Howard Hughes Medical Institute, Department of Embryology, Carnegie Institution of Washington, 115 West University Parkway, Baltimore, MD 21210, USA
*Author for correspondence (e-mail: spradling{at}ciwemb.edu
Accepted 29 May 2001
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SUMMARY |
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Key words: Drosophila, Oogenesis, Lar, Axis formation, Planar polarity, Follicle, Extracellular matrix, Actin
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INTRODUCTION |
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Once formed, the follicular epithelium extensively participates in
establishing both the anterior-posterior and dorsal-ventral egg axes (reviewed
by van Eeden and St Johnston,
1999). Posterior follicle cell
fates are defined after receipt of a local epidermal growth factor (EGF)
receptor-mediated signal from the oocyte nucleus. These follicle cells
communicate back to the oocyte and cause its microtubule cytoskeleton to
reorganize, a process essential for stable localization of the posterior
determinant Oskar, and for relocation of the oocyte nucleus to the future
dorsal anterior corner. Disrupting Notch signalling (Ruohola et al.,
1991
), Cadherin (Godt and
Tepass, 1998
; Gonzelez-Reyes
and St Johnston, 1998b) or Laminin A (Deng and Ruohola-Baker,
2000
) in the follicle cells,
or meiotic progression (Ghabrial et al.,
1998
) or cytoskeletal
structure (Emmons et al.,
1995
; Manseau et al.,
1996
; Schulman et al.,
2000
) in the oocyte interferes
with Oskar localization and downstream events. Subsequently, dorsal follicle
cells receive another round of EGF receptor-mediated signals from the
re-positioned germinal vesicle; they respond by defining the dorsal-ventral
axis and patterning additional sub-groups of follicle cells that will mediate
the final morphogenesis of the egg (Schnorr and Berg,
1996
; Wassermann and Freeman,
1998; Ghiglione et al., 1999
).
These terminal morphogenetic steps include the dumping of nurse cell contents
into the oocyte (see Matova et al.,
1999
), egg elongation (see
Edwards and Kiehart, 1996
) and
eggshell patterning (see Waring,
2000
), events that are also
controlled by germline steroid hormone levels (see Buszczak et al.,
1999
).
The structure and organization of the follicular epithelium are crucially
important for it to properly proliferate, migrate, change shape, acquire new
cell fates and to communicate with germ cells (Jackson and Berg,
1999; Bilder et al.,
2000
). Drosophila
follicle cells are organized along the apical-basal axis, much like other
epithelia (reviewed by Müller,
2000
; see
Fig. 1C). A prominent
extracellular matrix (ECM) is secreted from the basal surface, while septate
and adherens junctions hold neighboring cells together near their apical ends.
Several genes are known to be required to maintain epithelial polarity,
including armadillo (Peifer et al.,
1991
), DE-cadherin (Oda et
al., 1997
), egghead (Rubsam et
al., 1998
) and brainiac (Goode
et al., 1996
), while
-spectrin is needed to maintain the follicle cells in a monolayer (Lee
et al., 1997
; Zarnescu and
Thomas, 1999
). Mutations in
two junction-associated membrane proteins, Discs-large and Scribble, disrupt
follicle cell polarity, proliferation and development (Goode and Perrimon,
1997
; Bilder and Perrimon,
2000
).
Orthogonal to the apical-basal axis, many epithelial sheets are also
polarized in the plane of the tissue. For example, wing cells produce an
actin-rich hair and align them in parallel with their neighbors. Genetic
studies in imaginal disc epithelia have identified a genetic pathway that is
important for planar polarization (reviewed by Schulman et al.,
1998). Whether spherical
tissues such as ovarian follicles exhibit planar polarization has never been
addressed. Basal bundles of parallel microfilaments in the follicle cells of
stage 9-13 egg chambers have been described (Gutzeit,
1990
). Laminin, a major
component of the extracellular matrix, is also reported to occur in parallel
rows within the basement membrane that adjoins the basal surface of follicle
cells (Gutzeit et al., 1991
).
In females bearing kugelei mutations, the actin and laminin alignment
is lost and eggs frequently fail to elongate along the anterior-posterior
axis, leading the authors to propose that the basal cytoskeleton serves as a
molecular corset that resists the forces of elongation (Gutzeit et al.,
1991
). However, the molecular
nature of kugelei remains unknown and whether basal actin alignment
marks a follicle cell planar polarity has not been addressed.
The actin cytoskeleton provides the structural basis for cell polarity in
many systems. Recently, an important role has been found for protein complexes
containing Arp2/3, WASP/SCAR or ENA/VASP in controlling actin polymerization
(reviewed by Hu and Reichardt,
1999; Cooper and Schafer,
2000
). In outgrowing axons,
where the function of these complexes is well established, the `Leukocyte
antigen related' gene (Lar), which encodes a receptor-like tyrosine
phosphatase, signals to control the activity of these complexes (Krueger et
al., 1996
; Wills et al.,
1999a
; Wills et al.,
1999b
; reviewed by Lanier and
Gertler, 2000
). Recently, a
Drosophila homolog of the yeast cyclase-associated protein has been
identified (Benlali et al.,
2000
) and been shown to
regulate actin polymerization in developing egg chambers (Baum et al.,
2000
), suggesting that similar
complexes act in follicle cells.
We now identify female sterile mutations in Lar that disrupt multiple steps in follicle development and patterning. We show that Lar is required in posterior follicle cells to maintain Oskar localization in the oocyte. In addition, Lar plays an important role in aligning follicle cell actin relative to the polar cells to promote egg elongation along the anterior-posterior axis. We propose that Lar interacts closely with the extracellular matrix to facilitate the planar polarization of epithelial layers.
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MATERIALS AND METHODS |
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Antibodies and reagents
The following antisera were used at the indicated dilutions: rabbit
polyclonal anti-Laminin A 329 (Gutzeit et al.,
1991), rabbit polyclonal
anti-Oskar (Rongo et al.,
1995
) (1:3000), rabbit
polyclonal anti-ß-galactosidase antibody (Cappel) (1:3000) or mouse
monoclonal anti-ß-galactosidase antibody (Promega) (1:2000), and 7G10
anti-FasIII mouse monoclonal antibody (Patel et al.,
1987
) (1:10). Secondary
antibodies were goat anti-mouse or goat anti-rabbit IgG conjugated to
Alexa488 or Alexa568 (Molecular Probes) (1:400). Actin
was stained with Alexa488 or Alexa568 (Molecular Probes)
conjugated to phalloidin (1:200).
Antibodies against different segments of Lar protein were generated as
follows: 11 GST (pGEX, Amershan Pharmacia Biotech) fusion constructs were made
against pieces of Lar that varied from 150 to 200 amino acids in length.
Together the fusions spanned the entire protein. Fusion proteins were purified
according to Ausubel et al. (Ausubel et al.,
1987). Rats were immunized with
the material at Covance Research Products. Three antisera recognized a protein
of correct size on western blots (data not shown), but none specifically
labeled Lar protein in tissue.
Clonal analysis
Mitotic clones were generated according to Xu and Rubin (Xu and Rubin,
1993). One- to 3-day-old
females of the genotype hs-FLP; Lar5.5 or
Lar13.2/Arm-lacZ were heat shocked for 1 hour at 37°C
twice a day for 3 days. Flies were transferred to fresh yeasted food.
Armadillo-lacZ, which is strongly expressed in all cells in the
ovariole (Xie and Spradling,
1998
), was used as a clonal
marker. In all figures, the green cells are Lar+ and blank cells
are Lar-. All the clonal analysis data presented is from flies aged
7-14 days after heat shock regimen. This ensures that clones analyzed are stem
cell clones (Margolis and Spradling,
1995
).
Hs-hh induction of ectopic polar cells
Adult females containing the hsp70-hedgehog (hs-hh) transgene were
subjected to cycles of heat shock at 37°C followed as described by Forbes
et al. (Forbes et al., 1996). Ovaries were removed after 3 days of treatment
and the number of polar cells analyzed by staining with anti-FasIII
antibodies.
Actin labeling and immunofluorescence microscopy
For proper fixation of cytoskeleton, egg chambers were isolated in a
solution of equal osmolarity (Tilney et al.,
1996). For both dissection and
fixation, 1x Grace's medium (GIBCO BRL) was used. The presence of
phalloidin in the fixative also stabilizes F-actin. All steps were carried out
at room temperature. Ovaries were dissected in 1x Grace's medium and
immediately fixed for 20 minutes in freshly prepared F buffer (four parts
Grace's Medium, one part fresh EM grade 16% formaldehyde, 2% Triton-X-100, 1
U/ml of phalloidin). This was followed by two 20 minute washes in
phosphate-buffered saline (PBS) containing 0.2% Triton X-100 and 1 U/ml of
phalloidin, and another 20 minute wash in PBS containing 0.2% Triton X-100 but
without phalloidin. The ovaries were then rinsed in PBS alone, after which
they were ready for antibody staining. Immunofluorescence microscopy was
carried out as described previously (deCuevas et al.,
1996
). Stained ovaries were
mounted and analyzed using a Leica NTS-confocal microscope.
Scanning electron microscopy
Drosophila eggs were fixed in 3% glutaraldehyde/1% formaldehyde
0.1 M cacodylate pH 7.4 overnight. After an ethanol dehydration series,
specimens were stabilized in hexamethyldisilizane, coated with
platinum/palladium and imaged in JEOL SEM 35 microscope.
Molecular analysis of Larbola alleles
The location of the P insertions Larbola1 and
Larbola2 was determined by polytene chromosome in situ
hybridization as described (Karpen and Spradling,
1992) and mapped to 37F2-38A2.
Their location on the genomic sequence was determined by plasmid rescuing
flanking genomic DNA as described (Karpen and Spradling,
1992
), and sequencing across
the junction from the 5' P element end. Both flanking sequences gave a
unique match to genomic DNA sequences from region 37F2-38A2, at nucleotides
149,050 and 175,640, respectively, of contig AE03663.1 (version 1). The
annotated 5' end of the Lar transcript in the same coordinate system is
at 170,056 and the end of the first exon is at 170,203.
Northern blot analysis
Total ovary RNA was obtained, size fractionated and blotted as described by
Schneider and Spradling (Schneider and Spradling,
1997), except that
TRIzolTM (Gibco BRL) was used as extraction buffer. The blot was
probed with a 1.5Kb EcoRI fragment from Dlar55 cDNA (Streuli et al.,
1989
), corresponding to the
5' region of the gene. The blot was simultaneously probed with the 3.4
kb EcoRI-SacI fragment isolated from the full-length cDNA of
the cup gene (Keyes and Spradling,
1997
) as a loading
control.
Whole-mount in situ
Single strand digoxigenin-labeled sense and antisense cDNA probes from the
clone Lar55 (Streuli et al.,
1989) were generated by single
strand PCR (Patel et al., 1992). Hybridization was carried out according to
Suter and Steward (Suter and Steward,
1991
).
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RESULTS |
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To distinguish the roles of actin filaments in dumping and egg elongation, we identified two allelic female sterile mutations, termed bola1 and bola2. In mutant females 20-70% of the follicles fail to elongate, despite transferring their nurse cell contents normally (see Fig. 2A (inset),C). Unelongated follicles develop into eggs that are nearly round, resembling a soccer ball (`bola' in Portuguese), and contain unelongated follicle cells (Fig. 2C,D), but enclose a normal volume (112%±29% of wild type n=40). When stage 10A bola follicles were stained with Alexa-phalloidin to visualize actin filaments, normal microfilament bundles were observed in nurse cells, consistent with the ability of these eggs to normally transfer their cytoplasmic contents (Fig. 2G). Consequently, we analyzed the bundles of actin filaments located near the basal cortex of stage 8-13 follicle cells to determine if these structures were affected by the mutations.
Although basal actin filament bundles are known to be present and to be
aligned perpendicular to the anterior-posterior (AP) axis of the egg chamber
by stage 9 (Gutzeit, 1990),
their behavior earlier in oogenesis has not been reported previously. Taking
care to preserve actin morphology (see Materials and Methods), we fixed and
stained wild-type and homozygous bola mutant ovaries, and studied the
behavior of actin filaments in somatic cells. In the germarium, large actin
bundles are located along the basal cortex of somatic cells that line region
2b and 3 (Fig. 2E). These
filaments are aligned in an organized fashion such that they run around the
ovariole perpendicular to its AP axis. At the time a new follicle buds from
the germarium, this orientation is lost
(Fig. 2E, arrow). Newly formed
stage 2 chambers typically retain some organized actin near their poles, but
in the middle of the follicle, actin filaments swirl along the AP axis. Stage
4 egg chambers show no evident polarity or organization of their actin
filaments (Fig. 2F). By stage
7, however, the basal actin of all the follicle cells is aligned perpendicular
to the AP axis (Fig. 2H,J).
In bola mutant follicles, the actin filaments were present at normal levels and their orientation was indistinguishable from wild type in the germarium and stage 2-4 egg chambers. However, in 30-90% of follicles, the filaments never became aligned perpendicular to the AP axis, although intracellularly the number and size of actin bundles appeared normal (Fig. 2I,K). Thus, a failure to globally polarize the microfilament arrays within individual follicle cells correlated with the failure of bola mutant follicles to elongate normally. The organized circumferential bands of actin filaments in wild-type chambers might resist expansion perpendicular to the AP axis, channeling expansionary forces to elongate individual cells and the egg in the AP direction. These observations establish a role for actin filaments in egg elongation that is independent of their role in nurse cell dumping.
bola mutations are alleles of Lar
A single P element insertion in each bola mutation was mapped to
cytogenetic position 38A by polytene chromosome in situ hybridization (data
not shown). Genomic DNA flanking both alleles was recovered by plasmid rescue
and sequenced. The insertions in bola1 and bola2 were
located 20 kb upstream and 5 kb downstream from the previously reported
5' end of Lar, a previously characterized gene in region
37F2-38A2 (Fig. 3A). As the
Lar transcription unit is known to extend 128 kb downstream from the
5' end, rescue of the mutation with a genomic construct was impractical.
However, excision of the bola1 element reverted all the associated
phenotypes (not shown). In addition, we obtained two previously characterized
lethal Lar alleles, Lar5.5 and
Lar13.2, that are caused by stop codons at amino acid
positions 551 and 1055, respectively (Krueger et al.,
1996). These EMS-induced
mutations have been generated independently using a different genetic
background from that of Larbola1 and
Larbola2. Transheterozygotes between sterile and lethal
alleles, or between sterile alleles and known deficiencies that uncover
Lar produced sterile females whose follicles showed the same defects
in egg elongation and actin alignment as Larbola1 and
Larbola2 homozygotes (see Materials and Methods). Northern
analysis of RNA extracted from whole wild-type ovaries using a Lar cDNA probe
revealed a single band of 8.4 kb, the same as the previously characterized Lar
mRNA from nervous tissue (Kreuger et al.,
1996
). Lar transcripts were
undetectable in a corresponding preparation of ovarian RNA from
Larbola1 or Larbola2 homozygous
females (Fig. 3B, and data not
shown). We conclude that Larbola1 and
Larbola2 are new alleles of the Lar gene.
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To investigate Lar expression during oogenesis, we carried out
whole-mount in situ hybridization to wild-type ovaries. Strong Lar
expression was first observed in the somatic follicle cell precursors located
in region 2b of the germarium (Fig.
3C). Weaker expression was apparent in germ cells at these stages
as well. Lar expression continued as egg chambers budded from the
germarium, and was robust in both somatic and germline cells of older
follicles (Fig. 3C,D), in
agreement with a previous survey (Fitzpatrick et al.,
1995). Several antibodies were
prepared against both intracellular and extracellular Lar domains (see
Materials and Methods). Like the experience of other investigators, however,
we found that none of these antisera would specifically label Lar in tissue
(data not shown).
Polar cells and egg chamber planar polarity
A pair of specialized polar follicle cells lie at the anterior and
posterior poles of ovarian egg chambers beginning at stage 3, and possibly
earlier (Brower et al., 1981). Polar cells can be visualized by virtue of
their high Fasciclin III (FasIII) expression levels (Patel et al.,
1987) in stage 3 and later
follicles. Polar cells must be specified even earlier, as they cease division
in the germarium (Margolis and Spradling,
1995
; Tworoger et al.,
1999
); however, markers
specific for early polar cells are not available. To investigate whether polar
cells define the poles of the basal actin pattern, we fixed and labeled
wild-type egg chambers with Alexa-conjugated phalloidin and with anti-FasIII
antibodies to reveal actin filaments and polar cells. These studies showed
that the actin circles at the poles of stage 7 and later follicles were
precisely organized around the polar cell pairs
(Fig. 4A). In addition, the
actin polarization was observed to arise gradually during egg chamber
development proceeding from the poles toward the center. As a result, in stage
5-6 egg chambers, the actin was organized in the polar regions
(Fig. 4B, arrow) but still not
organized toward the center of the follicle
(Fig. 4B, lower right). The
observations that the polar cell pairs act as polarization centers, and that
polarization proceeds from the poles towards the center of the follicle,
suggests that the polar cells produce or stimulate production of a polarizing
signal that is received by nearby cells and used to re-orient their planar
cell polarity.
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We have carried out similar studies on mutant egg chambers to investigate whether abnormalities in polar cell differentiation or function might be responsible for the failure of actin filaments to polarize in a large fraction of mutant egg chambers. Nearly half (47%) of the Larbola follicles contained extra polar cells (Fig. 4F). Instead of possessing exactly two FasIII-positive polar cells at each end, bola follicles contain an average of 2.3 anterior polar cells and 2.6 posterior polar cells (n=91). Moreover, 3.3% of the follicles contained an ectopic polar cell cluster of from 2-12 cells (Fig. 4E). All the follicles with extra polar cells, whether located at the poles or ectopically, failed to undergo actin alignment. However, the converse was not true. Many follicles were observed in which the number and location of polar cells was normal, but the actin fibers were not aligned (Fig. 4C). When actin alignment was observed in the mutant, it was always centered on the polar cells, as in wild type (data not shown). The correlation between extra polar cells and actin misalignment supports the idea that polar cell defects are responsible for the actin misalignment.
To further investigate whether polar cells act directly to organize actin
polarity, we studied the effects of ectopic polar cells on actin organization.
Misexpression of hedgehog protein (Hh) causes excess follicle cell
proliferation and the production of ectopic polar cells (Forbes et al.,
1996a: Forbes et al.,
1996b
; Zhang and Kalderon,
2000
). We subjected flies
carrying a heat shock promoter-hedgehog fusion gene to periodic heat shocks
and analyzed their egg chambers for the presence of ectopic polar cells by
FasIII staining and for actin polarity. As expected, egg chambers bearing
extra FasIII-positive cell pairs were found, and actin fibers sometimes
circled around these ectopic cells, despite their location away from the true
poles of the egg chamber (Fig.
4D). However, many other FasIII-positive cell pairs were found
within fields of normally polarized actin filaments
(Fig. 4G). The presence of
these extra FasIII-positive cells did not usually cause a general disruption
of actin organization, like that observed in egg chambers from bola
mutant females. Most of the Hh-induced ectopic polar cells may not be fully
differentiated, and consequently will lack actin polarizing activity.
bola mutations affect egg chamber production and oocyte
polarity
Polar cells are normally specified in the germarium (Margolis and
Spradling, 1995; Tworoger et
al 1999
). The presence of
extra polar cells suggested that bola mutations might affect the
process of egg chamber formation. Germaria from Larbola
mutant females were found to frequently be abnormal
(Fig. 4I,K). When wild-type
germline cysts enter region 2b, they become lens-shaped and stretch to span
the entire width of the germarium (see Fig.
1B). In 57% of germaria from Larbola2 females,
the cysts in region 2b remain two-across and never become lens-shaped
(Fig. 4I,K, bracket). Egg
chamber budding is slowed and occurs abnormally
(Fig. 4K, arrow), frequently
causing additional round cysts to accumulate in region 3 of the germarium (see
below), and occasionally producing composite follicles containing two cysts
(not shown). Thus, defects caused by Larbola mutations are
apparent as follicle cells begin to associate with germline cysts, and
continue throughout the budding process These changes are likely to be related
to the polar cell abnormalities and actin misalignment that become apparent
later.
We suspected that egg chambers produced by Larbola females might also contain defects in oocyte polarity. To test this hypothesis, we stained egg chambers with antibodies specific for Oskar, whose localization at the posterior pole of stage 9-10 egg chambers represents a crucial step in posterior patterning and germ cell development. Strong staining of Oskar at the oocyte posterior was observed in all wild-type egg chambers (Fig. 4H), but in 20-30% of chambers from Larbola females, Oskar protein was present in one or more aggregates at other positions in the oocyte cytoplasm (Fig. 4J). The effects of the mutations on embryonic development could not be determined because the eggs produced by Larbola mothers never initiated embryonic development. The GV in Larbola follicles moved normally to the dorsal-anterior region and there were no defects in Gurken localization (data not shown), indicating that the defects were limited to the AP axis and possibly to the process of Oskar localization itself.
Lar function is required autonomously in posterior follicle cells for
Oskar localization
To determine which cells require Lar function we carried out
clonal analysis. Patches of marked Lar mutant cells were generated
using FLP-mediated somatic recombination (see Materials and Methods) and the
effects of different mutant cell groups on egg chamber development and
polarity were determined. The effects of Lar mutation on Oskar localization
are summarized in Fig. 5.
|
Oskar localization was found to depend on the genotype of the posterior follicle cells. When the approximately 70 posterior follicle cells were Lar positive, Oskar was always localized normally, regardless of the status of Lar function in the germline or in more anterior follicle cells (Fig. 5A). By contrast, when the posterior follicle cells lacked functional Lar, Oskar was normal only 15% of the time. In such chambers, a significant fraction of the Oskar protein was present as a central aggregate (Fig. 5B,C) or was absent altogether (Fig. 5D). We observed a number of follicles in which Oskar protein trailed from the posterior to a central aggregate (Fig. 5C), suggesting that posterior localization may have occurred normally, but not been maintained.
To address whether the action of posterior follicle cells in localizing or maintaining posterior Oskar was autonomous at the cellular level, we examined chambers that contained a clonal boundary within posterior follicle cells. In more than 70% of such cases, Oskar protein was found below the cells that were Lar+, but was strongly reduced or absent under cells that bore a Lar mutation (Fig. 5F,G). The boundary of localized Oskar varied no more than one cell diameter from the position of the Lar+ border.
Deng and Ruohola-Baker (Deng and Ruohola-Baker,
2000) recently described
similar effects of LanA clones in posterior follicle cells (Deng and
Ruohola-Baker, 2000
). As LanA
has been reported to be a ligand for Lar in mammalian cells (O'Grady et al.,
1998
), we examined the
distribution of LanA protein in developing egg chambers
(Fig. 6J). LanA was abundant in
the basal region of the follicle cells of stage 2-9 egg chambers.
(Fig. 6J, arrowhead), but was
not detected between the follicle cells and the oocyte (Gutzeit et al.,
1991
). Females mosaic for a
lethal null allele of LanA, LanA9-32, generated round eggs
at a low frequency (Fig. 6J, inset), further supporting a link between LanA and Lar.
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A possible explanation for the lack of correlation between stage 7-8 follicle cell genotype and actin polarity is that Lar is required in cells that interact with polar cell precursors much earlier in development. Consequently, we examined the effects of Lar mutant germline or somatic cell clones in the germarium on cyst shape and follicle budding. When the germline lacked Lar function, region 2b cysts never failed to become lens-shaped and region 3 never contained more than one round follicle in the process of budding. By contrast, when most of the follicle cells in region 2b and 3 were mutant (a relatively rare genotype), then 62% of the germaria contained such defects (Fig. 6G,H). Abnormal germaria frequently lacked Lar function in intercyst cells and contained an increased number of such cells (Fig. 6G,H). We suspected that polar cells were differentiating abnormally in such germaria, but this could not be directly addressed as markers that specifically label polar cell precursors in the germarium have not been found. Nonetheless, our experiments showed that lack of Lar function in region 2b-3 somatic cells frequently disrupts follicle formation and probably generates follicles with abnormal polar cells that fail to polarize normally during stage 6. By stage 7-8, the descendents of the crucial Lar mutant cells, which are not polar cell precursors themselves, may not occupy a consistent region of the follicle surface and cannot therefore be correlated with defective actin alignment.
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DISCUSSION |
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There are several possible reasons for actin polarity to arise gradually. The follicle cells that surround a newly budded chamber derive from multiple sources, and have spent varying times in association with germ cells at the time of budding. Some follicle cells contact a germline cyst near the region 2a/2b junction or are born from them later by division and remain associated with the cyst cells. Other follicle cells likely migrate from the intercyst cell population, as it contains many more cells late in region 2b than the final number of stalk and polar cells (H. F. and A. S., data not shown). Some time may be required to globally polarize the cytoskeletal actin of follicle cells with these diverse origins. In addition, all follicle cells, except polar cells, continue to divide actively in early egg chambers, but neither their cell cycles nor division planes are synchronized or spatially coordinated. Consequently, full polarization might not be possible until the disruptive effects of mitotic divisions cease during stage 6.
Our observation that polar cells define the polarization axis of follicular
actin suggests an additional reason that actin alignment develops gradually
and after a substantial delay. The differentiation of polar cells themselves
appears to be a gradual process. Polar cell precursors are specified in
germarium region two, where they cease division long before the progenitors of
main-body follicle cells (Margolis and Spradling,
1995; Tworoger et al.,
1999
). However, polar cell
genes such as FasIII are not expressed specifically in these cells until much
later, during stage 3. Polar cell differentiation may be an ongoing process in
early follicles and they may not acquire the capacity to organize actin
filaments until stage 5, when alignment begins locally in nearby follicle
cells. The behavior of ectopic polar cells generated by misexpressing Hedgehog
was consistent with this interpretation. Some of the polar cell pairs induced
local actin polarization while others had no effect
(Fig. 4). Polar cell pairs that
were able to orient actin in nearby cells may have been older, or more fully
developed, than those that were ineffective.
Polarized follicle cell actin directs egg elongation
The molecular mechanisms that late in oogenesis cause eggs in many insect
species to elongate along their anterior-posterior axis have long been a
matter of speculation. Studies in Drosophila have suggested that egg
elongation results when circumferential bands of follicle cell actin filaments
(Gutzeit et al., 1991) cause
forces generated at the time of nurse cell dumping to stretch the egg along
the AP axis. `Dumpless' mutations that disrupt the bundling of actin filaments
(reviewed by Robinson and Cooley, 1995) or functioning of myosin II (Edwards
and Kiehart, 1996
; Wheatley et
al., 1995
) block both the
transfer of nurse cell contents into the oocyte and egg elongation.
bola mutations allow these effects to be dissociated. Nurse cell
dumping, which depends on static actin bundles located within nurse cells, and
probably requires actin-mediated contraction to drive nurse cell contents into
the oocyte, is unaffected. However, follicles lacking somatic bola
function frequently fail to elongate, indicating that oriented follicle cell
actin filaments are crucially important for elongation. Eggs with an
unpolarized follicular epithelium may fail to elongate because contractile
forces are directed isotropically, blocking any net shape change. Consistent
with this model, the volume of the bola eggs was normal, and their shape
changed little during or after dumping. Disruption of basal actin filament
polarization in kugelei mutants also is associated with round eggs
(Gutzeit et al., 1991
),
further supporting a functional role for actin polarization as a `molecular
corset' that directs egg elongation along the anterior-posterior axis.
Identifying genes that cause round egg production in somatic cell clones is
likely to define additional genes required for this polarization pathway.
Follicle cell actin polarization and planar cell polarity
Sheets of epithelial cells become polarized along both the apical-basal
axis and in the plane of the epithelium (reviewed by Tepass,
1997). Planar cell polarity in
epithelial tissues such as in the imaginal discs involves signals transmitted
by an unknown ligand(s) that differentially activate the Frizzled receptor in
nearby cells (reviewed by Schulman et al.,
1998
; Bray,
1999
). For example, in the eye
and wing discs, the planar polarity signal is sent by cells lying along the
dorsal-ventral border. A gradient of Frizzled activity in cells lying
different distances from the source cells transduces polarity information via
a distinct pathway within the cell layer. In the eye disc, this involves
differential activation of Notch signalling (Cooper and Bray,
1999
; Tomlinson and Struhl,
1999
). Differences in Notch
activation control cell fate decisions that result in an inverted polarity of
photoreceptor clusters with respect to the dorsalventral border (equator).
Biologically, the alignment of follicle cell basal actin filaments described here represents an analogous process of planar polarity to those studied previously in imaginal disc derivatives. However, it remains to be determined to what extent these events use the same genetic pathways described in disc tissues. Polarizing a spherical tissue such as an ovarian follicular epithelium is likely to involve planar polarity signals from particular subregions whose receipt by follicle cells conveys positional information. Our data suggest that polarity signals are likely to derive from the poles of the follicle, rather than from a line of cells. Follicle cells may align their actin filaments towards other cells that receive the polarizing activity at an identical level. Our data make it unlikely that Lar acts as a primary receptor for polarity information in stage 6 and up follicles. Clones of Lar5.5 and Lar13.2 alleles produced ectopic veins in the wing, but there was little effect on the orientation of wing hairs (data not shown). Thus, the Lar pathway is redundant in this tissue.
Lar requirement for Oskar localization suggests a role for the
ECM
The structure and properties of Lar that have been revealed by studies of
both the Drosophila and mammalian counterparts suggest how it might
act as a polarity transducer. Lar is a receptor-like tyrosine phosphatase that
is important for axon pathfinding in Drosophila (reviewed by Lanier
and Gertler, 2000). A family
of closely related Lar-like phosphatases also exists in the mouse. Mouse Lar
is required for the development of the mammary epithelium (Schaapveld et al.,
1997
), while the related
PTPsigma functions in neuronal and epithelial development (Wallace et al.,
1999
). The Lar extracellular
domain contains three immunoglobulin and several fibronectin type III domains
and is thought to transduce signals via its cytoplasmic tyrosine phosphatase
domains after activation by adhesion to the ECM or by small protein ligands.
However, physiological Lar ligands have not been documented. In mammalian
cells, binding of the ECM laminin-nidogen complex to a specific fibronectin
domain in mammalian Lar causes changes in the actin cytoskeleton, suggesting
that Lar can transduce information from the ECM (O'Grady et al.,
1998
).
A clue to the mechanism of Lar action comes from our studies on
its role in Oskar localization. Posterior follicle cells must express
Lar to ensure that Oskar is localized properly at the oocyte
posterior (Fig. 5). Previously,
Deng and Ruohola-Baker (Deng and Ruohola-Baker,
2000) reported that when
posterior follicle cells lack the ECM component Laminin A (LanA), Oskar
localization is usually disrupted. They suggest that LanA and ECM
mediate the posterior follicle cell-oocyte signal. As Lar has been reported to
bind to the laminin-nidogen complex (O'Grady et al.,
1998
), Lar might act as the
LanA receptor in this pathway.
However, it remains less clear how a signal initiated by an interaction between LanA in the ECM and Lar on a posterior follicle cell would be transduced into the oocyte. Deng and Ruohola-Baker propose that some LanA-containing ECM resides between the apical surface of the posterior follicle cells and the oocyte, and that LanA interacts directly with the oocyte surface. We propose an alternative model. We only observed LanA on the basal side of the follicle cells, and observed that LanA clones could induce round eggs (Fig. 6J). These observations and the follicle cell autonomous requirement of Lar for Oskar localization argue that the LanA signal is received by Lar on the basal surface of the follicle cells and leads to some change in the receiving cells that is transduced to the oocyte. This could be via a secondary signal, or by changes in the structural or adhesive properties of the cells that can locally affect the oocyte surface with which they come into contact. Lar mutation did not affect the apical basal polarity of follicle cells, as the apical-basal asymmetry of actin staining was maintained and multiple-layered follicle cells were never observed.
Polar cells are likely to play a key role in polarizing actin in
stage 5-6 follicle cells
Several of our observations support the idea that polar cells organize the
actin planar polarity. Actin polarity focuses around both the anterior and
posterior polar cell pairs, and spreads from the poles towards the equator of
the follicle. Additionally, ectopic polar cells induced by Hh expression
sometimes have actin polarizing activity. These findings suggest that polar
cells send a signal that orients the actin alignment in circumferential
direction. In follicles where actin fails to become aligned, the polar cell
signal may have been blocked or reduced, despite the presence of
morphologically recognizable polar cells. Not all the ectopic polar cells
induced by Hh expression affected the actin alignment of nearby follicle
cells, supporting the idea that polar cell can express differentiation
markers, but still be incompetent as polarizing centers. However, Lar was not
required in polar cells, because we observed follicles with normally oriented
actin, despite the presence of a Lar mutant polar cell pair.
The apparent non-autonomy of Lar on actin polarity suggests an
indirect action on polar cell differentiation
The autonomy of the effects of Lar on Oskar localization contrasts with its
apparently non-autonomous action on follicle cell planar polarity at stages
6-8. There was no relationship between actin alignment and the Lar genotype of
particular somatic cells in stage 7 or later follicles. This observation can
be rationalized by postulating that Lar acts on a subset of cells that is
required for polar cell specification and differentiation. Lar is required in
region 2b of the germarium, a time when polar cells are not yet fully
specified, and in its absence somatic cell behavior and follicle formation is
compromised. We propose that the intercyst cells
(Fig. 1B) interact with the
polar cell precursors in a Lar-dependent manner. Intercyst cells mostly become
main body follicle cells and do not remain a recognizable subpopulation;
hence, it is not possible to infer their genotype from later stage egg
chambers and compare it with the state of actin polarization. We did note a
correlation between mutant intercyst cells and pinching defects
(Fig. 6G,H). Thus, a
relationship may exist between the intercyst cell genotype and actin
polarization that cannot be followed with existing markers.
One possibility is that Lar acts in a similar manner in intercyst cells and in posterior follicle cells. The germarium contains peripheral somatic cells whose basal actin fibers are aligned perpendicular to the AP axis (Fig. 2E). Integrin is aligned in a similar manner, suggesting that the basement membrane is correspondingly organized (H. F and A. S., unpublished). As budding proceeds, Lar may be required to interpret polarity information from the basement membrane as part of the process that partitions cells into main body and polar cells. Alternatively, Lar may act in a different manner within these intercyst cells to assist in polar cell specification and differentiation.
The Lar requirement for polar cell determination provides an explanation
for another interesting fact. The phenotypic effects of Lar mutations we
observed are very similar to weak mutations in Notch or other Notch-pathway
genes. Notch mutants, like those in Lar, cause the production of extra polar
cells, interfere with egg chamber budding and disrupt the anterior-posterior
axis of the oocyte (Ruohola et al.,
1991). Notch signalling is
required for polar cell specification (Ruohola et al.,
1991
; Tworoger et al.,
1999
). Mutations in either Lar
or Notch may cause similar disruptions in polar cell differentiation and hence
similar downstream effects on egg chamber development and patterning.
The ECM may transiently store polarity information
Our experiments suggest a novel function for the ECM during follicle cell
development the storage of patterning information for later use.
Somatic cells maintain an ECM that surrounds the germarium; when a follicle
buds off, it contains a portion of this ECM in the basement membranes of its
component cells. Our studies emphasize that this ECM may be a repository of
polarity information that is used at critical times when polarization and cell
specification are taking place. During follicle budding, the AP axis of the
new chamber is correlated with the differentiation of two pairs of polar cells
at each terminus. An interaction between posterior follicle cells and the
oocyte ensures that the germline AP axis will correspond to the somatic axis
(Godt and Tepass, 1998;
Gonzelez-Reyes and St Johnston,
1998
). We suggest that at
about the same time, somatic cell interactions ensure that exactly four
correctly positioned polar cells differentiate per follicle. This requires
Lar-dependent readouts from the same ECM that the interacting cells contribute
to polarizing. In stage 8, posterior follicle cells are likewise guided to
maintain localized Oskar over an appropriately sized polar region. In both
cases, cells that helped synthesize an ordered ECM later use it in a
Lar-dependent manner for additional and possibly more refined patterning. The
interactions we have studied may serve as a model for the roles of the ECM and
of Lar signalling in the development of other epidermal and neural cells.
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ACKNOWLEDGMENTS |
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