1 Department of Molecular Biology, Howard Hughes Medical Institute, Princeton
University, NJ 08544, USA
2 Harvard Medical School, 25 Shattuck Street, Boston, MA 02115, USA
3 Department of Biological Sciences, Stanford University, CA 94305
4 Wellcome Trust/Cancer Research UK Gurdon Institute and Department of Anatomy,
University of Cambridge, Cambridge CB2 1QN, UK
* Author for correspondence (e-mail: rhoang{at}haverford.edu)
Accepted 13 June 2006
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SUMMARY |
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Key words: Drosophila, Myosin, Gastrulation, Fog, Morphogenesis, Rho-kinase, RhoGEF, Arm
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Introduction |
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A rare opportunity to bridge these categorizations is offered by folded
gastrulation (fog) a gene involved in
Drosophila gastrulation. Mutations in fog disrupt the
movement of mesodermal and endodermal precursor cells into the interior of the
embryo (Costa et al., 1994;
Oda and Tsukita, 2001
).
fog is not itself a patterning gene, as cell fates are unaltered in
fog mutants, but fog is a direct target of a well described
patterning gene, twist (Costa et
al., 1994
). In addition, fog encodes a novel protein that
is thought to be the ligand for a signaling cascade that controls changes in
cell shape. fog may therefore function in morphogenesis by
interfacing between the patterning gene twist and the machinery that
produces cell shape change.
fog protein is required for the earliest visible changes in cell
shape that mark the onset of gastrulation. First, the prospective mesodermal
cells on the ventral side of the embryo flatten and then constrict their
apical surface. This shifts the cells towards the interior of the embryo in a
structure called the ventral furrow. In fog mutant embryos, these
cell shape changes are disorganized and proceed in an uncoordinated manner
(Costa et al., 1994). Second,
similar cell shape changes initiate internalization of the prospective
endoderm on the dorsal posterior surface of the embryo (the posterior midgut
primordium). In fog mutants these cell shape changes are blocked in
all but a few cells. Furthermore, a heat-shock-activated form of fog
has been shown to elicit the apical flattening of cells in other areas of the
embryo (Morize et al., 1998
).
fog therefore plays an important role in controlling the flattening
and constriction of the apical surface of cells and is the primary pathway
controlling these cell shape changes in the posterior midgut primordium,
whereas a second parallel pathway additionally contributes to these cell shape
changes in the ventral furrow. In all these studies fog function has
been analyzed with respect to the outward appearance of cells. Nothing is
known about the molecular remodeling of the cytoarchitecture that must
underlie fog function.
Fragments of a pathway have started to emerge for the function of
fog during gastrulation. Embryos lacking the gene product
concertina (cta) show the same disruptions to ventral furrow
and posterior midgut formation as seen in fog mutants
(Parks and Wieschaus, 1991).
Furthermore, cta has been positioned genetically downstream of
fog as the effects of a heat-shock-activated form of fog are
blocked in cta mutants and conversely, activated cta has
effects that are independent of fog
(Morize et al., 1998
).
cta is known to encode a G-protein alpha subunit of the
G
12/13 class. In addition to its own role in the
fog pathway, cta therefore also implicates a role for an as
yet unidentified G-protein-coupled receptor. Another gene that disrupts both
ventral furrow and posterior midgut formation is RhoGEF2, a guanine
nucleotide exchange factor that promotes Rho activation and thus also
implicates Rho signaling in this process
(Barrett et al., 1997
;
Hacker and Perrimon, 1998
).
RhoGEF2 mutants have been shown to be able to interact genetically
with a fog transgene during early embryonic development
(Barrett et al., 1997
).
However, the ventral furrow phenotype of RhoGEF2 mutants affects all
cells and is therefore much more severe than that of fog and
cta mutants. The way in which the fog and RhoGEF2
pathways interact therefore remains unclear and the relevant downstream
signaling components are unknown.
A likely target of the fog pathway is non-muscle myosin II (herein
referred to as myosin). Myosin is expressed at the right time and place to be
involved (Young et al., 1991)
and removal of RhoGEF2 lowers myosin levels at gastrulation
(Nikolaidou and Barrett,
2004
), though the extent and significance of this disruption is
not clear. Furthermore, myosin is known to play an important role in driving
many cell shape changes in a wide variety of organisms. One of the best
understood of all myosin-based processes is cytokinesis. During cell division,
myosin localizes to the cleavage furrow, an assembly of proteins responsible
for physically separating newly formed daughter cells. Myosin is thought to
function in this process by contributing force in a contractile actin-myosin
ring, though much remains to be understood about how this force is coupled to
the physical changes in cell shape
(Glotzer, 2001
;
Wang, 2001
). The ability of
myosin to function as an actin-based motor that provides contractile force is
also thought to underlie the role of myosin in many other morphogenetic
processes. However, the full range of myosin function is likely to be more
complicated, with reports of myosin also involved in downregulating adherens
junctions and serving as a spatial cue for cell wall formation during fission
yeast cell division (Rajagopalan et al.,
2003
; Sahai and Marshall,
2002
). Myosin may therefore contribute to Drosophila gastrulation
in a number of different ways.
Consistent with its involvement in a wide range of morphogenetic processes,
myosin is also widely expressed. Therefore, in the context of a developing
organism, myosin functions in multiple different processes sometimes within
the very same cell. As Drosophila gastrulation initiates myosin
localization is highly dynamic, being lost from the basal side of mesodermal
cells, where it functioned during cellularization, and accumulating apically
(Royou et al., 2004;
Young et al., 1991
) (this
study). Therefore, little is known about the specific role of myosin in
gastrulation and more generally about how different myosin-based processes,
such as cellularization and ventral furrow formation, are related to one
another during development. To begin to address these issues, it becomes
important to understand the precise dynamics, localization and regional
activities of myosin within individual cells over time.
In this paper, we address many of these issues through our analysis of fog function. We find that fog signal is apically polarized and that this in turn directs the apical localization of myosin. We show that the mechanism driving this apical localization of myosin requires interaction/contractility with actin, distinguishing it from the mechanism driving myosin to the basal side of the same cells. Furthermore, we demonstrate that fog is both necessary and sufficient for localization of myosin apically, and that this pathway of myosin activation requires RhoGEF2 and the downstream effector Rho kinase. Finally, we show that once localized apically, myosin continues to contract and the resulting force is translated into physical changes at the cell surface through a tethering of the action-myosin cytoskeleton by apical adherens junctions. We therefore provide a mechanism of fog function that takes us all the way from the patterning gene twist, to physical changes in cell shape at the onset of gastrulation.
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Materials and methods |
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Germline clones were produced from the following crosses using standard
techniques (Chou and Perrimon,
1992).
Arm: arm043A01 FRT101/FM6 x ovoD1
FRT101/Y; hs-flp 138 produced heat shocked
arm043A01 FRT101/ovoD1
FRT101 females used to collect embryos
(Tolwinski and Wieschaus,
2001).
DRhoGEF2: y w hs-flp;FRT42BG13
DRhoGEF21.1/CyO virgin females x
FRT42BG13 ovoD1/CyO males produced heat shocked
FRT42BG13 DRhoGEF21.1/FRT42BG13
ovoD1 females that were crossed to w males to collect embryos
(Barrett et al., 1997).
Drok alleles (Winter et al.,
2001) were used to make the following stocks and crosses: w
rok1or2 FRT18D/FM7 x ovoD2
FRT18D/Y; hs-flp138 produced heat shocked
wrok1or2 FRT18D/ovoD2
FRT18D females from which embryos were collected.
UASfog expressing embryos are from mat67;mat15 virgins x UASfog
males. Three UASfog lines were used: UASfog6 (III),
UASfog12 (II) and UASfog18/TM3Ser.
UASnullo-expressing embryos are from mat67;mat15 virgins x
UASnulloN39 males
(Hunter et al., 2002). Embryos
expressing UASmYFP-myosin IIDN are from mat67;mat15 virgins x
w; UAS mYFP-myosin IIDN males to produce w; mat67/UAS
mYFP-myosin IIDN;mat15/+ virgins backcrossed to w; UAS
mYFP-myosin IIDN to collect embryos.
Embryology, histology and image analysis
All embryos were heat-methanol fixed
(Muller and Wieschaus, 1996)
except those stained with anti-GFP or anti-Fog antibody, which were
formaldehyde fixed (Zallen and Wieschaus,
2004
). SEM analysis was as described previously
(Morize et al., 1998
). Stained
embryos were cross-sectioned by hand in 70% glycerol/PBS (using a 26-gauge
hypodermic needle), mounted in Aquapolymount (Polysciences, Warrington, PA)
and imaged using a Zeiss LSM-510 confocal microscope (Thornwood, NY).
Reagents used were Hoechst 33342 (Molecular Probes), rabbit anti-Myosin II antibody (1:1250, gift of C. Field), sheep anti-Dorsal antibody (1:500, gift of R. Stewart), mouse anti-Armadillo (1:50, N2-7A1 Developmental Studies Hybridoma Bank, DSHB), mouse anti-Neurotactin (1:10, BP106 from DSHB), guinea pig anti-Runt (1:500, gift of C. Alonso and J. Reinitz) and rabbit anti-GFP (1:2000, Torey Pines). Primary antibodies were detected with Alexa-conjugated secondary antibodies (Molecular Probes). Anti-Fog antibody was produced from a fusion of 6xHis, followed by three novel residues, then residues 28 to 303 of fog cDNA, then six novel residues. This protein was purified and injected into rabbits. Sera were purified using protein-A-conjugated beads.
Myosin intensity was measured in embryos stained for myosin II and cut by hand into cross-sections. Intensity measurements were taken from the monochrome myosin channel of confocal images, using IPLab software to average pixel intensities along a hand-drawn line on the cellularization front (basal) or apical edge of the embryo (apical) over 10 cell widths. Measurements were taken on both ventral and lateral sides of the embryo and normalized relative to the background pixel intensity (inside nuclei). Each sample contained measurements from at least 100 cells including at least five different embryos.
Myosin dynamics in fog mutants: fog114 females were crossed to sqhGFP males. F1 females carrying both fog114 and sqhGFP were backcrossed to the sqhGFP stock producing both fog mutant and control embryos expressing sqhGFP. Time-lapse movies were taken using confocal microscopy with images collected every 30 seconds. fog mutants were identified by emergence of the fog phenotype.
Heat-shock inactivation of shibirets: embryos were collected for 1 hour, allowed to age, dechorionated and transferred to a damp piece of paper towel that was placed on a 35°C heat block for 30 minutes, then fixed for 25 minutes in a 50:50 mixture of 4% formaldehyde in 0.1 M PIPES, 2 mM EGTA, 1 mM MgSO4 (pH 6.9) and heptane.
Constructs
pUAST-mYFP-myosin-IIDN
The central region of myosin II-coding region was PCR amplified from
pBS-Zipper (Kiehart et al.,
1989) (gift from D. Kiehart), to introduce a KpnI site
and a glycine linker, and to maintain the XhoI site. In a separate
reaction, the myosin N terminus was removed from pBS-Zipper by XhoI
digestion and self-ligation to give pBS-myosin-II-C-terminus. The PCR
KpnI-XhoI central myosin fragment was inserted into the
KpnI/XhoI-digested pBS-myosin-II-C-terminus backbone, to
create myosin-IIDN. This construct contains a dominant-negative
form of myosin II, with the head removed at precisely the same point as an
equivalent Dictyostelium myosin IIDN [head-neck junction:
amino acid 831 in Drosophila (QWWR) and position 809 in
Dictyostelium (PWWK)] (Burns et
al., 1995
; Zang and Spudich,
1998
). mYFP (Haseloff,
1999
) (gift from J. Haseloff) was PCR amplified to introduce
KpnI and NotI sites. The KpnI cut mYFP PCR product
was inserted into KpnI cut pBS-myosin-IIDN and correct
orientation of the insert selected by position of the NotI site. A
NotI-mYFP-myosin-IIDN-NotI cassette was excised
from pBS and inserted into NotI-digested pUAST
(Brand and Perrimon, 1993
). The
correct orientation of the insert was selected by position of the
XhoI site. This produced the construct
pUAST-mYFP-myosin-IIDN.
pUAST-fog
Full-length fog cDNA was obtained from plasmid pB26H
(Costa et al., 1994) by
digestion with NheI and purification of the fog-containing fragment.
pUAST (Brand and Perrimon,
1993
) was digested with XbaI and ligated with the
compatible NheI ends of the fog fragment. Correct orientation of the
insert was screened by PCR.
Transgenic flies were generated using standard techniques
(Barros et al., 2003).
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Results |
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To understand the molecular basis of the control of the cytoskeleton by
fog, we have investigated changes in myosin II dynamics in
fog mutant embryos. Analysis of myosin dynamics is easiest in the
posterior midgut where fog is the primary pathway controlling cell
constriction and the geometry of the egg enables visualization of a myosin
lightchain-GFP fusion (sqhGFP) in time-lapse movies of living
embryos. During gastrulation myosin localizes to the apical side of cells
throughout the posterior midgut primordium of control embryos
(Fig. 1G). However, in
fog mutant embryos of the same age, the apical localization of myosin
is severely disrupted and is restricted to just a few cells underlying the
pole cells (Fig. 1I). Analysis
of myosin localization in fixed embryos also reveals a disruption to apical
localization, both in the posterior midgut and the ventral furrow of
fog mutants. This is consistent with previous data showing that
myosin is also disrupted in cta mutants
(Nikolaidou and Barrett,
2004). In the ventral furrow only a subset of cells (39%,
n=62) localize myosin apically in fog114 mutant
embryos (Fig. 1H,J). The
fog114 allele is a an RNA null (see Materials and
methods). The patchiness of this defect in the ventral furrow of
fog114 mutants therefore probably reflects the redundancy
with additional pathways that control cell shape change in these cells
(Costa et al., 1994
) and/or a
small maternal contribution of fog.
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The first effects of fog expression are seen at the onset of gastrulation. In embryos uniformly expressing fog the apical localization of myosin now occurs in all cells instead of being restricted to a ventral domain (Fig. 2H,K,N). In the ventral cells of these fog-overexpressing embryos, the apical localization of myosin precedes the apical localization in the more lateral and dorsal cells and reaches a higher level. It is also a higher level than in the ventral cells of control embryos. It is unclear whether this reflects higher levels of fog expression in the ventral cells (owing to both endogenous and UASfog expression) or whether it reflects an earlier or increased competence of these ventral cells to react to fog signal. The apical localization of myosin in the lateral and dorsal cells of fog-overexpressing embryos continues throughout gastrulation (Fig. 2I,L,O) and occurs without any concomitant reduction in levels of basally localized myosin (Fig. 2N,O). This raises the possibility that the apical and basal localizations of myosin may be independently controlled.
Not all fog-overexpressing embryos show the same degree of ectopic apical myosin localization in lateral and dorsal cells. Furthermore, limited apical myosin staining is occasionally seen in control embryos. We quantified this variability over five separate experiments. During cellularization, onset of gastrulation and later gastrulation 0% (n=54), 73% (n=66) and 84% (n=25) of fog-overexpressing embryos show ectopic apical myosin compared with 0% (n=33), 8% (n=26) and 22% (n=23) of controls respectively.
In wild-type embryos myosin accumulates apically in all cells after the
completion of ventral furrow invagination, at the onset of germ band extension
(Bertet et al., 2004;
Zallen and Wieschaus, 2004
).
Therefore, apical accumulation of myosin in dorsal and lateral cells of
apparently gastrulating embryos may occur as the result of a delay in ventral
furrow formation. To investigate this possibility, we followed time-lapse
movies of gastrulating embryos and examined morphology in precisely timed
embryo collections. In both cases, we found a slight delay in the completion
of ventral furrow formation in fog-overexpressing embryos compared
with controls. In equivalently aged collections, only 32% (n=82) of
control embryos were undergoing ventral furrow formation compared with 45%
(n=145) of fog-overexpressing embryos. This implies that
fog-overexpressing embryos take about 1.4 times longer to complete
ventral furrow formation than control embryos. However, this is considerably
less than the
3.5 times delay that would be required to account for the
large difference seen in apical myosin localization between the
UASfog-expressing embryos and controls. Assuming that if the process
were to take twice as long in UASfog embryos this would account for
50% of the embryos showing apical myosin simply because they are in fact
older, we estimate that the process would have to be
3.5 times as long to
account for the actual increased numbers of embryos we see (73% versus
8%).
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Functionally distinct modes of myosin localization
It is possible that fog provides a signal to localize or transport
myosin apically, and myosin is then activated to interact and contract with
actin. An intriguing alternative, however, is that fog itself may be
activating myosin contractility, initiating an active motor-driven mechanism
of myosin localization. To help distinguish between these two possibilities,
we constructed a form of myosin that is no longer able to interact or contract
with actin and asked if this form of myosin was still able to localize
normally.
Myosin is a hexamer comprising two myosin heavy chains (MHCs), two
essential light chains and two regulatory light chains (RLCs). It is the
globular head domain of the MHC subunits that interacts directly with actin
and contains the region of ATPase activity that drives this actin-based motor.
In addition the ATPase activity and strength of actin binding can be modified
through phosphorylation of the regulatory light chains, while the coiled-coil
tail domains of the MHCs are required for assembly of multiple myosin
molecules into organized filaments (Tan et
al., 1992).
We constructed a myosin-YFP transgene (mYFP-myosin IIDN) in
which the YFP moiety has replaced the actin-binding motor head domain of the
myosin heavy chain, zipper (Fig.
3). Based on equivalent modifications in Dictyostelium,
mYFP-myosin IIDN homodimers should completely lack actin binding
and contractility, and the `single headed' wild-type myosin/mYFP-myosin
IIDN heterodimers should have severely decreased actin binding and
contractility (Burns et al.,
1995; Uyeda and Yumura,
2000
; Zang and Spudich,
1998
). Consistent with this, we find that YFP-containing myosin
isolated from mYFP-myosin IIDN expressing Drosophila
embryos shows reduced actin binding when compared with wild-type myosin in a
standard spin down assay (see Fig. S1 in the supplementary material). However,
we do not detect any dominant-negative activity of this transgene during
embryogenesis, presumably because of the high levels of endogenous myosin.
To analyze the localization of this mYFP-myosin IIDN, we used
the Gal4 system to express the transgene uniformly in embryos that also carry
wild-type copies of zipper. For comparison we examined: (1) a fully
functional myosin-GFP fusion, in which GFP is fused to the myosin light chain,
sqhGFP (Royou et al.,
2004); and (2) the endogenous myosin II of wild-type embryos. We
found no differences between the localization patterns of sqhGFP and
endogenous myosin, and herein refer only to the endogenous.
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The first differences between functional and non-functional myosin are observed at the onset of gastrulation. Unlike endogenous myosin (Fig. 3D), mYFP-myosin IIDN fails to localize apically at the onset of ventral furrow formation (Fig. 3J) and throughout later stages of apical constriction (Fig. 3E,K) and invagination (Fig. 3F,L). The ability of these cells to undergo normal ventral furrow formation despite a lack of apically localized mYFP-myosin IIDN presumably reflects the activity of endogenous zipper. Both endogenous myosin and mYFP-myosin IIDN are lost from the basal side of the invaginating ventral furrow cells. This basal loss is slightly delayed (Fig. 3E,K) and patchy (Fig. 3F,L) for mYFP-myosin IIDN, but otherwise proceeds normally.
The requirement for actin binding and subsequent actin-dependent
contractile activity therefore appears to distinguish two functionally
different modes of myosin localization: an actin-independent mode of
localization during cellularization and cytokinesis, and a second mode during
gastrulation where localization to the apical side of the cell is dependent
upon actin binding/contractility. It is possible that the mYFP-myosin
IIDN is defective in ways other than its ability to interact with
actin. However, equivalent constructs in Dictyostelium do not effect any other
aspects of myosin function, including RLC phosphorylation or filament assembly
(Burns et al., 1995;
Uyeda and Yumura, 2000
;
Zang and Spudich, 1998
).
Therefore, although such secondary effects can not be entirely ruled out here,
the defects seen are most likely a result of the inability to interact with
actin and at the very least distinguish two different types of myosin
localization to the apical and basal side of the cell. They also highlight the
potential importance of actin-myosin interaction and contractility as a target
for fog signaling.
RhoGEF, Rho-kinase and the fog pathway of myosin localization
The components acting downstream of fog to mediate its effects on
the cytoskeleton are largely unknown. One candidate, RhoGEF2 (a guanine
nucleotide exchange factor that promotes Rho activation) has been shown to be
required for ventral furrow formation and can genetically interact with a
fog transgene (Barrett et al.,
1997; Hacker and Perrimon,
1998
). However, embryos mutant for RhoGEF2 have a much
more severe disruption of ventral furrow formation than embryos mutant for
fog and the point at which the products of these two genes interact
on a mechanistic or subcellular level is unknown. Recent studies have shown a
requirement for RhoGEF2 in controlling actin dynamics/stability
during cellularization (Grosshans et al.,
2005
; Padash Barmchi et al.,
2005
) and have also shown a disruption to myosin localization at
gastrulation (Nikolaidou and Barrett,
2004
). We therefore examined the re-localization of myosin during
cellularization and gastrulation in RhoGEF2 mutants and extended
previous studies by looking at a potential downstream effector of
RhoGEF2 signaling.
|
However, despite the defects during early cellularization, RhoGEF2
mutant embryos that reach the end of cellularization look remarkably normal
(Fig. 4C,F,G,J). The
irregularity of the cellularization front recovers, particularly in the
ventral cells, and both the increased cell depth and basal loss of myosin
occur normally in these cells. However, in RhoGEF2 embryos, precisely
staged for the onset of gastrulation, there is an absolute failure to
re-localize myosin to the apical side of the ventral cells
(Fig. 4H,K), despite a normal
loss of myosin from the basal side of these cells
(Fig. 4H-L). This is consistent
with independent mechanisms controlling the basal loss and apical accumulation
of myosin during gastrulation and demonstrates an absolute requirement for
RhoGEF2 in apical myosin localization. It also confirms the previous report of
RhoGEF2 being required for apical myosin in ventral furrow cells
(Nikolaidou and Barrett,
2004).
RhoGEF2 interacts with myosin in other systems through the Rho-kinase
family of Ser/Thr kinases that inhibit myosin phosphatase and also directly
phosphorylate myosin (Amano et al.,
1996). Both these activities lead to activation of actin binding
by myosin and increased actomyosin based contractility. Additional myosin
activators include MLCK and citron kinase but the extent to which these
different activators play specific or overlapping roles with Rho-kinase is
unclear (Matsumura et al.,
2001
), and the role of any of these myosin activators during
Drosophila gastrulation is not known.
|
Despite these defects, many Drok mutant embryos complete cellularization (Fig. 5F) and though the increased depth of cellularization in ventral cells is difficult to discern, basal loss of myosin proceeds normally (Fig. 5C,F,G,J). However, Drok mutant embryos show a complete failure to localize myosin to the apical side of the ventral cells at the onset of gastrulation (n=14) (Fig. 5F). At later stages of gastrulation, the outer layer of wild-type embryos consists of a single cell layered epithelium that folds in specific locations during germband extension (Fig. 5H). In Drok mutant embryos this morphology is severely disrupted and the outer epithelium becomes multilayered and irregular, containing large often rounded cells (Fig. 5K). Drok is therefore required to maintain epithelial integrity.
Both Drok and RhoGEF2 mutant embryos show defects during cellularization and then fail to localize myosin to the apical side of ventral cells at gastrulation. However, it is unlikely that the earlier cellularization defects are what prevent the later apical myosin localization as many other cellularization mutants, such as nullo, display severe cellularization defects but still go on to localize myosin to the apical side of ventral cells at the onset of gastrulation (Fig. 5I,L). The failure of Drok and RhoGEF2 mutant embryos to localize myosin apically during gastrulation therefore probably reflects a direct requirement for both these genes in the apical localization of myosin. Despite these disruptions to gastrulation, Drok embryos do still produce fog protein that is as punctate and apically concentrated as in wild-type embryos (Fig. 1F). This is therefore consistent with a model whereby Drok driven activation of myosin contractility drives myosin apically in response to fog and RhoGEF2 signaling.
|
This connection is likely to require adherens junctions that anchor the
actin-myosin cytoskeleton to the cell membrane and hold the cells of an
epithelium together. Previous studies have concentrated on the role of
junctions in cell polarity and maintaining integrity of epithelial sheets, or
in cell rearrangements that do not involve changes in cell shape
(Bertet et al., 2004;
Tepass et al., 2001
). Much
less is understood about the role of adherens junctions in specific aspects of
cell shape change.
We therefore analyzed the behavior of adherens junctions in fog-overexpressing embryos. At the completion of cellularization the embryo consists of a single layered epithelium with the basal junctions of cellularization located just apical to the myosin rich cellularization front and the newly forming adherens junctions located about 6 µm in from the apical surface of the embryo (Fig. 7A). At the onset of ventral furrow formation, adherens junctions in the ventral most region of the embryo shift to a completely apical location as the cell surfaces flatten, whereas the junctions in more lateral cells maintain their sub-apical position (Fig. 7B). These relative positions are maintained during the phase of apical constriction (Fig. 7C) as basal junctions gradually disappear. When fog is expressed throughout the embryo the apical shift of adherens junctions occurs normally in the ventral furrow region (Fig. 7D,F), but now also occurs in more lateral and dorsal cells (Fig. 7E,G) and these junctions are more tightly condensed than the equivalent junctions of control embryos (Fig. 7G). The apical localization of myosin seen in fog-overexpressing embryos therefore correlates with an apical shift in adherens junctions.
The adherens junctions are possibly being pulled into an apical position because of forces generated by contractile myosin that has been apically re-localized in response to fog signal. To investigate the connection between myosin contractility and adherens junctions, we looked at myosin localization in embryos that lack adherens junctions.
It is not possible to examine embryos totally lacking junctional components
such as Armadillo (Arm) as the maternally supplied components are required
earlier during oogenesis. To get around this problem we have made use of the
effects of nullo protein. Expression of nullo during late
cellularization completely blocks the formation of apical spot junctions
(Hunter et al., 2002;
Hunter and Wieschaus, 2000
).
To confirm that results using this technique are due to the lack of adherens
junctions and not to additional effects of nullo expression we
repeated the analysis with embryos made from arm043A01
germline clones. The arm043A0 allele is of the `medium
class' of arm alleles, lacking the last few Arm repeats and the
entire C terminus. Germline clones of this class of alleles produce sufficient
levels of Arm function to enable a few eggs to complete oogenesis but
subsequent function of Arm in the embryo is severely compromised and these
embryos fail to assemble apical adherens junctions
(Cox et al., 1996
;
Tolwinski and Wieschaus,
2004
). We find the same results using both techniques.
In both cases, myosin localizes normally to the basal cellularization front (Fig. 7H) (A. Sokac, unpublished) and to the apical surface of cells in the ventral furrow (Fig. 7I,J). This implies that functional Arm-containing junctions are not required for myosin to become localized within the cell. However, subsequent events are affected. As ventral furrow cells of wild-type embryos undergo apical constriction, myosin is seen throughout the apical surface of cells (Fig. 7K) but in embryos lacking junctions to tether the actin-myosin network myosin appears to contract into the center or side of the cell forming a tight `ball' of presumably contracted myosin (Fig. 7L,M). The most likely explanation of these results is that myosin contractility is normal in cells lacking adherens junctions but when myosin is no longer tethered to junctions it contracts without being able to exert force on the plasma membrane. As a result, these cells are unable to flatten or constrict their apical surfaces. These results suggest that apically localized myosin is contractile and that this contractility alone is not sufficient to result in changes in cell shape but must be tethered to the apical adherens junctions to elicit apical flattening and constriction.
Adherens junctions are also known to play an important role in establishing
and maintaining apicobasal polarity in epithelial cells
(Nelson, 2003). However, our
results demonstrate that the polarizing signal for the apical activation of
myosin is not dependent upon any polarizing influence emanating from intact
apical adherens junctions. This is consistent with the idea that it is the Fog
protein, through its apical secretion and reception, that provides the
polarizing signal for myosin activation and that this process is independent
of intact adherens junctions.
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Discussion |
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We also demonstrate that apical myosin localization requires the ability of myosin to interact and/or contract with actin. Furthermore, we show that fog signaling results in a shift of adherens junctions from their usual apicolateral position to a more apical position and that these junctions are necessary to translate contractile forces into physical changes in cell shape.
Taken together these data provide us with the following model
(Fig. 8). Expression of the
patterning gene twi in the prospective mesoderm cells results in
activation of fog transcription. The resulting fog protein
is then secreted from the apical surface of the cells and this signal
activates fog receptors. The degree to which this activation is
paracrine versus autocrine has yet to be determined. The apically activated
receptors trigger a transduction pathway involving the G-alpha subunit,
Concertina, and the Rho activator RhoGEF2. A downstream target of this pathway
is Rho-kinase, which in turn activates the ability of myosin to interact and
contract with actin in this sub-apical region of the cell. A localized source
of activated actin-myosin contractility initiates an active motor-driven
mechanism of myosin localization which concentrates contractile myosin to the
apical side of the cell. This actin-myosin network is tethered to the cell
surface through adherens junctions. Contraction of this network therefore puts
tension on the junctions, pulling them into a completely apical location and
flattening the domed apical surface in the process. Continued contraction
exerts further tension and ultimately pulls the junctions together so much
that the entire apical cell surface constricts. Intriguingly, RhoGEF2 protein
can associate with the tips of microtubules in cultured cells
(Rogers et al., 2004). The
extent to which this may add to a polarization of the fog pathway
during gastrulation and how this ties in with the above model will therefore
be interesting avenues for further investigation. It will also be important to
examine any changes to the actin and microtubule organization of these
cells.
The effects of fog overexpression also reveal some interesting
features of fog function. Rates of apical flattening show more
variability with the UASfog construct than reported for heat-shock
fog embryos (Morize et al.,
1998). It is unclear whether this reflects a dose sensitivity of
the fog signaling pathway or whether the heat shock itself produces
certain changes in cytoarchitecture that alter the ability of the cell to
respond to fog signal. Furthermore, the fog-overexpressing
embryos are already making significant levels of fog protein at the
onset of cellularization but myosin localization during cellularization
remains unaffected. It is only after the completion of cellularization that
the embryo becomes receptive to fog signaling and apical myosin
localization. The ventral cells are then more receptive than dorsolateral
cells. The nature of this receptivity, whether it simply requires the timely
transcription of an additional component of the signaling cascade or whether
it reflects a more complex aspect of cell states will be interesting to
determine.
Two distinct mechanisms of myosin localization
In the transition between the morphogenetic processes of cellularization
and gastrulation, myosin localizes in an intriguing pattern. In the dorsal and
lateral regions of the embryo, myosin is restricted to the basal side of the
cells. However, in the ventral-most cells this myosin is lost basally and
accumulates apically. At first glance, this pattern of myosin localization is
suggestive of a functional link between the basal loss of myosin and its
appearance apically. However, by closely following myosin dynamics throughout
cellularization, we find that these two events are temporally separable
(R.E.D.-H., unpublished), with decreased levels of basally localized myosin
significantly preceding the appearance of myosin on the apical side of the
cell.
Here, we show that not only can the basal loss and apical accumulation of myosin be temporally separated, they can also be functionally separated. In embryos that express the fog signal ubiquitously, myosin accumulates apically in all cells but the basal loss of myosin is still restricted to the ventral cells. This demonstrates that a loss of myosin from the basal side of the cell is not required to localize myosin to the apical side. Conversely, in embryos mutant for RhoGEF2 we find that myosin fails to accumulate apically but does still decrease basally in the ventral cells. The source of apically localized myosin (e.g. unlocalized cytoplasmic pool versus de novo synthesis) will therefore be interesting to determine.
Further evidence for functional differences between apical and basal localization comes from the differing requirements for junctional components and actin binding. Embryos made from arm germline clones have significantly reduced levels of functional Arm to incorporate into the basal junctions during cellularization and the apical adherens junctions at gastrulation. However, myosin is still able to correctly localize to the cellularization front, and to the apical side of ventral cells, but in the absence of junctions subsequent aspects of apical myosin localization become abnormal. The untethered myosin appears to contract into a tight ball in the center of the cell and fails to mediate the cell shape changes of apical flattening and constriction. Finally, we demonstrate that a YFP-myosin fusion protein that is compromised in both its actin binding and contractility still localizes to the cellularization front but fails to localize apically at gastrulation. This implies a requirement for actin binding and/or contractility in the mechanism by which myosin localizes apically during gastrulation, but not for its correct localization basally during cellularization. The fact that this YFP-myosin localizes correctly to cytokinetic furrows indicates that both myosin localization during cellularization and cytokinesis are mechanistically more closely related than the mechanisms of apical and basal myosin localization that co-exist in the same cells at the onset of ventral furrow formation.
Multiple pathways controlling ventral furrow formation
It has been proposed that multiple pathways contribute to ventral furrow
formation during Drosophila gastrulation. The ventral furrow of
fog mutant embryos is disrupted but does still form, presumably
through a parallel pathway of cell shape change
(Costa et al., 1994). Embryos
expressing fog from a transgene show a genetic interaction with
RhoGEF2 during gastrulation
(Barrett et al., 1997
) but
RhoGEF2 mutant embryos show a much more severe ventral furrow
phenotype than fog mutants
(Barrett et al., 1997
;
Hacker and Perrimon, 1998
).
Inhibition of RhoA function at later stages of development has a wide range of
phenotypic consequences including disruption of apically localized myosin
(Bloor and Kiehart, 2002
).
Removal of RhoGEF2 also lowers levels of apical myosin
(Nikolaidou and Barrett,
2004
), though the degree and mechanism of this effect are less
clear. How these multiple pathways interact and combine to direct cell shape
changes during ventral furrow formation has therefore been unclear.
Here, we demonstrate that fog signaling directs the localization of myosin to the apical side of the cell, but in embryos lacking fog many cells of the ventral furrow still localize myosin apically. The ability of these cells to localize myosin in the absence of fog reveals that the parallel, fog-independent pathway also functions to control the apical localization of myosin. Furthermore, we demonstrate that RhoGEF2 is absolutely required for apical localization of myosin in all cells of the ventral furrow. The difference in apical myosin localization between fog and RhoGEF2 mutants is consistent with the fact that embryos lacking RhoGEF2 display more severe ventral furrow defects than fog mutants. These results therefore reveal that the fog pathway and the fog-independent pathway probably converge at the level of RhoGEF2 signaling of apical myosin localization.
In addition, the phenotypes of embryos lacking the Rho effector Rho-kinase (Drok) were distinct from those of RhoGEF2 mutant embryos. Although both genes are required for apical myosin localization during gastrulation, the Drok embryos have more severe cellularization defects than RhoGEF2 and some of these defects (particularly those involving nuclear morphology and fall-out) may even precede cellularization. This result is consistent with the idea that the RhoGEFs involved in activating Rho signaling tend to show more specificity for individual processes than the effectors downstream of Rho.
Furthermore, our results indicate that the role of RhoGEF2 signaling in the process of apical myosin localization is most likely through activation of actin-myosin binding/contractility. We show that the apical myosin pathway requires Rho-kinase, a RhoGEF2 effector known to directly activate myosin contractility. We also show that apical myosin localization is blocked when myosin contractility is impaired and that RhoGEF2 signaling is absolutely required for this contractility based form of myosin localization. Conversely, the form of myosin localization that is independent of myosin contractility (during cellularization) is also independent of RhoGEF2 signaling. Interestingly many mutant backgrounds that severely disrupt cellularization are still capable of localizing apical myosin in ventral cells at the onset of gastrulation (E.F.W., unpublished). This not only provides further evidence of the differences between these two mechanisms of myosin function, but also strengthens the significance of the RhoGEF2 results, as RhoGEF2 has only a mild cellularization phenotype but completely blocks apical myosin localization at gastrulation.
Parallels to other systems
Rho signaling has been shown to control myosin activation in a number of
different systems from cytokinesis in C. elegans to stress fiber
formation in vertebrates
(Etienne-Manneville and Hall,
2002; Piekny and Mains,
2002
; Ridley and Hall,
1992
). A variety of different types of receptors have been shown
to activate Rho signaling (Wettschureck
and Offermanns, 2002
) and one particularly important pathway of
Rho activation is through guanine nucleotide exchange factors (RhoGEFS), which
catalyze the exchange of GDP for GTP on Rho GTPases, thus activating them.
Signaling from Rho through the effector Rho-kinase then results in
phosphorylation of myosin RLC and this cascade has been conserved in a variety
of morphogenetic processes in Drosophila, including dorsal closure
and bristle orientation in the adult epidermis
(Bloor and Kiehart, 2002
;
Mizuno et al., 2002
;
Tan et al., 2003
;
Winter et al., 2001
). Our
results during gastrulation expand the role of this signaling cascade in
Drosophila to include activation of myosin contractility as a means
of controlling the subcellular localization of myosin and draws interesting
parallels to the activators of this cascade in other systems.
In vertebrate cells, lysophosphatidic acid (LPA) activates a
G-protein-coupled receptor. The alpha subunit of this G protein is a member of
the G-alpha12/13 subclass, signaling through which leads to
activation of Lsc/p115RhoGEF and the downstream cascade of Rho and Rho kinase
activities that lead to myosin activation and formation of stress fibers and
focal adhesions (Kozasa et al.,
1998; Sawada et al.,
2002
). An additional Rho effector acting in this pathway is Dia,
the interactions of which with both actin and microtubules may play a role in
mediating alignment of microtubules and microfilaments during stress fiber
formation (Tsuji et al.,
2002
). In the fog ventral furrow pathway, a number of
direct homologies can be drawn to the LPA pathway of vertebrates. Fog
signaling is thought to involve the G-protein alpha subunit cta,
which also belongs to the G-alpha12/13 subfamily, though the
identity of the receptor is not known
(Parks and Wieschaus, 1991
).
We show here that the pathway of myosin localization in the ventral furrow
also involves RhoGEF2, a Drosophila homologue of
Lsc/p115RhoGEF and that the consequence of this activation of Rho signaling,
is the Drok-mediated activation/localization of myosin, just as seen
in stress fiber formation. To extend this comparison further, it will be
interesting to address the role of Dia in ventral furrow formation.
Interesting parallels can also be drawn to myosin regulation in
Dictyostelium. Peculiarities of the situation in Dictyostelium (including the
lack of a Rho homologue, the importance of heavy rather than light chain
phosphorylation and the fact that cytokinesis can occur in the absence of
certain aspects of myosin phosphorylation and function) have led to the
suggestion that Dictyostelium has developed a very different and derived
system for regulating myosin activity (see
Matsumura et al., 2001). Our
results, however, highlight some striking similarities. A myosin-GFP fusion
protein that disrupts the ability of the head domain of MHC to interact with
actin has been produced in Dictyostelium and just as we have found in
Drosophila this compromised form of myosin is still able to localize
correctly during cytokinesis (Zang and
Spudich, 1998
) but is unable to do so during cAMP-activated
chemotaxis (Levi et al.,
2002
). As in Drosophila, ventral furrow formation this
cAMP activated chemotaxis involves recruitment of myosin to the cell cortex
and is a G-protein-coupled process. Our results therefore raise the
possibility of distinct mechanistic parallels between the myosin based
morphogenetic processes of these two organisms. Understanding how the multiple
pathways controlling myosin dynamics in Drosophila are integrated
into a developmental context, including how they interface with one another,
with their downstream effectors and with the upstream patterning genes that
control cell fate, may therefore provide insights that will extend beyond
Drosophila to a diverse range of myosin-based processes.
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ACKNOWLEDGMENTS |
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Footnotes |
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Supplementary material for this article is available at http://dev.biologists.org/cgi/content/full/132/18/4165/DC1
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