular signaling plays a
major role in generating the appropriate density and pattern of cortical actin
assembly in the early embryo.
It therefore becomes important to determine the sites of actin assembly at the cell surface and the way the G-protein-coupled receptors control the amount and pattern of actin assembly, so as to generate the appropriate shape and mechanical rigidity of the whole embryo.
Recent elegant experiments using epithelial cells in culture have shown
that during initial contacts between epithelial cells the cytoplasmic domains
of transmembrane cadherins are sites of actin assembly
(Ehrlich et al., 2002;
Jamora and Fuchs, 2002;
Kovacs et al., 2002a;
Kovacs et al., 2002b;
Vaezi et al., 2002).
E-cadherin engagement in cultured cells causes activation of Rac1 and Cdc42,
both of which are involved in actin polymerization and organization
(Betson et al., 2002;
Kim et al., 2000;
Kovacs et al., 2002a;
Nakagawa et al., 2001;
Noren et al., 2001). Actin
nucleation proteins such as members of the Arp2/3 complex and formin 1 are
associated with nascent cadherin-mediated adhesive contacts
(Kobielak et al., 2004;
Kovacs et al., 2002b;
Verma et al., 2004), as are
nucleation-promoting factors such as Ena/Vasp and cortactin
(Helwani et al., 2004;
Scott et al., 2006;
Vasioukhin et al., 2000).
However, the detailed mechanism by which cortical actin assembles at the
cadherin complex is still unknown (Adams
and Nelson, 1998; Drees et
al., 2005; Perez-Moreno et
al., 2003; Yamada et al.,
2005).
Could cadherins be the sites of assembly of cortical actin in the
Xenopus blastula? It is known that the early blastomeres are held
together by calcium-dependent adhesion
(Nomura et al., 1988;
Turner et al., 1992). The
calcium-dependent adhesion protein C-cadherin (also known as EP-cadherin)
(Choi et al., 1990;
Ginsberg et al., 1991) is
expressed during the egg-to-blastula stages and has been shown, by
antisense-mediated mRNA depletion, to be absolutely required for cell adhesion
at this stage (Heasman et al.,
1994).
In this paper, we show that C-cadherin is essential for the assembly of the dense cortical network seen in interphase blastomeres, but not for the sparse network seen during cell divisions. First, we show that C-cadherin proteins are organized as punctae on the surfaces of the blastomeres and that the actin filament network is associated with these punctae. Second, we show by both gain- and loss-of function experiments that the level of C-cadherin expression is both necessary and sufficient for the dense cortical actin network. Increasing the level of C-cadherin expression leads to increased cortical actin density, whereas decreasing C-cadherin expression, by depletion of its mRNA, reduces it. We also show that the juxtamembrane domain of C-cadherin, which binds p120 catenin, is essential for its control of actin assembly. Consistent with this, we show that p120 catenin is required for dense actin network assembly and that a mutant form of p120 catenin that lacks the binding site for C-cadherin fails to increase actin assembly. These data show that the cadherin complex is both necessary and sufficient for assembly of the normal pattern and density of cortical actin in the Xenopus blastula, and that this activity requires p120.
If cortical actin assembly depends on the expression level of C-cadherin, then LPA signaling, and signaling through the G-protein-coupled receptor Xflop, could act either at the level of actin nucleation, or at the level of C-cadherin presentation on the cell membrane (or both). We show here that both signaling pathways control the level of C-cadherin expression on the cell surface. These data show for the first time how C-cadherin levels on the cell surface are controlled in the Xenopus embryo and the consequences of this control for actin assembly.
|
MATERIALS AND METHODS |
|---|
|
TOPSUMMARYINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
|---|
).
DNA constructs and RNA
To generate a p120 catenin mRNA to which the MO could not bind, we
made a cDNA construct with three third-base alterations using the following
primer pair for PCR cloning: forward,
5'-CGGAATTCTATGGACGAACCTGAGTCTGAAAGTCCG-3'
(underlined are the EcoRI site for subcloning, the ATG start codon,
and the three substitutions at the third base of each codon, respectively);
reverse, 5'-TACGCAGGCAACCTGTAGTG-3'. Xflop R112A mutant was
generated by PCR-based mutagenesis, using the following PCR primers
(underlined nucleotides are designed to substitute arginine with alanine):
Xflop5'-BglII, 5'-AGCAGATCTATGGCGTGTAATCAGAGCTGTGAATAC-3';
R112A r, 5'-CACTGTAGCCACAAAAGCATCCATGGCTATACAGCT-3';
R112A f, 5'-TGTATAGCCATGGATGCTTTTGTGGCTACAGTGTTC-3'; and
Xflop3'-EcoRI, 5'-GTTGAATTCTATCCTGTCCTTTTTGATGACCTCCTTC-3'.
The primer pair of Xflop5'BglII and R112A r was used to generate fragment 1 using wild-type Xflop cDNA as template, whereas the primer pair of R112A f and Xflop3'EcoRI was used to generate fragment 2 using wild-type Xflop cDNA as template. The primer pair of Xflop5'BglII and Xflop3'-EcoRI was then used to generate the full-length Xflop R112A mutant using the purified fragments 1 and 2 as template. The full-length Xflop R112A was inserted into BamHI and EcoRI-cut pCS107 vector. All PCR products were verified by sequencing.
A Xenopus tropicalis C-cadherin construct was isolated from an
arrayed cDNA library constructed in vector pCS107 (a gift from Aaron Zorn,
University of Cincinnati, OH), linearized with Asp718, then
transcribed with SP6 RNA polymerase. Xenopus C-cadherin mutants
[C-cad(G-A)-HA (referred to below as C-cad G-A), C-cad 
CBD]
(Paulson et al., 2000) were
synthesized with SP6 RNA polymerase from a pCS2+ vector linearized with
NotI. Xenopus p120 iso1, p120 iso1 MO-resistant and the Arm1
deletion mutant (p120 Arm1) were constructed in pCS2-MT vector with a
Myc tag at the N-terminus of p120. For RNA synthesis, we linearized the vector
with NotI and transcribed from the SP6 promoter. pCS107 Xflop and
Xflop R112A and LPA2 were each linearized with Asp718 and transcribed
with SP6 RNA polymerase. Message Machine (Ambion) Kits were used for all in
vitro transcriptions.
Oligonucleotides
The antisense oligodeoxynucleotides (designated AS below) or morpholino
oligonucleotides (designated MO below) used were (asterisks indicate
phosphorothioate-modified residues):
C-cad AS61 (Heasman et al.,
1994),
5'-C*C*T*CTCCAGCTCCCT*A*C*G-3';
P120 catenin AS1, 5'-TGCATCCCTCCATCCTGT-3' (no modification);
P120 catenin MO1 (Fang et al.,
2004), 5'-ACTCTGGCTCATCCATATAGAAAGG-3';
Xflop oligonucleotides (Tao et al.,
2005) 1s,
5'-A*A*G*GGAACACTGTAG*C*C*A-3'
and 5s,
5'-G*T*T*GTACGTTTTGGC*T*G*G-3';
LPA1 MO (Lloyd et al.,
2005), 5'-TTCACTTCAGATGTCAGTCATGCTG-3'; and
LPA2 MO (Lloyd et al.,
2005), 5'-ACCTCCAATGTTACAGCGCAGCCTC-3'.
Immunofluorescence and F-actin staining
For F-actin single staining, caps were excised at the late blastula stage
(St9) (Nieuwkoop and Faber 1967) and fixed with 3.7% formaldehyde, 0.25%
glutaraldehyde in PBS, 0.1% Tween 20 (FG fix) for 10 minutes. Caps were then
washed three times for 10 minutes each and stained with 5 U/ml Alexa
488-conjugated phalloidin (Molecular Probes, Oregon) for 3 hours at room
temperature or overnight at 4°C. For anti-C-cadherin/F-actin double
staining, the phalloidin-stained caps were blocked with 10% normal goat serum
at room temperature for 1 hour and then incubated with 5 ng/ml anti-C-cadherin
monoclonal antibody (6B6, Developmental Study Hybridoma Bank, Iowa) overnight
at 4°C, followed by extensive washing with PBS, 0.3% Triton X-100. A
Cy5-conjugated goat anti-mouse IgG (Jackson Laboratory, 1:300) was then added,
followed by 2 hours incubation at room temperature. In every experiment, 3-5
caps were also incubated with secondary antibody only as negative controls.
For anti-C-cadherin single staining, 2% TCA in water was used to fix animal
caps for 30 minutes at room temperature. After anti-C-cadherin staining, caps
were dehydrated in a methanol series and cleared in Murray's Clear [from
Murray and Kirschner, as cited by Dent and Klymkowsky
(Dent and Klymkowsky, 1989)],
before examination by confocal microscopy.
Confocal microscopy and data acquisition
A Zeiss LSM 510 confocal microscope was used. A Fluar UV/20xNA0.75
objective was used for lower magnification imaging, and a C-Apochromat
63x/NA1.3 water lens or a Plan-Neofluar40x/NA1.3 oil lens was used
with 1.5-2x digital zoom for higher magnification imaging. All images
were gained with the size of 512x512 pixels (8 bit for experiments
represented in Figs 6,
7 and
8, 12 bit in experiments
represented in Figs 2,
3,
4 and
5,
9). The intensity of phalloidin
staining of each pixel measured by LSM510 software was used to quantify the
levels of F-actin. The level of F-actin for each imaged area was the average
(Mi) of intensity measurements from all 512x512 pixels. The level of
F-actin for each experimental treatment group was the average of 5-10 animal
caps (Mt=
Mi/i, where i=number of imaged areas). The linear range of pixel
intensity measurement was from 0 to 255 for 8-bit images or from 0 to 4095 for
12-bit images. In all experiments, the confocal settings were optimized to
allow more than 90% of pixels to have intensity measurement within the linear
range, which might introduce variations among control groups from experiment
to experiment. Each histogram shown in Figs
2,
3,
4,
5,
6,
7,
8 and
9 represents the mean pixel
intensity (Mt) ±s.d. from 5-10 animal caps. The two-tailed
t-test was used to generate P values.
Western blotting
To detect the total protein levels of p120 or C-cadherin, five embryos were
homogenized with 250 µl of ice-cold PBS, 1% Triton X-100 containing 1 mM
PMSF and a 1:100 dilution of protease inhibitor cocktail (PIC, Sigma P8340),
and cleared by centrifugation at 750 g for 10 minutes at
4°C. The supernatants were then transferred into precooled Eppendorf
tubes. An equal volume of 4x sample buffer was added to the cleared
supernatants.
|
Two to three embryo equivalents of membrane protein, or one embryo
equivalent of total protein, were boiled for 5 minutes and separated by 6% SDS
PAGE for 2 hours at 80 volts. Gels were blotted onto nitrocellulose membranes
using a semi-dry apparatus (BioRad). All membranes were subsequently blocked
(1 hour at room temperature) with 5% non-fat dry milk in PBS, 0.1% Tween 20
and incubated in primary antibodies overnight at 4°C. Antibody conditions
were as follows, all diluted in blocking buffer: primary antibody
anti-C-cadherin (Developmental Study Hybridoma Bank, 6B6) 1:500 with secondary
antibody GaM-HRP (Jackson ImmunoResearch) 1:2500; primary antibody anti-human
-catenin (BD Transduction laboratory, C26220) 1:500 with secondary
antibody GaM-HRP 1:2500; primary antibody anti-ß-catenin (Sigma, C2206)
1:2000 with GaR-HRP (Jackson ImmunoResearch) 1:2500; primary antibody
anti-p120 (rabbit polyclonal) (Fang et
al., 2004) 1: 5000 with GaRb-HRP (Jackson ImmunoResearch) 1:2500.
Membranes incubated with HRP-conjugated secondary antibodies were detected
with ECL developing solution (Amersham) and exposed to X-ray films (Hyperfilm,
Amersham) at variable times to obtain unsaturated bands.
|
RESULTS |
|---|
|
TOPSUMMARYINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
|---|
; Levi et al.,
1991). Cadherin clusters have been reported to be present as
punctae in the membranes of epithelial cells during their initial contact,
before becoming resolved into adherens junctions
(Adams et al., 1998;
Adams et al., 1996;
Vasioukhin et al., 2000;
Yonemura et al., 1995). To
analyze the pattern of expression of C-cadherin in Xenopus blastulae,
we followed the method described in Kofron et al.
(Kofron et al., 2002) (see
also Fig. 1A). Dissected animal
caps from late blastulae (Stage 9) were fixed for 30 minutes in 2%
trichloroacetic acid and stained as whole-mounts with a monoclonal antibody
raised against C-cadherin (6B6) (Choi et
al., 1990). Animal caps were cleared in Murray's Clear and
examined in a series of optical slices, starting at the blastocoelic surface.
When individual 1-1.5 µm optical slices were assembled into stacks of
30-70, C-cadherin appeared to extend in a continuous line around each adjacent
cell surface, as previously seen in conventional histological sections
(Heasman et al., 1994).
However, when individual 1.2 µm optical slices were examined, C-cadherin
could be seen as a series of punctae around adjacent plasma membranes
(Fig. 1B). Individual punctae
were measured by pixel counting and had areas of 0.7±0.3
µm2 spaced by distances of 1.9±0.99 µm. This can be
seen particularly clearly in grazing sections through cell surfaces (arrows in
Fig. 1C). When animal caps were
dissociated in calcium- and magnesium-free buffered saline, C-cadherin was
lost from the cell surfaces (Fig.
1D) (see also Hausen and
Riebesell, 2002). When cells were returned to calcium- and
magnesium-containing saline and allowed to contact each other, punctae of
C-cadherin rapidly assembled at the sites of membrane apposition
(Fig. 1E).
|
), this suggested the hypothesis that the dense actin network
might assemble on punctae of C-cadherin. To test this hypothesis, we fixed
animal caps from late blastulae in formaldehyde-glutaraldehyde fixative (see
Materials and methods), stained them with Alexa 488-coupled phalloidin to
reveal the cortical actin network, and then stained them as whole-mounts with
anti-C-cadherin antibodies. Fig.
1F,G show cortical actin filament bundles associated with
C-cadherin-containing punctae (arrows), interspersed with regions of
cadherin-containing membrane not associated with actin.
C-cadherin levels control the dense, but not the sparse, cortical actin networks in Xenopus blastulae
These data suggested that C-cadherin-containing complexes might be required
for cortical actin assembly at the membranes in Xenopus blastulae,
and that the pattern and density of cortical actin in the embryo could be
controlled by the expression of C-cadherin itself, in addition to other
possible levels of control. To test this, we first depleted the maternal mRNA
encoding C-cadherin, by injecting an antisense deoxyoligonucleotide (oligo)
complementary to part of this mRNA, into cultured full-grown oocytes as
described by Heasman et al. (Heasman et
al., 1994). The oocytes were matured in vitro using 1 µM
progesterone and fertilized by the host transfer method
(Heasman et al., 1991).
Fig. 2A,B (left panels) show
C-cadherin protein levels on the cell surfaces in control and
C-cadherin mRNA-depleted embryos at the late blastula stage. There is
a dramatic reduction in the level of staining, consistent with previous
findings (Heasman et al.,
1994). The density of the cortical actin network was
correspondingly decreased in the C-cadherin-depleted embryos (right panels of
Fig. 2A,B). The normal, dense
actin network characteristic of interphase blastomeres was replaced by a
sparser network similar to that seen in dividing or dissociated cells. Second,
C-cadherin mRNA (250-1000 pg per oocyte) was injected into oocytes
before maturation. These were matured in vitro and fertilized as above and the
affects on cortical actin assembly examined at late blastula stage.
Fig. 2C shows that the level of
C-cadherin staining (left panel) and the density of cortical actin (right
panel) were both increased in C-cadherin-overexpressing animal caps compared
with the controls (Fig. 2A).
Pixel intensities from 5-10 caps were measured using the Zeiss LSM 510
software. Statistically significant quantitative changes in actin staining
confirmed the results seen in the individual images
(Fig. 2D). These data show that
the dense actin network is cadherin-dependent, whereas the sparse actin
network seen in dividing or dissociated cells is cadherin-independent.
|
). Two
major functional domains of the cytoplasmic tail are the juxtamembrane region
(JMR) and the distal ß-catenin/plakoglobin (-catenin)-binding
domain (CBD). We used mutant forms of C-cadherin to test the roles of each of
these domains in actin assembly. First, we injected mRNA encoding a mutant
Xenopus C-cadherin in which the residues GGG (aa 731-733) in the JMR
were replaced with AAA. This mutant form is unable to bind p120 catenin
(Thoreson et al., 2000).
Unlike wild-type C-cadherin, overexpression of this mutant form did not
increase the density of the cortical actin
(Fig. 3A, middle lower panel);
in fact, it consistently decreased it (Fig.
3B). A mutant form lacking the distal catenin-binding domain
(C-cad CBD, deletion of aa 839-896) did not significantly alter the
density of cortical actin (Fig.
3A, right lower panel, Fig.
3B). Immunostaining shows that both mRNAs were expressed
(Fig. 3A, upper panels). These
data suggest that both the JMR and CBD domains are required for C-cadherin to
assemble the dense actin network.
In this experiment, the endogenous cadherin was still present and might
have affected trafficking or presentation of the mutant cadherins. To control
for this, we injected the two mutant mRNAs into embryos whose endogenous
C-cadherin had been depleted by antisense oligo injection into the oocytes.
Fig. 4A shows that
C-cadherin mRNA depletion dramatically reduced the level of cadherin
protein on the cell surface. Both mutant forms of cadherin were efficiently
translated, as shown by immunocytochemical staining with anti-C-cadherin
antibody. C-cad G-A was expressed on the cell surface and rescued cell
adhesion (Fig. 4A). However, it
did not rescue the cortical actin network. The only sites of actin
polymerization were in dense membrane projections at cell boundaries (arrowed
in Fig. 4B). However, over the
rest of the cell cortex, there was no dense cortical actin network in these
cells. C-cad CBD was not expressed at the cell surface, but instead
accumulated in large cytoplasmic vesicles
(Fig. 4A,B). This explained the
fact that it did not rescue cell adhesion and precluded an assessment of the
function of this mutant in actin assembly. Actin-rich cell-surface projections
were also seen in these cells, but there was no rescue of the cortical actin
network over the rest of the cell surfaces. The overall levels of polymerized
actin were quantitated by pixel intensity, as shown in
Fig. 4C. Neither of the mutant
forms of cadherin rescued the loss of polymerized cortical actin caused by
C-cadherin depletion. These data show that the p120-binding site of C-cadherin
is required for C-cadherin to assemble a dense cortical actin network.
|
), was
injected into cultured Xenopus oocytes (10-40 ng per oocyte), which
were subsequently matured in vitro and fertilized. The amount of p120 catenin
protein was assessed by western blotting and found to be reduced in a
dose-dependent manner by the mid-blastula stage
(Fig. 5B). The density of the
cortical actin network was also significantly reduced by p120 catenin
depletion (Fig. 5A,C). This
experiment was repeated with an antisense deoxynucleotide oligo, complementary
to a different region of the target mRNA (AS1, 7-9 ng per oocyte), which had
the same effect (data not shown). The effects of both the MO and the antisense
oligo were rescued by reintroducing p120 catenin mRNA (shown for the
antisense oligo experiment in Fig.
5C), showing that in both cases, the effects were specific for
p120 catenin depletion. Thus, p120 catenin is required for cortical actin
assembly at the blastula stage. Next, synthetic p120 catenin mRNA in doses of 250 to 1000 pg was injected into oocytes and the cortical actin examined at the late blastula stage. Fig. 5A,D show that the density of the cortical actin network was increased by the overexpression of p120 catenin. We conclude that p120 catenin is both necessary and sufficient to generate a dense cortical actin network.
p120 catenin is thought to regulate the steady-state levels of cadherins on
the cell surface (Chen et al.,
2003; Davis et al.,
2003; Elia et al.,
2006; Ireton et al.,
2002; Perez-Moreno et al.,
2006; Xiao et al.,
2003). We therefore asked whether p120 catenin controls cortical
actin assembly by controlling the level of C-cadherin on the surface of each
blastomere. Animal caps from p120 catenin-depleted blastulae were stained with
6B6 monoclonal antibody against Xenopus C-cadherin.
Fig. 5E shows that depletion of
p120 catenin caused a significant reduction in C-cadherin on the cell surface
at the late blastula stage and, conversely, that injection of p120
catenin mRNA into the early embryo caused increased levels of C-cadherin
at the cell surface (Fig. 5F).
These data show that p120 catenin plays an essential role in controlling the
steady levels of C-cadherin in the Xenopus embryo and, through this,
controls the level of cortical actin assembly.
To confirm that interaction between C-cadherin and p120 catenin is required
for their function in controlling the dense actin assembly, we generated a
p120 catenin mutant that lacks the cadherin-binding domain (spanning the first
Arm repeat, p120 Arm1). The first Arm repeat has been shown to be
necessary to target p120 catenin to cadherin on the cell membrane, but not to
affect p120 catenin regulation of Rho GTPases
(Anastasiadis et al., 2000;
Yanagisawa et al., 2004).
Overexpression of this construct did not increase either the density of
cortical actin or the level of C-cadherin on the cell surface in
Xenopus embryos (Fig.
5G,H), showing that the dense cortical actin assembly requires
interaction between p120 catenin and C-cadherin.
LPA and Xflop signaling control actin assembly by controlling the amount of C-cadherin expression
We have shown previously that the dense actin network assembly in the
Xenopus blastula is controlled by two maternally expressed
G-protein-coupled receptors, LPA1 and Xflop
(Lloyd et al., 2005;
Tao et al., 2005). We have
shown above that the C-cadherin-p120 catenin complex is a site of control of
the dense cortical actin network. We therefore explored the possibility that
one mechanism of Xflop and LPA function might be through regulation of
C-cadherin levels. First, mRNAs encoding LPA1 (400 pg per embryo) or Xflop
(250 pg per embryo) were injected into the animal cytoplasm at the 2-cell
stage, and the embryos allowed to develop to the late blastula stage before
fixation and staining for C-cadherin. Fig.
6A-C show that both mRNAs caused upregulation of C-cadherin
levels. To distinguish between increased total protein and altered cellular
distribution, we assayed the total C-cadherin levels by western blotting.
Fig. 6D shows that LPA or/and
Xflop overexpression increased the total level of C-cadherin at the late
blastula stage.
|
),
signal transduction must be occurring. However, this has not been shown for
Xflop, as its ligand(s) is unknown. We therefore created the point mutation
R112A, which has been shown to block the activation of G proteins by the motif
of aa 111-113 (DRF) in the second intracellular loop of GPCRs
(Rovati et al., 2007), mRNA
encoding Xflop R112A was injected into each cell of the 2-cell embryo in a
wide range of concentrations, up to 1 ng. In no case did it mimic the
wild-type Xflop in increasing the level of cortical actin
(Fig. 7). These data confirm
that signaling through the LPA and Xflop receptors controls the level of
C-cadherin at the late blastula stage.
The effects of LPA receptor and Xflop depletion on the dense cortical actin network can be rescued by the overexpression of C-cadherin
If the LPA and Xflop signaling pathways control cortical actin assembly by
regulating the amount of C-cadherin on the cell surface, then overexpression
of C-cadherin should rescue the depletion of these receptors. We therefore
injected 500 pg of C-cadherin mRNA into oocytes depleted of either
LPA1 or Xflop by injection of antisense oligos into cultured oocytes. In both
cases, the dense cortical actin network was restored
(Fig. 8B-E). Depletion of
either Xflop or LPA reduced both the membrane
(Fig. 8A,B,C) and total
(Fig. 8D,E insets) levels of
C-cadherin, further supporting the notion that Xflop and LPA receptors control
the level of C-cadherin at the late blastula stage in Xenopus.
|
). This suggest that LPA signaling, at least at higher than
physiological levels, can control the assembly of actin by other mechanisms in
addition to its role in cadherin expression.
|
|
DISCUSSION |
|---|
|
TOPSUMMARYINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
|---|
). It is
therefore of great interest to understand the mechanism of cortical actin
assembly in the early embryo. Previous work also showed that dissociated
cells, in which cell contact and signaling are prevented, lose their dense
cortical actin, and that the G-protein-coupled receptors LPA1 and LPA2, and
Xflop, are required for cortical actin assembly in vivo
(Lloyd et al., 2005;
Tao et al., 2005). However,
the mechanism of action of these receptors, and whether other aspects of cell
contact are required, have not been explored. Here, we show that there is a
direct relationship between the expression of C-cadherin on the cell surface
and the assembly of cortical actin in the cytoplasm, and that the level of
cell-surface cadherin is controlled by signaling through GPCRs. This
interacting system results in the level of cell cohesion and rigidity seen at
the late blastula stage.
It is already known that cadherin-containing punctae form at sites of
initial contact of epithelial cells in culture
(Helwani et al., 2004;
Kobielak et al., 2004;
Kovacs et al., 2002b;
Scott et al., 2006;
Vasioukhin et al., 2000;
Verma et al., 2004), and that
these associate with actin filaments and actin assembly components such as
Arp2/3 (Kovacs et al., 2002b;
Verma et al., 2004), formin 1
(Kobielak et al., 2004) and
Ena/Vasp (Scott et al., 2006).
Here we show, in a developing system in vivo, that such punctae and their
associated actin filament assemblies form the skeleton of the early embryo.
The presentation of C-cadherin on the cell surface is clearly a rate-limiting
step of this process, as its overexpression increases the dense cortical actin
network, whereas its depletion causes loss of the dense cortical actin.
|
|
;
Elia et al., 2006;
Perez-Moreno et al., 2006) and
this proved to be the case here. p120 overexpression caused increased amounts
of both C-cadherin and cortical actin, whereas depletion caused the reverse.
In addition, a p120 mutant lacking the cadherin-binding site failed to
increase either C-cadherin expression on the surface, or cortical actin
assembly, showing that the level of C-cadherin expression on the cell surface
requires its interaction with p120. It is possible that p120 controls cortical
actin assembly at other levels, in addition to cadherin presentation. Our data
suggest that p120-mediated cortical actin assembly requires its interaction
with C-cadherin. The fact that a cadherin construct lacking the p120-binding
site was expressed at high levels on the cell surface, but caused loss of
cortical actin filaments, supports this hypothesis. p120 catenin has been
shown to bind signaling intermediates in actin assembly such as Rac1 and Cdc42
(Noren et al., 2000), so it
will be interesting to see whether these interactions are also required for
actin assembly in Xenopus. A cadherin construct lacking the
ß-catenin-binding domain and that does not bind ß-catenin
(Pokutta and Weis, 2002)
failed to significantly increase cortical actin assembly. However, in the
absence of endogenous C-cadherin, this construct was not transported to the
cell membrane and did not rescue cell adhesion. The role of the CBD in
cortical actin assembly could therefore not be tested with this mutant.
However, our observations do suggest a potential role for the CBD in protein
trafficking and membrane presentation of cadherin. The high level of membrane
expression of the cadherin mutant lacking the p120-binding site might also be
due to the intact CBD in this mutant.
Once it became clear that the late blastula cells somehow titrate the
assembly of cortical actin against the level of C-cadherin on the surface,
this immediately raised the novel possibility that intercellular signaling
through GPCRs might control the density and pattern of cortical actin in the
blastula at the level of cadherin expression. This indeed proved to be the
case. Depletion or augmentation of the receptors LPA1 and LPA 2, and Xflop,
previously shown to be required for cortical actin assembly
(Lloyd et al., 2005;
Tao et al., 2005), caused
corresponding loss and increase of C-cadherin expression on the cell surface.
Furthermore, overexpression of C-cadherin by mRNA injection was able to rescue
cortical actin assembly after depletion of either LPA1 or Xflop in the
blastula. These data implicate the two signaling pathways in controlling the
level of expression of cadherin on the cell surface. It will be of major
interest to discover whether these signaling pathways are the sites of action
of cell cycle components, as the dense cortical actin network is lost during
cell division.
One interesting and unexplained fact in these studies is that there appears
to be two actin-containing networks in the blastula cells. Cells that are
dividing in vivo, or dissociated in calcium- and magnesium-free saline, or
depleted of C-cadherin, all lose their dense cortical actin network, but
retain a sparse network, even in the absence of cadherin on the cell surface
(Fig. 2) (see also
Lloyd et al., 2005). This
sparse network is clearly cadherin-independent and is independent of the dense
network. We do not know if the reverse is the case, as we have not found any
independent way of removing the sparse network. It is possible the LPA and
Xflop signaling also control the sparse network, although experimental details
are still lacking. We also do not know how the sparse network is associated
with the cell cortex. There are several known sites of actin insertion into
the cell membrane, in addition to cadherins. For example, 
-spectrin and
moesin, members of FERM (band 4.1/ezrin/radixin/moesin) proteins are
maternally expressed in Xenopus oocytes
(Carotenuto et al., 2000;
Thorn et al., 1999), and could
play a role in attaching the sparse actin network to the cell membrane. Some
integrins (ß1, for example) are present at the blastula stage
(Gawantka et al., 1992) and
could be sites of actin assembly.
Many more questions remain. We do not know the mechanism by which LPA and Xflop signaling control the level of C-cadherin on the cell surface. It is also possible that LPA and Xflop control actin assembly at the cadherin complex at levels other than via C-cadherin expression. They might also control the sparse network through mechanisms that do not involve cadherin expression. The Xenopus blastula represents a simple in vivo system with which to address these questions.
|
ACKNOWLEDGMENTS |
|---|
|
Footnotes |
|---|
|
REFERENCES |
|---|
|
TOPSUMMARYINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
|---|
Adams, C. L. and Nelson, W. J. (1998). Cytomechanics of cadherin-mediated cell-cell adhesion. Curr. Opin. Cell Biol. 10,572 -577.[CrossRef][Medline]
Adams, C. L., Nelson, W. J. and Smith, S. J.
(1996). Quantitative analysis of cadherin-catenin-actin
reorganization during development of cell-cell adhesion. J. Cell
Biol. 135,1899
-1911.
Adams, C. L., Chen, Y. T., Smith, S. J. and Nelson, W. J.
(1998). Mechanisms of epithelial cell-cell adhesion and cell
compaction revealed by high-resolution tracking of E-cadherin-green
fluorescent protein. J. Cell Biol.
142,1105
-1119.
Anastasiadis, P. Z., Moon, S. Y., Thoreson, M. A., Mariner, D. J., Crawford, H. C., Zheng, Y. and Reynolds, A. B. (2000). Inhibition of RhoA by p120 catenin. Nat. Cell Biol. 2, 637-644.[CrossRef][Medline]
Betson, M., Lozano, E., Zhang, J. and Braga, V. M.
(2002). Rac activation upon cell-cell contact formation is
dependent on signaling from the epidermal growth factor receptor.
J. Biol. Chem. 277,36962
-36969.
Carotenuto, R., Vaccaro, M. C., Capriglione, T., Petrucci, T. C. and Campanella, C. (2000). alpha-Spectrin has a stage-specific asymmetrical localization during Xenopus oogenesis. Mol. Reprod. Dev. 55,229 -239.[CrossRef][Medline]
Chen, X., Kojima, S., Borisy, G. G. and Green, K. J.
(2003). p120 catenin associates with kinesin and facilitates the
transport of cadherin-catenin complexes to intercellular junctions.
J. Cell Biol. 163,547
-557.
Choi, Y. S., Sehgal, R., McCrea, P. and Gumbiner, B.
(1990). A cadherin-like protein in eggs and cleaving embryos of
Xenopus laevis is expressed in oocytes in response to progesterone.
J. Cell Biol. 110,1575
-1582.
Davis, M. A. and Reynolds, A. B. (2006). Blocked acinar development, E-cadherin reduction, and intraepithelial neoplasia upon ablation of p120-catenin in the mouse salivary gland. Dev. Cell 10,21 -31.[CrossRef][Medline]
Davis, M. A., Ireton, R. C. and Reynolds, A. B.
(2003). A core function for p120-catenin in cadherin turnover.
J. Cell Biol. 163,525
-534.
Dent, J. A., Polson, A. G. and Klymkowsky, M. W. (1989). A whole-mount immunocytochemical analysis of the expression of the intermediate filament protein vimentin in Xenopus. Development 105,61 -74.[Abstract]
Drees, F., Pokutta, S., Yamada, S., Nelson, W. J. and Weis, W. I. (2005). Alpha-catenin is a molecular switch that binds E-cadherin-beta-catenin and regulates actin-filament assembly. Cell 123,903 -915.[CrossRef][Medline]
Ehrlich, J. S., Hansen, M. D. and Nelson, W. J. (2002). Spatio-temporal regulation of Rac1 localization and lamellipodia dynamics during epithelial cell-cell adhesion. Dev. Cell 3,259 -270.[CrossRef][Medline]
Elia, L. P., Yamamoto, M., Zang, K. and Reichardt, L. F. (2006). p120 catenin regulates dendritic spine and synapse development through Rho-family GTPases and cadherins. Neuron 51,43 -56.[CrossRef][Medline]
Fang, X., Ji, H., Kim, S. W., Park, J. I., Vaught, T. G.,
Anastasiadis, P. Z., Ciesiolka, M. and McCrea, P. D. (2004).
Vertebrate development requires ARVCF and p120 catenins and their interplay
with RhoA and Rac. J. Cell Biol.
165, 87-98.
Gawantka, V., Ellinger-Ziegelbauer, H. and Hausen, P. (1992). Beta 1-integrin is a maternal protein that is inserted into all newly formed plasma membranes during early Xenopus embryogenesis. Development 115,595 -605.[Abstract]
Ginsberg, D., DeSimone, D. and Geiger, B. (1991). Expression of a novel cadherin (EP-cadherin) in unfertilized eggs and early Xenopus embryos. Development 111,315 -325.[Abstract]
Hausen, P. and Riebesell, M. (2002). A simple flow-through micro-chamber for handling fragile, small tissue explants and single non-adherent cells. Methods Cell Sci. 24,165 -168.[CrossRef][Medline]
Heasman, J., Holwill, S. and Wylie, C. C. (1991). Fertilization of cultured Xenopus oocytes and use in studies of maternally inherited molecules. Methods Cell Biol. 36,213 -230.[Medline]
Heasman, J., Ginsberg, D., Geiger, B., Goldstone, K., Pratt, T., Yoshida-Noro, C. and Wylie, C. (1994). A functional test for maternally inherited cadherin in Xenopus shows its importance in cell adhesion at the blastula stage. Development 120, 49-57.[Abstract]
Helwani, F. M., Kovacs, E. M., Paterson, A. D., Verma, S., Ali,
R. G., Fanning, A. S., Weed, S. A. and Yap, A. S. (2004).
Cortactin is necessary for E-cadherin-mediated contact formation and actin
reorganization. J. Cell Biol.
164,899
-910.
Ireton, R. C., Davis, M. A., van Hengel, J., Mariner, D. J.,
Barnes, K., Thoreson, M. A., Anastasiadis, P. Z., Matrisian, L., Bundy, L. M.,
Sealy, L. et al. (2002). A novel role for p120 catenin in
E-cadherin function. J. Cell Biol.
159,465
-476.
Jamora, C. and Fuchs, E. (2002). Intercellular adhesion, signalling and the cytoskeleton. Nat. Cell Biol. 4,E101 -E108.[CrossRef][Medline]
Kim, S. H., Li, Z. and Sacks, D. B. (2000).
E-cadherin-mediated cell-cell attachment activates Cdc42. J. Biol.
Chem. 275,36999
-37005.
Kobielak, A., Pasolli, H. A. and Fuchs, E. (2004). Mammalian formin-1 participates in adherens junctions and polymerization of linear actin cables. Nat. Cell Biol. 6, 21-30.[CrossRef][Medline]
Kofron, M., Heasman, J., Lang, S. A. and Wylie, C. C.
(2002). Plakoglobin is required for maintenance of the cortical
actin skeleton in early Xenopus embryos and for cdc42-mediated wound healing.
J. Cell Biol. 158,695
-708.
Kovacs, E. M., Ali, R. G., McCormack, A. J. and Yap, A. S.
(2002a). E-cadherin homophilic ligation directly signals through
Rac and phosphatidylinositol 3-kinase to regulate adhesive contacts.
J. Biol. Chem. 277,6708
-6718.
Kovacs, E. M., Goodwin, M., Ali, R. G., Paterson, A. D. and Yap, A. S. (2002b). Cadherin-directed actin assembly: E-cadherin physically associates with the Arp2/3 complex to direct actin assembly in nascent adhesive contacts. Curr. Biol. 12,379 -382.[CrossRef][Medline]
Levi, G., Ginsberg, D., Girault, J. M., Sabanay, I., Thiery, J. P. and Geiger, B. (1991). EP-cadherin in muscles and epithelia of Xenopus laevis embryos. Development 113,1335 -1344.[Abstract]
Lloyd, B., Tao, Q., Lang, S. and Wylie, C.
(2005). Lysophosphatidic acid signaling controls cortical actin
assembly and cytoarchitecture in Xenopus embryos.
Development 132,805
-816.
Nakagawa, M., Fukata, M., Yamaga, M., Itoh, N. and Kaibuchi, K. (2001). Recruitment and activation of Rac1 by the formation of E-cadherin-mediated cell-cell adhesion sites. J. Cell Sci. 114,1829 -1838.[Abstract]
Nomura, K., Tajima, T., Nomura, H., Shiraishi, H., Uchida, M. and Yamana, K. (1988). Cell to cell adhesion systems in Xenopus laevis, the South African clawed toad. II: Monoclonal antibody against a novel Ca2+-dependent cell-cell adhesion glycoprotein on amphibian cells. Cell Differ. 23,207 -212.[CrossRef][Medline]
Noren, N. K., Liu, B. P., Burridge, K. and Kreft, B.
(2000). p120 catenin regulates the actin cytoskeleton via Rho
family GTPases. J. Cell Biol.
150,567
-580.
Noren, N. K., Niessen, C. M., Gumbiner, B. M. and Burridge,
K. (2001). Cadherin engagement regulates Rho family GTPases.
J. Biol. Chem. 276,33305
-33308.
Paulson, A. F., Mooney, E., Fang, X., Ji, H. and McCrea, P.
D. (2000). Xarvcf, Xenopus member of the p120 catenin
subfamily associating with cadherin juxtamembrane region. J. Biol.
Chem. 275,30124
-30131.
Perez-Moreno, M., Jamora, C. and Fuchs, E. (2003). Sticky business: orchestrating cellular signals at adherens junctions. Cell 112,535 -548.[CrossRef][Medline]
Perez-Moreno, M., Davis, M. A., Wong, E., Pasolli, H. A., Reynolds, A. B. and Fuchs, E. (2006). p120-catenin mediates inflammatory responses in the skin. Cell 124,631 -644.[CrossRef][Medline]
Pokutta, S. and Weis, W. I. (2002). The cytoplasmic face of cell contact sites. Curr. Opin. Struct. Biol. 12,255 -262.[CrossRef][Medline]
Rovati, G. E., Capra, V. and Neubig, R. R.
(2007). The highly conserved DRY motif of class A GPCRs: beyond
the ground state. Mol. Pharmacol.
71,959
-964.
Scott, J. A., Shewan, A. M., den Elzen, N. R., Loureiro, J. J.,
Gertler, F. B. and Yap, A. S. (2006). Ena/VASP proteins can
regulate distinct modes of actin organization at cadherin-adhesive contacts.
Mol. Biol. Cell 17,1085
-1095.
Tao, Q., Lloyd, B., Lang, S., Houston, D., Zorn, A. and Wylie,
C. (2005). A novel G protein-coupled receptor, related to
GPR4, is required for assembly of the cortical actin skeleton in early Xenopus
embryos. Development
132,2825
-2836.
Thoreson, M. A., Anastasiadis, P. Z., Daniel, J. M., Ireton, R.
C., Wheelock, M. J., Johnson, K. R., Hummingbird, D. K. and Reynolds, A.
B. (2000). Selective uncoupling of p120(ctn) from E-cadherin
disrupts strong adhesion. J. Cell Biol.
148,189
-202.
Thorn, J. M., Armstrong, N. A., Cantrell, L. A. and Kay, B. K. (1999). Identification and characterisation of Xenopus moesin, a Src substrate in Xenopus laevis oocytes. Zygote 7,113 -122.[CrossRef][Medline]
Turner, A. P., Brown, D., Heasman, J., Cook, G. M., Evans, J., Vickers, L. and Wylie, C. C. (1992). Involvement of a neutral glycolipid in differential cell adhesion in the Xenopus blastula. EMBO J. 11,3845 -3855.[Medline]
Vaezi, A., Bauer, C., Vasioukhin, V. and Fuchs, E. (2002). Actin cable dynamics and Rho/Rock orchestrate a polarized cytoskeletal architecture in the early steps of assembling a stratified epithelium. Dev. Cell 3,367 -381.[CrossRef][Medline]
Vasioukhin, V., Bauer, C., Yin, M. and Fuchs, E. (2000). Directed actin polymerization is the driving force for epithelial cell-cell adhesion. Cell 100,209 -219.[CrossRef][Medline]
Verma, S., Shewan, A. M., Scott, J. A., Helwani, F. M., den
Elzen, N. R., Miki, H., Takenawa, T. and Yap, A. S. (2004).
Arp2/3 activity is necessary for efficient formation of E-cadherin adhesive
contacts. J. Biol. Chem.
279,34062
-34070.
Xiao, K., Allison, D. F., Buckley, K. M., Kottke, M. D.,
Vincent, P. A., Faundez, V. and Kowalczyk, A. P. (2003).
Cellular levels of p120 catenin function as a set point for cadherin
expression levels in microvascular endothelial cells. J. Cell
Biol. 163,535
-545.
Yamada, S., Pokutta, S., Drees, F., Weis, W. I. and Nelson, W. J. (2005). Deconstructing the cadherin-catenin-actin complex. Cell 123,889 -901.[CrossRef][Medline]
Yanagisawa, M., Kaverina, I. N., Wang, A., Fujita, Y., Reynolds,
A. B. and Anastasiadis, P. Z. (2004). A novel interaction
between kinesin and p120 modulates p120 localization and function.
J. Biol. Chem. 279,9512
-9521.
Yonemura, S., Itoh, M., Nagafuchi, A. and Tsukita, S. (1995). Cell-to-cell adherens junction formation and actin filament organization: similarities and differences between non-polarized fibroblasts and polarized epithelial cells. J. Cell Sci. 108,127 -142.[Abstract]
This article has been cited by other articles:
|
|
 |
|
  P. Skoglund, A. Rolo, X. Chen, B. M. Gumbiner, and R. Keller Convergence and extension at gastrulation require a myosin IIB-dependent cortical actin network Development, July 15, 2008; 135(14): 2435 - 2444. [Abstract] [Full Text] [PDF]
|
|
| |||||||
