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First published online 4 October 2006
doi: 10.1242/dev.02588
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1 Instituto de Medicina Molecular, Faculdade de Medicina de Lisboa,
Portugal.
2 Instituto Gulbenkian de Ciência, Oeiras, Portugal.
3 Department of Zoology, University of Cambridge, Downing Street, Cambridge CB2
3EJ, UK.
4 Centro Andaluz de Biología del Desarrollo, Universidad Pablo de
Olavide, Sevilla, Spain.
5 Department of Physiology, School of Medical Sciences, University of Bristol,
University Walk, Bristol BS8 1TD, UK.
* Author for correspondence (e-mail: ajacinto{at}fm.ul.pt)
Accepted 18 August 2006
 |
SUMMARY |
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 TOP SUMMARY INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Key words: Cell invagination, Myosin II, Actin cytoskeleton, Small GTPase Rho1, RhoGEFs, RhoGAPs, Posterior spiracles, Drosophila
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INTRODUCTION |
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TOPSUMMARYINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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;
Symons and Settleman, 2000).
The GTPase switch GTP/GDP is determined by the action of two main classes of
regulatory proteins: the Guanine nucleotide exchange factors (GEFs), which act
as GTPase activators by releasing GDP and allowing its replacement by GTP
(Rossman et al., 2005), and
the GTPase activating proteins (GAPs), which promote the GTPase inactive state
by stimulating its GTP hydrolysing capacity
(Moon and Zheng, 2003).
Spatial and temporal restriction of small RhoGTPase activity inside a cell
is fundamental, for example, to the regulation of movements and cell-cell
contacts that are required for morphogenesis. During the formation of
epithelia, Rac and Cdc42 are specifically recruited to the cadherin-cadherin
contact sites formed between neighbouring cells, where they drive the
formation of filopodia and/or lamellipodia that will contribute to generate
intimate cell-cell contacts (Braga,
2000). Another example of spatially restricted RhoGTPase activity
has been found during migration of single cells, in which Rac-GTP accumulates
at higher levels at the leading edge
(Kraynov et al., 2000), where
it induces Actin polymerisation and integrin adhesion complex assembly.
Evidence that the spatial and temporal control of small RhoGTPase function
must be correlated with the activity of Rho regulators during tissue
morphogenesis is suggested from studies on cell shape changes occurring during
gastrulation and neurulation. In Drosophila gastrulation, a secreted
factor Folded gastrulation (Costa et al.,
1994) initiates a signal through interaction with a G
protein-coupled receptor (still unknown) and a heterotrimeric G protein
subunit, Concertina (Parks and
Wieschaus, 1991) to activate DRhoGEF2, the small GTPase Rho1 and
subsequently Myosin II (Barrett et al.,
1997; Nikolaidou and Barrett,
2004). This pathway is essential to induce constriction of the
apical surface of mesodermal cells, leading to their invagination. In
vertebrates, apical constriction of neuroepithelial cells, mediated by
reorganisation of the Actin cytoskeleton, contributes to closure of the
anterior neural tube. Interestingly, mice mutant for p190 RhoGAP show defects
in anterior neural tube closure, due to failure in apical constriction and in
intracellular re-organisation of the Actin cytoskeleton of neuroepithelial
cells (Brouns et al., 2000).
Together, these observations suggest that Rho acts apically during
constriction of (neuro)epithelial cells and its function is tightly regulated
by RhoGEF and RhoGAP activities.
Despite this, it is still unclear how spatially localised activation of
small RhoGTPases is achieved and, with few exceptions
(Bement et al., 2005), no tools
exist that allow the direct visualisation of their activity during embryonic
development. Additionally, little is known about how Rho regulators spatially
and temporally control their targets in vivo, during the complex morphogenetic
movements that shape the embryo.
Cell invagination is a widespread movement during embryogenesis and it is
commonly dependent on Rho function (Pilot
and Lecuit, 2005). Using the posterior spiracles of the
Drosophila embryo as a model, we show that apically localised Myosin
II and Rho1 are essential to control this morphogenetic movement. By
expressing a GFP-based probe that allows the visualisation of active Rho1, we
present evidence that during cell invagination this RhoGTPase is exclusively
activated on the apical membrane of epithelial cells. Correlating with this
local activation, we found that two Rho1 activators (RhoGEF2 and RhoGEF64C)
are upregulated and localise apically in the invaginating cells, while a Rho1
inhibitor, the RhoGAP Cv-c (Crossveinless-c), occupies the complementary
basolateral membrane domain. This differential distribution of Rho regulators
is required for the correct function of Rho1, which drives invagination of
epithelial cells.
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MATERIALS AND METHODS |
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TOPSUMMARYINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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);
SqhAX3; ubi-Sqh-GFP
(Royou et al., 2002); ems-GAL4
(Merabet et al., 2002);
UAS-Cat-GFP (Oda and Tsukita,
1999); UAS-GFP-Actin
(Verkhusha et al., 1999);
UAS-mRFP-Actin#30 (this study); UAS-GFP (Bloomington Stock Center); UAS-RhoN19
(Strutt et al., 1997);
UAS-PNKG58AeGFP (this study); UAS-RhoV14
(Fanto et al., 2000);
UAS-RacN17 (Luo et al., 1994);
UAS-DRhoGEF64C FL and UAS-DRhoGEF64C
Dbl
(Bashaw et al., 2001);
UAS-GEF64C5'3'UTR (this study); UAS-Cv-c
(Denholm et al., 2005);
UAS-Venus-Cv-c-93 (this study); rho11B/CyO
(Magie and Parkhurst, 2005);
rho172R/CyO (Strutt et
al., 1997); RpII140wimp/TM3 Sb1 (Bloomington
Stock Center); gef64C1/TM3 and
gef64C29/TM3 (Bashaw et
al., 2001); P[FRT(whs)]G13
DRhoGEF2I(2)04291/CyO
(Hacker and Perrimon, 1998);
yw P[FRT(whs)]101 sqh1/FM7
(Karess et al., 1991);
cv-c7/TM3 (Denholm et
al., 2005).
Germline clones of DRhoGEF2I(2)04291 and
sqh1 were generated using the FLP-DFS system
(Chou and Perrimon, 1996). yw
P[FRT(whs)]101 sqh1/w*
P[FRT(whs)]101 ovoD1 females carrying germline clones
were crossed with w; ems-GAL4 UAS-GFP-Actin males. Maternal and zygotic
mutants for rho11B were generated using the wimp
strategy (Magie et al.,
1999).
Live imaging
Embryos expressing ubi-DE-CadGFP or GFP-Actin with the ems-GAL4 driver were
prepared for live imaging and analysed as described
(Woolner et al., 2005).
Molecular biology and transgenic lines
To express PKNG58AeGFP, we generated pUAST-PKNG58AeGFP. The N-terminus of
Drosophila PKN (first 339 amino acids), including the point mutation
G58A, was PCR amplified from pCasperPKNG58A (kind gift from Jeffrey
Settleman). This PCR fragment was cloned into the EcoRI/XbaI
sites of pUAST (Brand and Perrimon,
1993). Next, eGFP was PCR amplified from pEGFP-N1 (Clontech) and
cloned into the XbaI site of the previous construct.
Venus-Cv-c was generated by cloning the full-length cDNA of cv-c
(Denholm et al., 2005) into
the pENTR directional TOPO cloning vector (pENTR/D) and subsequently
transferred into pTVW (UASt promoter, N-terminal Venus tag - obtained from
DGRC) using the Gateway technology (Invitrogen).
mRFP-Actin was made by fusing the 700 bp fragment of mRFP
(Campbell et al., 2002) with
1100 bp of Drosophila Actin 5c, and cloned into pUAST.
pUAST-RhoGEF64C5'3'UTR was obtained by digestion of
pUASTGEF64C FL (Bashaw et al.,
2001) with EcoRI and NheI and subcloning the
resulting fragment into pUAST. Transgenic flies were generated using standard
procedures.
S2 cell culture, RNAi and transfection
Drosophila Schneider S2 cells were cultured and RNAi was performed
according to Clemens et al. and Schneider
(Clemens et al., 2000;
Schneider, 1972). Templates
for in vitro transcription of Rho1 were generated as described below
for RhoGEF64C, and contained 527 bp from position 655 (CG8416-RC).
Expression of PKNG58AeGFP in S2 cells was obtained by co-transfection of 20 ng
of pUAST-PKNG58AeGFP with 100 ng of pAc5.1/V5-HisB-GAL4 in six well plates. In
some cases, 100 ng of the active forms RhoV14, RacV12, Cdc42V12 or for the
dominant-negative, RhoN19, RacN17 or Cdc42N17 cloned in pUAST were also
co-transfected by using the CellFectin method according to the manufacturer's
specifications (Invitrogen Life Technologies).
GTPase-binding assays
S2 cell extracts expressing PKNG58AeGFP were prepared 24 hours
post-transfection, in lysis buffer [50 mmol/l Tris (pH 7.8), 150 mmol/l NaCl,
1% Nonidet-40], and were precleared twice with protein A-Sepharose beads for
30 minutes (4°C). Drosophila Rho1, Rac1, Rac2 and Cdc42-GST
proteins were prepared according to standard methods and pre-incubated with
GDP or GTP
S according to (Lu and Settleman, 1999). GTPase binding to
PKNG58AeGFP was next assayed by SDS-PAGE, followed by immunoblotting with an
anti-GFP monoclonal antibody (Roche).
RNA interference in embryos
For the synthesis of dsRNA for RhoGEF64C, a region of 773 bp from
position 1237 (CG32239-RA) was PCR amplified from genomic DNA with primer
pairs containing the T7 promoter sequence at the 5' end. The PCR
products were used as templates for the T7 transcription reactions with the T7
Ribomax Large Scale Production Kit (Promega). The dsRNA was dissolved in
injection buffer at a final concentration of 2.2 µg/µl and injected into
0- to 1-hour-old embryos derived from the cross between w; ems-GAL4
UAS-GFP-Actin females and yw males.
In situ hybridisation, immunohistochemistry and cuticle preparations
Whole-mount in situ hybridisation was carried out using standard methods
(Lehmann and Tautz, 1994) with
a digoxigenin-labelled probe generated by transcription of DRhoGEF64C
(CG32239-RA) DNA region 1237-2010 (nucleotide positions).
For immunohistochemistry, embryos were dechorionated in commercial bleach,
fixed in a 1:1 mix of formaldehyde 4% in PBS (Ca2+ free): n-heptane
for 30 minutes at room temperature and hand devitellinised. The following
primary antibodies were used: mouse anti-Armadillo (N2 7A1, Hybridoma Bank)
1:50; rabbit anti-GFP (AbCAM #290) 1:2000; mouse anti-Dlg (4F3, Hybridoma
Bank) 1:250; mouse anti--Spectrin (3A9, Hybridoma Bank), 1:10; mouse
anti-GEF64C (Bashaw et al.,
2001) 1:50; rabbit anti-DRhoGEF2
(Padash Barmchi et al., 2005)
1:1000; rabbit anti-Myosin II heavy chain
(Bloor and Kiehart, 2001)
1:500; mouse anti-Rho1 (P1D9) (Magie et
al., 2002) 1:50 and Phalloidin Alexa-594 (Molecular Probes) 1:200.
All dilutions were done in PBT (BSA 1% and Triton X-100 0.1% in PBS).
Secondary antibodies were coupled to Alexa488 or
Alexa594 (Molecular Probes), and diluted in PBT at 1:400. Cuticle
preparations were done as described (Hu
and Castelli-Gair, 1999).
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RESULTS |
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TOPSUMMARYINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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70 epithelial cells localised
posteriorly to the eighth abdominal tracheal pit start to invaginate into the
embryo (Hu and Castelli-Gair,
1999). We studied spiracle cell behaviour during the invagination
process by performing time-lapse analysis in live Drosophila embryos
expressing DE-Cadherin fused to GFP (Oda
and Tsukita, 2001). At stage 11, when the embryonic germ band is
fully extended, the future spiracular chamber cells were still localised
superficially and were indistinguishable from the neighbouring cells
(Fig. 1A, t=0 minutes,
see Movie S1 in the supplementary material and
Fig. 1B, part i). As the germ
band started to retract, the most anterior cells rearranged their relative
positions around the tracheal opening (Fig.
1B, part ii, arrow) until they met, forming a lumen in continuity
with the tracheal dorsal branch (Fig.
1B, part iii). During this rearrangement, all the cells in the
spiracle primordium constricted apically, resulting in an inward depression of
the tissue around the luminal opening (Fig.
1A, t=134 minutes). This invagination occurred in a
spatially and temporally controlled manner, as the cells adjacent to the
tracheal opening invaginated first (forming a lumen), followed in turn by the
more distal (posterior) cells (Fig.
1B, parts iii,iv; see also the diagram in
Fig. 1E). Apical constriction
and ordered invagination continued until the end of germ band retraction, by
which time all the spiracle chamber cells had invaginated into the embryo,
forming a tube continuous with the trachea
(Fig. 1A, t=232
minutes, Fig. 1B, part v, and
Fig. 1D).
|
)
(Fig. 3A).
Myosin II is apically enriched and essential for spiracle invagination
Apical constriction and invagination of epithelial cells are generally
linked to recruitment of both F-Actin and Myosin II to the apical side
(Young et al., 1991). Using
transgenic embryos expressing the regulatory light chain of Myosin II [MRLC,
encoded by the spaghetti squash (sqh) gene] fused to GFP
(Royou et al., 2002), or
antibodies against the heavy chain of Myosin II (Zip), we confirmed that this
Myosin was apically enriched in the cells forming the spiracular chamber (see
Fig. S1A in the supplementary material, wild type). Furthermore, this
enrichment correlated with the onset of apical constriction. At stage 11,
before cell invagination started, Myosin II was distributed along the entire
lateral membrane, co-localising with the basolateral marker Dlg (Discs large)
(Fig. 2A, stage 11). As the
spiracle cells began constricting apically, puncta of Myosin II appeared on
their apical side (Fig. 2A,
stage 12). At stage 13, when the entire primordium had invaginated, Myosin II
was highly enriched apically, in a largely non-overlapping domain relative to
Dlg (Fig. 2A, stage 13). This
localisation, together with the observation that Actin also accumulates
apically, co-localising with Myosin II, suggests that a contractile force
based on Actin and Myosin II is acting during the process of invagination.
To test this hypothesis, we analysed the morphology of the spiracular
chamber in embryos with a strong reduction in the maternal and zygotic levels
of MRLC, by making germline clones of a hypomorphic mutation in sqh,
sqh1 (Karess et al.,
1991). This mutation, as opposed to null mutations in Myosin II,
does not abolish egg laying (Barros et al.,
2003) and allows embryonic development to proceed beyond spiracle
formation. Consistent with a role for apical Myosin II in exerting the driving
force during cell invagination, spiracle cells failed to invaginate and to
form a lumen in sqh1GLC embryos
(Fig. 2B, part i). In addition,
clusters of cells were seen detached from the main group
(Fig. 2B, part i, arrowhead),
suggesting that Myosin II is also important in coordinating cellular movement
during invagination. Although Actin was still apically enriched in most of the
spiracle cells in these mutants, failure in apical constriction was
occasionally observed (Fig.
2Bi'). Embryos that were rescued by the paternal
contribution of MRLC exhibited a milder phenotype, in which spiracle cells
were able to invaginate but failed to form a lumen, showing a disrupted apical
Actin cytoskeleton (Fig. 2B,
part ii, arrow; compare with wild type in
Fig. 2B). These observations
demonstrate that apical Myosin II, together with Actin, is essential to
correctly drive epithelial cell invagination and lumen formation.
|
). Thus,
apical enrichment of F-Actin and Myosin II during spiracle cell invagination
suggests the involvement of Drosophila Rho1 in this process. To test
this hypothesis, we analysed spiracle morphology in both zygotic and
maternal/zygotic mutants for Rho1, using the wimp strategy
(Magie et al., 1999). In both
cases, a large majority of Rho11B null embryos showed a
dilated or irregular Filzkörper, especially at the most distal part of
the tube, symptomatic of disrupted invagination (75.8% in zygotic
Rho11B mutants, n=116 and 69% in
Rho11B maternal/zygotic mutants, n=185)
(Fig. 3B; compare with wild
type, Fig. 3A). These defects
correlated with a partial disruption of the cortical Actin cytoskeleton in the
spiracular chamber of late embryos (Fig.
3D). Additionally, in these experiments about 4% of zygotic
Rho11B mutants and 13% of maternal/zygotic mutants showed
a complete failure in spiracle cell invagination
(Fig. 3C), with the
Filzkörper forming on the surface of the embryo. These observations
suggest that Rho1 activity is essential for proper epithelial cell
invagination and tubulogenesis and controls the assembly of apical Actin
during this process. Similar results were obtained by spiracle-specific expression of a dominant negative Rho protein (RhoN19), which led to a block of the invagination process, causing the spiracle cells to remain on the surface of the embryo (Fig. 3E,F). These uninvaginated cells showed a disrupted cortical Actin cytoskeleton at their apical side, correlating with a non-uniform pattern of apical Myosin II (Fig. 3H, arrowhead; compare with wild type, Fig. 3G, and see Fig. S1A in the supplementary material). Nevertheless, as observed in Rho1 mutants, elongation of the basolateral membrane could still be observed, indicating that Rho1 function is not required for this process (Fig. 3H, arrow). Knocking down Rho1 activity also affected the establishment of proper apical cell-cell adhesion, as assessed by the defects seen in the adherens junctions architecture (see Fig. S1B in the supplementary material).
Rho1 activity is apically restricted during spiracle cell invagination
Loss of Rho1 function led to apical but not basolateral defects in the
invaginating spiracle cells, suggesting that the activity of this GTPase is
required apically during such morphogenetic movement. This prediction is
further supported by two lines of evidence. First, immunofluorescence against
Rho1 protein revealed that, despite being ubiquitous, Rho1 was strongly
enriched on the apical side of the invaginating spiracle cells, surrounding
the lumen of the spiracular chamber (Fig.
4A, parts i,ii). Second, we developed a GFP-based probe (herein
termed PKNG58AeGFP), which recognizes the active, GTP-bound form of Rho1 and
that can be expressed in vivo using the UAS/GAL4 system (see Fig. S2 in the
supplementary material). To follow Rho1 activation and, simultaneously, Actin
distribution, during invagination and tube formation, we expressed this probe
together with mRFP-Actin in the spiracle cells. At stage 11, before the onset
of epithelial cell invagination, PKNG58AeGFP appeared diffuse throughout the
spiracle primordium (Fig. 4B,
early stage 11). Slightly later, a distinct accumulation of the GFP signal was
seen in the first invaginating cells adjacent to the last tracheal pit, and
this was accompanied by a higher accumulation of mRFP-Actin in these cells
(Fig. 4B, late stage 11).
During apical constriction and spiracle cell elongation, our probe accumulated
exclusively at the most apical side of the invaginating cells, where
mRFP-Actin was also found (Fig.
4B, b.c.). After invagination, PKNG58AeGFP and mRFP-Actin were
still detected apically, surrounding the lumen of the spiracular chamber
(Fig. 4B, stage 13 and stage
17). Similar results were observed using six independent transgenic lines of
UAS-PKNG58AeGFP, without any obvious phenotypic defects in spiracle
morphogenesis.
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RhoGEF64C mRNA and protein are apically localised in the spiracle cells
To further understand the mechanism by which Rho1 activity is apically
restricted, we searched for Rho activators that are expressed during spiracle
invagination. We found that RhoGEF64C, previously described to
regulate Rho1 in the embryonic central nervous system
(Bashaw et al., 2001), was
specifically expressed by the invaginating spiracle cells from early stage 12
until the spiracular chamber was fully formed
(Fig. 5A). This RhoGEF is also
present in other multicellular tubes, including the salivary glands, foregut
and hindgut and at low levels in the epidermis.
Immunostaining of RhoGEF64C with a specific monoclonal antibody
(Bashaw et al., 2001) showed
that this RhoGEF localised apically, overlapping with the polymerised F-Actin
surrounding the lumen of the spiracular chamber
(Fig. 5B). Strikingly, we
observed that RhoGEF64C mRNA was also apically enriched in the
invaginating spiracle cells (Fig.
5C) and in the hindgut (Fig.
5D), indicating that this mRNA can be apically transported in
epithelial cells. These observations suggest the possibility that RhoGEF64C
activity is spatially controlled by apical targeting of its mRNA, followed by
local translation.
To find the sequence elements responsible for the apical localization of
RhoGEF64C mRNA, we overexpressed several truncations of RhoGEF64C
cDNA in the posterior spiracles and assessed for mRNA localisation. Neither
truncating the 5'UTR alone (UAS-RhoGEF64C FL, full length)
(Bashaw et al., 2001) nor both
the 5'UTR and the 3'UTR (UAS-RhoGEF64C 5'3'UTR)
affected the apical localisation of RhoGEF64C mRNA. However, a deletion of 1.8
kb from the 3' region, including the 3'UTR, the Dbl domain (Rho
interacting domain) and a putative PH domain (Pleckstrin Homology)
(UAS-RhoGEF64CDbl) (Bashaw et al.,
2001), disrupted apical mRNA localisation in the posterior
spiracles (Fig. 5E). We
conclude that the apical localising element(s) of RhoGEF64C mRNA are situated
within the C-terminal coding region, which includes the Dbl and PH domains of
this RhoGEF.
|
Dbl, did not revert the RhoN19
phenotype. We conclude that RhoGEF64C acts as a Rho1 activator during the
formation of the spiracular chamber.
|
) (data not
shown).
As the loss of function of RhoGEF64C did not cause invagination
defects, as observed after the complete loss of Rho1 activity, we predicted
that other RhoGEF(s) could be acting together with RhoGEF64C to activate Rho1
apically. A probable candidate is the ubiquitous and apically localised
RhoGEF2, which is Rho1-specific and regulates epithelial cell invagination
during development (Nikolaidou and
Barrett, 2004; Padash Barmchi
et al., 2005). We confirmed the expression and found an apical
enrichment of DRhoGEF2 before and during invagination of the spiracle cells,
overlapping with apical Actin (Fig.
6B and see Fig. S3 in the supplementary material). Furthermore,
34% of maternal and zygotic mutants for the null allele
DRhoGEF2l(2)04291 (DRhoGEF2 MZ)
(Hacker and Perrimon, 1998)
failed spiracle invagination, 26% presented lumen defects and 3.5% showed both
defects (n=201) (see Fig. S3 in the supplementary material). DRhoGEF2
MZ embryos that were also heterozygous or homozygous (1:1) for the
Rhogef64C1 null allele showed a mild increase in the
frequency of spiracle defects (invagination defects, 31%; lumen defects, 35%;
both defects, 8%; n=86). Taken together, these results show that at
least two apical RhoGEFs, RhoGEF2 and RhoGEF64C, contribute to activate Rho1
apically during spiracle cell invagination.
RhoGAP Cv-c is localised basolaterally and is required for proper apical Rho1 activity
One emerging theme in small Rho GTPase regulation studies is that cycling
between their GTP- and GDP-bound states might be required for effective signal
flow in order to elicit downstream biological functions
(Moon and Zheng, 2003).
RhoGAPs are important players in this balance, as they accelerate the return
of RhoGTPases to their inactive state, and thus may act as positive regulators
of Rho function (Symons and Settleman,
2000). Consistent with this view, previous work has implicated the
RhoGAP Cv-c in spiracle cell invagination
(Denholm et al., 2005).
Zygotic cv-c mutants (cv-c7) showed partial or
complete invagination defects in their posterior spiracles, with a strongly
disorganised pattern of apical Actin (Fig.
6A). As the vertebrate homologues of cv-c, p122/DLC-1 and
DLC-2, were shown to act on RhoA, the mammalian homologue of
Drosophila Rho1 (Leung et al.,
2005; Wong et al.,
2003), we hypothesised that Cv-c could also regulate Rho1 cycling
during spiracle morphogenesis.
|
-Spectrin
(Fig. 6Bi-iii). Double
labelling of RhoGEF2 and Cv-c confirmed that these two classes of Rho
regulators essentially occupied non-overlapping domains in the membrane of the
invaginating cells, the apical and basolateral domains, respectively
(Fig.
6Bi'-iii').
To test whether Cv-c controls Rho1 activity during cell invagination, we
overexpressed this RhoGAP in the spiracle cells. An increased expression of a
RhoGAP is expected to downregulate the activity of the target RhoGTPase(s),
thus mimicking the phenotype produced by the expression of their dominant
negative forms. Gain of function of Cv-c [using UAS-Cv-c
(Denholm et al., 2005)] caused
phenotypic defects similar to Rho1 loss of function (Rho1 mutants and
UAS-RhoN19), but not to Rac loss of function (UAS-RacN17, see also Fig. S1B in
the supplementary material): irregular Filzkörpers at the distal end and
partially uninvaginated spiracles (compare
Fig. 6C with wild type in Fig.
S1B in the supplementary material). Frequently, the distal cells of the
spiracular chamber failed to invaginate and showed decreased Rho1 activity, as
confirmed by the lower levels of apical PKNG58AeGFP fluorescence
(Fig. 6D). Furthermore, these
cells maintained an elongated shape, but their apical Actin cytoskeleton was
highly disrupted (Fig. 6C), in
a manner similar to that observed upon expression of RhoN19
(Fig. 3H). Expression of
stronger transgenic lines of UAS-Venus-Cv-c led to completely uninvaginated
spiracles and fully phenocopied the defects caused by RhoN19 (data not shown).
Together, these results indicate that Cv-c acts as a Rho1-GAP, being mainly
excluded from the apical membrane domain in which Rho1 is active.
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|
DISCUSSION |
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TOPSUMMARYINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Using a probe that allows the visualisation of Rho1 activity in the course
of normal development, we present evidence that this GTPase is active at the
apical side during the process of cell invagination. In the spiracles Rho1
activity is essential to control this movement, similarly to that previously
shown during Drosophila gastrulation, when mesodermal cells fail to
invaginate after inhibition of Rho1 function
(Barrett et al., 1997;
Hacker and Perrimon, 1998). We
also observed an apical enrichment of Myosin II, a possible target of
activated Rho1, analogous to that reported in other tissues of the fly embryo
where this type of movement occurs, such as the mesoderm and the salivary
glands (Nikolaidou and Barrett,
2004). Inhibition of Rho1 activity results in a disorganised
pattern of apical Myosin II and F-Actin in the spiracle cells. We suggest that
concentration of active Rho1 at the apical side organises the Actin
cytoskeleton and promotes high Myosin II accumulation/activity in this region,
leading to a contractile Actin-Myosin based force to produce a wedge-shaped
cell.
Our data show that spatial restriction of Rho1 activity is achieved by distinct mechanisms. First, albeit ubiquitous, Rho1 protein is strongly enriched on the apical side of the invaginating spiracle cells. Second, to ensure that this GTPase is active exclusively on that side of the cell, opposing Rho regulators are differentially distributed in two distinct membrane domains: two Rho activators, RhoGEF64C and RhoGEF2, are apically localised, whereas a Rho inhibitor, the RhoGAP Cv-c, occupies the complementary, basolateral domain. As we have shown, cell shape changes and inward cell movements driving invagination are impaired if Rho1 becomes activated in a spatially unrestricted manner. These observations stress the importance of finely tuning Rho1 localisation and activation during normal tissue morphogenesis.
Several mechanisms might be at work to achieve the specific localisation of
the Rho regulators that direct cell invagination. In the case of RhoGEF64C we
show that its mRNA and protein are apically localised, suggesting that apical
transport of RhoGEF mRNA followed by local translation is a mechanism to
activate Rho1 in a spatially restricted manner. Recent studies show that the
mRNA of RhoA can also be transported and locally translated in the axons and
growth cones of embryonic rat neurons, where RhoA controls growth cone
collapse in response to Semaphorin 3A (Wu
et al., 2005). This shows that intracellular mRNA transport of Rho
GTPases and of their regulators may be an important mechanism to control
spatial GTPase activation.
Loss of function of the RhoGEFs involved in spiracle invagination leads to variable apical defects, which are compatible with a partial loss of Rho1 function: knocking out RhoGEF64C resulted in a mild disruption of cortical Actin without blocking invagination, while the absence of RhoGEF2 could result in a complete failure of the invagination process. These results suggest that several RhoGEFs are required to properly activate Rho1 during spiracle cell movement and organ shaping.
One interesting observation from our studies is the fact that mutants for
the RhoGAP Cv-c did not show ectopic activated Rho1 on the basolateral
membrane where this RhoGAP was localised. Thus, several mechanisms must be at
work to ensure that Rho1 activity is excluded from the basolateral domain
during cell invagination: the presence of at least one RhoGAP on the basal
membrane, the apical restriction of RhoGEFs and the existence of low levels of
Rho1 protein on the basolateral side of the cells. In addition, we also
observed that spiracles from severe cv-c mutants showed lower levels
of apical Rho1-GTP than their wild-type counterparts, correlating with the
disruption of their apical Actin. Defects in apical Actin/Myosin II were also
reported during invagination of the tracheal pits in cv-c mutants
(Brodu and Casanova, 2006).
Taken together, these observations suggest that GTP hydrolysis is a necessary
step in the regulation of Rho1 function during cell invagination and the
RhoGAP Cv-c may help to maintain a steady state level of apical Rho1-GTP.
Based on the differential distribution of Rho1 GEFs and GAPs, we propose a model in which Rho1 must shuttle back and forth between two membrane compartments, being GTP-bound on the apical cell membrane and GDP-bound on the basolateral side (Fig. 7). Thus, during tissue morphogenesis, epithelial cells can couple their apical-basal polarity to the spatial control of small RhoGTPase function.
RhoGTPases act as dynamic switches in many developmental and cellular contexts. In order to understand how they orchestrate these dynamic processes, their activity states needs to be visualised over time. We anticipate that this work and the tools described will provide a basis for studying Rho1 activity in vivo. It will be interesting to extend this analysis to other contexts in which Rho GTPases are known to act - such a dorsal closure, neurulation, wound healing - and to identify the Rho regulators involved in each case, relating their spatial/temporal distribution with the patterns of Rho GTPase activity.
Supplementary material
Supplementary material for this article is available at
http://dev.biologists.org/cgi/content/full/133/21/4257/DC1
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ACKNOWLEDGMENTS |
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