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First published online 13 December 2006
doi: 10.1242/dev.02728
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Instituto de Biología Molecular de Barcelona, Consejo Superior de Investigaciones Científicas, Parc Cientific de Barcelona, Josep Samitier 1-5, Barcelona 08028, Spain.
Author for correspondence (e-mail:
embbmc{at}cid.csic.es)
Accepted 2 November 2006
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SUMMARY |
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 TOP SUMMARY INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Key words: Drosophila, Metamorphosis, Histoblasts, Morphogenesis, Abdomen, Cell death, Cell replacement
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INTRODUCTION |
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TOPSUMMARYINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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The movement and fusion of epithelial cells during development is an
essential and general morphogenetic event. Many different morphogenetic
processes, both in vertebrates and invertebrates, and the related processes of
adult and embryonic vertebrate wound healing, could be included in this
category (reviewed by Martin and Wood,
2002
). However, although extensive morphological descriptions of
these processes represent classical paradigms of embryology, the genetic basis
and the cellular behaviour underlying these events remain poorly understood.
Thorough studies have shown that to undertake these processes epithelial cells
can employ two alternative approaches: first, leading edge cells facing a free
cellular space become specified and promote the migration of epithelial sheets
acting as the main driving force for migration (in most cases in the absence
of cell proliferation), e.g. Caenorhabditis elegans ventral enclosure
(Williams-Masson et al., 1997)
and imaginal discs fusion in Drosophila
(Pastor-Pareja et al., 2004).
Second, overall synchronous action of the whole epithelial cell population can
induce global changes in its motility, conforming tissue shape changes, e.g.
teleost epiboly (Trinkaus et al.,
1992) and extension of the neural plate in Xenopus
(Keller et al., 2000).
Two models of unbounded epithelial expansion, in which epithelial sheets
with a free leading edge advance, meet and fuse to equivalent epithelial
sheets in order to seal the surface of the animal, have been described in
Drosophila. These models are dorsal closure and the fusion of
imaginal discs. Dorsal closure is a late embryonic event that begins with the
elongation of the epidermal cells and finishes when the dorsalmost cells fuse
at the midline (reviewed by Harden,
2002). The dimensions of the epidermal layer are not controlled by
proliferation but are the result of changes in cell adhesion and cell shapes.
Cells at the leading front that are planary polarized guide the migration of
the epithelium, express specific markers and have a very active cytoskeleton.
Dorsal closure depends on at least two signalling pathways - the JNK signal
transduction pathway and the signalling pathway activated by Dpp
(Martin-Blanco et al., 1998;
Riesgo-Escovar and Hafen,
1997; Kockel et al.,
1997). These two pathways are also involved in the expansion and
merging of excorporated imaginal tissues (imaginal discs) at the expense of
larval cells during metamorphosis (reviewed by
Martin-Blanco and Knust,
2001). The imaginal discs evert and later on differentiate to give
rise to the external structures of the adult. In this process the peripheral
cells of the disc expand over the larval cells and gradually displace them.
Finally, the discs recognize and fuse with each other to complete the
continuous epithelium that will give rise to the head and the thorax of the
adult.
Much less is known about the mechanisms controlling the movements of
bounded epithelia without a free edge (e.g. the invagination of the basal
region during sea urchin gastrulation, extension of the neural plate,
rearrangement of scale precursors in the wings of insects, etc.) (see
Bard, 1990). A potential model
for the study of the coordinated expansion of this type of epithelia is the
development of the adult abdomen of Drosophila during metamorphosis.
The adult epidermis is formed by cells descending from histoblasts, founder
imaginal cells specified during embryonic stages as small incorporated groups
organized in nests. Each adult abdominal segment forms from four pairs of
histoblast nests: the anterior and posterior dorsal pairs (which produce the
tergites), the ventral pairs (which produce the sternites and pleurites) and
the spiracular pairs (which form the spiracle and the surrounding pleurite
tissues). Each anterior dorsal and ventral nest is composed of approximately
16 cells; each posterior dorsal nest consists of approximately five cells; and
each spiracle nest has approximately three cells
(Fig. 1A)
(Guerra et al., 1973;
Roseland and Schneiderman,
1979; Madhavan and Madhavan,
1980). During larval stages histoblasts are mitotically quiescent
and arrested in G2 (Garcia-Bellido and
Merriam, 1971; Madhavan and
Schneiderman, 1977; Roseland
and Schneiderman, 1979). At the onset of metamorphosis histoblasts
undertake a rapid proliferation that allows them to expand and fuse, at the
expense of the preexisting surrounding polyploid larval epidermal cells (LECs)
that commit programmed cell death
(Madhavan and Madhavan,
1980).
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MATERIALS AND METHODS |
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TOPSUMMARYINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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); Esg-Gal4 Ay+Gal4 UAS-GFP UAS-FLP (y, w; NP5130,
Act FRT y+ FRT Gal4, UAS-GFP/CyO; UAS-FLP/TM6B); Hs-Gal4
(w; Hsp70 Gal4.PB 89-2-1) (Bloomington Stock Center 1799); UAS-FLP
(y, w; UAS-FLP.Exel 3) (Bloomington Stock Center 8209); 056-Gal4 -
Enhancer trap ubiquitously expressed in the epidermis (y, w; NP0056)
(NIG-FLY Stock Center); Ay+Gal4 UAS-GFP (y, w; Act FRT y+ FRT
Gal4, UAS-GFP) (Bloomington Stock Center 4411); Srp-Gal4 UAS-GFP
[srpHemo-Gal4#16 A3B, UAS-Src-EGFP (II)]
(Bruckner et al., 2004);
Sqh-GFP (y w sqhAV3 cv; sqh-GFP42)
(Royou et al., 2004);
UAS-RokCAT (Winter et
al., 2001); UAS-P35 [w; UAS-p35.H BH1 (II)] (Bloomington
Stock Center 5072); UAS-Chickadee (L. Cooley, Yale University Medical School,
Department of Genetics, New Haven, CT, USA); UAS-MbsN300
(Lee and Treisman, 2004);
UAS-Dap (UAS-Dap II.3) (Lane et
al., 1996); UAS-EcR-RNAi (UAS-EcR-AB RNAi)
(Schubiger et al., 2005);
UAS-EcRDN (w; UAS-EcR.B1-DeltaC655.W650A) (Bloomington
Stock Center 6872); UAS-RacV12 (y, w;
UAS-Rac1V12) (Bloomington Stock Center 6291); Stb-YFP (w;
Act Stb-YFP/TM3) (Strutt et al.,
2002); H2-YFP(III) [His2avD-EYFP(II)]
(Rebollo et al., 2004); Hs-FLP
Ay+Gal4 UAS-GFP H2YFP (hsp70-flp; Act FRT y+ FRT Gal4,
UAS-GFP/CyO; H2YFP/TM2); Hs-FLP Ay+Gal4 UAS-GFP Stb-YFP (hsp70-flp;
Act FRT y+ FRT Gal4, UAS-GFP/CyO; Act Stb-YFP/TM2);
UAS-Src-GFP (w; UAS-SrcEGFP M7E) (Bloomington Stock Center 5432);
UAS-Actin-GFP (w; UAS-Act5C.GFP 2) (Bloomington Stock Center 7310);
DE-Cadherin-GFP [Ubi-DE-Cad-GFP (II)]
(Oda and Tsukita, 2001);
UAS-Tau-GFP (w; UAS-GFP.btau)
(Brand, 1995).
Targeted expression in histoblasts
The Gal4/UAS system (Brand and Perrimon,
1993) was used for targeted expression in the histoblasts. The
Esg-Gal4 was used as a histoblast-specific driver, although expression of this
driver can also be detected in salivary glands, wing discs, eye discs and
gut.
Permanent expression of UAS constructs in histoblasts
The Esg-Gal4 driver expression declines during late pupal stages. To
overcome this problem we took advantage of the UAS-FLP/Actin-FRT-y+-FRT-Gal4
system (Struhl and Basler,
1993) by raising flies with the genotype y, w; NP5130, Act FRT
y+ FRT Gal4, UAS-GFP/CyO; UAS-FLP/TM6B. The larval expression
of Esg-Gal4 triggers activation of the UAS-FLP transgene, which promotes the
cis-recombination of FRT sites in the Actin-FRT-y+-FRT-Gal4 cassette
generating a FLP-out event in histoblast cells. As a result, the Gal4 gene
comes under the control of the Actin promoter, which promotes its permanent
expression in histoblasts.
Time- and tissue-specific expression of UAS constructs in LECs
To achieve a tissue- and time-specific expression of UAS constructs in LECs
we used a combination of the Gal4/UAS system and the FLP/FRT system by using
the hsp70-FLP; Act FRT y+ FRT Gal4, UAS-GFP/CyO; MRKS/TM2
strain (Ito et al., 1997).
This system has been used for the generation of positively marked clones in
diploid cells where the frequency of recombination is low
(Struhl and Basler, 1993). The
larval epidermal cells are polyploid and contain multiple copies of the
genome. As a result the recombination frequency between FRT sites is far
greater compared with diploid cells. A short heat shock treatment of 8-10
minutes leads to recombination, and hence the activation of a UAS-GFP
reporter, in 95-100% of the larval epidermal cells 5-7 hours after the heat
shock pulse. Heat shocks were performed during late larval stages (wandering
larva) when histoblasts are mitotically arrested. In this way, we were able to
express distinct UAS constructs in LECs although only small fractions of
histoblast were affected (data not shown).
Immunohistochemistry
Primary antibodies used were: rabbit anti-laminin A antiserum at 1:100
dilution (Fessler et al.,
1987), mouse anti-Fasciclin III (7G10-Hybridoma Bank) at 1:1000,
mouse anti-GFP at 1:500 (Cell Signalling), rabbit anti-active Caspase 3 at
1:100 (Cell Signalling), mouse anti-EcRB1 at 1:10 (AD4.4 Hybridoma Bank) and
mouse anti-EcRA at 1:10 (15G1A-Hybridoma Bank). Secondary antibodies were
anti-mouse or anti-rabbit FITC, Cy3 or Cy5 conjugated (Molecular Probes) at
1:250 dilutions. Immunohistochemistry was performed using standard procedures.
For pupal staging, white pupae [0 hours after puparium formation (APF)] were
used as reference. After selection, the white puparium prepupa were
transferred to fresh vials and kept at 25°C and standard humidity up to
dissection. Whole pupae were bisected along the anteroposterior axis in
sterilized 1x PBS (pH 7.4). The internal organs were cleaned from the
epidermis by flushing with 1x PBS using a P10 pipette. The epidermis was
detached from the pupal case using forceps and transferred to an ependorff
tube on ice. Fixation was performed for 10 or 15 minutes in 4%
paraformaldehyde. After fixation, the epidermis was rinsed three times in
1x PBS and permeabilized in sterilized PBT (0.3% Triton in 1x PBS)
(3x15 minutes). After permeabilization, the tissue was blocked for 1
hour using PBTB [1% bovine serum albumin (BSA) in PBT]. Primary antibodies
were incubated overnight at 4°C with gentle shaking. The epidermis was
rinsed in 1x PBS, and washed 3x15 minutes in PBTB. After 1 hour
blocking in PBTB, the secondary antibody was incubated for 3 hours at room
temperature. After rinsing in 1x PBS, the tissue was stained using DAPI
(1 ng/µl) to mark the nuclei and Rhodamine-coupled Phalloidin (Molecular
Probes) at a dilution of 1:1000 from a 1 µg/µl stock solution was used
to visualize polymerized actin. Finally the tissue was washed 3x15
minutes in 1x PBS, equilibrated in Vectashield (Vector) and mounted on
coverslips. Actin staining using Phalloidin alone was performed as above after
10 minutes fixation and omitting the blocking steps.
In vivo imaging and time-lapse microscopy
Staged pupae and prepupae were washed in PBS. Early pupae were directly
imaged through the transparent pupal case. For late pupal imaging (after 12
hours APF), a small window was opened into the pupal case on top of abdominal
segments 2 and 3 by careful surgery with a fine needle. At this stage the
pupal case is detached from the epidermis and can be removed without
disturbing the underlying epidermis. The animals were positioned on a glass
bottom microwell dish (MatTek) in a small drop of Voltaleff oil to improve
optics and to avoid desiccation. Images were captured at different time
intervals using an inverted Leica TCS 4D confocal microscope or an inverted
Leica AOBS confocal microscope. Laser intensity was kept at a minimum to avoid
photobleaching and to minimize phototoxicity. Each movie was repeated at least
three times. In most cases, the animal survived the dissection and data
acquisition and developed to adult stages.
In vivo quantification of cell proliferation rate
Cell proliferation was recorded using confocal time-lapse imaging in the
posterior dorsal histoblast nest from 17 hours APF pupae of the genotypes
Hs-FLP; Ay+Gal4 UAS-GFP; H2-YFP or Hs-FLP; Ay+Gal4
UAS-GFP/UAS-P35; H2YFP/+, where the expression of the P35 transgene was
activated by heat shock during late larval stages. To minimize stress during
image acquisition the pupal case was not removed and imaging was performed
directly through the puparium at 10 minute intervals. Individual cells were
followed from mitosis to mitosis using the ubiquitously expressed Histone2-YFP
fusion protein as a marker. The mitosis of mother cells were taken as time 0
and the consecutive division of one of the two daughter cells as time 1. Cell
doubling time was calculated for a time window of 6 hours (17-23 hours APF).
Doubling times for 10 cells in each experiment were counted.
Flow cytometry
To measure the cell cycle phasing of the abdominal histoblasts, whole pupal
cuticles (50 animals for each condition) were dissected, cleaned and then
subjected to an incubation in 9x Tripsin, 1x PBS, with 1 mg/ml
Hoechst 33342 for 3 hours at room temperature. Histoblasts were positively
marked by Esg-Gal4 expression driving UAS-GFP. Prepupae animals were staged as
less than 12 hours APF and greater than 3 hours APF. Early pupae animals were
determined as greater than 12 hours APF and less than 24 hours APF. In order
to compare cell cycle profiles, samples of each time point were prepared and
ran simultaneously. We used a MoFlo flow cytometer (DakoCytomation, Fort
Collins, Colorado, USA). Excitation was performed with an argon-ion laser of
Coherent Enterprise II and the optical alignment obtained with fluorescent
particles of a diameter of 10 µm (Flowcheck, Coulter Corporation, Miami,
Florida, USA). Different populations were defined combining green (GFP) and
blue (Hoechst 33342) emissions and the refringency parameters FSC and SSC.
Quantification of cell size
Cell size was calculated by dividing the number of cells in a histoblast
nest over the total area of the nest. The area of the nest was calculated from
z-stack projections covering the whole depth of the nest. Histoblasts
were visualized using Esg-Gal4 driving the expression of a UAS-nuclear-GFP and
UAS-cytoplasmic-GFP. Area measurements were carried out using ImageJ (NIH
Image).
Image analysis
Image analysis was performed with Leica Confocal Software, Imaris 5D
(Bitplane) software was used for 3D reconstruction of time-lapse movies,
ImageJ (NIH Image) for cell tracking and mounting of time-lapse movies in AVI
format, Photoshop 7.0 (Adobe Corporation) for data processing and Corel
R.A.V.E. for conversion of movies to QuickTime format.
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RESULTS |
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TOPSUMMARYINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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). Upon pupariation, histoblasts divide rapidly
but remain confined to their original dimensions. A second, slower histoblast
proliferation phase begins about 15 hours APF and lasts until about 36 hours
APF. During this period, the histoblast nests enlarge, and the histoblasts
replace the adjacent LECs. By time-lapse analysis, we validated these distinct
phases. We found that during prepupal stages, histoblasts underwent very fast
synchronous cell divisions, with a cell doubling time of around 2 hours
(Fig. 1B; see Movie 1 in the
supplementary material) and preferentially oriented spindles (Tau-GFP
labelling, see Fig. 1C). This
early stage lasted for three cycles and ran between 4 and 12 hours APF. A
characteristic feature of the prepupal proliferation of histoblasts is the
lack of cell growth between cycles, resulting in a rapid decrease in cell size
(Madhavan and Madhavan, 1980)
(Fig. 1E, left panel). We found
by FACS analysis that the short cell doubling time and lack of growth of
histoblasts isolated at this stage was associated with a very reduced G1 phase
(Fig. 1F).
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Histoblast nest expansion requires the active planar intercalation of histoblasts into the larval epithelia
During the prepupal highly proliferative stage the area occupied by each
histoblast nest did not significantly increase; as a consequence of this, as
revealed by cytoskeleton (Actin) and adherens junction (DE-Cadherin) markers,
histoblast nests became organized into pseudo stratified monolayers (see Fig.
S1A in the supplementary material). Subsequently, from 15 hours APF onwards,
in synchrony with histoblast cell cycle slowness, nests initiated expansion
and invaded the territories occupied by polyploid LECs. As an outcome, they
rapidly rearranged into unstratified epithelia (see Fig. S1B in the
supplementary material). The anterior and posterior dorsal nests fused into a
single hemitergite nest between 15 and 18 hours APF (see Fig. S1C and Movie 2
in the supplementary material). Hemitergite nests from adjacent segments began
to fuse at about 18 hours APF. The hemisternite (ventral) histoblast nest and
the spiracular anlagen joined up between 18 and 22 hours APF, while the
hemitergite histoblast nest and the spiracular anlagen joined up between 22
and 26 hours APF. The process was completed upon the fusion of left and right
nests at the dorsal midline by 36 hours APF
(Madhavan and Madhavan, 1980)
(and data not shown).
To gain insight into the mechanisms involved in histoblast nest expansion, we monitored this process in vivo. This analysis revealed that at its onset, nest spreading proceeded through the intercalation of guiding histoblasts into the surrounding larval epidermal palisade (Fig. 2A; see Movie 3 in the supplementary material). To do so, invading peripheral histoblasts extended dynamic cellular protrusions in between neighbouring LECs that, by anchoring and shrivelling, promoted traction and the forward movement of histoblast cell bodies (Fig. 2B; see Movie 4 in the supplementary material). These structures were actin-rich and developed by effective actin polymerization at their tips, resembling actin comets (Fig. 2C,D; see Movie 5 in the supplementary material). The initial planar intercalation was followed by the coordinated expansion of the whole nest epithelia. Every cell at the edge of the nests weakly, but reproducibly, downregulated adherens junction markers (see Fig. S1C and Movie 2 in the supplementary material) and emitted both apical and basal filopodia and lamellipodia, which expanded over the surface of the LECs (Fig. 2E) and actively advanced over the underlying extracellular matrix (ECM) in the direction of migration (Fig. 2F). Nest expansion progressed centrifugally for several hours up to the merge of the adjacent ipsilateral and contralateral nests (data not shown). Finally, the sealing of the epithelia proceeded by the assembly of an apical purse string that brought together the apices of the leading cells (Fig. 2G; see Movie 6 in the supplementary material).
Altogether, these results reveal a complex, multistep process that involves
the invasion of the larval epithelia by histoblasts (planar intercalation)
mediated by active cellular actin-rich protrusions. To test this model we
interfered with actin polymerization in histoblasts. During
Drosophila bristle morphogenesis, the activities of the products of
the cpb (Capping protein beta) and chickadee (Profilin)
genes hold a tight balance between actin depolymerization and assembly
(Hopmann and Miller, 2003).
Loss of Cpb or excessive Profilin activity resulted in actin accumulations and
abnormal bristles. In this light, we overexpressed Profilin in histoblasts
under the control of a permanent Esg-Gal4 driver (see Materials and methods).
The overexpression of Profilin disrupted bristle morphogenesis, as expected,
and caused the inefficient expansion of histoblast nests, provoking abdominal
clefts (Fig. 3A). In this
condition, actin aggregated within histoblasts (compare
Fig. 3B with 3C) and, in
contrast to wild-type nests, where numerous long protrusions formed during
expansion, Profilin-overexpressing histoblasts lacked such structures and just
exhibited rare short filopodia and lamellipodia
(Fig. 3D; see Movie 7 in the
supplementary material). Moreover, the transition of histoblast nests from
pseudo stratified epithelia to an unstratified monolayer was delayed (data not
shown). Thus, the inability of histoblasts to resort to long cellular
protrusions resulted in the delay of their expansion and migration, suggesting
that the actin-mediated planar intercalation of histoblasts between LECs is
essential for their proper expansion.
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What is the mechanism for basal extrusion of LECs? Are the histoblasts
pushing away the polyploid LECs (Fig.
2B)? Or are pulling contractile forces from the LECs driving nest
expansion? Or a combination of both? To analyse these issues we evaluated
actomyosin dynamics during the expansion of histoblast nests. Myosin II
regulatory light chain (Spaghetti squash)
(Royou et al., 2004)
accumulated as perimetral rings in the apical domain of LECs. Assessment of
video time-lapse recordings revealed that LEC extrusion was initiated by
apical constriction, apparently mediated by actomyosin contraction
(Fig. 4C; see Movie 9 in the
supplementary material). The apical constriction of LECs seemed to be a
cell-autonomous event, as it could be observed occasionally in LECs without
direct contact with the expanding histoblasts but several cell diameters
away.
Non-muscle myosin II is a hexamer composed of two of each of three
subunits: the heavy chain, the regulatory light chain (MRLC) and the essential
light chain (Korn and Hammer,
1988). The force-generating activity of actomyosin is mainly
controlled through the phosphorylation of the MRLC
(Craig et al., 1983). Two
kinases, Rho-kinase (Rok; also known as Rock)
(Winter et al., 2001) and
Myosin light chain kinase (MLCK)
(Totsukawa et al., 2000)
phosphorylate MRLC in both mammals and Drosophila and activate myosin
contraction. By contrast, dephosphorylation of MRLC by myosin light chain
phosphatase (MLCP) inhibits myosin activity. This serine/threonine protein
phosphatase is a heterotrimer consisting of a catalytic subunit, a 20 kDa
protein of unknown function, and the myosin binding subunit (MBS) that targets
MLCP to MRLC (Fukata et al.,
1998). Phosphorylation by Rok of a specific threonine within a
conserved motif in MBS has been shown to inhibit MLCP activity, suggesting
that Rok activates MRLC both by direct phosphorylation and also by inhibition
of MBS (Kawano et al.,
1999).
To test whether myosin contraction was required for cell delamination, we
overexpressed in LECs a truncated constitutively active form of the
Drosophila MBS orthologue (MbsN300) that cannot be phosphorylated by
Rok (generalized clonal expression of UAS-MbsN300; see Materials and methods)
and inhibits Myosin contractility (Lee and
Treisman, 2004). In this condition, a significant proportion of
LECs did not extrude and remained within the abdominal epidermis at
postmetamorphosis periods (Fig.
4D). As a consequence, cuticular clefts were observed in the
abdomen (Fig. 4E). Further, we
found that the overexpression of a constitutively active form of Rok
(RokCAT) (Winter et al.,
2001), which upregulates myosin contractility, led to premature
delamination of LECs. In wild-type animals, LECs extrude out progressively at
a very stereotyped speed. Thus, out of 70±2 LECs found in the dorsal
posterior compartment of abdominal segments in third instar larvae, only
14±2 cells remained in place at 24 hours APF. The targeted
overexpression of RokCAT in these cells
(En-Gal4/UAS-RokCAT) accelerated their delamination and, in the
same period and starting from the same number of original cells, only three to
five remained within the epidermal layer. Altogether, these results strongly
suggest that the apical constriction of LECs plays a significant role at the
initiation of the cell replacement process and that apical actomyosin
contractility is both necessary and sufficient for the extrusion of LECs.
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). In
Drosophila, the phagocytic function is performed by circulating
haemocytes (Tepass et al.,
1994), and we found that Serpent-expressing haemocytes densely
populated the basal lamina of the abdominal epidermis
(Fig. 5A). Is the
actomyosin-driven apical contraction the only significant activity resulting
in the rapid elimination of LECs? Close examination of the process of
replacement revealed that, during their apical constriction, LECs became
surrounded on their basal side by bright fluorescent motile bodies, which we
identified as haemocytes (Fig.
5B). Double-labelling video time-lapse using Srp-Gal4 driving GFP
expression in haemocytes and a Stb-YFP apical membrane epithelial marker
ubiquitously expressed showed that the extrusion of LECs was a coordinated
process in which apical constriction was immediately followed by encapsulation
from the basolateral surface by haemocytes
(Fig. 5C; see Movie 10 in the
supplementary material). No cell blebbing was observed before extrusion, but
LEC nuclei collapsed and then became fragmented upon engulfment, turning
TUNEL-positive (data not shown). Is thus corpse removal by haemocytes required
for the expansion of histoblast nests and abdominal morphogenesis? To assess
this issue we genetically blocked the migratory capacity of haemocytes by
expressing a constitutively active form of the small GTPase Rac
(RacV12) (Paladi and Tepass,
2004). The expression of RacV12 under the Srp-Gal4
driver resulted in a 100% penetrant absence of Srp-Gal4/UAS-GFP-expressing
haemocytes from the abdomen at 24 hours APF (data not shown). The inability of
haemocytes to migrate into the abdomen generated late pupal lethality;
nonetheless, the morphogenesis of the adult abdomen proceeded relatively
normally (data not shown). Overexpression of Ricin or a dominant-negative form
of Pvr, which is necessary for haemocyte survival, led to similar phenotypes,
albeit not 100% penetrant. These results suggest that haemocytes are
dispensable for the expansion of the histoblast nests at the expense of
LECs.
Ecdysone signalling is required for both the proliferation of histoblasts and the death of LECs
The larval-prepupal transition of Drosophila is associated with a
peak of ecdysteroids that terminates larval feeding, initiates premetamorphic
behaviours and commits larval tissues to a pupal fate. A second Ecdysone
pulse, approximately 12 hours APF, defining the prepupal-to-pupal transition,
follows this larval peak (Riddiford,
1993). In Drosophila, the response of tissues to Ecdysone
is mediated by a heterodimer of the Ecdysone receptor (EcR) and Usp, both
nuclear receptor proteins (Koelle et al.,
1991; Yao et al.,
1992). Ecdysone receptors directly induce the transcription of
primary-response genes. Some of these early genes encode transcription factors
that regulate large batteries of secondary-response late genes (reviewed by
Thummel, 2001). The
EcR gene encodes three isoforms: EcR-A, EcR-B1 and EcR-B2, which
share common DNA- and ligand-binding domains but differ in their N-terminal
sequences. During larval stages, imaginal discs express high levels of EcR-A,
whereas most larval tissues and the imaginal histoblasts predominantly express
EcR-B1 (Talbot et al., 1993).
This pattern changes upon pupariation, and histoblasts and LECs then express
both EcR-A and EcR-B1 isoforms (see Fig. S2 in the supplementary
material).
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) or a
truncated version of its receptor (Cherbas
et al., 2003) either in histoblasts (Esg-Gal4) or in the LECs at
the onset of metamorphosis. Contrary to published reports
(Bender et al., 1997), blocking
autonomously Ecdysone signalling in histoblasts completely abolished their
proliferation up to the time of nest spreading
(Fig. 6A), when cell division
resumes. This yielded viable adults with variable abdominal defects (data not
shown). We interpret this variability as being a consequence of the decay of
the Esg-Gal4 driver expression (data not shown), which might fail to sustain
EcR inhibition. Conversely, clonal autonomous expression of a truncated
(dominant-negative) form of EcR in LECs completely blocked their ability to
apically constrict, extrude and undergo apoptosis. As a result, abdominal
morphogenesis did not succeed, and LECs were still present in the abdomen of
pharate adult flies, forming part of the epidermis
(Fig. 6B). We conclude that
Ecdysone signalling triggers the onset of proliferation in the histoblasts
upon pupariation and primes the death of the larval cells in coordination with
histoblast nest spreading.
Coordination of proliferation and cell death
During the early stages of histoblast proliferation, cell division takes
place in the absence of cell growth. By contrast, during late stages,
histoblasts couple proliferation with growth. Interestingly, the shift in the
histoblast cell cycle precedes the onset of LEC death, suggesting that
histoblast growth might influence non-autonomously the triggering of LEC
apoptosis.
To explore whether cell growth and cell death during cell sheet replacement
are coordinated and interdependent processes, we performed two types of
analysis. First, we blocked the proliferation of histoblasts and monitored the
non-autonomous effects on LECs. Strong inhibition of proliferation by
interfering with Ecdysone signal reception (see above) or by permanent
overexpression of Dacapo, an inhibitor of Cyclin E-Cdk2 (Cdc2c - FlyBase)
complexes and G1 to S progression (de
Nooij et al., 1996), in histoblasts
(Fig. 7A) resulted in a
profound delay in LEC death (Fig.
7D,E). In none of these cases was the developmental timing of
pupariation and unrelated processes, such as head and imaginal disc eversion,
affected (data not shown). Pharate adults escapers showed multiple
differentiation and morphogenetic abdominal defects (data not shown).
In addition, we prevented the death of LECs by interfering in the caspase cascade. The clonal expression of the apoptosis inhibitor P35 in LECs (see Materials and methods) resulted in LEC survival, impaired extrusion due to the partial inhibition of LEC apical constriction (Fig. 7B) and failure in the recruitment of haemocytes and the engulfment process. Those few LECs able to undergo extrusion in the presence of P35 remained viable cells (as judged by their nuclear and cellular morphology) under the epithelial layer (Fig. 7C).
In this condition, we found a significant decrease in the progression of histoblast nest spreading and the average number of histoblasts (Fig. 7F). These non-autonomous effects were a consequence of the delamination and death of numerous histoblasts (Fig. 7G), which does not occur in wild-type animals (data not shown). Histoblast death compensates and surpasses a weak non-autonomous enhancement of their mitotic index (Fig. 7H). Pharate adults could be recovered, but they showed abdominal cuticle clefts with many LECs still present (data not shown). These results strongly support a mechanism coordinating proliferation and spreading of histoblasts with the programmed death of LECs. Whether this mechanism relies on mutual signalling events, mechanical forces or cell competition for survival factors remains to be determined.
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DISCUSSION |
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TOPSUMMARYINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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), the
difficulty in visualizing morphogenetic changes in vivo and interfering with
signal reception in individual cells has become a major impediment in our
understanding of Ecdysone actions during metamorphosis.
|
). Other studies
have uncovered the ecdysteroid dependence of multiple differentiative and
maturational responses (Schubiger et al.,
1998). Nonetheless, little is known about ecdysteroid control of
cell proliferation. A revealing analysis in Manduca showed that
proliferating cells of the optic lobe reversibly arrest in G2 whenever the
concentration of ecdysteroid drops below a critical threshold
(Champlin and Truman, 1998).
Furthermore, earlier studies had shown that the number of histoblasts is
reduced in hypomorph mutants for EcR isoforms
(Bender et al., 1997). Whether
this was the result of lack of proliferation or cell death had not been
defined. Here, we show in vivo a direct role for Ecdysone in cell
proliferation (see Fig. 8). The
rapid cell cycles experienced by abdominal histoblasts at the end of the third
larval instar halt if Ecdysone-signalling reception is cell-autonomously
compromised (Fig. 6A).
Histoblasts remain quiescent in G2 and competent to resume proliferation in
response to late Ecdysone pulses. Our observations suggest that Ecdysone
signalling controls the cell cycle by regulating the expression of genes
involved in the G2-to-M transition (N.N., M. Grande and E.M.-B.,
unpublished).
The destruction of larval tissues in Drosophila also results from
a major transcriptional switch triggered by Ecdysone. The anterior larval
muscles and larval midgut (Lee et al.,
2002) and the head and thoracic LECs (N.N. and E.M.-B.,
unpublished) degenerate during the first half of prepupal development
(prepupal Ecdysone peak), while the larval salivary glands
(Jiang et al., 2000),
abdominal muscles and abdominal LECs (see
Fig. 5) histolyze after the
second Ecdysone pulse (pupation). Given that the exposure to Ecdysone is
systemic, the stage-specific cell death responses of different cell types to
Ecdysone must be differentially regulated.
The death of abdominal LECs shows apoptotic characters and proceeds in two steps: the basal extrusion of cells initiated by the contraction of an apical actomyosin ring, and their removal by haemocytes (see Figs 5, 6 and 7). The cell-autonomous inhibition of EcR activity in LECs led to abortive extrusion and cell survival (Fig. 6B). Thus, the death of LECs share with other obsolete larval cells a common priming hormonal (Ecdysone) input (Fig. 8).
It is still not clear how cell proliferation and cell death are differentially controlled by Ecdysone. The trigger of histoblast proliferation seems to be directly dependent on Ecdysone signalling. However, we still do not know how the onset of LEC death is set. In other words, how do LECs distinguish between the late larval and the pupal Ecdysone pulses? In a plausible scenario, to avoid detrimental epithelial gaps at the surface, signalling clues from `matured' histoblasts (after their rapid proliferation in response to the initial prepupal Ecdysone pulse), could assist Ecdysone signalling to instruct LECs to die. Indeed, LECs do not die in response to the pupal Ecdysone pulse if histoblast proliferation (and hence, `maturation') has been experimentally delayed (Fig. 7). The identification and characterization of this putative signal awaits further genetic and molecular analysis. Thus, Ecdysone signalling is necessary, but not sufficient, for LEC death.
The control of the cell cycle
The developmental control of cell cycle dynamics and diversity represents a
key regulatory mechanism that directs cell size, cell number and ultimately
the organ size of adult individuals. Despite numerous elegant experiments, the
details of how cell division is regulated and coupled to cell growth remain
poorly understood.
During abdominal morphogenesis, the trigger of cell proliferation occurs
simultaneously in all histoblast nests within each segment. Cell counting
reveals that up to eight cell divisions are required to build the complete
adult hemitergite (Merriam,
1978). The same proportions apply to the ventral and spiracular
nest. We have found that the first three histoblast divisions during
pupariation are synchronous and extremely fast, skip the G1 phase and resemble
the early embryonic blastoderm divisions
(Fig. 1). In this early stage,
histoblast cleave and progressively reduce their size. We found that Ecdysone
signalling is involved in the initiation of the proliferation programme (see
above). But, how is the histoblast cell cycle regulated to achieve fast
proliferation in the absence of cell growth? Does it rely on the storage of
preexistent control molecules, as in early embryos
(Knoblich et al., 1994), or is
it linked to signals impeding their growth? While this issue remains to be
unravelled, the extreme growth of histoblasts during previous larval stages
makes plausible the accumulation of G1 regulators, which, upon Ecdysone
signalling, could allow a fast transition through G1 phase. Indeed, we have
found that Cyclin E concentration (which regulates entry into S phase) in
histoblasts builds up during the larval period. The observed deceleration of
histoblast proliferation could then be the consequence of the exhaustion of
the entire stock of Cyclin E (N.N., M. Grande and E.M.-B., unpublished).
Still, the implication of growth control mechanisms in the regulation of
histoblast proliferation cannot be ruled out. Multiple cell types, such as the
animal-cap blastomeres from Xenopus embryos
(Wang et al., 2000), change
their cell cycles from size-independent to size-dependent after they become
smaller than a critical cell size. Histoblasts might sense size in an
analogous way. Thus, pathways that regulate growth, such as insulin-mediated
signalling, Myc and Ras oncoproteins and the products of the Tuberous
sclerosis complex 1 and 2 genes (reviewed by
Jorgensen and Tyers, 2004),
should be explored to evaluate their potential roles in the coupling mechanism
linking growth and cell cycle progression.
|
).
The comet-like protrusions of guiding histoblasts break through the LEC
epithelial barrier, leading to planar intercalation of histoblast cell bodies
(Fig. 2; see Movie 4 in the
supplementary material). They account for the capacity of histoblasts to
achieve migration within the bounded epithelial layer. Indeed, electron
micrographs reveal that the advancing histoblasts form junctions with
non-adjacent LECs before the adjacent LECs histolyze, thus insuring the
continuity of the epidermis (Roseland and
Reinhardt, 1982). Time-lapse observations (see Movie 5 in the
supplementary material) suggest that these protrusions grow by sequential
addition of actin molecules at their forward end (see
Bershadsky, 2004). In this
sense, they resemble, although being considerably slower, the actin tails
employed by Listeria to propel through the cytoplasm of infected
cells (Skoble et al., 2001),
or the actin-rich pseudopodia extended by neutrophils in response to
chemoattractants (Weiner et al.,
1999). Proper actin cytoskeleton dynamics appear to be essential
to build up these protrusions and the full repertoire of activities leading to
the expansion of histoblast nests. The equilibrium between actin
polymerization and depolymerization activities should be exquisitely
regulated, and the forced polymerization of actin by Profilin overexpression
not only blocks the cytoskeletal dynamics of single cells, but impedes the
spreading of the whole histoblast nest
(Fig. 3). Potential roles for
further actin dynamics regulators, the Arp2-Arp3 (Arp14D-Arp66B - FlyBase)
complex, Dynamin (Shibire - FlyBase), membrane polyphosphoinositides, Cdc42,
WASp-family proteins and other molecules (reviewed by
Machesky, 1999) in building up
these projections remain to be explored. Further, although these protrusions
appear to have a mechanical role, they also seem to be involved in the
recognition of guidance cues, as they follow stereotyped paths. Indeed,
gradients of cell affinity have been described for the patterning of the
Drosophila abdomen (Lawrence et
al., 1999), and it would be of major interest to understand how
these cells interpret the larval landscape.
|
), other studies suggested that LECs are histolyzed and die by
autophagy (Juhasz and Sass,
2005). Our findings are conclusive in this respect. The death of
LECs involves a caspase-mediated apoptotic process that implicates
cytoskeletal remodelling and apical cellular constriction leading to
delamination. The actomyosin mediated contractile force of dying LECs
contributes in bringing together neighbouring histoblasts. Once the LECs
initiate extrusion, they become immediate targets for circulating haemocytes,
which extend membrane projections and engulf them. Finally, LECs are degraded
inside haemocytes (Figs 4 and
5).
Apical constriction is a process shared by multiple morphogenetic events,
e.g. Drosophila mesodermal cells accumulate myosin and apically
constrict during gastrulation under the control of the small GTPase Rho
(Nikolaidou and Barrett,
2004). Myosin activity is also sufficient to promote the apical
constriction and elimination of photoreceptor cells in the Drosophila
eye in response to the overexpression of an activated form of the Rok kinase
(Rosenblatt et al., 2001).
Indeed, we found that the apical contractility of LECs depends on the level of
phosphorylation of the MRLC and could be enhanced or abolished by modulating
the counteracting kinase and phosphatase activities of Rok and MLCP
(Fig. 4). As a consequence, LEC
delamination is either accelerated or delayed. How these regulatory activities
are themselves regulated remains to be established. Yet, the LEC extrusion
defects observed in weakened caspase cascade conditions after P35
overexpression (Fig. 7B,C)
strongly suggest that apoptotic signals could be involved in the trigger of
actomyosin contractility in LECs. Apical contraction would thus be an early
event in the LEC apoptotic process. Being particularly important to analyse
the differences that modulate the activity of myosin during apical
constriction of living cells and during extrusion of apoptotic cells, the
replacement of LECs could become an exceptionally suitable model to unravel
how myosin activity is regulated in apoptotic cells in vivo.
The recruitment of haemocytes to dying LECs during abdominal cell
replacement is extremely fast. The apical constriction of LECs takes about 2
hours, but the time that a haemocyte needs to fully engulf a LEC is less than
10 minutes. This entails a very reliable chemoattracting mechanism. In
mammals, caspase 3-dependent lipid attraction signals, released by dying
cells, induce the migration of phagocytes
(Lauber et al., 2003).
Furthermore, several receptors are implicated in corpse recognition, including
lectins, integrins, tyrosine kinases, the phosphatidylserine receptor (PSR)
and scavenger receptors (Krieser and
White, 2002). In Drosophila, the elements involved in
cell recognition by macrophages are mostly unknown. Haemocytes express
Croquemort, a scavenger receptor homologue, which is required for the uptake
of dead cells (Franc et al.,
1999), and Pvr, a homologue of the vertebrate PDGF/VEGF receptor
that seems to affect their motility (Cho et
al., 2002). Still, the signals that haemocytes recognize in dying
cells and the links between those signals and the apoptotic cascade are
essentially unknown.
As macrophages are responsible for much of the engulfment of dead cells in
developing animals, an important role for macrophages in tissue morphogenesis
has been suggested (Sears et al.,
2003). However, this is not the case during abdominal
morphogenesis, as the inhibition of haemocyte motility, which abrogates the
removal of LECs, does not affect their replacement by histoblasts. Our results
are consistent with studies showing that macrophage removal of cell debris is
not required for the regeneration of laser-induced wounds in
Drosophila (Stramer et al.,
2005).
Cell cooperation or cell competition in epithelial cell replacement
Histoblast nest expansion is tightly coordinated with LEC removal. A naive
view of the process of LEC extrusion suggests that their death is altruistic -
it would promote the expansion of histoblasts. However, several results
suggest that LECs do not execute this process autonomously. First, histoblast
nests initiate their expansion in the absence of LEC death. Second, histoblast
nests, during their spreading, grow, with no obvious planar orientation, by
stochastic cell divisions not restricted to their edges
(Fig. 1). Finally, and most
importantly, the inhibition of histoblast proliferation exerts non-autonomous
effects on both extrusion and removal of LECs
(Fig. 7). A working model in
which histoblast proliferation and LEC death are synchronized by a spatially
and temporally controlled exchange of signals (secreted ligands or
cell-to-cell communication modules) is thus strongly appealing. This potential
mechanism for replacement of LECs by histoblasts somewhat resembles the
elimination and death by anoikis of amnioserosa cells upon dorsal closure
completion during Drosophila embryogenesis
(Reed et al., 2004). Through
this process, physical contacts and intracellular signalling among epithelial
leading cells, the amnioserosa and the yolk sac coordinate the different
behaviour of these cell types, which is essential for the accurate progress of
both germ band retraction and dorsal closure. In this scenario, coordinated
extrinsic and intrinsic events, hormonal inputs, cell contacts and cell
signalling events will be responsible for the ordered proliferation and
expansion of histoblasts and the extrusion and death of LECs.
An alternative mechanism for the ordered cell substitution taking place
during abdominal morphogenesis involving cell competition could also be
proposed. Competition can be defined as an interaction between individuals
brought about by a shared requirement leading to a reduction in the
survivorship, growth and/or reproduction rates. Classical experiments in
Drosophila imaginal discs have shown that cells heterozygous mutant
for ribosomal protein genes (Minutes) placed beside wild-type cells are
outcompeted and eliminated from the epithelium
(Morata and Ripoll, 1975).
More recent work has shown that imaginal wild-type cells are outcompeted by
cells with growth advantage overexpressing the proto-oncogene Myc
(Moreno and Basler, 2004).
Cell competition does not just apply to the fight for survival of cells with
their `fitness' experimentally altered, but also applies to the homeostasis of
self-renewing cell pools such as lymphocytes
(Gett et al., 2003) or stem
cells (Oertel et al., 2006).
The substitution of LECs by histoblasts closely resembles cell competition.
Rapidly dividing and expanding histoblasts may become competent to displace
the surrounding less-metabolically-active LECs. During normal development,
having `weaker' neighbours, histoblasts do not compete against each other, and
cells from Minute clones in the abdomen are not eliminated in heterozygous
animals (Morata and Ripoll,
1975). However, when confronted with death-resistant LECs
(Fig. 7), `winner' histoblasts
may become `losers'. Histoblasts in an increasingly crowded environment will
compete against each other, and the less fit individuals (less competent in
signalling reception and transduction, or with slower proliferation rates)
would eventually become more sensitive to `killing' signals and would die.
Our findings here demonstrate that the replacement of LECs by histoblasts, independently of being driven by cooperative mechanisms, cell competition or both, represents an extremely amenable morphogenetic model for the study of the dynamic control of the cell cycle and cell death, of the coordination of cytoskeleton activities and cell adhesion, and for the study of cell invasiveness.
Supplementary material
Supplementary material for this article is available at
http://dev.biologists.org/cgi/content/full/134/2/367/DC1
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ACKNOWLEDGMENTS |
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Footnotes |
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REFERENCES |
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TOPSUMMARYINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Bard, J. (1990). Morphogenesis: The Cellular and Molecular Basis of Developmental Anatomy. Cambridge, New York: Cambridge University Press.
Bender, M., Imam, F. B., Talbot, W. S., Ganetzky, B. and Hogness, D. S. (1997). Drosophila ecdysone receptor mutations reveal functional differences among receptor isoforms. Cell 91,777 -788.[CrossRef][Medline]
Bershadsky, A. (2004). Magic touch: how does cell-cell adhesion trigger actin assembly? Trends Cell Biol. 14,589 -593.[CrossRef][Medline]
Brand, A. H. (1995). GFP in Drosophila. Trends Genet. 11,324 -325.
