|
|
|
|
|
|
|
||||
| Home Help Feedback Subscriptions Archive Search Table of Contents | ||||
First published online May 23, 2006
doi: 10.1242/10.1242/dev.02404
Department of Biochemistry and Howard Hughes Medical Institute, Stanford University School of Medicine, Stanford, CA 94305-5307, USA.
* Author for correspondence (e-mail: krasnow{at}cmgm.stanford.edu)
Accepted 12 April 2006
 |
SUMMARY |
|---|
 TOP SUMMARY INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
|---|
-integrins, also show the tendrils phenotype, and localization
of myospheroid ß-integrin protein is disrupted in
tendrils mutant terminal cells. The results provide evidence that
integrin-talin adhesion complexes are necessary to maintain tracheal terminal
branches and luminal organization. Similar complexes may stabilize other
tubular networks and may be targeted for inactivation during network
remodeling events.
Key words: Drosophila, Talin, Trachea, Integrin, Tube morphogenesis
|
INTRODUCTION |
|---|
|
TOPSUMMARYINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
|---|
; Lubarsky and Krasnow,
2003; O'Brien et al.,
2002). The tube walls are typically an epithelial sheet of cells
with the apical surface lining the lumen, in direct contact with the
transported substance. During development, the cells are shaped to create
tubes of different length, gauge, cellular architecture and branching pattern
that are tailored to organ function. Once formed, tubes are usually maintained
for long periods. However, at specific times during development or under
certain physiological conditions, some tubular networks rearrange or undergo
dramatic remodeling events. These include pruning or complete elimination of
branches, as occurs during metamorphosis of insect tracheal (respiratory)
systems (Whitten, 1957),
during conversion of primitive vascular plexi into mature blood vessel
networks during vertebrate development
(Risau and Flamme, 1995),
during mammary gland involution after weaning
(Green and Lund, 2005) and
following treatment of tumors with anti-angiogenesis drugs
(Inai et al., 2004). Although
progress has been made identifying mechanisms that control branch outgrowth
and tube morphogenesis, much less is known of the molecular events that
stabilize tubes and destabilize them during remodeling events. Here, we
describe a gene required for maintenance of terminal branches of the
Drosophila tracheal system.
The larval tracheal system of Drosophila is a network of thousands
of interconnected tubes that delivers oxygen to tissues
(Fig. 1A,B)
(Manning and Krasnow, 1993;
Rühle, 1932). Air enters
the network and diffuses through primary, secondary and terminal branches to
reach the tissues (Wigglesworth,
1972). Terminal branches (tracheoles) are fine gauge tubes,
ranging from 0.1-1.0 µm in diameter
(Fig. 1C,D)
(Wigglesworth and Lee, 1982).
These blind-ended tubes ramify extensively on and attach tightly to internal
tissues to facilitate gas exchange
(Manning and Krasnow, 1993;
Noirot and Noirot-Thimotee,
1982). The attachments are generally long lived, although under
certain conditions, cellular projections from oxygen-starved cells can bind to
and redistribute nearby terminal branches to satisfy their oxygen need
(Wigglesworth, 1959;
Wigglesworth, 1977).
The tracheal system arises during mid-embryogenesis from ten pairs of
epithelial sacs, each of which forms a hemi-segment of the larval tracheal
network (Samakovlis et al.,
1996). Small groups of tracheal cells migrate out from each sac to
form primary branches, guided by branchless FGF
(Sutherland et al., 1996).
Some cells at the tips of budding primary branches form unicellular secondary
branches sealed by an autocellular junction. These same cells go on to form
terminal branches by extending long cellular projections towards
oxygen-starved, branchless-expressing cells in the target tissues
(Guillemin et al., 1996;
Jarecki et al., 1999). A lumen
then forms within each projection by a poorly understood process that creates
an intracellular, membrane-bound channel without any associated cell junctions
(Fig. 1D)
(Guillemin et al., 1996;
Lubarsky and Krasnow, 2003).
This process of outgrowth and lumen formation is repeated many times during
the 5 days of larval life so that by the end of the third larval instar a
single terminal cell has formed dozens of terminal branches, each with a
single air-filled lumen, that are neatly matched to the oxygen needs of the
target (Fig. 1C,D). Although
signaling pathways and transcription factors that control branch sprouting and
outgrowth have been characterized (Affolter
et al., 2003; Ghabrial et al.,
2003; Uv et al.,
2003; Zelzer and Shilo,
2000), downstream effectors that mediate outgrowth, lumen
formation, substrate attachment and branch maintenance remain to be
identified.
Here, we describe a complementation group called tendrils
identified in a genetic mosaic screen for tracheal mutants. tendrils
mutations cause a dramatic alteration in terminal branch lumen organization
and a reduction in terminal branch number. We provide evidence that these
phenotypes arise late in development from a defect in branch maintenance.
tendrils is allelic to rhea, the gene that encodes the
Drosophila talin (Brown et al.,
2002), a large cytoskeletal protein that links integrin
cell-adhesion molecules to the cytoskeleton
(Calderwood and Ginsberg,
2003; Calderwood et al.,
1999; Garcia-Alvarez et al.,
2003). The major ß-integrin of Drosophila localizes
along the basal surface of terminal branches, and terminal cells lacking this
integrin or the -integrins Multiple edematous wings and Inflated
exhibit the tendrils phenotype. The results suggest that
integrin-talin adhesion complexes anchor mature terminal branches to their
substrata and maintain luminal organization. In the absence of these
complexes, lumens become disorganized and retract from branches, and lumenless
branches degenerate.
|
MATERIALS AND METHODS |
|---|
|
TOPSUMMARYINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
|---|
;
Prout et al., 1997);
myospheroidXG43 (mys)
(Bunch et al., 1992),
mysM2 and mysG4
(Jannuzi et al., 2002);
inflatedk27e (if), multiple edematous
wingsM6 (mew) and ifk27e
mewM6 double mutant; scab1; and
integrin linked kinase1 (ilk),
blistery3R-13 (by), vinculin1
(vinc) and steamer duck3R-17 (stck).
Mutations that were viable through the third larval instar were analyzed as
homozygotes. Others were recombined as necessary onto FRT bearing chromosomes
[FRT19A(X), FRT40A(2L),
FRTG13(2R), FRT2A(3L) or
FRT82B(3R) as appropriate] and analyzed in genetic
mosaics. Germline clones of rhea79 were generated by the
dominant female sterile technique (Chou
and Perrimon, 1996). For analysis of maternal-zygotic
tendrils mutant embryos, rhea79 eggs generated
this way were fertilized with tendrils13-8 or
tendrils6-66 sperm.
Mosaic analysis
Embryos harboring hsFLP122, btl-Gal4, UAS-GFP and/or
UAS-DsRed, and the mutation of interest on an FRT chromosome in trans
to a chromosome containing an identical FRT insertion and either
UAS-Gal80 (Lee and Luo,
1999) or GFP RNAi (UAS-GFPi; A.S.G., B.P.L. and M.A.K.,
unpublished) transgenes, were collected 0-4 hours after egg lay (AEL). Embryos
were subjected to a 38°C heat shock for 45-60 minutes to induce expression
of FLP recombinase, and then allowed to develop at 25°C for the times
noted. Homozygous mutant tracheal cell clones lack the dominant repressor
(UAS-Gal80 or UAS-GFPi) and express GFP (btl-Gal4,
UAS-GFP). For whole-mount analysis, larvae were mounted in 50% glycerol,
killed by heating on a 70°C block for 
5 seconds, and examined with an
Axiophot compound fluorescence microscope equipped with a CCD camera.
For quantification of phenotypes, mosaic wandering third instar larvae were analyzed. The number of lumen tips and branch tips in individual terminal cells were counted and used to calculate the lumen:branch ratio. Lumen tip number could be reliably ascertained up to approximately six tips per branch. Only mature branches in which a cytoplasmic process was visible by GFP fluorescence and a lumen was visible under bright-field optics were counted. To quantify branch loss, the number of branch tips in each mutant terminal cell was compared with the number in the contralateral wild-type cell.
Tracking development of mutant clones
Individual terminal cells marked with GFP were identified in early L3
mosaic larvae anesthetized with ether. The mutant cell was imaged as above,
and the larva was placed in a fresh vial and allowed to continue development
at 25°C. After 48 hours, the larva was heat killed, and the mutant cell
was imaged again. Contralateral wild-type terminal cells served as
controls.
Genetic mapping
Two rounds of meiotic recombination mapping of
tendrils13-8 were carried out using an isogenized mapping
chromosome (ru h th st cu sr e ca). Recombinants were tested for
complementation of tendrils15-39. SNP markers have been
described (Berger et al.,
2001). SNP 3L078A [5'ATCTTGTAACCT(A/C)GGGGGTTGCAC] is an
AvrII RFLP identified here that was amplified by PCR with primers
5'GCGGACCAAGAACCCAGTGACAAC and 5'GTTCCTCGAAGTGACCCGAATGTCC.
Complementation tests were carried out between
tendrils15-39 and chromosomal deficiencies Df(3L)ZP1,
Df(3L)66C-G28 and Df(3L) BSC13, and the mutations
rhea1, cblKG03080, hairy22,
l(3)DTS41, l(3)L0139, l(3)j8E8, l(3)neo13 and
Grunge03928.
DNA sequencing
tendrils alleles were rebalanced over TM3 Sb
twi-Gal4,UAS-GFP (FlyBase). Homozygous tendrils embryos were
identified by absence of GFP. DNA was extracted using Chelex beads
(Walsh et al., 1991). All
rhea exons and splice sites were amplified by PCR and sequenced.
Analysis of tendrils13-8 splicing defect
RNA was isolated with Trizol reagent (Invitrogen) from
tendrils13-8/rhea79 embryos derived
from rhea79 germline clones. cDNA was prepared using
PowerScript reverse transcriptase (BD Biosciences) and primer
5'CGCTGTGGCTCCGTCAGATTTAC. Talin cDNA was amplified by PCR using primers
5'TTACCCAAGGAAACGACGAC and 5'AAGGTAACGCCGTAGGTGG flanking the
fourth intron. PCR products were sequenced to determine the splicing
pattern.
Immunostaining
L3 larvae were filleted open along the ventral midline, dissected in 3.7%
formaldehyde/phosphate-buffered saline (PBS), fixed for 20 minutes at room
temperature, then transferred to 100% methanol at -20°C for at least 20
minutes. Samples were rehydrated into PBS with 0.3% Triton X-100 (PBST).
Antigen blocking was at room temperature for 30 minutes in PBST with 0.2% BSA
(PBSTB). Subsequent incubations were carried out in PBSTB as described
(Patel, 1994). Primary
antibody incubations were at room temperature for 2 hours or 4°C
overnight. Primary antibodies were anti-ßmys-integrin mAb
CF6G11 (Developmental Studies Hybridoma Bank; 1:1000 dilution) and rabbit
anti-GFP (Molecular Probes; 1:500). Secondary antibodies coupled to Cy2 and
Cy3 (Jackson ImmunoResearch) were used at 1:500. Samples were mounted in
ProLong media (Molecular Probes) and images acquired on a BioRad 1024 confocal
fluorescence microscope and adjusted using Adobe Photoshop. Embryo fixation
and staining was as described (Samakovlis
et al., 1996), except maternal-zygotic tendrils embryos
were stained with a rhodamine-conjugated chitin probe
(Devine et al., 2005).
Transmission electron microscopy
Genetic mosaic animals were generated as above, except clones were marked
with GFP and CD2-HRP (Larsen et al.,
2003; Watts et al.,
2004). Mosaic larvae were identified by GFP fluorescence,
dissected in PBS, and immediately fixed for 15 minutes at room temperature in
2.5% glutaraldehyde/PBS. Horseradish peroxidase (HRP) signal was enhanced
using biotin-labeled tyramide amplification (PerkinElmer) and avidin-HRP, and
developed with diaminobenzidine (DAB) and NiCl for 5-10 minutes at room
temperature (Vectastain Elite, Vector Laboratories). Clones were photographed,
and fillets were postfixed for 1 hour at room temperature in 1% osmium
tetraoxide and stained overnight in 0.5% uranyl acetate. After dehydration
into 100% ethanol, samples were infiltrated with EMbed 812 resin (Electron
Microscopy Sciences) and oriented in casts for sectioning. Sectioning with a
Leica UCT microtome and imaging on a Jeol TEM1230 electron microscope were
carried out as described (Watts et al.,
2004). Images were acquired with a CCD camera and adjusted using
Adobe Photoshop. Tracheal clones were identified by electron dense precipitate
at the plasma membrane, and their lumens by the ridged cuticle lining.
|
RESULTS |
|---|
|
TOPSUMMARYINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
|---|
|
|
In addition to the effects on lumen number, organization and size,
tendrils mutations reduced the number of terminal branches. Dorsal
terminal cells in wild-type third instar larvae contained 16±5 mature
branches (n=32) (Fig.
2A,B). Terminal cell clones homozygous for the weak
tendrils allele (tendrils15-39) had 70%
fewer terminal branches than normal, the moderate alleles
(tendrils14-69, tendrils6-66) showed a 90%
reduction in branches and the strongest allele
(tendrils13-8) displayed an almost complete loss of
terminal branches. The severity of terminal branch reduction paralleled the
strength of the multi-lumen phenotype (Fig.
2B,C).
Both the multi-lumen and terminal branch reduction defects were strictly cell autonomous. Cells neighboring tendrils clones, including other terminal cells, had normal morphology (Fig. 1N). tendrils mutations are recessive as terminal cell morphology in tendrils heterozygotes was indistinguishable from wild type.
The tendrils phenotype arises late in development from failure to maintain branches
The clonal analysis examined the tendrils effect on terminal cells
at the end of third larval instar, five days after fertilization. We analyzed
tendrils terminal cells at earlier stages of development to determine
when defects arise and how they progress. The first terminal branches sprout
late in embryogenesis, 13-20 hours after fertilization. These sprouts
developed normally in homozygous tendrils embryos, as assessed by
staining fixed embryos for a tracheal luminal antigen (2A12) and cytoplasmic
marker (btl-Gal4, UAS-GFP) (data not shown). This appears to be true
also in embryos lacking maternal as well as zygotic tendrils,
although this was difficult to ascertain rigorously owing to global
developmental defects in these embryos.
Because homozygous tendrils mutants did not survive beyond embryogenesis, we examined the tracheal phenotype of developing larvae in dorsal branch terminal cell clones of the strong tendrils13-8 mutant (Fig. 3A-C). By early second instar, tendrils13-8 mutant terminal cell clones had formed multiple terminal branches with normal lumen morphology and appeared grossly similar to neighboring wild-type terminal cells, although in some clones we noticed a slight reduction in the number of terminal branch sprouts (Fig. 3A). In early third instar, defects became prominent in tendrils13-8 clones. The number of terminal branches was significantly reduced relative to wild-type terminal cells, and the lumens of the branches in mutant terminal cells were convoluted (Fig. 3B). By late third instar larvae, the phenotype was dramatic. tendrils13-8 mutant terminal cells typically had just a single stubby branch packed with a tangle of convoluted lumens (Fig. 3C). Thus, early stages of terminal cell development proceed normally in tendrils mutant cells, and defects become prominent during the third instar, several days after terminal branches have begun sprouting and forming air-filled lumens.
Phenotypic progression was also assessed by examining individual terminal cell clones twice during third larval instar. This was carried out by characterizing GFP-labeled clones in live, anesthetized larvae, then placing the larvae in fresh vials and re-examining the same clones 48 hours later. Fig. 3D shows a representative tendrils13-8 mutant terminal cell clone in an anesthetized early third instar larva, which appears similar to the clones described above in heat-killed larvae of the same age: there were multiple terminal branches, each containing one or more convoluted lumens. The same mutant cell 48 hours later is shown in Fig. 3E,E'. Terminal branches 1, 2 and 3 have been lost and there is a corresponding increase in number of lumens in the remaining branch. The neighboring control tendrils+ terminal cell did not show any branch loss or change in lumen morphology during the same period (data not shown). When the moderate allele tendrils6-66 and the rhea79 allele described below were analyzed in the same way, a similar loss of branches and increase in lumen number in the remaining branches occurred during the 48-hour period, although the effects were less extensive and occasionally no branches were lost during the observation period (Fig. 3F,G; data not shown). In clones of the moderate alleles, we often detected thin terminal branches whose lumen was displaced into the proximal terminal branch during the 48 hour period (e.g. branches 2, 3 in Fig. 3F-G'). These branches lacking lumens are likely to be intermediates in the process of degenerating. Thus, the tendrils phenotype appears to result from a catastrophic event(s) late in terminal cell development that disrupts lumen organization, and destabilizes and ultimately destroys terminal branches.
|
tendrils is allelic to rhea, the Drosophila talin
To initiate molecular analysis of tendrils, we mapped the
tendrils locus and identified the gene
(Fig. 4A). Initial meiotic
recombination mapping using visible genetic markers placed
tendrils13-8 between roughoid and hairy
on chromosome III. Subsequent high-resolution recombination mapping with SNP
markers (Berger et al., 2001)
and mapping with chromosomal deficiencies in the region localized
tendrils to a 260 kb interval between cytological positions 66D2
and 66D8-9. All known lethal mutations in the candidate interval were tested
for their ability to complement tendrils mutations for viability. The
tendrils alleles all failed to complement mutations in rhea,
the gene encoding Drosophila talin
(Brown et al., 2002). Terminal
cell clones of rhea2 and rhea79 were
generated and showed the tendrils phenotype, similar in severity to
moderate tendrils alleles (Fig.
3I-J'; data not shown).
|
;
Calderwood and Ginsberg, 2003)
(Fig. 4D). Talins contain an
N-terminal head domain with a FERM motif required for binding to
ß-integrins, actin, PIPKI
and FAK, and a long
C-terminal rod with two higher affinity F-actin binding domains
(Fig. 4B)
(Brown et al., 2002;
Calderwood and Ginsberg,
2003). Sequencing the talin-coding region and RNA splice sites in
each tendrils mutant identified a single nucleotide change in each
allele that alters the coding region or a splice signal
(Fig. 4B).
tendrils14-69, tendrils6-66
and tendrils15-39 are nonsense mutations that truncate the
coding region after amino acids 933, 1249 and 2051, respectively, eliminating
regions of the C-terminal rod domain and one or two F-actin binding domains.
The strongest allele, tendrils13-8, is a G to A
substitution in the 3' splice site of the fourth intron
(Fig. 4C). Analysis of
tendrils13-8 mRNA by RT-PCR revealed that the mutation
causes mis-splicing and use of a cryptic 3' splice site located seven
nucleotides 3' of the normal site. This alters codon 268 and beyond,
which truncates talin in the head domain, removing much of the FERM motif and
the entire rod domain. We conclude that tendrils mutations are
alleles of rhea and that the tendrils alleles constitute a
C-terminal deletion series of talin, with the extent of truncation paralleling
the severity of the tendrils phenotype.
myospheroid ß-integrin and two -integrins function with talin to maintain terminal branches and luminal organization
Talin is required for integrin-mediated adhesion in Drosophila
(Brown et al., 2002;
Devenport and Brown, 2004),
but integrin-independent functions of talins have also been reported
(Becam et al., 2005;
Borowsky and Hynes, 1998). To
determine if talin functions in an integrin-dependent process in terminal
cells, we examined the localization and function in terminal cells of ßPS
[encoded by myospheroid (mys), hereafter called
ßmys-integrin], the major Drosophila ß-integrin
required for virtually all known integrin-mediated processes
(Brown et al., 2000;
Devenport and Brown, 2004).
Localization of ßmys-integrin in third instar larval terminal
cells was assessed by immunostaining fillets. ßmys-Integrin
was detected in punctae along the periphery of terminal branches
(Fig. 5A,A'). These may
represent focal adhesions attaching the branches to the underlying tissue.
|
). Mutant
clones of all three mys alleles shared the distinctive rhea
phenotype (Fig. 5E,F), with
substantial branch pruning and multiple convoluted lumens coursing through the
remaining terminal branches, similar to the null allele,
rhea79 (Fig.
5D). Ultrastructural analysis of mysXG43
mutant terminal cells confirmed that multiple lumens were present within
single terminal branches (Fig.
5J-L), like terminal cells that lack talin
(Fig. 1K-M). When individual
mysXG43 terminal cells were imaged in live early third
instar larvae and again 48 hours later
(Fig. 5H,I), terminal branches
were lost (branch 5) or collapsed (branches 1-4) as the multi-lumen defect
became more prominent, as in terminal cells that lack talin
(Fig. 3D-G'). Indeed, as
with moderate talin mutants, lumens were often displaced from branches before
the branch was completely lost (branches 1-4). We conclude that
ßmys-integrin is required along with talin to maintain
terminal branches and luminal organization in terminal cells. Consistent with
this, the punctate peripheral localization of ßmys-integrin
required talin (Fig.
5B,B'), as in the developing wing
(Brown et al., 2002).
Integrins are ß heterodimers. Five -integrin genes are
present in the Drosophila genome
(Brown et al., 2000;
Hynes and Zhao, 2000). Mutant
terminal cell clones were generated for if, mew and scb, the
three -integrin genes for which loss-of-function mutations are
available. None of the single mutations caused any detectable defects in
terminal cell morphology when homozygous (data not shown). However homozygous
mewM6, ifk27e double mutant clones exhibited
the tendrils phenotype (Fig.
5G,G') demonstrating that these two -integrins are
genetically redundant and function along with ßmys-integrin in
terminal cells. Terminal cell clones mutant for the integrin complex genes
ilk, by, vinc and stck were also examined and showed normal
morphology, suggesting that only rudimentary integrin complexes are required
for maintenance of terminal branches (data not shown).
|
DISCUSSION |
|---|
|
TOPSUMMARYINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
|---|
;
Calderwood and Ginsberg, 2003;
Liu et al., 2000).
ßmys-Integrin localizes in a talin-dependent manner to
discrete domains along the basal surface of terminal branches, and terminal
cells mutant for mys or doubly mutant for mew and
if -integrins show the same striking phenotype as cells
lacking talin. The results support a model in which integrins and talin form adhesion complexes in terminal cells, and these complexes are required late in larval life to anchor tracheal terminal branches to the underlying substratum and maintain lumen organization within the branches (Fig. 6). In the absence of these complexes, substrate adhesion is compromised, lumen organization is disrupted, lumens retract from the branches and the lumenless branches degenerate. We consider below the implications of these results and this model for the function of integrins and talin in tracheal branch outgrowth, maintenance and lumen organization, and discuss their significance for other tubular networks.
Integrins and talin are required for maintenance of tracheal terminal branches
Drosophila integrins and talin have been implicated in multiple
cell adhesion, migration and spreading events (reviewed by
Bokel and Brown, 2002;
Brown et al., 2000;
Brown et al., 2002), including
migration of dorsal epidermis, midgut and salivary glands
(Bradley et al., 2003;
Devenport and Brown, 2004). By
contrast, ßmys-integrin and talin were dispensable for the
initial sprouting and outgrowth of tracheal terminal branches. It is unlikely
that the other Drosophila ß-integrin,
ßv-integrin, functions in terminal branch budding and
outgrowth, as it is expressed specifically in the gut
(Devenport and Brown, 2004).
This suggests that the crucial role of integrins and talin in tracheal
terminal cell development is in branch maintenance rather than outgrowth.
A requirement for -integrins mew and if in
spreading of tracheal visceral branches along the gut during embryonic
development has been demonstrated (Boube et
al., 2001). This suggests that integrins may also play a role in
tracheal branch outgrowth and spreading, at least for this particular branch
at this stage of development. Alternatively, the visceral branch phenotype
could be an early manifestation of the proposed branch maintenance function of
integrins and talin: the gut undergoes substantial morphogenetic movements at
this stage that could stress its association with developing visceral terminal
branches and cause their detachment. Indeed, embryos lacking maternal and
zygotic talin exhibit a similar visceral branch phenotype as mew and
if mutants (B.P.L., unpublished). However, the overall morphology of
the visceral branch and its attachment to the gut are not compromised in the
presence of rhea, mys or mew if double mutant terminal cell
clones, presumably because wild-type terminal cells neighboring the clone are
sufficient to anchor the branch to the gut.
|
; Prokop et al.,
1998; Tepass et al.,
2000) (B.P.L., unpublished). Interestingly,
tendrils13-8 and other mutations that result in C-terminal
truncations of talin (Brown et al.,
2002) cause a more severe phenotype than deletion of the gene,
null mys mutations or mew if double mutations. This suggests
that the remaining N-terminal region of talin has antimorphic effects that
interfere with integrin-independent functions of talin, perhaps including its
interaction with other cell-adhesion systems. The postulated involvement of
two adhesion systems in terminal branch outgrowth could explain why the
requirement for ßmys-integrin and talin manifests only late in
development. Perhaps that is when the outgrowth adhesion system is
downregulated and the integrin-talin system becomes necessary to anchor the
branches stably to the hypoxic cells they grew out to supply.
Integrins and talin are also required for luminal organization
Surprisingly, the first obvious manifestation of the tendrils
phenotype in terminal branch development is not loss of branch adhesion but
alteration in lumen organization - lumens become convoluted. We also observed
lumens of some mutant terminal branches retracting into the parental branch,
while the branch it originally occupied was still present and associated with
its target. This explains why the residual branches have multiple lumens with
different diameters. The results suggest that, in addition to their substrate
adhesion function, talin and integrins play an important role maintaining
luminal organization within terminal branches. Although the molecular
mechanisms of lumen formation and maintenance are unknown, it is likely that
the cytoskeleton is crucial for directing assembly of the lumen and
maintaining it in a central position within the cytoplasm of each terminal
branch (Hogan and Kolodziej,
2002; Lee et al.,
2003; Lee and Kolodziej,
2002). Because talin is a crucial mediator of the interaction
between integrins and the actin cytoskeleton
(Brown et al., 2002;
Calderwood and Ginsberg, 2003;
Cram and Schwarzbauer, 2004),
we speculate that it plays a similar role in tracheal terminal cells and that
this interaction is required to support the lumen
(Fig. 4D,E). Moesin, a protein
that interacts with the microfilament cytoskeleton
(Speck et al., 2003),
localizes around the lumen in tracheal terminal cells, and this localization
is maintained in terminal cells lacking talin (B.P.L., unpublished). This
implies that components of the actin cytoskeleton and apicobasal polarity are
not grossly disrupted in the mutants, and suggests that talin may play a
specific role coupling the apical (luminal) actin network to the basal cell
surface, where integrins bind the ECM (Fig.
4E).
A prediction of this model is that mutations in components of the proposed linkage between the ECM and lumen, including other components of integrin-talin adhesion complexes, would also affect lumen organization. None of the mutations we analyzed in other integrin pathway genes had a tendrils phenotype, including strong or null mutations in ilk, stck, by and vinc. However, another gene identified in our tracheal screen has a tendrils-like phenotype, and other complementation groups display defects in lumen organization without affecting branch adhesion or maintenance (A.S.G., B.P.L. and M.A.K., unpublished). These mutants may help identify additional components of the terminal cell integrin complex and components that link it to the luminal membrane.
Terminal branch degeneration in the absence of integrins and talin
In addition to the striking luminal organization defects observed in
rhea, mys and mew if clones, the number of terminal branches
was substantially reduced. This reduction results largely from failure to
maintain terminal branches, including mature branches complete with air-filled
lumens. Absence of integrin-talin complexes in terminal cells thus not only
compromises branch adhesion and luminal organization, but also leads to
destabilization and destruction of branches. How are destabilized branches
eliminated? One possibility is that the branches are degraded, for example, by
autophagy or engulfment by phagocytes. Another possibility is that the
destabilized branches are subsumed by the parental branch, like cultured
fibroblasts that round up when released from their substratum or neuronal
processes that retract by a regulated actomyosin-dependent process
(Billuart et al., 2001). The
retraction mechanism is appealing because it would explain why the remaining
branches in mutant terminal cells lacking talin or integrins are thicker than
normal - the contents of the daughter branches are consolidated into the
parental branch - and why empty basement membranes are sometimes found in
positions distal to tendrils clones (B.P.L., unpublished).
Although terminal branches normally attach stably to their targets, under
hypoxic conditions cellular projections from oxygen-starved cells have been
shown to bind nearby terminal branches and redistribute them on the tissue
(Wigglesworth, 1959;
Wigglesworth, 1977) (E.
Johnson and M.A.K., unpublished). Thus, there is a physiological mechanism
that can release substrate attachments without leading to branch destruction
(Fig. 6, `remodeling'). It will
be interesting to investigate how integrin-talin adhesion complexes are
modified during this process and during the much more extensive remodeling of
the tracheal network that occurs during metamorphosis.
Branch and lumen maintenance in other tubular networks
Our finding that tracheal terminal branches are actively maintained by an
integrin-talin adhesion system raises the issue of whether branches and lumens
in other tubular networks also require active maintenance. A parallel is found
in aging C. elegans, where renal tubules that lose contact with their
substratum degenerate in a manner similar to that described here for mutant
terminal branches: the distal part of the branch retracts while the lumen is
retained as a convoluted tangle in the proximal part of the cell
(Buechner, 2002). Disruption
of integrin complexes and substrate adhesion in MDCK or endothelial cell in
vitro tubulogenesis assays can cause misplacement or absence of lumens
(Bayless et al., 2000;
Yu et al., 2005), and mutant
mice lacking specific integrins or ECM molecules have a variety of tubular
defects, some of which might be due to defects in branch or lumen maintenance
(McCarty et al., 2002;
Wang et al., 2006;
Zhu et al., 2002). It will be
important to determine the molecular mechanisms of tube maintenance in other
systems, and to define the roles that substrate adhesion systems play in
maintaining branches and their luminal organization, and destabilizing them
during network remodeling events.
|
ACKNOWLEDGMENTS |
|---|
|
REFERENCES |
|---|
|
TOPSUMMARYINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
|---|
Affolter, M., Bellusci, S., Itoh, N., Shilo, B., Thiery, J. P. and Werb, Z. (2003). Tube or not tube: remodeling epithelial tissues by branching morphogenesis. Dev. Cell 4, 11-18.[CrossRef][Medline]
Bayless, K. J., Salazar, R. and Davis, G. E.
(2000). RGD-dependent vacuolation and lumen formation observed
during endothelial cell morphogenesis in three-dimensional fibrin matrices
involves the alpha(v)beta(3) and alpha(5)beta(1) integrins. Am. J.
Pathol. 156,1673
-1683.
Becam, I. E., Tanentzapf, G., Lepesant, J. A., Brown, N. H. and Huynh, J. R. (2005). Integrin-independent repression of cadherin transcription by talin during axis formation in Drosophila. Nat. Cell Biol. 7,510 -516.[CrossRef][Medline]
Berger, J., Suzuki, T., Senti, K. A., Stubbs, J., Schaffner, G. and Dickson, B. J. (2001). Genetic mapping with SNP markers in Drosophila. Nat. Genet. 29,475 -481.[CrossRef][Medline]
Billuart, P., Winter, C. G., Maresh, A., Zhao, X. and Luo, L. (2001). Regulating axon branch stability: the role of p190 RhoGAP in repressing a retraction signaling pathway. Cell 107,195 -207.[CrossRef][Medline]
Bokel, C. and Brown, N. H. (2002). Integrins in development: moving on, responding to, and sticking to the extracellular matrix. Dev. Cell 3,311 -321.[CrossRef][Medline]
Borowsky, M. L. and Hynes, R. O. (1998).
Layilin, a novel talin-binding transmembrane protein homologous with C-type
lectins, is localized in membrane ruffles. J. Cell
Biol. 143,429
-442.
Boube, M., Martin-Bermudo, M. D., Brown, N. H. and Casanova,
J. (2001). Specific tracheal migration is mediated by
complementary expression of cell surface proteins. Genes
Dev. 15,1554
-1562.
Bradley, P. L., Myat, M. M., Comeaux, C. A. and Andrew, D. J. (2003). Posterior migration of the salivary gland requires an intact visceral mesoderm and integrin function. Dev. Biol. 257,249 -262.[CrossRef][Medline]
Brown, N. H., Gregory, S. L. and Martin-Bermudo, M. D. (2000). Integrins as mediators of morphogenesis in Drosophila. Dev. Biol. 223,1 -16.[CrossRef][Medline]
Brown, N. H., Gregory, S. L., Rickoll, W. L., Fessler, L. I., Prout, M., White, R. A. and Fristrom, J. W. (2002). Talin is essential for integrin function in Drosophila. Dev. Cell 3,569 -579.[CrossRef][Medline]
Buechner, M. (2002). Tubes and the single C. elegans excretory cell. Trends Cell. Biol. 12,479 -484.[CrossRef][Medline]
Bunch, T. A., Salatino, R., Engelsgjerd, M. C., Mukai, L., West, R. F. and Brower, D. L. (1992). Characterization of mutant alleles of myospheroid, the gene encoding the beta subunit of the Drosophila PS integrins. Genetics 132,519 -528.[Abstract]
Calderwood, D. A. and Ginsberg, M. H. (2003). Talin forges the links between integrins and actin. Nat. Cell Biol. 5,694 -697.[CrossRef][Medline]
Calderwood, D. A., Zent, R., Grant, R., Rees, D. J., Hynes, R.
O. and Ginsberg, M. H. (1999). The Talin head domain binds to
integrin beta subunit cytoplasmic tails and regulates integrin activation.
J. Biol. Chem. 274,28071
-28074.
Chou, T. B. and Perrimon, N. (1996). The autosomal FLP-DFS technique for generating germline mosaics in Drosophila melanogaster. Genetics 144,1673 -1679.[Abstract]
Cram, E. J. and Schwarzbauer, J. E. (2004). The talin wags the dog: new insights into integrin activation. Trends Cell. Biol. 14,55 -57.[CrossRef][Medline]
Devenport, D. and Brown, N. H. (2004).
Morphogenesis in the absence of integrins: mutation of both Drosophila beta
subunits prevents midgut migration. Development
131,5405
-5415.
Devine, W. P., Lubarsky, B., Shaw, K., Luschnig, S., Messina, L.
and Krasnow, M. A. (2005). Requirement for chitin
biosynthesis in epithelial tube morphogenesis. Proc. Natl. Acad.
Sci. USA 102,17014
-17019.
Garcia-Alvarez, B., de Pereda, J. M., Calderwood, D. A., Ulmer, T. S., Critchley, D., Campbell, I. D., Ginsberg, M. H. and Liddington, R. C. (2003). Structural determinants of integrin recognition by talin. Mol. Cell 11,49 -58.[CrossRef][Medline]
Ghabrial, A., Luschnig, S., Metzstein, M. M. and Krasnow, M. A. (2003). Branching morphogenesis of the Drosophila tracheal system. Annu. Rev. Cell Dev. Biol. 19,623 -647.[CrossRef][Medline]
Green, K. A. and Lund, L. R. (2005). ECM degrading proteases and tissue remodelling in the mammary gland. BioEssays 27,894 -903.[CrossRef][Medline]
Guillemin, K., Groppe, J., Ducker, K., Treisman, R., Hafen, E., Affolter, M. and Krasnow, M. A. (1996). The pruned gene encodes the Drosophila serum response factor and regulates cytoplasmic outgrowth during terminal branching of the tracheal system. Development 122,1353 -1362.[Abstract]
Hogan, B. L. and Kolodziej, P. A. (2002). Organogenesis: molecular mechanisms of tubulogenesis. Nat. Rev. Genet. 3,513 -523.[CrossRef][Medline]
Hynes, R. O. and Zhao, Q. (2000). The evolution
of cell adhesion. J. Cell Biol.
150,F89
-F96.
Inai, T., Mancuso, M., Hashizume, H., Baffert, F., Haskell, A.,
Baluk, P., Hu-Lowe, D. D., Shalinsky, D. R., Thurston, G., Yancopoulos, G. D.
et al. (2004). Inhibition of vascular endothelial growth
factor (VEGF) signaling in cancer causes loss of endothelial fenestrations,
regression of tumor vessels, and appearance of basement membrane ghosts.
Am. J. Pathol. 165,35
-52.
Jannuzi, A. L., Bunch, T. A., Brabant, M. C., Miller, S. W.,
Mukai, L., Zavortink, M. and Brower, D. L. (2002). Disruption
of C-terminal cytoplasmic domain of betaPS integrin subunit has dominant
negative properties in developing Drosophila. Mol. Biol.
Cell 13,1352
-1365.
Jarecki, J., Johnson, E. and Krasnow, M. A. (1999). Oxygen regulation of airway branching in Drosophila is mediated by branchless FGF. Cell 99,211 -220.[CrossRef][Medline]
Larsen, C. W., Hirst, E., Alexandre, C. and Vincent, J. P.
(2003). Segment boundary formation in Drosophila embryos.
Development 130,5625
-5635.
Lee, M., Lee, S., Zadeh, A. D. and Kolodziej, P. A.
(2003). Distinct sites in E-cadherin regulate different steps in
Drosophila tracheal tube fusion. Development
130,5989
-5999.
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]
Lee, T. and Luo, L. (1999). Mosaic analysis with a repressible cell marker for studies of gene function in neuronal morphogenesis. Neuron 22,451 -461.[CrossRef][Medline]
Liu, S., Calderwood, D. A. and Ginsberg, M. H. (2000). Integrin cytoplasmic domain-binding proteins. J. Cell Sci. 113,3563 -3571.[Abstract]
Lubarsky, B. and Krasnow, M. A. (2003). Tube morphogenesis: making and shaping biological tubes. Cell 112,19 -28.[CrossRef][Medline]
Manning, G. and Krasnow, M. (1993). Development of the Drosophila tracheal system. In The Development of Drosophila melanogaster. Vol. 1 (ed. M. Bate and A. Martinez-Arias), pp. 609-686. Cold Spring Harbor: Cold Spring Harbor Laboratory Press.
McCarty, J. H., Monahan-Earley, R. A., Brown, L. F., Keller, M.,
Gerhardt, H., Rubin, K., Shani, M., Dvorak, H. F., Wolburg, H., Bader, B. L.
et al. (2002). Defective associations between blood vessels
and brain parenchyma lead to cerebral hemorrhage in mice lacking alphav
integrins. Mol. Cell. Biol.
22,7667
-7677.
Noirot, C. and Noirot-Thimotee, C. (1982). The structure and development of the tracheal system. In Insect Ultrastructure. Vol. 1 (ed. R. C. King and H. Akai), pp. 351-381. New York: Plenum Press.
O'Brien, L. E., Zegers, M. M. and Mostov, K. E. (2002). Building epithelial architecture: insights from three-dimensional culture models. Nat. Rev. Mol. Cell Biol. 3,531 -537.[CrossRef][Medline]
Patel, N. (1994). Imaging neuronal subsets and other cell types in whole mount Drosophila embryos and larvae using antibody probes. In Methods in Cell Biology, Drosophila melanogaster: Practical Uses in Cell Biology (ed. L. S. B. Goldstein and E.A. Fyrberg), Vol. 44, pp.416 -487. New York: Oxford University Press.
Prokop, A., Martin-Bermudo, M. D., Bate, M. and Brown, N. H. (1998). Absence of PS integrins or laminin A affects extracellular adhesion, but not intracellular assembly, of hemiadherens and neuromuscular junctions in Drosophila embryos. Dev. Biol. 196,58 -76.[CrossRef][Medline]
Prout, M., Damania, Z., Soong, J., Fristrom, D. and Fristrom, J. W. (1997). Autosomal mutations affecting adhesion between wing surfaces in Drosophila melanogaster. Genetics 146,275 -285.[Abstract]
Risau, W. and Flamme, I. (1995). Vasculogenesis. Annu. Rev. Cell Dev. Biol. 11, 73-91.[CrossRef][Medline]
Rühle, H. (1932). Das larvale tracheensystem von Drosophila melanogaster Meigen und seine variabilität. Z. Wiss. Zool. 141,159 -245.
Samakovlis, C., Hacohen, N., Manning, G., Sutherland, D. C., Guillemin, K. and Krasnow, M. A. (1996). Development of the Drosophila tracheal system occurs by a series of morphologically distinct but genetically coupled branching events. Development 122,1395 -1407.[Abstract]
Speck, O., Hughes, S. C., Noren, N. K., Kulikauskas, R. M. and Fehon, R. G. (2003). Moesin functions antagonistically to the Rho pathway to maintain epithelial integrity. Nature 421, 83-87.[CrossRef][Medline]
Sutherland, D., Samakovlis, C. and Krasnow, M. A. (1996). branchless encodes a Drosophila FGF homolog that controls tracheal cell migration and the pattern of branching. Cell 87,1091 -1101.[CrossRef][Medline]
Tepass, U., Truong, K., Godt, D., Ikura, M. and Peifer, M. (2000). Cadherins in embryonic and neural morphogenesis. Nat. Rev. Mol. Cell Biol. 1, 91-100.[CrossRef][Medline]
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]
Walsh, P. S., Metzger, D. A. and Higuchi, R. (1991). Chelex 100 as a medium for simple extraction of DNA for PCR-based typing from forensic material. Biotechniques 10,506 -513.[Medline]
Wang, J., Hoshijima, M., Lam, J., Zhou, Z., Jokiel, A., Dalton,
N. D., Hultenby, K., Ruiz-Lozano, P., Ross, J., Jr, Tryggvason, K. et al.
(2006). Cardiomyopathy associated with microcirculation
dysfunction in laminin alpha4 chain-deficient mice. J. Biol.
Chem. 281,213
-220.
Watts, R. J., Schuldiner, O., Perrino, J., Larsen, C. and Luo, L. (2004). Glia engulf degenerating axons during developmental axon pruning. Curr. Biol. 14,678 -684.[CrossRef][Medline]
Whitten, J. (1957). The post-embryonic development of the tracheal system in Drosophila melanogaster. Q. J. Microsc. Sci. 98,123 -150.
Wigglesworth, V. B. (1959). The role of epidermal cells in the migration of tracheoles in Rhodnius prolixus (Hemiptera). J. Exp. Biol. 36,632 -640.[Abstract]
Wigglesworth, V. B. (1972). The Principles of Insect Physiology. London: Chapman & Hall.
Wigglesworth, V. B. (1977). Structural changes in the epidermal cells of Rhodnius during tracheole capture. J. Cell Sci. 26,161 -174.[Abstract]
Wigglesworth, V. B. and Lee, W. M. (1982). The supply of oxygen to the flight muscles of insects: a theory of tracheole physiology. Tissue Cell 14,501 -518.[Medline]
Yu, W., Datta, A., Leroy, P., O'Brien, L. E., Mak, G., Jou, T.
S., Matlin, K. S., Mostov, K. E. and Zegers, M. M. (2005).
ß1-integrin orients epithelial polarity via rac1 and laminin.
Mol. Biol. Cell 16,433
-445.
Zelzer, E. and Shilo, B. Z. (2000). Cell fate choices in Drosophila tracheal morphogenesis. BioEssays 22,219 -226.[CrossRef][Medline]
Zhu, J., Motejlek, K., Wang, D., Zang, K., Schmidt, A. and Reichardt, L. F. (2002). beta8 integrins are required for vascular morphogenesis in mouse embryos. Development 129,2891 -2903.[Medline]
CiteULike 
Complore 
Connotea 
Del.icio.us 
Digg 
Reddit 
Technorati 
Twitter What's this?
Related articles in Development:
This article has been cited by other articles:
|
|
 |
|
  M. Affolter and E. Caussinus Tracheal branching morphogenesis in Drosophila: new insights into cell behaviour and organ architecture Development, June 15, 2008; 135(12): 2055 - 2064. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 A. Khokhar, N. Chen, J.-P. Yuan, Y. Li, G. N. Landis, G. Beaulieu, H. Kaur, and J. Tower Conditional Switches for Extracellular Matrix Patterning in Drosophila melanogaster Genetics, March 1, 2008; 178(3): 1283 - 1293. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
M. M. Baer, A. Bilstein, and M. Leptin A Clonal Genetic Screen for Mutants Causing Defects in Larval Tracheal Morphogenesis in Drosophila Genetics, August 1, 2007; 176(4): 2279 - 2291. [Abstract] [Full Text] [PDF]
|
|
| |||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||
