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First published online 10 July 2006
doi: 10.1242/dev.02467
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1 The Gurdon Institute and Department of Physiology, Development and
Neuroscience, University of Cambridge, Tennis Court Road, Cambridge CB2 1QN,
UK.
2 Centre for the Molecular Genetics of Development, Research School of
Biological Sciences, Australian National University, Canberra ACT 0200,
Australia.
* Author for correspondence (e-mail: ahb{at}mole.bio.cam.ac.uk)
Accepted 30 May 2006
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SUMMARY |
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 TOP SUMMARY INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Key words: Drosophila, Fes, Fer, Tyrosine kinase, Src, Adherens junction, Dorsal closure, ß-catenin
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INTRODUCTION |
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TOPSUMMARYINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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).
Mammalian Fer has been implicated in the regulation of adherens junctions
(AJs) in tissue culture (Kim and Wong,
1995; Piedra et al.,
2003; Rosato et al.,
1998; Xu et al.,
2004). Adherens junctions (AJ) are responsible not only for
adhesion between neighbouring cells, but also for providing a link between the
plasma membrane and the actin cytoskeleton. AJs are rich in phosphotyrosine
residues, and several tyrosine kinases and phosphatases are known to regulate
their assembly, stability and function
(Daniel and Reynolds, 1997;
Muller et al., 1999;
Taddei et al., 2002).
Cell adhesion and cytoskeletal reorganisation are the driving force behind
many morphogenetic movements of embryonic development. For example, during
dorsal closure in the Drosophila embryo, the epidermal sheets on each
side of the embryo extend dorsally to meet and fuse at the dorsal midline,
thereby enclosing the amnioserosa and yolk sac (for reviews, see
Jacinto et al., 2002;
Noselli and Agnes, 1999). Cell
adhesion in the dorsal-most row of cells, the leading edge cells, is regulated
in part through the modification of adherens junctions.
Intercellular adhesion is mediated by the transmembrane protein E-Cadherin,
a homophilic adhesion molecule that binds to E-Cadherin on neighbouring cells.
ß-catenin binds to the cytoplasmic tail of E-Cadherin. ß-catenin can
also interact with the F-actinbinding protein 
-catenin (reviewed by
Perez-Moreno et al., 2003).
-catenin acts as a molecular switch regulating F-actin assembly
(Drees et al., 2005), binding
alternately to ß-catenin and to F-actin
(Yamada et al., 2005).
p120-catenin (p120ctn) binds to the juxtamembrane domain (JMD) of E-Cadherin
(Daniel and Reynolds, 1995),
and is thought to act as a regulator of adherens junction assembly and
disassembly.
Both ß-catenin and p120-catenin are regulated by phosphorylation. Fer
binds constitutively to p120-catenin (Kim
and Wong, 1995; Lilien et al.,
1999) and can phosphorylate it in vitro
(Kim and Wong, 1995).
Phosphorylation of p120-catenin increases its affinity for Cadherin
(Roura et al., 1999). When
overexpressed or activated, Fer can also phosphorylate ß-catenin and
thereby disrupt adhesion (Piedra et al.,
2003; Rosato et al.,
1998): phosphorylation on Tyr654 disrupts its association with
E-Cadherin (Roura et al.,
1999); phosphorylation on Tyr142 blocks interaction with
-catenin (Piedra et al.,
2003). In C. elegans, the Fer orthologue associates with
ß-catenin in vivo and depends upon ß-catenin for localisation to
cell-cell junctions (Putzke et al.,
2005). Fer can also stabilise the cadherin complex by
phosphorylating and activating the phosphatase PTP1B, which in turn keeps
ß-catenin (Tyr654) dephosphorylated
(Xu et al., 2004). Thus Fer
has the capacity to both positively and negatively regulate cadherin complex
stability.
Several of the putative substrates of Fer are also substrates of the Src
family of cytoplasmic tyrosine kinases (SFKs). Both p120ctn and Cortactin were
initially identified as being substrates of Src
(Kanner et al., 1991;
Wu et al., 1991). Like Fer,
the SFK Fyn binds constitutively to p120-catenin, and can phosphorylate
ß-catenin at Tyr142 (Piedra et al.,
2003). Src is able to phosphorylate ß-catenin at Tyr654
(Roura et al., 1999). Given
the overlap in substrate specificity, Fer and SFKs may play overlapping or
redundant roles in the regulation of cell adhesion and motility. Functional
redundancy between different SFKs has been demonstrated in mammals and in
insects. In mice, double mutations in SFKs lead to overt phenotypes, where
single mutations do not (Stein et al.,
1994). In Drosophila, members of the Src kinase family
cooperate to regulate JUN kinase (JNK) activity
(Takahashi et al., 2005;
Tateno et al., 2000): double
mutations in Src42A;tec29, and Src42A;Src64 give a dorsal
open phenotype, whereas single mutations do not.
Only a single member of the Fes/Fer family is found in Drosophila,
DFer (Fps85D -FlyBase). The canonical form of DFer, p92dfer, was identified by
similarity to other family members (Fig.
1B) (Katzen et al.,
1991), and shares equal homology with Fes and Fer. Subsequently, a
second cDNA encoding a short isoform, p45dfer, was discovered
(Paulson et al., 1997)
(Fig. 1B). Paulson et al.
showed that DFer can transform vertebrate cells
(Paulson et al., 1997),
suggesting that the molecular pathways through which Fer signals are likely to
be conserved.
Here, we show that DFer acts in conjunction with Src42A in the process of dorsal closure. dfer mRNA is specifically upregulated in the leading edge cells of the dorsal epidermis. DFer localises to adherens junctions and is required for the formation of the F-actin cable in leading edge cells, and for normal rates of dorsal closure. When mutations in dfer are combined with a mutation in Src42A, dorsal closure fails completely. We have isolated a gain-of-function dfer mutant (dfergof) that blocks dorsal closure and causes axon misrouting. The dfergof mutant expresses an N-terminally fused form of DFer, similar to oncogenic forms of Fer. ß-catenin phosphorylation levels are reduced in dfer loss-of-function mutants and increased in dfergof mutants. This supports a role for DFer in the regulation of AJs and cell-cell adhesion, and may begin to explain its role in oncogenesis.
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MATERIALS AND METHODS |
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TOPSUMMARYINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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), UAS-puc
(Martin-Blanco et al., 1998),
UAS-GFP-Actin (Verkhusha et al.,
1999), da-GAL4 (Wodarz et al.,
1995), Src42Amyri
(Tateno et al., 2000),
pucE69 lacZ
(Martin-Blanco et al., 1998).
The following stocks were generated in this study:
UAS-DFerRB;dfer
1/TM3,hb-lacZ,
daGAL4,dfer1/TM3,GFP,
Src42Amyri/CyO,GFP;dferex1,
Src42Amyri/CyO,GFP;dfer1/TM3,GFP,
UAS-puc,dfergof/TM3,GFP,
daGAL4,dfergof/TM3,GFP,
UAS-DFerRB;+;dfergof/TM3,GFP and
UAS-GFP-Actin;dfergof/TM3,GFP. Transgenic flies containing
UAS-DFerRB were generated as previously described
(Brand and Perrimon, 1993),
except that DNA was prepared using a Qiagen Midiprep Kit.
To generate DFer mutants by male recombination, w/Y;+;pGawBMZ465
homozygous males were mated en masse to yw hop6;+;ru st e ca/TM6B virgins at
27°C. Mosaic eyed yw hop6/Y;+;ru st e ca/pGawB F1 males were then crossed
to w+;+;mwh th st ri roe pp cu sr es virgins
en masse and the progeny screened for recombinant scarlet males. In a typical
crossing scheme, around 10,000 males were scored, to isolate 30 scarlet
recombinants. Recombinants were then crossed to yw;+;D gl/TM3 p[Kr-GAL4,
UAS-GFP] (Casso et al., 1999)
(hereafter called TM3,GFP) virgins to generate a stock.
dfergof was generated by imprecise excision. All fly
crosses and egg collections were performed at 25°C unless otherwise
stated.
Molecular biology
The 3.3 kb full-length dfer cDNA (pBS-p92)
(Katzen et al., 1991) was
digested with EcoRI and XbaI, and cloned into pUAST
(Brand and Perrimon, 1993) to
create pUAS-DFerRB.
In MZ465, pGawB is inserted between bases 156733 and 156734 in GenBank
genomic sequence AE003682 (TACTCGAAAC<pGawB>-ACTCGGGCCG); in
dferex1 mutants, bases
155504-156733 (ATTTGCTAAT...TACTCGAAAC) are deleted; in
dfer1 mutants, bases
102998-156733 (ATTTCACAAT...TACTCGAAAC) are deleted.
Analysis of isoforms DFerRA and DFerRC is based on Gadfly Release 4.2
annotations (Celniker and al.,
2002; Stapleton et al.,
2002) and independent sequencing of cDNA clones RH14840 and
AT17877. The existence of isoform DFerRD/p45dfer was originally proposed based
on a cDNA, 9C13 (Paulson et al.,
1997). However, comparison of 9C13 with the genomic sequence and
the full-length cDNA for DFerRA (RH14840) reveals several frameshift mutations
which, when removed, reveal an open reading frame upstream of the predicted
translational start. This, combined with a lack of 5' ESTs aligned with
the proposed transcriptional start site for DFerRD, and the fact that DFerRD
is not apparent on northern or western blots
(Paulson et al., 1997),
suggests that 9C13 is a partial cDNA of the long isoform DFerRA.
For RT-PCR, total RNA was prepared from dfergof
homozygous mutant or wild-type embryos. First strand cDNA libraries were
generated using the SMARTTM RACE Kit (Clontech), and analysed using
5' and 3' RACE with forward and reverse gene-specific primers.
RT-PCR analysis of dferex1
flies used the QIAshredder and RNeasy kits (QIAGEN) and SuperScript III RNase
H-Reverse Transcriptase (Invitrogen). Mutants were mapped using direct PCR,
inverse PCR and plasmid rescue on adult fly genomic DNA, and direct PCR on
single embryo DNA using standard methods.
Immunohistochemistry
To generate DFer antibody, the N terminus of DFer was amplified using
5'-GCGCTCGAGATATGGGCTTCTCATCAGC-3' and
5'-GCGGAATTCCTGGCGGCATAGGTCATCCTT-3' primers, and subcloned into
the XhoI and EcoRI sites of the pRSETC vector (Invitrogen).
DFer antibody (Eurogentec) was pre-adsorbed for several days at a dilution of
1:10 on dferex1 homozygous
mutant embryos and used at a final concentration of 1:100. Antibodies used in
this study were: MAb phospho-Tyrosine (Cell Signalling; 1:100); MAb 1D4
anti-Fasciclin 2 (1:10), MAb 2D5 anti-Fasciclin 3
(Patel et al., 1987) (1:2),
MAb 40-1a anti-ß-galactosidase (Developmental Studies Hybridoma Bank,
DHSB; 1:200), Rat anti-DECad, DCAD2 (Oda
et al., 1994) (1:50), Mouse anti-arm (N2 7A1; DHSB; 1:10) and
Rabbit anti-odd-skipped (Spana and Doe,
1996) (1:1000). F-actin in embryos was labelled with
Alexa568-Phalloidan as described previously
(Kaltschmidt et al.,
2002).
Immunoprecipitation and western blotting
Five hundred stage 14 to 17 embryos were homogenised in 50 mM HEPES pH 7.6,
1 mM MgCl2, 1 mM EGTA, 50 mM KCl, Roche complete EDTA-free protease
inhibitor cocktail, Sigma phosphatase inhibitor cocktail I and Sigma
phosphatase inhibitor cocktail II. Extracts were incubated for two hours at
4°C with Protein A beads coupled to anti-arm (N2 7A1). Immunoprecipitates
were spun down and washed five times with the above buffer containing 0.01%
Tween. The final immunoprecipitates were boiled in SDS-PAGE sample buffer and
equal loadings were separated on a 4-15% SDS-PAGE BIO-RAD Ready gel, then
transferred to PVDF membrane. Membranes were probed with mouse anti-arm (N2
7A1; 1:10) and mouse anti phospho-tyrosine (Cell Signalling; 1:10,000). For
detection, we used horseradish peroxidase (HRP)-conjugated secondary
antibodies and the ECL plus western blotting detection system (Amersham
Biosciences), or, for quantification, Alexa680-conjugated secondary antibodies
(Molecular Probes) and a LI-COR Odyssey Infrared Imager.
Whole embryo extracts for western blotting were prepared as for immunoprecipitation. Equal numbers of embryos were loaded onto 4-15%SDS-PAGE Ready gels, transferred as above and probed with anti-arm (N27A1; 1:10) or anti-DFer (1:2000). Loadings were confirmed by western blotting with anti-actin [Sigma (20-33); 1:1000].
Embryonic staging
To document the reduced rates of closure, we selected embryos at late stage
16. This was defined as the stage when the gut has undergone its two major
constrictions, but prior to the successive lateral movements that disrupt the
symmetry of the three gut sections. To ensure that
dfer1 mutants did not reach
this morphology prematurely, we selected dechorionated embryos that were
midway through germ band retraction (stage 12), and allowed them to develop to
late stage 16 on apple juice agar plates. In all cases,
dfer1 mutant embryos
(n=9) developed characteristic gut morphologies at similar rates to
control embryos (n=28).
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RESULTS |
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TOPSUMMARYINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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), and has
the canonical structure of Fes/Fer family kinases, consisting of an FCH motif
at the N terminus and three coiled-coil regions, an SH2 domain and a kinase
domain (Fig. 1C). The longest
isoform, DFerRA, shares the promoter and exon structure of DFerRB, but
includes an additional exon, which encodes a novel protein domain that lies
between the canonical N-terminal domain and the SH2 domain
(Fig. 1C). A short isoform,
DFerRC, transcribed off a second promoter, encodes a truncated form of DFerRB,
lacking most of the N-terminal domain but including the third coiled-coil
domain (Fig. 1C). A fourth
isoform, DFerRD/p45dfer, has been described
(Paulson et al., 1997), which
shares the SH2 and kinase domain, but has a distinct 10 residue N-terminal
domain (Fig. 1C). Recent genome
and EST sequence data, however, raise some doubt over the validity of this
isoform (see Materials and methods for details).
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), dfer expression is highly dynamic in a
wide range of tissues, including the epidermis
(Fig. 2A), central nervous
system (Fig. 2B-E) and
developing trachea (data not shown). In the epidermis, dfer is
strongly expressed in the leading edge cells
(Fig. 2A, arrowheads),
beginning at late stage 13 and continuing throughout dorsal closure to the end
of stage 15. In the CNS, dfer is expressed in the midline glia at
stage 13 (Fig. 2B), and later
in a segmentally repeated subset of cells
(Fig. 2C) and in the dMP2
neurons (Fig. 2D,E). To
investigate the distribution and localisation of DFer protein, we raised an
antibody against an N-terminal fragment of DFer that excludes the SH2 and
kinase domains. This fragment overlaps the predicted protein for the
intermediate isoform DFerRC by 30 residues. The antibody recognises ectopic
expression of DFer-RB using the GAL4/UAS system
(Brand and Perrimon, 1993), and
staining is lost in the dferex1
mutant (Fig. 7C), which
confirms the specificity of the antibody.
DFer protein is ubiquitously expressed at relatively uniform levels
throughout embryogenesis. Prior to dorsal closure, during stages 9-10, DFer is
predominantly localised to the cytoplasm, although some weak staining is seen
around the perimeter of epidermal cells (data not shown). During stages 11-12,
DFer becomes localised to cell-cell junctions in the epidermis
(Fig. 3A). DFer becomes
polarised in the leading edge cells as dorsal closure proceeds, as is seen for
several other cell-cell junction proteins [e.g. Canoe
(Kaltschmidt et al., 2002)].
Although initially present around the entire circumference of the cell, DFer
is lost from the border with the amnioserosa during stages 13-14
(Fig. 3B, arrow). DFer protein
is also apically enriched in other epithelial sheets such as the gut and
trachea (data not shown), and is expressed in a subset of cells within the CNS
(Fig. 7A, arrowheads).
DFer localises to the adherens junction in the dorsal epidermis
In vertebrates, Fer associates with the adherens junction components
p120-catenin and ß-catenin (Kim and
Wong, 1995; Rosato et al.,
1998). To test whether DFer also localises to adherens junctions,
we co-stained embryos for DFer and DE-Cadherin (Drosophila
E-Cadherin). In the dorsal epidermis (Fig.
3C-H), DFer extensively colocalises with DE-Cadherin, although the
distribution is not identical. At the leading edge, DE-Cadherin can be seen
around the apical circumference at stage 14, whereas at this stage DFer is
lost from the leading edge itself (Fig.
3C-E). In the amnioserosa, DFer is only occasionally detected at
cell-cell junctions, where again it colocalises with DE-Cadherin (data not
shown). At the leading edge, DFer is apical to the more basal septate junction
proteins, Fasciclin 3 (Fig.
3I,J) and Discs large.
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), a
P-element has inserted upstream of the first, non-coding, exon of
dfer (Fig. 1A). We
generated mutations in dfer by imprecise excision of the P-element
and by male recombination (see Materials and methods), and screened for loss
of DFer protein expression (e.g. Fig.
7C). Nine recombinant lines were generated. In one line,
dferex1, the deletion removes
only the promoter and first exon of the dfer gene
(Fig. 1A). In the other eight
lines, a region of over 50 kb is deleted, encompassing the entire
dfer locus and the four genes proximal to dfer (CG8129, a
threonine dehydratase; CG18473, a phosphotriesterase; CG33936, a zinc finger
protein; and CG33937, GadFly release 4.2;
Fig. 1A). One of these lines,
Df(3R)dfer1 (hereafter
dfer1), was selected for
further analysis. Both dfer1
and dferex1 still express
functional GAL4 in patterns similar to that of MZ465.
dfer1 homozygotes are embryonic
lethal in 53% of cases (n=180) with the remainder dying during larval
and pupal development. dferex1
homozygotes are both viable and fertile. All results in this study using
dferex1 are from embryos
derived from dferex1 homozygous
parents.
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1 is an RNA and protein
null for all DFer isoforms. In
dferex1 mutants, dfer
mRNA is lost from the CNS and leading edge, but tracheal expression is still
evident, as is a low level of ubiquitous staining in the epidermis. DFer
protein is not detected in the CNS (Fig.
7C), but some cell-cell junction staining is faintly visible in
the epidermis. The first, non-coding, exon and promoter are deleted in
dferex1 mutants; however, mRNA
transcripts starting at the second exon, which encodes the translational
start, are present. Western blots show that full-length DFer protein is
produced in dferex1 mutants
(see Fig. S1A in the supplementary material). Thus,
dferex1 is a hypomorphic
mutation in which dfer mRNA expression is lost in a subset of tissues
and DFer protein levels in the dorsal epidermis are reduced.
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) (Fig. 4G,
arrowhead). As closure continues, leading edge cells extend filopodial and
lamellipodial processes (Fig.
4D, down arrowheads) that zip up the epidermal sheets that meet at
the dorsal midline (reviewed by Jacinto et
al., 2002).
In dfer1 mutants, the
actomyosin cable is reduced and the leading edge maintains an irregular
profile during closure (Fig.
4E, up arrowhead). P-Tyr levels at the leading edge are also
decreased, consistent with the loss of a cytoplasmic tyrosine kinase
(Fig. 4H). These morphological
differences are accompanied by a slower rate of closure. In wild-type embryos,
dorsal closure is complete by the end of stage 15.
dfer1 mutant embryos are still
open dorsally three hours later, at the end of stage 16 (90% n=10,
Fig. 4B; see Materials and
methods for staging details). Closure eventually completes, although 2%
(n=180) of cuticles from
dfer1 mutants exhibit an
anterior hole (Table 1). By
contrast, dferex1 mutants
appear to close normally and exhibit normal leading edge morphology, F-actin
and P-Tyr staining.
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1 mutants are due to loss
of DFer, and not to the four other genes within the deletion, we used the
GAL4/UAS system to express the canonical form of DFer, DFerRB, ubiquitously
throughout the embryo in the
dfer1 background. In these
embryos, the morphology of leading edge cells is largely restored to normal
(Fig. 4F,I) and closure is
completed at rates similar to wild type. All late stage 16 embryos are closed
(n=10, Fig. 4C). Our
results show that DFer is required for the normal morphology of leading edge
cells and for normal rates of closure.
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) and the
Arp2/3 activator Cortactin (Kim and Wong,
1998; Wu and Parsons,
1993), and play similar roles in regulating cell-cell adhesion
(Frame et al., 2002;
Matsuyoshi et al., 1992;
Rosato et al., 1998). Src
family kinases are functionally redundant in several developmental processes,
including dorsal closure (Takahashi et
al., 2005; Tateno et al.,
2000): single mutations in Src42A, tec29A and
Src64C do not exhibit dorsal holes, whereas double mutants, such as
tec29A,Src42A, do (Tateno et al.,
2000). Therefore, we tested whether Src42A and DFer act together
in dorsal closure.
Src42A single mutants do not exhibit dorsal holes, but show
defects in mouthpart formation (Tateno et
al., 2000) and epithelial organisation following closure
(Takahashi et al., 2005). We
find that Src42A embryos also exhibit defects in leading edge cells
that are similar to, but less severe than,
dfer1 mutants: the actomyosin
cable is disrupted (compare Fig.
5D, arrowhead, with Fig.
4D), P-Tyr staining is weaker than in wild type (compare
Fig. 5G, arrowhead, with
Fig. 4G), and dorsal closure is
slightly defective. Eight percent (n=25) of embryos show a very small
dorsal hole at late stage 16 when analysed by confocal microscopy, and the
remainder show an irregular arrangement of epidermal cells that is reflected
later in the arrangement of dorsal hairs
(Fig. 5A). Embryonic lethality
is 63% (n=191), with 60% of the unhatched embryos showing malformed
mouthparts and a small anterior hole (Fig.
5M).
When the Src42A and
dferex1 mutants are combined,
leading edge cells have a more irregular profile
(Fig. 5E, arrowhead) and P-Tyr
staining is weaker (Fig. 5H,
arrowhead). When analysed by confocal microscopy, most embryos are still
undergoing dorsal closure by late stage 16 (85%, n=34;
Fig. 5B). Embryonic lethality
is 100% (n=152), and embryos have breaks and irregularities in the
dorsal hair pattern, and a small anterior hole near the mouthparts (95%,
n=152; Fig. 5K,
arrowhead). In the remaining embryos, dorsal closure fails completely, leaving
a large anterior hole (5%, n=152;
Fig. 5N, arrowhead). When the
dfer deficiency,
dfer1, and Src42A are
combined, these defects are further enhanced. The leading edge becomes highly
irregular with a complete loss of the F-actin cable
(Fig. 5F, arrowhead) and a
substantial reduction in P-Tyr staining
(Fig. 5I, arrowhead). When
analysed by confocal microscopy, embryos exhibit a large dorsal hole at late
stage 16 (n=5; Fig.
5C). Cuticle preparations from these embryos show a large anterior
hole (30%, n=102; Fig.
5O) or a small anterior hole (59%, n=102;
Fig. 5L, down arrowhead), often
with small scabs along the dorsal midline
(Fig. 5L, up arrowhead). The
remaining embryos do not secrete a cuticle (11%, n=102). Therefore,
when either DFer or Src42A expression is reduced, leading edge cell morphology
is compromised and closure is delayed, but when both are removed dorsal
closure fails completely.
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; Snyder and Theilen,
1969). In v-fes and v-fps, a fragment of the
viral GAG protein is fused to the N terminus of the endogenous protein. The N
terminus of Fes/Fer is implicated in the regulation of autophosphorylation
(Orlovsky et al., 2000), and
N-terminal fusions result in a constitutively active kinase. Activated kinases
have also been created by the introduction of an N-terminal myristoylation
sequence (Greer et al., 1994),
and by point mutations in the first coiled-coil domain
(Cheng et al., 2001). These are
thought to disrupt intramolecular autoregulatory interactions. We isolated a third dfer mutant, dfergof, which behaves as a gain-of-function mutant. Homozygous dfergof mutants are embryonic lethal and exhibit a number of embryonic defects, including a large dorsal hole (Fig. 6B) and an aberrant midline crossing of axons in the CNS (Fig. 7E, arrowhead). DFer protein is expressed at higher levels than in wild-type embryos (Fig. 7A,B), and when the levels of DFerRB are further increased in dfergof mutants, the midline-crossing defect is enhanced (Fig. 7F). By contrast, expression of DFerRB in a wild-type background has no effect on dorsal closure or CNS development. dfergof mutants express an N-terminally modified form of DFerRB, similar to the activated forms of Fes/Fer kinases, such as v-fps (see below). This appears as an extra band, slightly larger than the canonical DFer isoform (see Fig. S1B in the supplementary material).
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The amnioserosa is also affected in dfergof mutant embryos. In wild-type embryos, F-actin staining becomes increasingly strong at the perimeter of amnioserosal cells as they progressively contract (Fig. 6C, double-down arrowhead). In dfergof mutants, F-actin staining is much less concentrated at cell-cell junctions, and the cell cortices are irregular (Fig. 6D, double arrowhead). Accelerated contraction of isolated amnioserosal cells still occurs (Fig. 6D, arrow).
To characterise further dfergof mutants, we expressed
GFP-actin (Verkhusha et al.,
1999) ubiquitously in wild-type and dfergof
backgrounds. The leading edge actomyosin cable and filopodia are reduced in
dfergof mutants, and less GFP-Actin is concentrated at the
cell-cell junctions of amnioserosal cells. In addition, the amnioserosal cells
exhibit more lamellipodia (Fig.
6F).
In dfergof mutants the P-element, pGawB, has undergone
a rearrangement, duplicating and inverting the GAL4 gene, deleting
pBluescript, and all but the promoter and first exon of the mini-white gene
(Fig. 1D). As predicted from
this map, dfergof still expresses GAL4. In fact, GAL4 is
expressed at higher levels and in more tissues, such as the epidermis and the
amnioserosa, than in the original starting line, GAL4MZ465 (data
not shown). We detect three new fusion transcripts in which the first exon of
the mini-white gene is spliced to the beginning of the second exon of
dfer, (wex1-DFerRB), to the beginning of the third exon (wex1_stop1),
or to an alternate splice acceptor in intron2 (wex1_stop2)
(Fig. 1E). The second and third
of these transcripts encode short proteins comprising the first 24 residues of
White, followed shortly thereafter by stop codons. The first transcript
encodes a predicted fusion protein in which the first 24 residues of White
(MGQEDQELLIRGGSKHPSAEHLNN) are followed by 12 novel amino acids (RAATQIGSNESI)
and the entire DFerRB protein. This chimaeric protein is strikingly similar to
the oncogenic forms of Fes, such as the Fujinami sarcoma virus protein
GAG-Fps, in which part of the retroviral GAG sequence is fused to the N
terminus of the entire Fps gene (Shibuya
et al., 1980) (see also Discussion).
DFer cooperates with Src42A to activate dpp expression in leading edge cells
In wild-type embryos, dorsal closure is initiated by activation of the JNK
pathway in leading edge cells, resulting in the transcription of two JNK
pathway targets, decapentaplegic (dpp), a TGF-ß
homologue, and puckered (puc), a dual specificity
phosphatase (Glise and Noselli,
1997; Hou et al.,
1997; Martin-Blanco et al.,
1998; Riesgo-Escovar and
Hafen, 1997). Dpp signals to the neighbouring epidermal cells,
causing them to elongate dorsoventrally. Puc inhibits Jun kinase, initiating a
negative-feedback loop. Given that both Src and DFer function in dorsal
closure, and that Src is an upstream regulator of the JNK pathway, we tested
whether DFer also regulates the JNK pathway.
We first assayed whether the activity of the JNK pathway in leading edge
cells is altered in dfer loss-of-function mutants. In
dfer1 and
dferex1 mutants, dpp
expression levels appear normal. In Src42A mutants, dpp
expression is slightly reduced, becoming patchy from stage 13 onwards
(Fig. 8B). In
Src42A;dfer1 double
mutants, dpp expression in the leading edge is reduced further and is
almost abolished by stage 13 (Fig.
8C). This suggests that DFer facilitates Src42A-mediated JNK
signalling. However, neither DFerRB nor wex1-DFerRB is able to induce ectopic
expression of dpp, suggesting that DFer is not itself sufficient to
activate the pathway. Furthermore, JNK activation is normal in leading edge
cells of dfergof mutants
(Fig. 8E,G).
dfer transcription is upregulated in LE cells, as are dpp
and puc. Although dfer transcription occurs at a later stage
than that of dpp and puc, we tested whether dfer
might nonetheless be a transcriptional target of the JNK pathway. In
loss-of-function mutants for the Jun kinase kinase hemipterous
(hep) (Glise et al.,
1995), dpp is lost in leading edge cells (see Fig. S2B in
the supplementary material). dfer, however, is still expressed (Fig.
S2E in the supplementary material). When a constitutively active form of Hep,
UAS-hepCA (Adachi-Yamada et al.,
1999), is expressed in the engrailed pattern, dpp is
upregulated in the posterior half of each segment, but dfer is not
(see Fig. S2C,F in the supplementary material). Therefore, dfer is
not a JNK target.
|
|
Tyrosine phosphorylation of ß-catenin is altered in dfer mutants
DFer localises to AJs where it may regulate the phosphorylation state, and
hence stability, of AJ proteins. To test whether DFer regulates the
phosphorylation of ß-catenin, we determined the extent of ß-catenin
tyrosine phosphorylation in dfer mutants. ß-catenin was
immunoprecipitated from yw,
dfer1 and
dfergof embryos, and probed with anti-armadillo
(ß-catenin) and anti-phospho-tyrosine
(Fig. 9A,B). In
dfer1 embryos, ß-catenin
tyrosine phosphorylation is reduced nearly fivefold with respect to control
embryos (Fig. 9A). Conversely,
tyrosine phosphorylation is significantly increased in
dfergof embryos (Fig.
9B). Much less ß-catenin is recovered from
dfergof embryos, suggesting that hyperphosphorylated
ß-catenin is removed from AJs and degraded. This is confirmed by
whole-mount staining of dfergof embryos, where the levels
of ß-catenin are decreased in the epidermis of stage 13 embryos
(Fig. 9C). DE-Cadherin levels
may also be slightly reduced in stage 13 dfergof embryos,
but the result is more variable. Western blots on total extracts from late
stage dfergof embryos also show a reduction in the levels
of ß-catenin (Fig.
9D).
|
DISCUSSION |
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|
TOPSUMMARYINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
|---|
;
Myster et al., 2003). We find
that in dfer1 mutants
ß-catenin phosphorylation is reduced. Conversely, ß-catenin is more
highly phosphorylated in dfergof mutants, demonstrating
that the role of Fer in the phosphorylation of ß-catenin is conserved in
Drosophila. Interestingly, the overall level of ß-catenin at
cell-cell junctions is lower in dfergof mutants,
suggesting that phosphorylated ß-catenin is lost from AJs and
subsequently degraded.
dfer mutants also exhibit a disorganised and reduced F-actin cable
at the leading edge. Formation of the F-actin cable appears to depend on
adherens junctions, as F-actin nucleation begins at the level of the AJs and
the F-actin cable is disrupted in DE-Cadherin mutants
(Takahashi et al., 2005).
Recently it has been suggested that elevated levels of cytoplasmic
-catenin near stable AJs could favour the formation of F-actin bundles
(Drees et al., 2005). DFer may
contribute to the formation of the F-actin cable by phosphorylating
ß-catenin, reducing its affinity for -catenin, and thereby
increasing the local levels of cytoplasmic -catenin (see Fig. S3 in the
supplementary material). If DFer promotes stable F-actin bundles then the
regulated loss of DFer from the leading edge at stage 14 may enable the more
motile Arp2/3 regulated F-actin filopodia to form and complete dorsal closure
by `zipping up'.
We have shown that DFer and Src42A cooperate during dorsal closure. DFer
localises to AJs and regulates ß-catenin phosphorylation. In
Drosophila, Src42A binds and phosphorylates ß-catenin, although
this may not be direct (Takahashi et al.,
2005). Consequently, the more severe phenotypes seen in the
dfer;Src42A loss-of-function mutants are most likely due to a
combined loss of phosphorylation on at least two different tyrosine residues
of ß-catenin.
dfer mRNA is upregulated in leading edge cells. This, together
with reports that vertebrate v-Fps and Fes mediate JNK pathway activation
(Li and Smithgall, 1998),
suggested that dfer might activate the JNK pathway during dorsal
closure. Although DFer itself cannot induce dpp expression, it does
play a supporting role in the maintenance of dpp levels, as revealed
in the Src42A mutant background. A similar failure in the maintenance
of dpp, as opposed to its induction, is seen in mutants of the Wnt
pathway (Morel and Arias,
2004). Given the comparable phenotypes, and the fact that
phosphorylated ß-catenin is reduced in dfer mutants, it is
possible that DFer contributes to the maintenance of dpp via the Wnt
pathway.
We isolated a novel, gain-of-function mutation, dfergof, in which a fragment of the White protein is fused to the N terminus of Dfer. This protein, Wex1-DFerRB, is analogous to oncogenic forms of Fps in which part of the viral GAG protein is fused to the N terminus of the endogenous proto-oncogene, generating an activated kinase. Although dfergof mutants express DFer at higher levels, this alone seems unlikely to account for the observed defects, as overexpression of DFerRB gives no obvious embryonic phenotype (M.J.M. and A.H.B., unpublished).
In dfergof mutants, the leading edge cells fail to elongate and the F-actin-rich filopodia are greatly reduced. The overall levels of the AJ junction components DE-Cadherin and ß-catenin are reduced, and ß-catenin is hyperphosphorylated. This suggests that AJs are disrupted in dfergof mutants. By contrast, it is interesting that the morphology of amnioserosal cells is shifted to a more motile appearance: F-actin is reduced at the cortex and there is an increase in the number of filopodia, perhaps because of a disruption of cell-cell junctions. In vertebrates, Fer has the capacity to both positively and negatively regulate cadherin-complex stability. This dual function may reflect a difference in binding partners present at AJs in different tissues.
Although loss of dfer does not appear to affect axon guidance,
dfergof mutants have a clear CNS phenotype in which axons
aberrantly cross the midline. A similar phenotype is seen with overexpression
of the abelson tyrosine kinase, which antagonises the receptor Robo
(Bashaw et al., 2000).
dfergof mutants also disrupt axon guidance in the PNS,
with some general misrouting of motor nerves and some overly large synapses
(data not shown). In vertebrates, Fer associates with N-Cadherin in elongating
neurites, where it can coordinately regulate N-Cadherin and integrin adhesion
(Arregui et al., 2000). Fer has
been shown to be concentrated in growth cones of stage 2 hippocampal neurons
and is required for neuronal polarisation and neurite development
(Lee, 2005). Similar to the
leading edge, DFer may be required at growth cones to regulate filopodial
extensions. In chick retinal cells, the phosphatase PTP1B when phosphorylated
by Fer, localises to the catenin-binding domain of N-Cadherin
(Xu et al., 2004).
Interestingly, the Drosophila homologue of PTP1B, DPTP61F, is
expressed in the CNS and binds to the axon guidance molecule Dock
(Clemens et al., 1996;
Walchli et al., 2000).
Strikingly, all of the phenotypes associated with dfergof mutants are rescued by expression of the Puckered tyrosine phosphatase. Given that JNK pathway activity appears normal in dfergof mutants, Puckered may target DFer itself, or its substrates, at least one of which we have shown to be hyperphosphorylated in dfergof mutants. We have demonstrated a role for DFer during embryonic development in the regulation of AJ stability, in the formation of the contractile leading edge during dorsal closure, and in axon guidance. It cooperates with Src42A to regulate ß-catenin phosphorylation at AJs. We isolated a gain-of-function mutant with structural similarity to oncogenic forms of vertebrate Fer. Unregulated Fer activity leads to oncogenesis, possibly through unregulated epidermal to mesenchymal transition. We have shown that activated DFer, or loss of DFer together with Src42A, disrupts AJs. This may provide a model for studying oncogenesis in the whole organism.
Supplementary material
Supplementary material for this article is available at
http://dev.biologists.org/cgi/content/full/133/16/3063/DC1
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ACKNOWLEDGMENTS |
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