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First published online February 9, 2006
doi: 10.1242/10.1242/dev.02266
1 Institute of Genetics, Biological Research Center, Hungarian Academy of
Sciences, H-6726 Szeged, Temesvári krt. 62, Hungary.
2 Brookdale Department of Molecular, Cell and Developmental Biology, Mount Sinai
School of Medicine, One Gustave L. Levy Place, New York, NY 10029, USA.
3 European Molecular Biology Laboratory, Meyerhofstrasse 1, 69117, Heidelberg,
Germany.
* Authors for correspondence (e-mail: mihaly{at}brc.hu; marek.mlodzik{at}mssm.edu)
Accepted 28 December 2005
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SUMMARY |
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 TOP SUMMARY INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Key words: Formin, DAAM, Actin cytoskeleton, Tracheal cuticle, Src kinases, Drosophila
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INTRODUCTION |
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TOPSUMMARYINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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;
Wallar and Alberts, 2003).
Recent work suggested that formins catalyze the assembly of unbranched actin
structures and nucleate actin filaments directly
(Evangelista et al., 1997;
Evangelista et al., 2002;
Pruyne et al., 2002;
Sagot et al., 2002a;
Sagot et al., 2002b).
Although all members of this highly conserved family are defined by the
presence of the formin homology domain 2 (FH2), which is both necessary and
sufficient to nucleate actin in vitro
(Pring et al., 2003;
Pruyne et al., 2002;
Sagot et al., 2002b), formins
are multi-domain proteins that contain several other conserved sequences as
well. The FH2 domain is usually flanked on the N-terminal side by a
proline-rich FH1 domain that can serve as a docking site for the
G-actin-binding protein profilin (Chang et
al., 1997; Evangelista et al.,
1997; Imamura et al.,
1997; Watanabe et al.,
1997). In addition, the FH1 domain also binds WW domains and SH3
domains, including those of the Src family
(Chan et al., 1996;
Kamei et al., 1998;
Tominaga et al., 2000;
Vallen et al., 2000). A
distinct formin subfamily, the DRF group (Diaphanous related formins) has the
ability to interact with an activated Rho GTPase through an N-terminal
GTPase-binding domain (GBD) (Evangelista et
al., 1997; Imamura et al.,
1997; Kohno et al.,
1996; Watanabe et al.,
1997). This binding alleviates the autoinhibitory interaction
between the GBD and the short C-terminal DAD domain (Diaphanous autoregulatory
domain) (Alberts, 2001).
Crystallographic analysis of the N-terminal part of mouse Dia1 provided a
structural basis of this regulation and identified several other functional
domains within this part of the protein, including a dimerization domain (DD),
(Fig. 1B)
(Otomo et al., 2005a;
Rose et al., 2005). In
addition, the crystal structure of C-terminal formin domains reveals that the
FH2 domain also forms a dimer required for actin nucleation and processive
filament capping (Otomo et al.,
2005b; Shimada et al.,
2004; Xu et al.,
2004), strongly suggesting that native formins act in dimeric
forms.
Recently, a novel formin subtype, DAAM (Dishevelled-associated activator of
morphogenesis), has been identified and implicated in planar cell polarity
signaling during Xenopus gastrulation
(Habas et al., 2001). The
polarized orientation of cells within the plane of a tissue, or planar cell
polarization (PCP), is an important aspect of cellular differentiation and is
often necessary to the formation of functional organs. Genes controlling PCP
have been extensively studied in Drosophila, revealing a crucial role
for frizzled (fz) signaling through dishevelled
(dsh) during the course of PCP establishment. The downstream
components of the Fz/PCP pathway include RhoA, Drok, the JNK cascade and
several other genes, most of which act tissue specifically
(Adler, 2002;
Mlodzik, 2002). Recent work
has demonstrated that many of the genes involved in PCP signaling are also
required for polarized morphogenetic cell movements such as convergent
extension during early vertebrate embryogenesis
(Fanto and McNeill, 2004;
Mlodzik, 2002;
Strutt, 2003;
Wallingford et al., 2002).
Habas et al. (Habas et al.,
2001) suggested that the novel FH2 protein Daam1 is required for
convergent extension is Xenopus embryos and that Daam1 might function
as a bridging factor between Dsh and RhoA. Moreover, Wnt/Fz-mediated
activation of RhoA appeared to depend on Dvl (a Dsh homolog) and Daam1
(Habas et al., 2001). However,
in contrast to this model, previous work has provided evidence that formins
act as Rho effectors downstream of the Rho GTPases (reviewed by
Wasserman, 1998).
To gain further insights into the function of this novel class of FH2
proteins, we analyzed the single Drosophila member of the DAAM family
(DAAM). Phenotypic analysis of DAAM mutants showed that it
plays either no role or possibly a redundant role in PCP establishment in
Drosophila. However, we found evidence that DAAM is involved
in the regulation of the actin cytoskeleton in several different tissues,
including the tracheal system. The Drosophila tracheal network is one
of the best characterized model systems of branching morphogenesis
(Affolter and Shilo, 2000;
Uv et al., 2003). During the
first phase of tracheal development, the primordial cells invaginate in each
embryonic hemisegment from the epidermis, migrate and undergo cell shape
changes to form the primary branches
(Affolter and Shilo, 2000;
Manning, 1993; Uv et al.,
2003). Subsequently, some tracheal branches fuse with an adjacent
branch to build up a continuous tubular network. This process is mediated by
fusion cells located at the branch tips, which recognize each other in the
adjacent hemisegments and become doughnut shaped, forming a lumen that
connects the two branches (Samakovlis et
al., 1996). Before the end of embryogenesis, tracheal cells
secrete a cuticle on their apical, luminal surface that protects the larvae
from dehydration and infection. The tracheal cuticle is distinguished from the
epidermal cuticle by the presence of cuticle ridges, often called taenidial
folds that are thought to prevent the collapse of tracheal tubes while
allowing them to expand and contract along their length.
Here, we show that DAAM is required to organize an array of parallel running actin cables beneath the apical surface of the tracheal cells that define the taenidial fold pattern of the cuticle. We found that the actin ring pattern corresponds exactly to that of the taenidial fold pattern, and we propose that the actin rings organized by DAAM define the position of taenidial fold formation. Our genetic interaction and epistasis data are consistent with a model that DAAM activity is regulated by RhoA. In addition, we show that DAAM works together with the non-receptor tyrosine kinases Src42A and Tec29 to regulate the actin cytoskeleton of the Drosophila tracheal system.
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MATERIALS AND METHODS |
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TOPSUMMARYINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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For gain-of-function and epistasis analysis, UAS-C-DAAM/TM3,Sb,Kr-GFP; Src42AE1/CyO,Act-GFP; UAS-C-DAAM/TM3,Ser,Act-GFP; Tec295610<FRT40A/CyO,Act-GFP; UAS-C-DAAM/TM3, Ser,Act-GFP; RhoA72F/CyO,Act-GFP; UAS-C-DAAM/TM3,Ser,Act-GFP stocks were established and crossed to btl-GAL4 (on II) homozygous females. In co-expression experiments btl-GAL4<UAS-FL-DAAM13.59/CyO,Act-GFP was crossed to UAS-C-DAAM/TM3,Sb,Kr-GFP. The appropriate genotypes were selected by the loss of GFP expression. For live imaging, embryos were collected from the progeny of a btl-GAL4 to UAS-actin::GFP/CyO cross.
In addition, we used the following mutant alleles: w1118, Oregon-R, w,Df(1)AD11/FM7c (BL#1098), w,Df(1)AC7/FM7c (BL#1144), Tp(1;3)sta sta1:sssta/FM3 (BL#844), and y1,l(1)EFd5/Dp(1;Y)Sz280, y2/FM6, l(1)1111 (BL#6021)
DNA techniques and transgenic constructs
Breakpoints of the excisions were mapped by standard Southern blot and PCR
analysis. For overexpression and rescue experiments the UAS-FL-DAAM construct
has been created by inserting the full-length RE67944 cDNA into the pUASp
vector, while the UAS-C-DAAM construct carries a PCR amplified C-terminal
region of the cDNA in the same pUASp vector encoding amino acids 518-1153. The
polyclonal anti-DAAM antibody was raised in rabbit injected with a bacterially
produced and purified His-tagged C-DAAM protein. After four boosts, the crude
serum was used for immunostaining.
Immunohistochemistry and RNA in situ hybridization
For RNA in situ hybridization, probe was prepared by the DIG RNA Labelling
Kit (Roche). Fixation and whole-mount embryo hybridization was carried out
according to standard protocols. For immunofluorescence staining, the
following antibodies were used: rat anti-DCAD2 1:10 (DSHB), rabbit anti-DAAM
1:4000, rat anti-Elav 1:300, mouse anti-ß-gal 1:1000 (Promega), mouse
anti-GFP 1:1000, mouse anti-2A12 1:5 (DSHB) and Rhodamine phalloidin at 1:100
(Molecular Probes). Secondary antibodies were: anti-rabbit-Alexa546, 1:500;
anti-rabbit-Cy5, 1:300; anti-mouse-Alexa488, 1:500; anti-rat-Cy5, 1:300;
anti-rat-Alexa488, 1:500; anti-mouse-Alexa488, 1:500 (all from Molecular
Probes); and anti-mouse-biotinylated IgM 1:300 (Vector Laboratories). Embryos
and larval tracheas were mounted in glycerol/PBS 4:1. Fluorescent and DIC
images were collected using Zeiss LSM 510 and Olympus FV500 confocal
microscopes. Bright-field images were collected using Zeiss Axiocam on Zeiss
Axioskop MOT2. Images were assembled in Adobe Photoshop 7.0, GIMP 2.2.6 and
ImageJ 1.32.
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RESULTS |
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TOPSUMMARYINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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).
This analysis suggested that the DAAM subfamily clearly represents a distinct
class. This is consistent with the previous finding that the DAAM subfamily
exhibits extensive sequence similarity both within and outside of their highly
conserved FH1 and FH2 domains (Habas et
al., 2001). At present, however, very little is known about the in
vivo function of the DAAM subfamily. Sequence analysis revealed the presence
of a single DAAM ortholog in the Drosophila genome corresponding to
the annotated gene CG14622 (Fig.
1A). Although the integrated Drosophila database
(FlyBase) predicts that this gene might code for several different transcript
classes, we focused our work on the predicted CG14622-RB transcript that
appears to be encoded by the RE67944 EST clone
(Fig. 1A). Sequencing of this
full-length cDNA clone indicated a protein consisting of 1153 amino acids.
Further analysis revealed that the ORF contains several conserved domains
including an FH1, FH2, GBD and a putative DAD domain characteristic of
formins. Based on the overall homology level and the phylogenetic analysis of
FH2 domains (Higgs and Peterson,
2005), the DAAM subfamily appears to be most related to the Dia
subfamily, raising the possibility that similar to the DRFs, DAAM formins are
also regulated by an autoinhibitory mechanism that can be relieved upon RhoGTP
binding. To begin the functional analysis of DAAM, we generated loss-of-function mutant alleles by using two transposon insertions, EP(1)1336 and EP(1)1542, in the immediate vicinity of the DAAM locus (Fig. 1A). Remobilization of these insertions allowed us to isolate a set of deletion alleles, two of which were mapped to the 5' region of DAAM, and five of them were mapped to the 3' region. The large 5' deletions affect only 5' UTR exons and intronic sequences (Fig. 1A), while the 3' alleles carry smaller deletions (965-2538 bp), but affect the DAAM ORF, leading to C-terminal truncations that (depending on the allele) remove up to 457 amino acids of the predicted protein (Fig. 1B). All 5' and 3' deletion alleles are lethal as hemi- or homozygotes, with the exception of DAAMEx1, the smallest C-terminal deletion allele which is semiviable and fertile (viability is 17% when compared with wild type). The lethal 5' and 3' deletion alleles do not complement each other and the deficiencies uncovering this region [Df(1)AD11, Df(1)AC7 and Df(1)sta]. They are, however, complemented by Dp(1;3)sta and Dp(1;Y)Sz280, two transpositional duplications that carry the 1F2-3 cytological region, including DAAM. All lethal alleles are viable over DaamEx1, the weakest and homozygous viable allele, with the exception of the two largest C-terminal deficiency alleles (DaamEx68 and DaamEx36). To demonstrate that the lethality associated with these alleles is solely due to the lack of DAAM, transgenic flies carrying UAS-FL-DAAM (containing the full length RE67944 cDNA clone) were generated and tested in rescue experiments. When UAS-FL-DAAM was expressed under the control of Act-Gal4 or tub-Gal4 in a DAAM mutant background lethality was rescued to adult viability, even in case of the largest deletion allele DaamEx68. Taken together, these results demonstrate that all lethal alleles represent DAAM alleles and these mutations do not affect other essential genetic elements.
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). To test whether the Drosophila DAAM is
involved in PCP signaling, we carried out a loss-of-function analysis of the
gene. We have thus induced DAAM mutant clones in the wing and
compound eye, two PCP organization model tissues. Although several alleles
were tested, none of them exhibited PCP defects in the eye or wing (not
shown). The same was observed in adult escapers of the
DAAMEx1/DAAMEx249 allelic combination. We have also tested whether the PCP gain-of-function phenotypes induced by overexpression of PCP proteins can be modified by DAAM mutant alleles. To achieve this, we have compared the effects of Fz and Dsh overexpression in the wing or eye in wild-type and DAAMEx68 heterozygous mutant backgrounds. No significant difference between the wild-type and the DAAM mutant backgrounds was observed (not shown). Thus, the loss-of-function analysis of DAAM argues that the Drosophila DAAM ortholog is either not required for PCP signaling or at best plays a redundant role during PCP establishment.
DAAM regulates the formation of taenidial folds in the cuticle of the trachea
As the DAAM alleles are lethal, it encodes a gene with essential
function(s). To identify these, we examined DAAM mutant larvae, and
noticed that these animals display severe trachea defects. These include the
collapse and flattening of the tracheal tubes, in both the main airways and
the side branches (Fig. 2B,C).
Wild-type tracheal cells secrete a cuticle on their apical (luminal) surface,
which is continuous with the epidermal cuticle. Nevertheless, it is easily
distinguished by the presence of cuticle ridges called taenidial folds that
project into the lumen (Fig.
2A,B). These ridges form annular rings or run a helical course
around the lumen of the tubes. Tracheal tubes of DAAM mutant larvae
fail to secrete such a highly ordered cuticle. Instead, although some local
order is visible, we observed short and curvy apical ridges that rarely run
perpendicular to the tube axis (Fig.
2C,D). These striking cuticle pattern defects are detected
throughout the tracheal system independent of tube types, with the exception
of the fusion cells that secrete a different type of cuticle characterized by
a dotty pattern instead of taenidial folds (see below,
Fig. 5C). To confirm that the
trachea phenotype is caused by DAAM loss of function, we expressed a
UAS-FL-DAAM construct under the control of a trachea specific
btl-Gal4 driver. This significantly rescued the trachea disruptions
(Fig. 2E), just as a general
driver (e.g. Act-Gal4, not shown).
Next, we examined the expression of DAAM and confirmed that the mRNA is expressed in the developing embryonic tracheal system, starting from stage 13 of embryogenesis onwards (Fig. 3A). Similarly, immunostaining experiments (using a polyclonal anti-DAAM serum raised against the C-terminal half of the protein; see Materials and methods for details) revealed the presence of the DAAM protein in the developing embryonic tracheal cells in stage 13 and older embryos (Fig. 3B). In DAAMEx68 embryos, the antibody failed to detect protein in the trachea (Fig. 3D). Similarly, detectable protein was absent from mutant clonal tissue in eye-antennal imaginal discs, where DAAM is expressed during later stages of development (Fig. 3E-H). This confirmed that the antibody is specific to DAAM, and that DAAMEx68 is likely to be a protein null. Consistently, the trachea phenotype of DAAMEx68 homo- or hemizygous larvae is very similar to that of DAAMEx68/Df(1)AD11 larvae (Fig. 2E).
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The embryonic tracheal cells are rather small, and thus it was difficult to study the formation of these actin rings at the subcellular level. Nevertheless, it is clear that these actin rings are not only present when cuticle secretion begins, but are also detected through the third larval instar stage. In contrast to the remarkable actin organization found in wild-type tracheal cells, in DAAMEx68 mutants this organization is completely abolished. Whereas the absence of DAAM does not appear to reduce the apical F-actin level of the tracheal cells, actin cables formed are much shorter and thinner than in wild type (Fig. 4L,M). Additionally, actin bundles often appear to be crosslinked to each other instead of running parallel (Fig. 4L). Taken together, in DAAMEx68 the actin bundles in the tracheal cells display abnormal morphology and fail to be organized into parallel running actin rings under the apical surface of the cells.
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).
Activated DAAM interferes with cuticle patterning in tracheal cells and transforms the cuticle pattern of the fusion cells
Truncated formins consisting only of the C-terminal FH1 and FH2 domains
behave as constitutively active forms in many contexts
(Evangelista et al., 1997;
Habas et al., 2001;
Tominaga et al., 2000;
Watanabe et al., 1999). To
test whether DAAM activity is regulated in a similar manner,
UAS-C-DAAM (carrying a truncated DAAM encoding the
C-terminal 637 amino acids including the FH1 and FH2 domains;
Fig. 1B) transgenic flies were
generated. Strikingly, C-DAAM expressed in the trachea (with the
btl-Gal4 driver) caused distinct phenotypic effects in the fusion
cells and the regular tracheal cells.
After the cuticle is laid down in the tracheal system, fusion cells are easily distinguished from others because these narrow, doughnut-shaped cells have a characteristic granulated cuticle pattern that is very distinct from the parallel running apical ridges (Fig. 5C). The same difference can be found at the level of actin organization, as apical actin is also concentrated into a granular pattern (Fig. 5D). Significantly, anti-DAAM staining in wild type revealed that fusion cells, unlike the normal tracheal cells, do not express DAAM (Fig. 4F,G). When C-DAAM is expressed in fusion cells it induces a dramatic change in the actin organization and cuticle pattern, causing strong actin accumulation at the apical surface. However, most of the actin is found in largely unorganized bundles (Fig. 5F), although the typical granular pattern is barely visible. In addition, such fusion cells are often wider than their wild-type counterparts, and their granular cuticle pattern is transformed towards a stripy pattern, partly resembling the taenidial folds of normal tracheal cells (Fig. 5E). However, when full-length DAAM was expressed in the fusion cells (or elsewhere in the tracheal system) we could not detect any phenotypic effects (not shown).
Several conclusions can be drawn from these data. First, C-DAAM behaves as an activated formin, consistent with its domain structure and showing that the activity of DAAM is regulated by N-terminal sequences like the DRF subfamily. Second, although actin cables polymerized by C-DAAM failed to organize into such perfect rings as found in wild-type tracheal cells, C-DAAM plays an instructive role in cuticle secretion by polymerizing actin cables that are sufficient to change the cuticle pattern in a DAAM non-expressing cell towards the pattern characteristic of DAAM-expressing cells. Third, the mere presence of DAAM is not sufficient to change the cuticle pattern of fusion cells, only the activated form can do that, suggesting that fusion cells lack the activator(s) of this formin.
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; Otomo et al.,
2005a; Otomo et al.,
2005b; Rose et al.,
2005; Takeya and Sumimoto,
2003; Xu et al.,
2004; Zigmond et al.,
2003). It has also been demonstrated that homotypic protein
interaction of the mammalian formin, Fhos, is mediated by the FH2 domain, and
an FH1-FH2 (C-DAAM equivalent) Fhos fragment is able to bind the full-length
protein (Takeya and Sumimoto,
2003). Therefore, it was possible that the
btl-Gal4/UAS-C-DAAM phenotype is, at least partly, caused by an
antimorphic effect of C-DAAM by interfering with the wild-type protein through
dimerization. In that case, it is expected that increasing wild-type DAAM
levels will suppress the btl-Gal4/UAS-C-DAAM phenotype. To test this hypothesis, we examined btl-Gal4 driven co-expression of FL-DAAM and C-DAAM. This caused a strong suppression of the btl-Gal4/UAS-C-DAAM phenotype in regular tracheal cells (Fig. 7E), but not in fusion cells (not shown). These data indicate that C-DAAM indeed behaves as an antimorph in regular tracheal cells. In fusion cells, however, the lack of suppression by expressing FL-DAAM is consistent with DAAM having no endogenous function in these cells. Because the UAS-Gal4 system usually induces a robust expression, C-DAAM is likely to be in excess when compared with endogenous DAAM in btl-Gal4/UAS-C-DAAM tracheal cells. Thus, the tracheal phenotype in the regular tracheal cells is possibly the sum of two effects: an antimorphic effect through interference with the endogenous protein and a neomorphic effect through an excess of activated C-DAAM, which increases actin polymerization rate but fails to organize actin bundles properly. An alternative hypothesis to explain that C-DAAM overexpression leads to similar phenotypic effects to the loss of DAAM, could be that the amount of F-actin produced by C-DAAM exceeds the actin organizing capacity of endogenous DAAM. As we could not detect higher levels of apical actin in these tracheal cells, we favor the first hypothesis.
The precise mechanism by which C-DAAM impairs actin organization and cuticle patterning in tracheal cells is unclear. It is remarkable that although activated DAAM is lacking most of the formin regulatory domains, it was specific and only disrupted cuticle patterning without affecting other biological processes in the trachea (Fig. 5G). By comparison, expression of a constitutively activated form of dia (DiaCA), the most closely related Drosophila formin to DAAM, destroys the normal branching pattern of the tracheal tubes (Fig. 5H). In addition, dorsal trunk fusion is often blocked and tracheal cuticle is not secreted (Fig. 5H). Thus, it appears that DiaCA has an earlier effect on trachea development than C-DAAM. Together, these results suggest that DAAM carries a highly specific trachea function that is restricted to the assembly and organization of apical actin cables, which direct cuticle patterning, with most of the functional specificity coming from the C-terminal FH1-FH2 domains.
DAAM interacts with RhoA and Src family non-receptor tyrosine kinases
To gain insights into the regulation of DAAM and to identify potential
partners that work together with DAAM to organize apical actin in tracheal
cells, we carried out a genetic interaction test with a panel of cytoskeleton
regulators known to be expressed in the Drosophila respiratory
system. We used the weak hypomorphic DAAMEx1 allele that
exhibits a mild phenotype (Fig.
6A), and thus appears to be the most appropriate for dominant
genetic interaction assays. This approach identified RhoA, and two
Src family non-receptor tyrosine kinases, Src42A and Tec29,
as strong enhancers of the DAAMEx1 phenotype
(Fig. 6B-D). To test whether
these interactors have tracheal cuticle patterning phenotypes on their own, we
examined the trachea of their homozygous mutant embryos and larvae. In
agreement with the observation that btl-Gal4 driven expression of
dominant-negative RhoA (RhoAN19) blocks lumen
formation (Lee and Kolodziej,
2002), we found that absence of RhoA arrests tracheal
development before dorsal trunk fusion takes place (not shown). Therefore, it
was not possible to assess directly the effect of RhoA single mutants
on apical actin organization and/or cuticle structure. However, although some
of the very few hatching Src42A and Tec29 mutant larvae
displayed a tracheal necrosis phenotype, others displayed a superficially
normal-looking tracheal tree, where cuticle secretion has been completed.
These larvae die as first instar, and, strikingly, exhibit an abnormal
taenidial fold pattern similar to that of DAAM mutant larvae
(Fig. 6E-H). Although the
Tec295610 and Src42AE1 tracheal
phenotypes are weaker than that of the DAAM null allele, consistent
with our previous conclusion that apical actin organization determines the
taenidial pattern, these mutations also affect apical actin bundles in the
tracheal system (Fig. 6I,J).
Thus, these results indicate that Src42A and Tec29 are
involved in taenidial fold patterning and during this process probably act
together with DAAM. Significantly, the FH1 domain is known to bind
SH3 proteins (Kamei et al.,
1998; Tominaga et al.,
2000; Vallen et al.,
2000), and the Src family kinases have been implicated in several
forming-related processes (Gasman et al.,
2003; Tominaga et al.,
2000). Additionally, we found that the RhoA, Src42A and
Tec29 mutations not only enhance the tracheal cuticle defects caused
by DAAM, but also decreased the viability of the semilethal
DAAMEx1 allele (not shown), suggesting that these proteins
may work together in other tissues as well. The observation that RhoA
is an enhancer of DAAMEx1 further supports the model that
DAAM family formins are regulated by Rho GTPases, similar to the regulation of
the DRF subfamily. These data are also consistent with the previous report
demonstrating that RhoA is a direct binding partner of human DAAM1
(Habas et al., 2001).
Surprisingly, however, DAAMEx1 does not show a genetic
interaction with dsh, the Drosophila homologue of Dvl,
another protein found to bind human DAAM1
(Habas et al., 2001). Although
this result does not exclude the possibility that DAAM activation in the
tracheal cells normally requires Dsh binding, a detailed analysis of the
trachea function of wg signaling, including dsh, failed to
reveal a tracheal cuticle defect
(Llimargas, 2000), suggesting
that Dsh is not involved in DAAM regulation in the trachea.
To further investigate the regulatory relationship between DAAM
and its genetic interactors, we examined if the btl-Gal4/UAS-C-DAAM
phenotype was sensitive to the gene dose of RhoA, Src42A and
Tec29. This epistasis analysis indicated that while RhoA did
not modify the effect of C-DAAM expression, Src42A and Tec29
dominantly suppressed the respective cuticle defects
(Fig. 7A-D). These data are
consistent with the model that RhoA is an upstream regulator of
DAAM, whereas the non-receptor tyrosine kinases act downstream of or
in parallel to it. Hence, these results further support the view that the DAAM
family formins are regulated by Rho GTPases. With respect to non-receptor
tyrosine kinases, our data support similar conclusions to those of previous
reports, demonstrating that mouse Dia1 and mouse Dia2 act upstream of Src in
the regulation of actin dynamics (Tominaga
et al., 2000), and that human DIA2C is required for Src activation
during the process of endosome regulation
(Gasman et al., 2003).
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DISCUSSION |
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TOPSUMMARYINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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The DAAM gene encodes a novel member of the formin family of proteins, involved in actin nucleation and polymerization. Consistent with this, DAAM is colocalized with apical actin in the tracheal cells, and the activated form of DAAM is able to induce actin assembly when expressed in tracheal cells (this work) and in other cell types (T.M. and J.M., unpublished). In DAAM mutant tracheal cells, apical actin is still detected, albeit at a somewhat lower level than in wild type, but the bundles formed in the mutant are much shorter and thinner than in wild type, and often appear to be crosslinked to each other. Most strikingly, global actin organization is almost completely lost, although some local order can still be detected. Remarkably, the cuticle pattern in mutant tracheal cells still follows the underlying aberrant actin pattern. Overall, in DAAM mutants, both the tracheal cuticle and the apical actin pattern resemble a mosaic of locally ordered patches that failed to be coordinated and aligned with each other and the axis of the tracheal tubes. We thus propose that the apical actin bundles play a key role in patterning the tracheal cuticle by defining the place of taenidial fold formation. Regarding the function of DAAM, our results suggest that the major role of this formin in the tracheal cells is to organize the actin filaments into parallel running actin rings or spirals instead of simply executing the well characterized formin function related to actin assembly. However, whether this is a direct effect on actin organization, and thus represents a novel formin function, needs to be further elucidated. An alternative model could be that DAAM is primarily required for actin polymerization but tightly coupled to an actin `organizing' protein. In such scenario, the polymerization activity should be a redundant requirement, whereas the link to the organizing protein would be a DAAM-specific function, thereby explaining the presence of unorganized actin bundles in DAAM mutant tracheal cells.
In the case of the main tracheal airways, which are multicellular along their periphery, it is striking that in wild type the run of the actin bundles is perfectly coordinated across cell boundaries. In addition, the run is always perpendicular to that of the tube axis. How does DAAM ensure the coordination of these two aspects of actin organization? Because the DAAM protein and the apical actin cables are both found at the level of the adherens junctions, it is possible that DAAM regulates the coordination of the actin cables at the cell boundaries through a direct interaction with junctional protein complexes. However, other explanations are also possible, and further experiments will be required to elucidate the molecular mechanism of this regulatory function. The fact that actin cables normally run perpendicular to the tube axis seems to suggest that tracheal cells are able to sense a global orientation cue and align their actin bundles accordingly. The nature and source of this cue is unknown, as is the mechanism by which DAAM is involved in the read-out of this signal. Nevertheless, it is interesting that in DAAM and btl-Gal4/UAS-C-DAAM mutant trachea, the main pattern of the cuticle phenotype is often changing from one segment to the other, suggesting that the effect of the `global' orientation cue is limited to metameric units.
Regulatory interaction between RhoA and DAAM
Sequence comparisons of FH2 proteins suggest a close phylogenetic
relationship between the DRF, FRL and DAAM subfamilies
(Higgs and Peterson, 2005).
Members of these three subfamilies have a high level of conservation in the
FH2 domain, and importantly, also in the region of the GBD and DAD domains,
suggesting that the FRL and DAAM family formins are also regulated by
autoinhibition and RhoGTPases, like the DRFs. We present further evidence in
support of this view. First, DAAM and RhoA display a strong
genetic interaction. Second, C-DAAM (an N-terminally truncated form of DAAM)
behaves like an activated form much the same way DRF family formins behave.
Third, epistasis experiments with C-DAAM and RhoA suggest that RhoA
acts upstream of DAAM. Thus, our data support the model in which DAAM, at
least in the Drosophila tracheal system, is regulated by
autoinhibition that can be relieved by RhoGTPases.
This conclusion, however, contradicts the observation that human DAAM1 is
required for Wnt/Fz/Dvl dependent RhoA activation in cultured cells and that
xDaam1 appears to mediate Wnt-11 dependent RhoA activation in Xenopus
embryos (Habas et al., 2001).
These results suggested that DAAM functions upstream of RhoA in non-canonical
Wnt/Fz-PCP signaling. An explanation for these distinct conclusions might be
related to the fact that DAAM, in contrast to xDaam1, does not appear to be
required for Fz/Dsh-PCP signaling. Hence, it is possible that the
Drosophila ortholog is regulated in the same way as the DRF formins,
while the vertebrate family members can be regulated in a different way, once
bound by Dsh/Dvl and recruited into PCP signaling complexes.
DAAM cooperates with Src42A and Tec29 to regulate the tracheal cuticle pattern
Genetic interactions with the hypomorphic DAAMEx1
allele identified two non-receptor tyrosine kinases, Src42A and
Tec29, as strong interacting partners. Although both of these kinases
play multiple roles during embryogenesis, we demonstrate that single mutants
for both affect the tracheal cuticle pattern in a similar way to
DAAM. These results suggest that DAAM and the Src family
kinases work together to regulate the actin cytoskeleton and cuticle pattern
in tracheal cells. Although it is not known whether DAAM physically binds
Src42A and/or Tec29, it has been established that the FH1 region of DRFs and
other formins can bind SH3 domains, including those of the Src family kinases
(Gasman et al., 2003;
Tominaga et al., 2000;
Uetz et al., 1996). In
agreement with our data that DAAM acts upstream of Src42A and Tec29
in tracheal cells, cytoskeleton remodeling and SRF activation mediated by
mouse Dia1 and mouse Dia2 requires Src activity
(Tominaga et al., 2000).
Moreover, a recent report suggests that RhoD and human DIA2C regulate endosome
dynamics through Src activation, proposing that Src activity is stimulated via
human DIA2C dependent recruitment to early endosomes
(Gasman et al., 2003).
Similarly, the Limb deformity protein (a formin) interacts with Src on the
perinuclear membranes of primary mouse fibroblasts
(Uetz et al., 1996). Based on
these examples, we speculate that in Drosophila tracheal cells the
RhoA/DAAM/Src module may not only be required to organize apical actin
bundles, but additionally it might represent a link to secretory vesicles and
to the regulation of exocytosis. Future studies will be required to test this
hypothesis, and to unravel the mechanisms whereby DAAM family formins and Src
family kinases contribute to cytoskeletal remodeling in the
Drosophila tracheal system and in other tissues.
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ACKNOWLEDGMENTS |
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REFERENCES |
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TOPSUMMARYINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
|---|
Adler, P. N. (2002). Planar signaling and morphogenesis in Drosophila. Dev. Cell 2, 525-535.[CrossRef][Medline]
Affolter, M. and Shilo, B. Z. (2000). Genetic control of branching morphogenesis during Drosophila tracheal development. Curr. Opin. Cell Biol. 12,731 -735.[CrossRef][Medline]
Alberts, A. S. (2001). Identification of a
carboxyl-terminal diaphanous-related formin homology protein autoregulatory
domain. J. Biol. Chem.
276,2824
-2830.
Chan, D. C., Bedford, M. T. and Leder, P. (1996). Formin binding proteins bear WWP/WW domains that bind proline-rich peptides and functionally resemble SH3 domains. EMBO J. 15,1045 -1054.[Medline]
Chang, F., Drubin, D. and Nurse, P. (1997).
cdc12p, a protein required for cytokinesis in fission yeast, is a component of
the cell division ring and interacts with profilin. J. Cell
Biol. 137,169
-182.
Evangelista, M., Blundell, K., Longtine, M. S., Chow, C. J.,
Adames, N., Pringle, J. R., Peter, M. and Boone, C. (1997).
Bni1p, a yeast formin linking cdc42p and the actin cytoskeleton during
polarized morphogenesis. Science
276,118
-122.
Evangelista, M., Pruyne, D., Amberg, D. C., Boone, C. and Bretscher, A. (2002). Formins direct Arp2/3-independent actin filament assembly to polarize cell growth in yeast. Nat. Cell Biol. 4,260 -269.[CrossRef][Medline]
Evangelista, M., Zigmond, S. and Boone, C.
(2003). Formins: signaling effectors for assembly and
polarization of actin filaments. J. Cell Sci.
116,2603
-2611.
Fanto, M. and McNeill, H. (2004). Planar
polarity from flies to vertebrates. J. Cell Sci.
117,527
-533.
Gasman, S., Kalaidzidis, Y. and Zerial, M. (2003). RhoD regulates endosome dynamics through Diaphanous-related Formin and Src tyrosine kinase. Nat. Cell Biol. 5,195 -204.[CrossRef][Medline]
Habas, R., Kato, Y. and He, X. (2001). Wnt/Frizzled activation of Rho regulates vertebrate gastrulation and requires a novel Formin homology protein Daam1. Cell 107,843 -854.[CrossRef][Medline]
Higgs, H. N. and Peterson, K. J. (2005).
Phylogenetic analysis of the formin homology 2 domain. Mol. Biol.
Cell 16,1
-13.
Imamura, H., Tanaka, K., Hihara, T., Umikawa, M., Kamei, T., Takahashi, K., Sasaki, T. and Takai, Y. (1997). Bni1p and Bnr1p: downstream targets of the Rho family small G-proteins which interact with profilin and regulate actin cytoskeleton in Saccharomyces cerevisiae. EMBO J. 16,2745 -2755.[CrossRef][Medline]
Kamei, T., Tanaka, K., Hihara, T., Umikawa, M., Imamura, H.,
Kikyo, M., Ozaki, K. and Takai, Y. (1998). Interaction of
Bnr1p with a novel Src homology 3 domain-containing Hof1p. Implication in
cytokinesis in Saccharomyces cerevisiae. J. Biol.
Chem. 273,28341
-28345.
Kohno, H., Tanaka, K., Mino, A., Umikawa, M., Imamura, H., Fujiwara, T., Fujita, Y., Hotta, K., Qadota, H., Watanabe, T. et al. (1996). Bni1p implicated in cytoskeletal control is a putative target of Rho1p small GTP binding protein in Saccharomyces cerevisiae. EMBO J. 15,6060 -6068.[Medline]
Lee, S. and Kolodziej, P. A. (2002). The plakin Short Stop and the RhoA GTPase are required for E-cadherin-dependent apical surface remodeling during tracheal tube fusion. Development 129,1509 -1520.[Medline]
Li, F. and Higgs, H. N. (2003). The mouse Formin mDia1 is a potent actin nucleation factor regulated by autoinhibition. Curr. Biol. 13,1335 -1340.[CrossRef][Medline]
Llimargas, M. (2000). Wingless and its signalling pathway have common and separable functions during tracheal development. Development 127,4407 -4417.[Abstract]
Manning, G. and Krasnow, M. A. (1993). Development of the Drosophila tracheal system. In The Development of Drosophila melanogaster, vol. I (ed. M. Bate and A. Martinez-Arias), pp. 609-686. Plainview (NY): Cold Spring Harbor Laboratory Press.
Mlodzik, M. (2002). Planar cell polarization: do the same mechanisms regulate Drosophila tissue polarity and vertebrate gastrulation? Trends Genet. 18,564 -571.[CrossRef][Medline]
Otomo, T., Otomo, C., Tomchick, D. R., Machius, M. and Rosen, M. K. (2005a). Structural basis of Rho GTPase-mediated activation of the formin mDia1. Mol. Cell 18,273 -281.[CrossRef][Medline]
Otomo, T., Tomchick, D. R., Otomo, C., Panchal, S. C., Machius, M. and Rosen, M. K. (2005b). Structural basis of actin filament nucleation and processive capping by a formin homology 2 domain. Nature 433,488 -494.[CrossRef][Medline]
Pring, M., Evangelista, M., Boone, C., Yang, C. and Zigmond, S. H. (2003). Mechanism of formin-induced nucleation of actin filaments. Biochemistry 42,486 -496.[CrossRef][Medline]
Pruyne, D., Evangelista, M., Yang, C., Bi, E., Zigmond, S.,
Bretscher, A. and Boone, C. (2002). Role of formins in actin
assembly: nucleation and barbed-end association.
Science 297,612
-615.
Rose, R., Weyand, M., Lammers, M., Ishizaki, T., Ahmadian, M. R. and Wittinghofer, A. (2005). Structural and mechanistic insights into the interaction between Rho and mammalian Dia. Nature 435,513 -518.[CrossRef][Medline]
Sagot, I., Klee, S. K. and Pellman, D. (2002a). Yeast formins regulate cell polarity by controlling the assembly of actin cables. Nat. Cell Biol. 4, 42-50.[Medline]
Sagot, I., Rodal, A. A., Moseley, J., Goode, B. L. and Pellman, D. (2002b). An actin nucleation mechanism mediated by Bni1 and profilin. Nat. Cell Biol. 4, 626-631.[Medline]
Samakovlis, C., Manning, G., Steneberg, P., Hacohen, N., Cantera, R. and Krasnow, M. A. (1996). Genetic control of epithelial tube fusion during Drosophila tracheal development. Development 122,3531 -3536.[Abstract]
Shimada, A., Nyitrai, M., Vetter, I. R., Kuhlmann, D., Bugyi, B., Narumiya, S., Geeves, M. A. and Wittinghofer, A. (2004). The core FH2 domain of diaphanous-related formins is an elongated actin binding protein that inhibits polymerization. Mol. Cell 13,511 -522.[CrossRef][Medline]
Strutt, D. (2003). Frizzled signalling and cell
polarisation in Drosophila and vertebrates.
Development 130,4501
-4513.
Takeya, R. and Sumimoto, H. (2003). Fhos, a
mammalian formin, directly binds to F-actin via a region N-terminal to the FH1
domain and forms a homotypic complex via the FH2 domain to promote actin fiber
formation. J. Cell Sci.
116,4567
-4575.
Tanaka, H., Takasu, E., Aigaki, T., Kato, K., Hayashi, S. and Nose, A. (2004). Formin3 is required for assembly of the F-actin structure that mediates tracheal fusion in Drosophila. Dev. Biol. 274,413 -425.[CrossRef][Medline]
Tominaga, T., Sahai, E., Chardin, P., McCormick, F., Courtneidge, S. A. and Alberts, A. S. (2000). Diaphanous-related formins bridge Rho GTPase and Src tyrosine kinase signaling. Mol. Cell 5,13 -25.[CrossRef][Medline]
Uetz, P., Fumagalli, S., James, D. and Zeller, R.
(1996). Molecular interaction between limb deformity proteins
(formins) and Src family kinases. J. Biol. Chem.
271,33525
-33530.
Uv, A., Cantera, R. and Samakovlis, C. (2003). Drosophila tracheal morphogenesis: intricate cellular solutions to basic plumbing problems. Trends Cell Biol. 13,301 -309.[CrossRef][Medline]
Vallen, E. A., Caviston, J. and Bi, E. (2000).
Roles of Hof1p, Bni1p, Bnr1p, and myo1p in cytokinesis in Saccharomyces
cerevisiae. Mol. Biol. Cell
11,593
-611.
Wallar, B. J. and Alberts, A. S. (2003). The formins: active scaffolds that remodel the cytoskeleton. Trends Cell Biol. 13,435 -446.[CrossRef][Medline]
Wallingford, J. B., Fraser, S. E. and Harland, R. M. (2002). Convergent extension: the molecular control of polarized cell movement during embryonic development. Dev. Cell 2, 695-706.[CrossRef][Medline]
Wasserman, S. (1998). FH proteins as cytoskeletal organizers. Trends Cell Biol. 8, 111-115.[CrossRef][Medline]
Watanabe, N., Madaule, P., Reid, T., Ishizaki, T., Watanabe, G., Kakizuka, A., Saito, Y., Nakao, K., Jockusch, B. M. and Narumiya, S. (1997). p140mDia, a mammalian homolog of Drosophila diaphanous, is a target protein for Rho small GTPase and is a ligand for profilin. EMBO J. 16,3044 -3056.[CrossRef][Medline]
Watanabe, N., Kato, T., Fujita, A., Ishizaki, T. and Narumiya, S. (1999). Cooperation between mDia1 and ROCK in Rho-induced actin reorganization. Nat. Cell Biol. 1, 136-143.[CrossRef][Medline]
Xu, Y., Moseley, J. B., Sagot i Poy, F., Pellman, D., Goode, B. L. and Eck, M. J. (2004). Crystal structures of a Formin Homology-2 domain reveal a tethered dimer architecture. Cell 116,711 -723.[CrossRef][Medline]
Zigmond, S. H., Evangelista, M., Boone, C., Yang, C., Dar, A. C., Sicheri, F., Forkey, J. and Pring, M. (2003). Formin leaky cap allows elongation in the presence of tight capping proteins. Curr. Biol. 13,1820 -1823.[CrossRef][Medline]
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