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First published online 10 July 2006
doi: 10.1242/dev.02482
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Institut de Biologia Molecular de Barcelona (IBMB), CSIC, C/Josep Samitier 1-5, 08028 Barcelona, Spain.
* Author for correspondence (e-mail: mlcbmc{at}cid.csic.es)
Accepted 7 June 2006
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SUMMARY |
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 TOP SUMMARY INTRODUCTIONMATERIALS AND METHODSRESULTSREFERENCES |
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Key words: Epithelial integrity, Egfr, Tracheal system, ERK type MAPK pathway, Mkp3, Cell adhesion, DE-cadherin, Cortical actin, Drosophila
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INTRODUCTION |
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TOPSUMMARYINTRODUCTIONMATERIALS AND METHODSRESULTSREFERENCES |
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; Pilot and Lecuit,
2005); however, the signals and mechanisms that ensure sufficient
cell adhesiveness during this process remain poorly understood. Loss of tissue
integrity occurs during epithelial-mesenchymal transition (EMT), a process by
which epithelial stationary cells acquire a migratory phenotype. EMT occurs
during normal development (e.g. gastrulation or neural tube formation), and
importantly, it is a key step in malignant transformation
(Kang and Massague, 2004;
Shook and Keller, 2003;
Thiery, 2003). Given the
importance of tissue integrity maintenance, both during normal development and
during invasive malignancies, it is crucial to understand the molecular
mechanisms and the signalling pathways that ensure it. Here, by studying
Drosophila tracheal development, we unveil a new role for the Egfr
pathway in maintaining epithelial integrity.
The tracheal (respiratory) system of the fly has become one of the most
amenable and best-documented models for tubulogenesis (reviewed by
Affolter et al., 2003;
Hogan and Kolodziej, 2002;
Lubarsky and Krasnow, 2003).
The tracheal tree consists of a network of epithelial tubules that oxygenate
the tissues. It arises from clusters of cells in the ectoderm, known as
tracheal placodes, which are specified at mid-embryogenesis and invaginate.
Subsequently, and by processes of directed cell migration, changes in cell
shape and rearrangement of the cells within the tissue, tracheal cells undergo
stereotyped processes of branching and branch fusion that generate the mature
structure by the end of embryogenesis. Convergent efforts have identified the
tracheal requirements of several genes and signalling pathways (reviewed by
Ghabrial et al., 2003).
However, the links between the genetic pathways that govern tracheal
development and the cellular responses remain elusive.
As tracheal development occurs in the absence of cell proliferation and
cell death (Samakovlis et al.,
1996), tracheal cells undergo tremendous rearrangements to give
rise to fully extended branches without compromising tube continuity. Tracheal
development is, therefore, an ideal model with which to tackle the analysis of
tissue remodelling and tissue integrity maintenance in vivo. To date, few
elements have been shown to regulate tracheal epithelial integrity. The
nuclear zinc-finger protein Hindsight (Hnt; Pebbled-FlyBase)
(Wilk et al., 2000) and a
component of the luminal extracellular matrix
(Jazwinska et al., 2003),
Piopio (Pio), are required for maintaining tracheal integrity. Other
epithelial integrity regulators impinge on the cellular junctions that attach
epithelial cells to one another and that establish their apico-basal polarity.
Mutants for Lachesin (Lac) and other septate junction (SJ)
components produce luminal breaks that result from an insufficient adhesion
between tracheal cells (Llimargas et al.,
2004; Wu and Beitel,
2004). The small GTPase Rac has been shown to regulate epithelial
integrity by modulating the assembly and disassembly of E-Cadherin (DE-cad) at
the adherens junctions (AJs) (Chihara et
al., 2003). Regulation of the activity of AJs lies at the basis of
many of the coordinated cellular changes that occur during extensive tissue
remodelling (Lecuit, 2005;
Pilot and Lecuit, 2005).
Indeed, the remodelling of AJs is required for proper tubulogenesis and
intercalation in the developing trachea
(Ribeiro et al., 2004).
Here, we unveil a new role for Egfr through the ERK-type MAPK pathway (hereafter referred to as the MAPK pathway) in maintaining epithelial integrity. We show that the downregulation of this pathway results in a loss of tissue continuity as branches extend. Conversely, upregulation of the pathway produces defects in branch extension consistent with excess stiffness. Remarkably, Egfr- promoted epithelial integrity does not require the nuclear transcription factor Pointed (Pnt), which otherwise mediates most of the Egfr pathway outcomes. In contrast to Egfr, MAPK pathway regulation by Breathless (Btl, another tyrosine kinase receptor known to trigger the pathway in the trachea) seems not to affect epithelial integrity. The defects we observe in tracheal integrity correlate with subtle differences in the accumulation of DE-cad and of an apical actin belt. Consistently, mutants impairing AJs or actin cytoskeleton assembly [shotgun (shg), encoding DE-cad, and crossveinless c (cv-c), encoding a RhoGAP involved in actin dynamics, respectively] are also required for integrity of the tissue.
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MATERIALS AND METHODS |
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TOPSUMMARYINTRODUCTIONMATERIALS AND METHODSRESULTSREFERENCES |
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88,
pnt737, Egfrf2, UAS-EgfrDN, UAS-
top,
btlLG19, btlH8211, UAS-btlDN,
UAS-ßGal, Df(3L)H99, shg2, Mkp35J4,
UAS-RasN17, UAS-RafDN, rhoP5,
UAS-tauGFP, UAS-DE-cadGFP and EP3142. The following Gal4 drivers were used:
btlGal4 to drive expression in all tracheal cells from invagination onwards,
ptcGal4, armGal4 and 69B. Mkp3M76-R2b and Mkp3M76 have
been described previously (Gomez et al.,
2005), as has cv-c7
(Denholm et al., 2005).
UAS-p35 was obtained from S. Campuzano (CBM, Madrid), btlGFP and
hsflp;btlenhancer>y+>Gal4, UAS-GFPactin; btlenhancer-mRFP-moe from M.
Affolter (Biozentrum, Basel), and GS10283 from the Drosophila Gene Search
Project (DGSP). To recognise the chromosomes carrying the desired mutations,
we used second or third blue balancers.
To induce the expression of UAS-EgfrDN or
UAS-top in small groups of tracheal cells, we
crossed the lines to hsflp;btlenhancer>y+>Gal4, UAS-GFPactin;
btlenhancer-mRFP-moe (Ribeiro et al.,
2004). Embryos were collected at 25°C for 5-6 hours, heat
shocked for 35 minutes at 36°C, and transferred to 25°C before
fixation. Raising the embryos at 29°C (with or without heat shock) induced
btlGal4 in virtually all tracheal cells. Therefore, embryos could not
be raised at 29°C after the heat shock to achieve the maximal efficiency
of the Gal4/UAS system, and therefore the strongest and more
penetrant phenotypes with the transgenes.
Molecular analysis
The GS element in line 801 was mapped by inverse PCR techniques
following standard protocols (Berkeley Drosophila Genome Project, please
contact authors for details).
Antibody stainings and in situ hybridisation
Embryos were staged according to Campos-Ortega and Hartenstein
(Campos-Ortega and Hartenstein,
1985) and stained following standard protocols. Immunostainings
were performed on embryos fixed in 4% formaldehyde for 20 to 30 minutes,
except for DCAD2 stainings, for which embryos were fixed for 10 minutes. The
following antibodies were used: anti-GFP (Molecular Probes and Roche), mAb2A12
(Developmental Studies Hybridoma Bank, DSHB), anti-Pio (from M. Affolter),
anti-Hnt (from R. Wilk, University of Toronto, Toronto), anticleaved caspase-3
(Cell Signaling Technologies), anti-DSRF (2-161 from Cold Spring Harbor
Laboratories), anti-Kni (developed by J. Reivitz and provided by M.
Ruiz-Gomez, CBM, Madrid), anti-DE-cad (DCAD2, Developmental Studies Hybridoma
Bank), anti-actin (MP Biomedicals), anti-Trh (made by N. Martín in J.
Casanova's Laboratory, IBMB, Barcelona) and anti-ß-Gal (Cappel and
Promega). Biotinylated or Cy3-, Cy2- and Cy5-conjugated secondary antibodies
(Jackson ImmunoResearch) were used at a dilution of 1/300. For HRP
histochemistry, the signal was amplified with the Vectastain-ABC kit. For
fluorescent stainings, the signal was amplified with TSA (NEN Life Sciences)
when required. Photographs were taken in a Nikon Eclipse 80i microscope.
Confocal images were obtained with a Leica TCS-SP1- or TCS-SP2-AOBS system,
Leica DM IRE2 microscope and LCS software.
Unless otherwise stated, in all panels labelled `GFP' the embryos carried btlGal4 UAS-tauGFP, and these were stained with anti-GFP to highlight the shape of tracheal cells. btlGal4 also drove the expression of the indicated UAS constructs. We used mAb2A12 or CBP to visualise the lumen.
In situ hybridisation was performed according to standard protocols with digoxigenin-labelled RNA probes prepared from the Mkp3 cDNA clone LD02618.
Western blot
Fifty embryos of the selected genotypes were ground up in 100 ml lysis
buffer [125 mM Tris (pH 6.8), 21% Glycerol, 5% SDS, 10% bromophenol blue and
ß-mercaptoethanol]. For better protein extraction, samples were boiled
five times at 91°C for 1 minute, and sonicated for 1 second. Proteins were
separated by SDS-PAGE (10% polyacrylamide) and transferred onto a
nitrocellulose membrane (Schleicher and Schuell). The membrane was blocked in
PBT-5% milk overnight at 4°C, incubated with primary antibody (anti-GFP
1/750, or anti-
-Tubulin 1/500, Sigma) and, subsequently, with
HRP-linked secondary antibody (1/10000, Amersham). Proteins were visualized by
an enhanced chemiluminescence (ECL) detection system (Amersham). The blots
were reprobed sequentially with both antibodies to obtain the specific
proteins levels of each sample. Intensities of the bands were quantified by
densitometric scanning of the film exposed to chemiluminescence using the
QuantityOne program (BioRad).
Time lapse
Embryos carrying btlGal4 UAS-tauGFP;801 were collected at 29°C
and dechorionated for 2 minutes with sodium hypochlorite diluted 1/100. They
were glued to a coverslip and mounted in 10S Voltalef oil with the hanging
drop method in an oxygen-permeable chamber. Images were collected from stage
14 at 21°C on a Leica TCS-SP2-AOBS system, Leica DM IRE2 microscope and
LCS software. The 488 nm emission line of an Argon laser was used for
excitation and sections were recorded every 6 minutes over a 3-hour period.
TIFF projection images were processed into 3D and 4D LCS software, and the
movie was assembled using ImageJ.
Analysis and quantification of DE-cad levels
For DCAD2 stainings, the mutant and control populations were collected,
fixed and immunostained together. Mutant and control sibling populations were
obtained by crossing btlGal4 UAS-tauGFP/CyO to
UAS-top, UAS-EgfrDN, or line 801
(identifying the mutant and control population by the presence or absence of
GFP), or from a
rhoP5/TM3
ftzlacZ stock (identifying the mutant and control population by the
presence or absence of anti-ß-Gal staining). We compared the levels of
DCAD2 staining between the two populations within each experiment in a high
number of embryos (n>20). We note that there was variability in
the accumulation of DCAD2 in each experiment, both within the mutant
population and within the internal control population, but the differences we
report in the Results between controls and mutants, although subtle, were
consistently and reproducibly seen. In Fig.
5, we show representative, although extreme, examples from the
different genotypes.
To quantify DE-cad levels, eight to 11 embryos of each genotype were scanned with a 63x objective zoomed 4x, using the same laser power for each experiment. Projections of 0.5 µm confocal sections of the posterior region of the tracheal tree were analysed. We quantified the pixel intensity of staining by taking four measurements along the dorsal branches (DBs) and six measurements of the contour of DT cells in each embryo with a freeline tool in ImageJ 1.25 (Rasband, http://rsb.info.nih.gov/ij). We calculated the average of the four measurements of the DBs and six measurements of the DT for each embryo, and compared the mutant and control populations. The levels for each experiment depended on the staining and the laser power used, and therefore different experiments cannot be directly compared. However, to facilitate the visualisation of the differences, the average levels of the control population of each single experiment were normalised to 100%, and the level of staining of the mutant population is expressed as the percentage of the level of the control population.
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RESULTS |
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TOPSUMMARYINTRODUCTIONMATERIALS AND METHODSRESULTSREFERENCES |
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) to generate an original collection of lines (C.C. and M.L.,
unpublished). We selected a line containing a GS element, termed 801,
on the basis of its embryonic tracheal phenotype when crossed to the
btlGal4 driver, which is expressed in all tracheal cells from
invagination onwards. Tracheal development of such embryos proceeded normally
up to stage 13. At stage 14, we detected defects in branch formation and
outgrowth, as several dorsal branches (DBs) and ganglionic branches (GBs)
never formed, or contained only one or two cells
(Fig. 1B). In addition to these defects in primary branching, we detected many branch breaks when luminal markers were used (not shown). This phenotype was highly penetrant (100% of embryos, n=98) and commonly observed in thin unicellular branches, such as the most ventral (lateral trunk posterior, LTp-GB) and dorsal branches (Fig. 3J). The branches could be broken at any position, giving rise to cells completely isolated from the rest of the tracheal tree, or to cells abnormally separated and only attached to the stalk by thin cytoplasmic extensions or bridges (Fig. 1C,D). Branch breaks increased with time and they were very conspicuous at late stages of embryogenesis, when sometimes almost no branches, not even the dorsal trunk (DT), appeared continuous (Fig. 1B,C). In vivo imaging showed that tracheal cells start to pull apart, establishing long cytoplasmic extensions that eventually can completely break (see Movie in the supplementary material). We interpret these phenotypes as resulting from a loss of tissue integrity that is likely to be needed to counteract the pulling forces driving branch extension. In addition, the tracheal cells appeared more rounded or cuboidal when compared with the wild type (Fig. 1C,D).
This phenotype was also observed when we used Gal4 drivers that induce a more general and earlier expression, such as armGal4, 69B or ptcGal4 (data not shown). We reasoned that the over- or misexpression of the gene(s) near the GS insertion caused the loss of epithelial integrity that maintains tracheal cell attachments to one another.
Overexpression of Mkp3 is responsible for the branch integrity phenotype
We mapped the GS element insertion of line 801 to 75F6. The two
closest genes are MAP Kinase Phosphatase 3 (Mkp3; positioned
21 bp 5' of the GS element) and Misexpression Suppressor of Ras
6 (MESR6; at 6.8 kb 3' of the element;
Fig. 2A). Two independent GS
lines, GS10283 [Drosophila Gene Search Project (DGSP)] and
Mkp3M76 (Gomez et al.,
2005), located near our insertion, produced the same phenotype
when crossed to btlGal4 (Fig.
2B,C), whereas the EP3142 line that drives MESR6
expression did not produce a consistent phenotype (data not shown). In
addition, a null mutation in Mkp3, Mkp3M76-R2b
(Gomez et al., 2005), reverted
the integrity defects of Mkp3M76 tracheal overexpression
(Fig. 2D). Altogether these
results point to a major contribution of over- or misexpression of
Mkp3 in the tracheal cells to the phenotype.
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) [BDGP in situ
(http://www.fruitfly.org/cgi-bin/ex/insitu.pl)].
Mkp3 accumulated in the tracheal pits at stage 11
(Fig. 2E), and we also observed
expression in tracheal branches at later stages
(Fig. 2F). As expected,
Mkp3 was overexpressed in the tracheal cells of btlGal4-801
embryos (Fig. 2G).
We studied the tracheal requirement for Mkp3. The null mutations
Mkp3M76-R2b and Mkp35J4 exhibited a
mild delay in branching, as if the loss of Mkp3 caused increased
tissue stiffness. To quantify the delay, we visualised the tip cells with a
DSRF antibody (Affolter et al.,
1994) and analysed the number of DBs between metameres 4 to 8 that
have reached the dorsal midline at stage 14-15 (not shown). We found a delay
in 12% of DBs (n=190) in Mkp35J4 mutants compared
with 0.3% in wild type (n=50). Increased stiffness may produce
defects in cell intercalation. To test this hypothesis, we analysed DBs of
late embryos stained with a Trachealess (Trh) antibody to determine whether
tracheal cells were properly positioned end-to-end, and with an AJ marker
(DCAD2 antibody) to reveal the cell intercalation state
(Ribeiro et al., 2004). We
detected defects in cell intercalation with both markers
(Fig. 3K,N) in
Mkp35J4 mutants. The lack of a stronger tracheal phenotype
could indicate that Mkp3 is not absolutely required during embryonic
stages, or that it shares redundant functions with other MAP Kinase
Phosphatases (Mkps). Alternatively, the zygotic requirement might be rescued
by the maternal contribution. Homozygous females for the null
Mkp3M76-R2b allele are sterile and only lay unfertilised
eggs, precluding us from determining the tracheal requirement of the
maternally provided protein.
The MAPK pathway plays a role in the maintenance of tracheal branch integrity, but not through the nuclear effector Pnt
Mkp3 has been shown to act as a specific negative regulator of the ERK-type
MAPK (Kim et al., 2002;
Rintelen et al., 2003). ERK is
a central element of the Ras/MAPK pathway whose activity is regulated by
phosphorylation (Roux and Blenis,
2004). Upon receptor activation, the signal is transduced through
activation of the small GTPase Ras and several serine-threonine protein
kinases. We therefore investigated whether this pathway is required to
maintain tracheal epithelial integrity. Indeed, we found that tracheal
expression of dominant-negative versions of Ras and Raf
(Fig. 3A; not shown) produced
branch integrity defects in most embryos (95%, n=20, and 100%,
n=27, respectively) and a similar expressivity
(Fig. 3J). Similarly, the
zygotic absence of ERK, encoded by rolled (rl),
gave rise to defects in branch continuity
(Fig. 3B), and to branch
patterning defects. Conversely, the constitutive activation of rl
(UAS-rlsem) in the tracheal cells produced a branching
extension delay in 27% of DBs (n=90), and incomplete or impaired cell
intercalation (Fig. 3L,O).
These defects are consistent with an excess of tissue stiffness as opposed to
a loss of tissue integrity upon MAPK pathway downregulation. These results
indicate that the MAPK pathway regulates epithelial integrity, unveiling a new
role of this pathway during tracheal development.
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The ETS protein Pnt acts as a nuclear positive effector of the MAPK pathway
in many developmental contexts (Rebay,
2002). We therefore investigated whether pnt is also
required to maintain tracheal integrity. As previously described, null or
hypomorphic mutations in pnt,
pnt88 or
pnt737, show an absence of secondary branching and
migration defects in most branches (Myat
et al., 2005; Samakovlis et
al., 1996). However, we did not detect breaks in the branches, as
we did when the MAPK pathway is downregulated
(Fig. 3D). This result
indicates that pnt is not involved in branch integrity maintenance
and suggests that the MAPK pathway regulates this aspect directly by
modulating cytoplasmic targets, or through other nuclear effectors.
The Egfr, not the btl, signalling pathway is required to maintain epithelial branch integrity
MAPK pathway is activated by tyrosine kinase receptors
(Rebay, 2002). During tracheal
development, two tyrosine kinase receptors have been shown to activate the
pathway, Egfr and Btl (reviewed by
Ghabrial et al., 2003). We
investigated whether either or both receptors are responsible for maintaining
branch integrity.
Null mutants for Egfr produced embryos with a strong morphological
phenotype, nevertheless, we could also distinguish clear branch integrity
defects (Fig. 3E).
Consistently, 96% of embryos (n=37) expressing a dominant-negative
version of the receptor, EgfrDN, displayed defects such as
branch breaks or cells abnormally separated and attached by cytoplasmic
bridges (Fig. 3G,J), as in line
801 overexpression. But in contrast to btlGal4 801 embryos,
the general pattern and outgrowth of DBs and GBs was not grossly affected in
btlGal4 UAS-EgfrDN. Accordingly, we detected a normal
pattern of kni expression (Fig.
4F) and the correct number of cells in the DBs, indicating that
primary branching proceeded normally in these embryos. Conversely, a
constitutively active form of the receptor, top, gave rise to
delayed tracheal extension in 35% of DBs (n=120;
Fig. 3M). In addition, we also
detected incomplete cell rearrangements and confirmed the lack of cell
intercalation in some DBs by the absence of autocellular AJs
(Fig. 3P,
Fig. 5E). These phenotypes are
consistent with an excess of tissue stiffness.
rhomboid (rho) encodes a peptidase involved in the
secretion of the Egf ligand (Urban et al.,
2001). Apart from other tracheal defects associated with
invagination, null
rhoP5 mutants showed
a similar phenotype to that of the overexpression of line 801
(Fig. 3F). Furthermore, the
haploinsuficiency of Egfrf2 or
rhoP5 in btlGal4
801 embryos significantly increased the branch integrity defects
(Fig. 3J; data not shown).
Altogether, the data assign a new role for Egfr signalling in
maintaining the epithelial integrity of tracheal branches through the
activation of the MAPK pathway.
In contrast to Egfr mutants, null mutants for btl
(btlLG19) did not visibly display tissue integrity defects
(not shown), suggesting that btl is not required to maintain branch
integrity. However, a btl integrity requirement could be masked by
the absence of extended branches in null mutants. Therefore, we used other
btl alleles. A dominant-negative form of the receptor
(Reichman-Fried and Shilo,
1995), UAS-btlDN, expressed in tracheal cells
did not produce branch integrity defects, although it produced mild defects in
fusion and terminal branching (Fig.
3H). Additionally, the hypomorphic mutation
btlH8211, which
allows some branch outgrowth, did not cause a branch integrity phenotype
(Fig. 3I). Moreover, halving
btl dose in btlGal4 801 embryos did not significantly affect
the penetrance of the branch integrity phenotype, but resulted in a
significant increase of branches that do not form
(Fig. 3J). These results
indicate that btl does not play an essential role in maintaining
integrity of the tracheal tissue, although it is essential for proper
branching pattern, as has been already shown
(Klambt et al., 1992). Hence,
we propose that the branching defects caused by MAPK pathway downregulation
might be mainly attributed to btl, whereas the branch integrity
defects might depend on Egfr.
|
). However, the Egfr pathway has been shown to
promote cell survival in several tissues
(Cabernard and Affolter, 2005;
Kurada and White, 1999). We
therefore investigated whether cell death precedes and accounts for the
tracheal defects of Egfr downregulation. A caspase-3 antibody
revealed scattered cell death throughout the tracheal tree of btlGal4
801 embryos (Fig. 4A), and
accordingly we detected few apoptotic cells, and the presence of macrophages
(see Movie in the supplementary material). This suggests that, although it
could contribute in part to it, cell death does not play a major role in
tissue integrity. Consistently, protecting the tracheal cells of btlGal4
801 embryos from dying by expressing at the same time the viral
antiapoptotic p35 protein (Hay et
al., 1994) still produced branch integrity and primary branching
defects (Fig. 4B). In addition,
tracheal cells, even those present in fragmented branches, showed a normal
(although more rounded) shape, instead of the characteristic condensed shape
of apoptotic cells (Fig. 1).
Similarly, preventing cell death with p35 or Df(3L)H99, a
deficiency that suppresses reaper-dependent apoptosis
(White et al., 1994), in
btlGal4 UAS-EgfrDN or Egfrf2 embryos,
respectively, did not rescue the branch integrity phenotype
(Fig. 4C,D). These results
indicate that cell death is not the primary cause of the branch integrity
phenotype.
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5 embryos
expressed general markers, such as btl
(Fig. 4A,B,D,E), the lumen
marker 2A12 (Fig. 4C),
trh (Fig. 5B,G),
hnt, pio and others (not shown), spatially restricted markers, such
as kni (Fig. 4F), and
later markers, like DSRF (Fig.
4E). Additionally, most branch fusions occurred (not shown),
indicating that cells underwent normal differentiation. These results rule out
that misspecification of tracheal fates is the cause of the branch integrity
phenotype.
Downregulation of the MAPK pathway impinges on Cadherin-based cell adhesion
Epithelial cells are attached to one another by several types of junctions
composed of different protein complexes that occupy the most apical region of
the lateral membrane. As a consequence, the cells show a marked apico-basal
polarity (Knust and Bossinger,
2002; Tepass et al.,
2001). The rounded shape of the tracheal cells and the loss of
epithelial integrity upon Egfr pathway downregulation are reminiscent of
non-polarised cells. We analysed markers for different junctional complexes
and did not detect defects in the localisation of most of them (not shown; see
below), indicating that tracheal cells did not loose their general apico-basal
polarity.
DE-cad, encoded by the shg gene, is a classical cadherin that
represents a major constituent of AJs
(Tepass et al., 2001). When
analysing the accumulation of DE-cad in
rhoP5, btlGal4
UAS-EgfrDN or btlGal4 801 embryos, we detected a
reproducible, but mild, decrease in the levels, and a loss in the sharpness of
staining when compared with the internal control embryos
(Fig. 5B-D; see also Fig. S1 in
the supplementary material). The effects were more conspicuous in thin
unicellular branches, such as the DBs, where the staining that appears as a
line lining the lumen in the wild type
(Fig. 5A) was sometimes lost or
very reduced. Nevertheless, the decrease was detected in all branches, even in
the DT, where the characteristic mesh-like staining that reflects the apical
outline of the cells was, in extreme examples, lost or became spotty around
some cells (Fig. 5A-D).
Conversely, high levels of DE-cad were observed in embryos expressing a
constitutively activated Egfr (Fig.
5E; Fig. S1 in the supplementary material), and occasionally we
observed abnormally straight junctions compared with the wavy wild type ones,
as if the cells were subjected to higher tension.
To further prove the modulation of DE-cad levels upon modulation of the
Egfr pathway, we downregulated or overactivated Egfr signalling in small
groups of tracheal cells by inducing the expression of
EgfrDN (Fig.
6A-D) or top
(Fig. 6E-G). In spite of the
fact that we had to perform these experiments under conditions of moderate
activation of these transgenes (see Materials and methods), we could detect,
respectively, a mild decrease or increase in DE-cad levels in several
examples, validating the above results.
The relationship and interdependence of AJs and the actin cytoskeleton have
been extensively documented (Bershadsky,
2004; Carthew,
2005; Gates and Peifer,
2005; Goodwin and Yap,
2004; Zhang et al.,
2005). For this reason, we analysed the actin cytoskeleton in our
mutant conditions. In the wild type, we observed a prominent cortical actin
bundle in the tracheal tubes by late embryogenesis
(Fig. 5F). In loss-of-function
conditions of the Egfr pathway, we detected thinner accumulation of cortical
actin in most embryos when compared with the internal controls
(Fig. 5G-I). Conversely, when
the Egfr is constitutively activated in tracheal cells, we observed an
enrichment of apical actin (Fig.
5J). Altogether, our results establish a correlation between the
levels of DE-cad and cortical actin, and the activity of the Egfr pathway
during tracheal development.
|
). The
antibody used to detect DE-cad recognises the endogenous protein, including
the maternally provided protein. To analyse the effect of the Egfr pathway on
the newly synthesized protein, we made use of a functional DE-cadGFP
(Oda and Tsukita, 1999)
expressed under the control of heterologous promoters. In otherwise wild-type
conditions, this protein is incorporated into the cell junctions
(Chihara et al., 2003). We
first used btlGal4 to express DE-cadGFP in the tracheal
system, and found lower levels of GFP signal, when compared with wild type,
under several conditions of Egfr pathway downregulation
(Fig. 7A-C; not shown). We
confirmed these results in western blot experiments
(Fig. 7F). Similarly, when we
used widely expressed drivers, such as 69B or ptcGal4, we also
detected lower levels of DE-cadGFP in several tissues, such as the trachea
(Fig. 7D,E), the salivary
glands (Fig. 7G,H) or hindgut
(not shown) upon downregulation of Egfr pathway. These results show that the
Egfr pathway can regulate DE-cad levels posttranscriptionally in various
contexts.
Mutants affecting cadherin-based cell adhesion display a tracheal branch integrity phenotype
If the branch integrity phenotype of Egfr pathway downregulation is due to
a decrease in cell-cell adhesion, we would expect a similar phenotype when the
integrity of AJs is compromised. Mutants for shg had been described
as being defective in tracheal branch fusion and they display interruptions in
the tubes (Uemura et al.,
1996). In addition to these phenotypes, we observed that tracheal
cells show a rounded shape and that branches tend to break, leaving cells
isolated from the rest of the tree, as occurs when the Egfr pathway is
downregulated (Fig. 8A).
Furthermore, the levels of cortical actin decreased in shg mutants
(Fig. 8C).
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) and, accordingly, we observed that it is required for the
proper accumulation of cortical actin in tracheal tubes (not shown).
cv-c mutants showed a mild branch integrity phenotype that consists
of cells abnormally separated and sometimes attached by cytoplasmic bridges
(Fig. 8B), and we detected a
mild decrease in DE-cad accumulation (Fig.
8D). These results indicate that compromising AJs or the actin
cytoskeleton, which can impinge on one another, results in tracheal integrity
defects.
|
A new role of the Egfr signalling pathway during tracheal development
The requirement for Egfr signalling in the developing tracheae has been
already studied by us and by others. The pathway plays a pivotal role during
tracheal invagination and also in primary branching
(Bradley and Andrew, 2001;
Llimargas and Casanova, 1997;
Llimargas and Casanova, 1999;
Wappner et al., 1997). In
fact, it was suggested that the defects observed in branching could be a
consequence of an abnormal invagination of the cells
(Bradley and Andrew, 2001;
Llimargas and Casanova, 1999).
This or these requirements seem to depend on a peak of Egfr activity
observed prior to invagination, and correlate with a peak of ERK
phosphorylation, as visualised with a dpERK antibody
(Gabay et al., 1997).
Here, we document a new role for the Egfr pathway in the regulation of
tissue integrity. This new requirement could depend on the described early
peak of Egfr activity (Gabay et
al., 1997), which would be sufficient to prevent defects at later
stages. However, we propose that Egfr-promoted epithelial integrity
depends on a later, or continuous but lower, or basal activity of the pathway
that does not correlate with detectable ERK phosphorylation. Consistent with
this hypothesis, we find that downregulation of the pathway by overexpressing
801 or UAS-EgfrDN with btlGal4, which is
expressed after the early peak of ERK phosphorylation, produces a conspicuous
branch integrity phenotype. In any case, tissue integrity defects are mainly
observed in the most dorsal and ventral tracheal branches, which are subjected
to stronger pulling forces as development proceeds, and, therefore, it is
precisely at late stages when defects in tissue integrity are expected.
AJs connecting epithelial cells dynamically disassemble and reassemble,
thereby allowing tissue remodelling. Tracheal tissue remodelling might require
the fine-tuning of cell adhesion properties, as tracheal cells need to be able
to change their relative position (probably by loosening cell adhesion) while
maintaining epithelial continuity. Our data indicates that the Egfr pathway is
a modulator of this balance, not only in the tracheal system, but also in
other tissues undergoing extensive remodelling, such as the salivary glands,
where we find a similar regulation of DE-cad and actin levels upon modulation
of Egfr signalling (C.C. and M.L., unpublished). Conversely, we do not find
such a regulation in more static tissues, like the ectoderm (C.C. and M.L.,
unpublished), whose maintenance was proposed to depend on the maternally
provided DE-cad protein (Uemura et al.,
1996). We suggest that the Egfr pathway plays a role in the
modulation of cell adhesion in tissues that undergo dramatic morphogenetic
events, which might need the zygotic DE-cad contribution and a more dynamic
regulation of cell adhesion. Our results indicating a modulation of junctional
complexes and/or the actin cytoskeleton by the Egfr pathway establish a link
between a developmental pathway required for many biological events and cell
biology in terms of cell adhesiveness and cell shape.
The activity and activation of the MAPK signalling pathway during tracheal development
Our results show that downregulation of several intracellular elements of
the MAPK pathway produce defects in branch integrity, whereas a constitutively
activated form of rl (rlsem) rescues the
phenotype of btlGal4 801 embryos. This suggests that the conserved
MAPK cassette is required to maintain branch integrity.
Two tyrosine kinase receptors, Egfr and Btl, activate the MAPK pathway
during embryonic tracheal development. However, we find that the two
receptors, acting through the same intracellular cascade, elicit different
responses. The MAPK pathway requirement in primary branching is likely to
depend on input by btl, whereas the tissue integrity requirement is
likely to depend on input by Egfr. How does the same MAPK pathway
trigger distinct outcomes depending on the receptor that activates it? A
temporal and/or spatial differential activation of the MAPK pathway could
account for the different outcome. In addition, differences in the composition
of the intracellular cascade due to specific transducers for one type of
receptor, such as downstream of FGFR (dof;
stumps-FlyBase) (Petit et al.,
2004), could contribute. Finally, quantitative and/or qualitative
differences in the activation of the intracellular transducers by the
different receptors could also underlie the outcome diversity.
Similar to our observations, air sac development in Drosophila has
been recently reported to require both Btl and Egfr, and each receptor seems
to elicit different responses. Furthermore, as we find during embryonic
tracheal development, an uncoupling of the MAPK cassette and pnt has
been observed during air sac development
(Cabernard and Affolter, 2005).
These parallels suggest a common mechanism for generating different responses
from the same intracellular transduction pathway.
Regulation of tissue integrity by the Egfr pathway
The loss of tissue continuity and cell detachment observed in Egfr
downregulation conditions may be due, at least in part, to a decrease in cell
adhesion. Accordingly, we detect a mild, but reproducible, decrease in the
accumulation of DE-cad and cortical actin. As inferred from the phenotypes,
such a mild decrease could cause a loss of cell adhesion during tracheal
remodelling, while not grossly affecting other processes requiring
DE-cad-based cell adhesion, such as branch fusion
(Tanaka-Matakatsu et al.,
1996; Uemura et al.,
1996). As expected, we find that compromising AJ assembly or the
actin cytoskeleton also gives rise to defects in tracheal tissue
integrity.
Cadherins have been shown to support cell cohesion and participate in
morphogenetic events. The actin cytoskeleton also plays an important role in
shaping the cell architecture and in many morphogenetic processes. AJs and the
actin cytoskeleton are intimately coupled, and their formation and maintenance
is interdependent (Bershadsky,
2004; Carthew,
2005; Gates and Peifer,
2005; Goodwin and Yap,
2004; Zhang et al.,
2005). We also observe such interdependence in the tracheal
system.
Cadherin-based cell-cell adhesion can be regulated at transcriptional and
posttranscriptional levels. The modulation of a DE-cadGFP chimaera
driven by heterologous promoters shows that, in our case, DE-cad regulation is
posttranscriptional. Several posttranscriptional mechanisms of DE-cad
regulation have been proposed
(D'Souza-Schorey, 2005), and
we can envisage a role for the Egfr pathway in each of them. A first mechanism
is at the level of DE-cad endocytic trafficking. In this context, the Egfr
pathway could modulate the balance between recycling to the plasma membrane of
internalised DE-cad or lysosomal targetting and degradation. A second
mechanism of cell-cell adhesion regulation is posttranslational modifications
of AJ components, such as phosphorylation or ubiquitination. Finally, another
possible mechanism of regulation is through the cytoskeleton. The Rho family
of small GTPases plays a key role in actin cytoskeleton regulation, and growth
factor receptors such as Egfr have been reported to regulate their activity
(Burridge and Wennerberg,
2004). Remarkably, the Egfr pathway has been recently shown to
regulate the expression of the rhoGAP cv-c in the tracheal placodes
(Brodu and Casanova, 2006), and
we find that cv-c mutants display tracheal integrity defects,
although they are milder than those seen upon downregulation of the Egfr
signal. We therefore propose that cv-c is at least one of the
effectors of Egfr-mediated modulation of DE-cad levels and tracheal tissue
integrity. Further analysis will be needed to disentangle the exact molecular
mechanisms and to find other possible mediators of the Egfr signal.
The decrease of cadherin activity upon activation of the Egfr pathway has
been extensively reported in the literature
(Comoglio et al., 2003;
Dumstrei et al., 2002;
Lilien and Balsamo, 2005).
Here, we report the opposite: that Egfr pathway downregulation correlates with
a decrease of cadherin-based cell adhesion. Although this is not the first
example of such a relationship (Brown and
Freeman, 2003), it illustrates the versatility and complexity of
the interactions occurring between signalling pathways and adhesion molecules,
and establishes another model with which to analyse how cell adhesion is
modulated.
Supplementary material
Supplementary material for this article is available at
http://dev.biologists.org/cgi/content/full/133/16/3115/DC1
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ACKNOWLEDGMENTS |
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REFERENCES |
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