The Origin of Asymmetry: Early
Polarisation of the Drosophila
Germline Cyst and Oocyte
Review
Jean-René Huynh1 and Daniel St Johnston2
The anterior–posterior axis of Drosophila is established
before fertilisation when the oocyte becomes
polarised to direct the localisation of bicoid and oskar
mRNAs to opposite poles of the egg. Here we review
recent results that reveal that the oocyte acquires
polarity much earlier than previously thought, at the
time when it acquires its fate. The oocyte arises from
a 16-cell germline cyst, and its selection and the initial
cue for its polarisation are controlled by the asymmetric
segregation of a germline specific organelle
called the fusome. Several different downstream
pathways then interpret this asymmetry to restrict
distinct aspects of oocyte identity to this cell. Mutations
in any of the six conserved Par proteins disrupt
the early polarisation of the oocyte and lead to a
failure to maintain its identity. Surprisingly, mutations
affecting the control of the mitotic or meiotic cell
cycle also lead to a failure to maintain the oocyte fate,
indicating crosstalk between the nuclear and cytoplasmic
events of oocyte differentiation. The early
polarity of the oocyte initiates a series of reciprocal
signaling events between the oocyte and the somatic
follicle cells that leads to a reversal of oocyte polarity
later in oogenesis, which defines the anterior–posterior
axis of the embryo.
Introduction
In many invertebrates and vertebrates, one or more of
the body axes are already set up in the egg [1]. For
instance, the anterior–posterior and dorsal–ventral
axes in Drosophila are established during oogenesis by
the asymmetric localisation of bicoid, oskar, and gurken
mRNAs within the oocyte (the future egg) [2,3]. The
localisation of these transcripts depends on the
polarised organization of the oocyte cytoskeleton, and,
therefore, on the polarity of the oocyte itself. Recent
data show that the first asymmetry that leads to this
polarisation of the oocyte can be traced back to earlier
and earlier stages of oogenesis, ultimately leading to
the very first steps at which the oocyte is determined.
The determination of the Drosophila oocyte has
puzzled biologists for more than a century, because it
arises from a syncytium that is formed by four rounds
of incomplete division of a single germ cell, producing
a germline cyst of 16 sister cells that share the same
cytoplasm [4,5]. Only one cell will become the oocyte,
however, while the other 15 cells differentiate as nurse
cells, which provide the oocyte with nutrients and cytoplasmic
components. This means that the selection of
the oocyte can be viewed as the polarisation of the cyst
cytoplasm and membrane [6]. In addition, the oocyte
may inherit some of its polarity from this cyst asymmetry.
The determination and polarisation of the oocyte
are, therefore, two intimately linked issues, which will
be the main focus of this review.
Early Oogenesis in Drosophila
The Drosophila ovary is composed of 16–20 ovarioles,
each of which contains a chain of progressively more
and more mature egg chambers [7]. New egg chambers
are generated at the anterior of the ovariole in a region
called the germarium, which has been divided into four
regions according to the developmental stage of the
cyst (Figure 1). Oogenesis begins in region 1, when a
germline stem cell divides asymmetrically to produce a
posterior cystoblast, and a new germline stem cell,
which remains attached to the neighboring somatic
cells at the anterior (for reviews see [8,9]). The cystoblast
then undergoes precisely four rounds of mitosis
with incomplete cytokinesis to form a cyst of 16
germline cells, which are interconnected by stable cytoplasmic
bridges called ‘ring canals’. During these divisions,
a cytoplasmic structure called the fusome
anchors one pole of each mitotic spindle (see movie 1
and 2 in supplemental data) and, therefore, ensures that
cells follow an invariant pattern of division [10]. This
leads to the formation of a symmetric cyst comprising
two cells with four ring canals, two with three ring
canals, four with two and eight with one. This invariant
pattern is important, as the oocyte always differentiates
from one of the two cells with four ring canals, which
are, therefore, called the pro-oocytes [11].
Once the 16 cell cyst has formed, it enters region 2a
of the germarium. At this stage, all the cells of one cyst
look the same, but by the time it reaches region 2b, one
cell will have differentiated as an oocyte. This differentiation
can be followed with several types of marker
(Figure 1): First, oocyte-specific proteins, such as BicD,
Orb, Btz and Cup and mRNAs, such as osk, BicD and
orb first concentrate in the two pro-oocytes, and come
to lie on either side of the largest ring canal that connects
them. By the end of region 2a, they accumulate
just in the oocyte [12–17]. Live imaging of GFP-labeled
mitochondria reveals that they show a similar pattern of
localisation to one cell [18]. Second, microtubules are
initially diffusely distributed throughout the cyst in association
with the fusome, but their minus ends gradually
become restricted to the future oocyte [19,20]. Third,
the centrioles appear to be inactivated after the last
mitotic division, and they migrate along the fusome into
Current Biology, Vol. 14, R438–R449, June 8, 2004, ©2004 Elsevier Ltd. All rights reserved. DOI 10.1016/j.cub.2004.05.040
1Institut Jacques-Monod, CNRS, Universités Paris 6 et 7, 2,
Place Jussieu, F-75251 Paris, Cedex 05, France.
E-mail: huynh@ijm.jussieu.fr
2The Wellcome Trust/Cancer Research UK Gurdon Institute
and Department of Genetics, University of Cambridge,
Tennis Court Rd, Cambridge CB2 1QR, UK.
the pro-oocytes, and then into the oocyte [18–20].
Fourth, although the oocyte is the only cell to go
through meiosis, electron microscopic studies show
that the other pro-oocyte also enters meiotic prophase,
and reaches the pachytene stage before becoming a
nurse cell, while the two cells with 3 ring canals reach
the zygotene stage [21]. This can be followed with antibodies
against components of the synaptonemal
complex, a proteinaceous structure that forms between
paired homologous chromosomes [22–24].
In region 2b, the cyst changes shape and becomes a
one cell-thick disc that spans the whole width of the
germarium. Oocyte-specific factors now become concentrated
in the oocyte and a microtubule organising
centre (MTOC) is clearly detectable. At the same time,
somatic follicle cells start to migrate and surround the
cyst. As the cyst moves down to region 3 (also called
stage 1), the mitochondria, centrosomes, Golgi vesicles,
proteins and mRNAs organize at the anterior of
the oocyte to form a typical Balbiani body [18–20]. At
the same time, the cyst rounds up to form a sphere with
the oocyte always lying at the posterior pole. The cyst
then leaves the germarium and enters the vitellarium.
The oocyte becomes polarised, as proteins, mRNAs,
centrosomes and a subset of the mitochondria move to
the posterior cortex, its DNA becomes highly condensed
to form a structure called the karyosome,
whereas the nurse cells start to polyploidize [25,26].
Formation of a Polarised Cyst
The Fusome and Determination of the Oocyte
Two main models have been proposed to explain how
the oocyte is selected. One model is based on the symmetrical
behavior of the two pro-oocytes until mid-late
region 2a, and proposes that there is a competition
between the two pro-oocytes to become the oocyte
[21,27]. The ‘winning’ cell would become the oocyte,
while the ‘losing’ cell would revert to the nurse cell fate.
A second model suggests that the choice of the oocyte
is biased by the establishment of some asymmetry as
early as the first cystoblast division, which is maintained
until the overt differentiation of the oocyte [6,28].
The role played by the fusome and the analysis of its
formation provide the stongest evidence in support of
this model [28–32].
The fusome arises from a spherical structure called
the spectrosome in the germline stem cell, which is
made of small membranous vesicles kept together by
components of the sub-membraneous cytoskeleton,
such as αα-spectrin, ββ-spectrin, and Hts (an adducin-like
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Figure 1. Drosophila early oogenesis.
Each ovariole is made of a chain of progressively
more mature egg chambers
toward the posterior (p). An egg chamber
comprises 16 germline cells surrounded
by a monolayer of follicle cells. The egg
chambers are produced in a specialized
structure, called the germarium, at the
anterior (a) of the ovariole. The germarium
is divided into four morphological regions
along the anterior–posterior axis. The
germline stem cells reside at the anterior
tip of the germarium (left) and divide to
produce cystoblasts, which divide four
more times in region 1 to produce 16 cell
germline cysts that are connected by ring
canals. The stem cells and cystoblasts
contain a spectrosome (red circles), which
develops into a branched structure called
the fusome, which orients each division of
the cyst. In early region 2a, the synaptonemal
complex (red lines) forms along
the chromosomes of the two cells with
four ring canals (pro-oocytes, yellow) as
they enter meiosis. The synaptonemal
complex then appears transiently in the
two cells with three ring canals, before
becoming restricted to the pro-oocytes in
late region 2a. By region 2b, the oocyte
has been selected, and is the only cell to
remain in meiosis. In region 2a, cytoplasmic
proteins, mRNAs and mitochondria
(green), and the centrosomes (blue
circles) progressively accumulate at the
anterior of the oocyte. In region 2b, the
minus-ends of the microtubules are
focused in the oocyte, and the plus-ends
extend through the ring canals into the
nurse cells. The follicle cells (gray) also
start to migrate and surround the germline
cells. As the cyst moves down to region 3, the oocyte adheres strongly to the posterior follicle cells and repolarises along its anterior–posterior
axis, with the microtubule minus-ends and specific cytoplasmic components now localized at the posterior cortex.
1 2a 2b 3
+
+
+
+
+
--
+
++
+
Selection of
the oocyte
Selection of the
pro-oocytes
Stem
cell
Cystoblast
divisions
--
Germarium Stage 3 Stage 6
Germline cyst
Follicle
cells
a p
gurken mRNA
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Review
R440
protein) [10,33]. The cystoblast inherits one third of the
fusome from the GSC [30]. When the cystoblast
divides, one pole of the spindle is anchored by the
inherited fusome (the ‘original’ fusome), while a new
fusome plug forms in the arrested ring canal, at the
opposite pole of the cell [29,32]. The two fusomes then
come together to fuse, so that one cell contains the
‘original’ fusome plus half of the plug, whereas the
other cell only retains the other half of the fusome plug
(Figure 2). This asymmetric behaviour of the fusome is
then repeated during the next three divisions [32]. The
oldest cell, therefore, retains the ‘original’ fusome and
accumulates three more fusome plugs. Thus, this cell
has more fusome than all of the other cells and can be
identified throughout the divisions.
Unfortunately, most of the fusome has already
degenerated by the time oocyte-specific proteins
such as BicD or Orb accumulate in a single cell in late
region 2a. However, the preferential accumulation of
the centrosomes as well as of osk and orb mRNAs in
one cell can be detected earlier in region 2a, and this
is always the cell with the most fusome [18,19]. This is
particularly obvious in egl and BicD mutants, in which
the fusome perdures longer, and the centrosomes
clearly accumulate in the cell with the largest piece of
fusome remnant [20]. These data strongly suggest
that the ‘original’ fusome marks the future oocyte,
thus supporting the second model. It does not rule out
the possibility that both pro-oocytes can become the
oocyte, but shows that if there is a competition, it is
strongly biased.
What is the link between the asymmetric inheritance
of the fusome and the specification of the oocyte? The
simplest model is that an oocyte-determining factor is
asymmetrically distributed at each division with the
‘original’ fusome into the future oocyte. It has been
proposed that one of the cystoblast centrioles could
stay in contact with the fusome during each division,
and, because of the semi-conservative replication of
the centrosome, could be inherited by the oocyte [6].
Consequently, oocyte determinants could co-segregate
with this centriole. Indeed, it has recently been
shown that such a mechanism mediates the segregation
of dpp and eve mRNAs into specific cells during
the asymmetric divisions of the early embryo of the
snail Ilyanassa obsoleta [34]. Alternatively, the oocyte
could inherit more of some protein or activity associated
with the fusome, and this early bias could initiate
a feedback loop that induces the transport of oocyte
determinants toward this cell.
Figure 2. Formation of the fusome.
During the first incomplete division, the
spectrosome (red) of the cystoblast interacts
with one of the centrosomes (green
and blue spheres) to anchor one pole of
the mitotic spindle (green lines). A fusome
plug (red) forms in the arrested furrow or
ring canal (blue). The spectrosome (or
‘original’ fusome) and the fusome plug
come together to fuse. The direction of
these movements is not known. The same
mechanism is repeated for the second,
third and fourth division: first, one pole of
each mitotic spindle is anchored by the
fusome and a new fusome plug forms in
each ring canal. Then the ring canals
move centripetally for the fusome plugs to
fuse with the central fusome (black
curved arrows). This behavior has several
crucial consequences: cystocyte 1 has
more fusome than the other cystocytes;
the same centrosome (green sphere)
could be inherited by cystocyte 1 from the
first division through the fourth division;
and the fusome always marks the anterior
of cystocyte 1, after the clustering of the
ring canals.
1
?
1 2
?
1 23 4
109
877
1 2
9
77 8
10
88
5 6
3 4 88884447777 333
13 11 12 14
1 2
1 2
43
877
1 2
77 888
5 6
3 4
Division
First
division
Second
division
Third
division
Fourth
division
Clustering
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The Fusome and the Polarisation of the Oocyte
It is not known what drives the movement of the
fusome plugs, but several lines of evidence suggest
that it is microtubule-dependent. Mutations in the
microtubule minus-end directed motor, Dynein, or in its
associated regulator, Lis1, affect the formation of the
fusome, which appears less branched, and mutant
cysts contain fewer than 16 cells [20,31,35]. It is not yet
clear, however, if these mutations affect the movement
and formation of the plugs, or the anchoring of the
mitotic spindles to the fusome, or both. Because of the
centripetal gathering of the ring canals, the cyst adopts
a ‘rosette’ shape [4]. In this configuration, the distance
between the ring canals is minimized [32]. This shape
is later on stabilized by the formation of adherens junctions
around the ring canals, which can be visualized
in EM sections and by the localisation of Armadillo, ECadherin
and Bazooka/Par-3 proteins ([36] and A.P.
Mahowald, personal communication).
It is reasonable to assume that as early as the twocell
stage, the central fusome occupies a fixed position,
while the plugs move towards it. The fusome, therefore,
marks the ‘inner’ part of each cell. In particular in region
2b, the fusome remnant is always located at the anterior
of the oocyte, where all the cytoplasmic components
accumulate to form a Balbiani body [18]. The fusome
thus marks the future anterior side of the oocyte, suggesting
that the anterior–posterior polarity of the oocyte
is inherited from the polarised cyst divisions.
In conclusion, the fusome marks the future oocyte
and also the future anterior side of the oocyte, strongly
suggesting that it plays a direct role in the specification
and polarisation of the oocyte. This has been difficult to
prove, however, because of the earlier functions of the
fusome. For example, hts and αα-spectrin mutants lack
a fusome and often fail to specify an oocyte, but the
divisions also become asynchronous and randomly oriented,
resulting in cysts with a variable number of cells,
and this latter defect could be the primary cause of the
failure in oocyte determination [10,33,37].
The Read-Out of Fusome Polarity: Nurse Cell
Versus Oocyte Differentiation
Although the oocyte appears to be selected early in
region 1, its identity only becomes obvious two days
later, in late region 2a. The differentiation of the cyst in
region 2a is gradual and twofold. In the cytoplasm, the
oocyte accumulates specific components and
organelles, and in the nucleus, it enters meiosis and
arrests in prophase I. Recent results have revealed that
there are at least three pathways to restrict different
aspects of oocyte identity to one cell.
Cytoplasmic Differentiation
One of the main features of cyst differentiation is the
formation of a microtubule array that is polarised toward
the oocyte and extends through the ring canals into the
other cells of the cyst [38]. As the microtubules form
along the fusome, this polarisation has been suggested
to be a direct readout of fusome polarity [19]. Indeed,
microtubules are essential for the determination of the
oocyte, as treatment with the microtubule depolymerising
drug, colchicine, results in egg chambers with 16
nurse cells and no oocyte [39]. Furthermore, none of the
oocyte-specific proteins or mRNAs is asymmetrically
localized within these cysts [38]. Thus, the transport of
these proteins and mRNAs is microtubule-dependent,
suggesting that the formation of the polarised microtubule
cytoskeleton precedes and predicts the specification
of the oocyte.
Although colchicine disrupts the localisation of proteins
and mRNAs to the oocyte, it does not disrupt the
migration of the centrioles or the restriction of meiosis
to one cell [20,22]. Because microtubule-destabilizing
drugs like colchicine only affect dynamic microtubules,
one possibility is that the centrioles migrate along
stable microtubules that are not affected by these
treatments. In support of this view, it has recently been
found that antibodies against acetylated tubulin, a
marker for stable microtubules, label a population of
microtubules associated with the fusome [40]. Furthermore,
mutations in the Drosophila spectroplakin Shot
disrupt these stable microtubules without affecting the
fusome itself, and this blocks the migration of the centrioles
as well as of mRNAs and proteins into one cell.
Shot is a component of the fusome, and contains a
GAS-domain, which has been shown to bind and stabilize
microtubules. This suggests a model in which
Shot assembles stable microtubules on the fusome,
along which the centrioles migrate. The situation is less
clear for the restriction of meiosis, as it is only partially
disrupted in shot mutant germline clones. This raises
the possibility that a third microtubule-independent
pathway directly reads the fusome polarity to control
entry into meiosis (Figure 3).
Independent evidence for three distinct pathways
that restrict different aspects of oocyte identity comes
from the analysis of the phenotypes of Bicaudal-D
(BicD) and egalitarian (egl) mutants. Egl and BicD proteins
co-purify with each other, and null mutations in
either gene give rise to egg chambers with 16 nurse
cells and no oocyte, without affecting the asymmetric
distribution of the fusome [20,32,41–43]. Oocyte-specific
proteins and mRNAs remain evenly distributed
within mutant cysts, and this defect correlates with a
failure to polarize the microtubule cytoskeleton, as
observed with GFP-tubulin and several microtubule
minus-end markers. However, the migration of the centrioles
is normal in BicD and egl null mutants, indicating
that these genes act downstream of the initial asymmetry
that selects the oocyte [20]. This provides further
evidence that the movement of the centrioles occurs by
a mechanism that is distinct from the transport of
mRNAs and proteins.
Although egl and BicD cause identical defects on
mRNA and protein localisation and MT organization,
they have opposite effects on meiosis. In egl mutants, all
16 cells of the cyst enter meiosis and reach the full
pachytene stage, before they revert to the nurse cell fate
[22,27]. In contrast, none of the cells enters meiosis in
BicD mutants, and they all become nurse cells directly
[22]. Thus, an upstream asymmetry must regulate the
activity of the BicD/Egl complex to promote meiosis in
the oocyte and inhibit it in the nurse cells.
In addition to its role in oocyte determination, the
BicD/Egl complex is required for the localisation of
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oskar, K10 and gurken mRNAs in the stage 9–10
oocyte, and for the apical transport of wingless and pair
rule transcripts in the syncytial blastoderm embryo
[43–45]. Injection experiments in the embryo have
demonstrated that BicD and Egl are required to recruit
the dynein motor complex to mRNAs, so that it can
transport them to the apical cytoplasm [45,46]. Consistent
with this, a conserved amino-terminal domain of
the mammalian homologs of BicD has been shown to
bind the dynamitin subunit of the dynein/dynactin
complex and to activate dynein motility, while a
carboxy-terminal region binds to GTP-Rab6, and acts
as an adaptor for the dynein-dependent transport of
Rab6 positive Golgi membranes [47–49].
Very recent results indicate that Egl also plays a role
in coupling cargoes to the dynein motor complex, as a
newly identified carboxy-terminal domain of the protein
has been shown to bind the dynein light chain [50]. Furthermore,
egl mutations that specifically disrupt this
binding site abolish mRNA and protein localisation to
the oocyte, as do mutants in the dynein light chain (dlc)
itself. These results suggest that Egl is an essential
adaptor that links the dynein motor complex to the
mRNAs and proteins that are transported into the
oocyte. Surprisingly, in contrast to the egl null mutations,
neither the egl alleles that disrupt the interaction
with the Dlc nor dlc mutants affect the restriction of
meiosis to one cell. These data strongly support the idea
that Egl has two separable functions in oocyte selection,
an early dynein-independent function that controls
meiosis, and a later function in the dynein-dependent
transport of mRNAs and proteins into the oocyte.
Germline clones mutant for specific alleles of the
dynein–dynactin complex, such as Dhc64C4-19, Lis1E415,
and dynamitinK16109 have similar phenotypes to BicD
and egl, as they give rise to egg chambers with 16 nurse
cells and no oocyte [20,31,35,51,52]. However, the
requirements for the BicD–Egl and dynein–dynactin
complexes are not identical during early oogenesis.
Firstly, the Dynein heavy chain (Dhc) and Lis1 are
required during the cyst divisions in region 1, whereas
BicD and Egl are not. Secondly, unlike BicD and Egl,
Dhc is required for the migration of the centrioles into
the oocyte, as well as the localisation of meiosis and
oocyte-specific mRNAs and proteins [20]. This observation
suggests that dynein may transport the centrioles
into the oocyte along stable microtubules, in addition to
moving mRNAs and proteins along colchicine-sensitive
microtubules. These results need to be interpreted with
caution, however, as Dhc mutants also disrupt the
fusome, which starts to fragment when the cyst enters
region 2a. This may be the primary cause of the total
lack of polarisation in dynein germline clones [20].
In summary, although it was originally thought that
the oocyte was specified by the transport of determinants
along a single polarised microtubule cytoskeleton,
recent results have uncovered a more complex
reality, in which multiple processes function in parallel
to restrict different aspects of oocyte identity to
one cell. However, all of these processes probably
depend on the initial polarity of the fusome, which
may act in three distinct ways to select the oocyte:
First, the fusome organizes a polarised network of
dynamic microtubules that direct the localisation of
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Figure 3. The early steps in the determination
and polarisation of the oocyte.
The early differentiation of the oocyte is a
multistep process. Genes involved at each
step are indicated though the list is not
exhaustive. Regulators of cytoplasmic differentiation
are shown in red, whereas regulators
of the cell cycle are shown in blue.
The top panel shows a 4-cell cyst (αα-spectrin,
in red, marks the fusome and anilin, in
green, marks the ring canals). One cell has
more fusome than the other cells (white
arrow). The panel below shows a 16-cell
cyst after the last division with one cell
having more fusome (white arrow). There
are three different pathways to restrict
oocyte identity to one cell (asterisk). The
left panels show the actin in red and the
synaptonemal complex in green. In the
middle panels, nuclear GFP is shown in
green, and Orb (an oocyte-specific cytoplasmic
protein) is labelled in red. On the
right panels, γγ-tubulin marks the centrosomes
in red, αα-spectrin marks the
fusome in blue, and nuclear GFP is shown
in green. Orb and the centrosomes are
clearly seen migrating from the anterior of
the oocyte to the posterior, revealing the
repolarisation of the oocyte.
Asymmetric divisions
of the cyst
Maintenance of the
fusome integrity
Dynein
α-spectrin, Hts,
Dynein, Lis-1
Klp61F
Bam, Bgcn,
CycA, Orb
Enc/Stwl/Dap
Stage1 checkpoint : Maintenance of the oocyte identity
∗
∗ ∗
∗
∗
∗
BicD and Egl
activity
Microtubules
Centrosome
migration
Proteins and mRNAs
localisationRestriction of meiosis
Cyclin E
Enc/Stwl/Dap
Anterior-posterior
repolarisation
Maintenance in
meiosis
Microtubules
BicD/Egl/Dynein
Baz/Par6/aPKC
Par-1,-5, LKB1
Shot
Dynein, stable
Microtubules ?
Dynein,
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oocyte-specific proteins and mRNAs into one cell, presumably
through the Dynein-dependent transport of
cargoes that are linked to the motor through BicD, Egl
and the Dynein light chain. Second, the fusome also
nucleates stable microtubules that are associated with
Shot, and the centrioles may migrate along these in a
process that could also involve Dynein. Finally, the
fusome appears to regulate a Dynein light chain and
microtubule-independent activity of the BicD/Egl
complex that controls entry into meiosis. Although the
molecular mechanisms are only emerging, these results
also suggest an exciting link between oocyte differentiation,
vesicular trafficking and mRNA transport.
Nuclear Differentiation
The differentiation of the oocyte is marked by several
cell cycle changes and as the future female gamete,
the oocyte is the only cell to complete meiosis. In contrast,
the nurse cells go through several rounds of
endoreplication without mitosis to become highly polyploid.
Interestingly, recent results show that cell cycle
regulators also control the identity of the oocyte.
Cell-Cycle Regulators and Maintenance of the
Oocyte Fate
The first link between the control of the cell cycle and
the identity of the oocyte came from the analysis of
mutations in encore, which encodes a novel protein
with a putative RNA binding domain [53,54]. At 25°C,
one group of alleles induces precisely one more division
in the germline, producing cysts with 32 cells and
one oocyte. At 18°C, a second group of alleles produce
cysts with 16 polyploid nurse cells and no oocyte.
However, in these cysts oskar mRNA initially accumulates
into only one cell, but then diffuses away [53]. This
demonstrates that the initial selection of the oocyte is
not affected by encore mutations, and defines a novel
step in the determination of the oocyte, which is the
maintenance of its fate. Encore interacts with a splicing
factor called Half-pint in yeast two-hybrid assays, and
mutations in half-pint produce cysts that go through
one less division and contain only 8 cells, which sometimes
all become nurse cells [55]. Interestingly, Half-pint
regulates the splicing of ovarian tumor (otu) mRNA, and
mutations in otu also affect the number of cyst divisions,
and the determination of the oocyte [56,57].
Although these genes are not direct regulators of the
cell cycle, they reveal a link between the regulation of
the number of cyst divisions and the determination of
the oocyte. The lack of an oocyte does not seem to be
a consequence of the abnormal number of divisions,
however, as the two phenotypes are produced by different
alleles or under different conditions. The same is
true for mutations in the transcription factor stonewall
(stwl). A high percentage of stwl mutant cysts contain
16 polyploid cells, whereas a lower percentage go
through one extra division to produce 32 cells cysts
[58]. Interestingly, stwl also affects the maintenance of
the oocyte fate rather than its selection, as orb mRNA
initially accumulates normally into one cell.
Direct regulators of the cell cycle also participate in
the determination of the oocyte. Overexpression of
string/cdc25 inhibits the last cyst division to produce
cysts with 8 cells, 50% of which lack an oocyte [59]. A
very similar phenotype is produced by loss-of-function
mutations in tribbles, which is a negative regulator of
string [59]. In contrast, overexpression of tribbles
induces a fifth division to produce cysts with 32 cells
and 2 oocytes. Surprisingly, these effects are the opposite
of those observed in other tissues, where overexpression
of string and mutants in tribbles induce extra
mitoses, suggesting that there is something different
about the regulation of mitosis in the female germ line.
Mutants in cyclin E, which regulates entry into Sphase,
can also affect the number of cyst divisions: a
subset of mutant egg chambers contain only 8 cells
(7 nurse cells and one oocyte), and another subset
have 16 cells, with two or three cells having oocyte-like
nuclear features, such as low ploidy and condensed
chromatin [60]. This phenotype is dominantly suppressed
by mutations in the cyclin-dependent kinase
inhibitor, dacapo (dap) [24]. Furthermore, egg chambers
homozygous mutant for dap contain 16 polyploid
nurse cells. Interestingly, these egg chambers initially
restrict SC, Orb and BicD into one cell, but proteins
and mRNAs diffuse away from the oocyte at stage 1,
instead of moving to the posterior pole. Thus, dap is
also required for the maintenance of the oocyte fate,
rather than its specification [24]. One possible explanation
for this phenotype is that the elevated levels of
cyclin E in dap mutants cause the oocyte to undergo
endocycles and become polyploid, and that the oocyte
loses its cytoplasmic differentiation as a consequence.
Thus, an abnormal cell cycle state blocks the nuclear
and cytoplasmic differentiation of the oocyte, and
induces it to revert to the nurse cell fate, much like the
losing pro-oocyte does normally.
The Spindle Genes: The Meiotic Checkpoint and
RNA Silencing
Mutants in the spindle group genes affect the formation
of the anterior–posterior and dorsal–ventral axes, and
delay the specification of the oocyte [61–64]. This delay
is characterised by an arrest in the formation of the
karyosome, a mispositioning of the oocyte, a failure to
localize the cytoplasmic components of the oocyte to
the posterior pole, and the differentiation of the losing
pro-oocyte as a second oocyte. These defects are transient,
however, and there is usually only one oocyte at
later stages, indicating that the selection of the oocyte
is not abolished but merely slowed down [22,64].
Despite the puzzling pleiotropy of phenotypes, the molecular
characterisation of some of the genes has
revealed a common cause. The spindle-class comprises
the 5 spindle genes, spn-A, -B, -C, -D, -E and
okra, aubergine, vasa, and maelstrom [41,61,65]. spnA,
-B, -D and okra encode homologs of yeast RAD51
and RAD54, which are involved in repairing dsDNA
breaks [66–68]. Furthermore, the patterning defects of
spn-A, -B, -C and -D and okra mutants can be suppressed
by mutations in meiW68, which block the formation
of the dsDNA breaks that initiate recombination,
and also by mutations in meiP41 or chk2, which are
components of the DNA damage checkpoint pathway
[68,69]. Thus, the primary defect in these spindle mutations
appears to be a failure to repair the dsDNA breaks
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formed during recombination, and the patterning
defects and most probably the delay in the oocyte differentiation
are a consequence of the activation of a
meiotic checkpoint pathway (reviewed in [70,71]).
Not all of the spindle genes are involved in the
repair of dsDNA breaks, and it has recently emerged
that a second class, comprising spindle-E, maelstrom,
aubergine, and the newly identified gene
armitage are all required for translational silencing by
micro-RNAs, and RNA interference [72–76]. Mutants
in these genes produce a similar spectrum of phenotypes
to the other spindle mutants, including a delay
in the selection and polarisation of the oocyte, while
spindle-E and maelstrom mutants have also been
found to cause premature cytoplasmic streaming in
the oocyte at stage 9 [77–80]. Interestingly, all of
these mutants also cause the precocious translation
of oskar mRNA in the nurse cells, strongly suggesting
that oskar is regulated by micro-RNAs [76]. These
results indicate that a number of key processes in
development of the germline cysts is regulated by
post-transcriptional gene silencing by microRNAs,
although the relevant mRNA targets in early oogenesis
have not yet been identified.
Early Polarisation of the Oocyte
When a germline cyst reaches region 2b, the oocyte
specific proteins and mRNAs, as well as the centrosomes
and mitochondria have been transported along
the fusome into the presumptive oocyte. These components
remain associated with the fusome remnants
and, therefore, accumulate at the anterior of the oocyte
to form a Balbiani body [18]. When the oocyte moves
through region 3, all of the components of the Balbiani
body disassociate and move around the oocyte
nucleus to form a tight crescent at the posterior cortex.
This movement is the first sign of anterior–posterior
polarity in the oocyte, and is a crucial step in the maintenance
of its identity [26,81].
The importance of this polarisation step was first
demonstrated by the analysis of a null mutation in the
par-1 gene, which specifically disrupts this process
[26]. PAR-1 is the Drosophila homolog of Caenorhabditis
elegans PAR-1, a serine/threonine kinase which is
required for the polarisation of the C. elegans zygote
[82]. In Drosophila, par-1 mutant egg chambers contain
16 nurse cells and lack an oocyte [26,83]. The oocyte
appears to be selected normally, as the centrosomes,
the SC, and Orb accumulate in one cell in region 2b/3
[26]. However, these components do not translocate to
the posterior of the oocyte in region 3 and the oocyte
de-differentiates as a nurse cell, by exiting meiosis and
becoming polyploid.
Anterior–posterior polarity in C. elegans also requires
a number of other PAR proteins: the PDZ-domain proteins
PAR-3, and PAR-6, which form a complex with
atypical protein kinase C at the anterior of the zygote;
the RING finger protein, PAR-2, which is required to
recruit PAR-1 to the posterior cortex, as well as the 14-
3-3 homolog, PAR-5, and the homolog of the mammalian
kinase LKB1, PAR-4, both of which are not
localized [84–89]. It has subsequently been found that
the Drosophila homologs of all of these proteins,
except for PAR-2, which is not conserved, are also
required for the polarisation of the Drosophila oocyte
and the maintenance of its fate [36,83,90,91]. Thus,
there is a striking homology between the earliest polarisation
of the AP axis in worms and flies.
The fact that mutations in the par genes, which are
widely involved in cell polarisation, and in cell cycle regulators
such as dacapo (see above) produce strikingly
similar phenotypes demonstrates that the nuclear and
the cytoplasmic development of the oocyte are intimately
linked, and that stage 1 (region 3) of oogenesis
is a crucial developmental checkpoint (Figure 3). If the
oocyte is not in the appropriate cell cycle state by this
stage or if it fails to polarize along its anterior–posterior
axis, it will revert to the nurse cell fate.
The Par proteins define and maintain complementary
anterior and posterior cortical domains in the C.
elegans embryo, with the PAR-3/PAR-6/aPKC complex
localising to the anterior cortex, and PAR-1 and PAR-2
to the posterior (reviewed in [92]). The spatial relationship
between the Par proteins in the Drosophila oocyte
appears more complex, as most antibodies and GFP
fusions place the PAR-3 homologue Bazooka (Baz) and
aPKC at the adherens junctions that form around the
ring canals, and PAR-1 on the fusome [36,93]. However,
isoform specific antibodies detect Baz at the anterior of
the oocyte and PAR-1 at the posterior pole [94]. Furthermore,
the localisation of these proteins appears to
be interdependent, as Baz extends to the posterior in
par-1 null cysts, and PAR-1 remains anterior in baz null
clones. Recent data have begun to elucidate the molecular
mechanisms that may underlie these mutually
exclusive localisations. PAR-1 has been found to phosphorylate
two sites in Baz to recruit 14-3-3 (PAR-5),
which then disrupts the Baz/PAR-6/aPKC complex by
blocking oligomerization of Baz and its interaction with
aPKC [95,96]. Furthermore, mammalian aPKC can
phosphorylate PAR-1 to inhibit its activity and localisation
to the plasma membrane [97]. Thus, the mutual
antagonism between the Baz/PAR-6/aPKC complex
and PAR-1/14-3-3 may provide a general mechanism to
establish complementary cortical domains in polarised
cells, including the early Drosophila oocyte.
One likely target of the PAR proteins in the
Drosophila oocyte is the microtubule cytoskeleton [98].
In oocytes of par null mutants, the microtubule minusends
fail to switch from the anterior of the nucleus to
the posterior cortex in region 3 [26,90,91,93,94]. Furthermore,
the polarity defects of the par mutants can
be phenocopied by mild colchicine treatments [94]. In
addition, several of the par proteins appear to control
microtubule organization later in oogenesis, because
stage 8–9 oocytes mutant for par-1, lkb1 and 14-3-3e
have a disorganised microtubule cytoskeleton, in
which the minus-ends lie all around the oocyte cortex,
and the plus-ends point toward the center of the
oocyte, instead of the posterior pole [90,91,98]. How
direct the regulation of microtubules by the Par-cassette
is remains unknown. Indeed, BicD, Egl or Dynein
could also be targets as they give very similar phenotypes
to par mutations. In oocytes mutant for hypomorphic
alleles of BicD or Dhc, proteins accumulate
into one cell, but fail to translocate to the posterior
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[22,94]. Similarly, in BicD and egl null mutant oocytes,
the centrosomes remain at the anterior of a pseudooocyte
[20]. It will, therefore, be very informative to
work out the relationships between the Par-cassette,
the BicD/Egl/Dynein complex and the microtubules.
Upstream regulators remain more elusive. In C.
elegans, the sperm entry provides an extrinsic cue that
triggers the Par-dependent polarisation of the embryo
[92]. This cannot be the case in the germarium, as fertilisation
only occurs at the end of oogenesis, about a
week later. In other cell-types, such as epithelial cells,
the localisation of the PAR3/PAR6/aPKC complex is
regulated by an apical complex of Crumbs and Stardust
proteins, as well as by the lateral proteins, Discs Large,
Scribble, and Lethal Giant Larvae [99,100]. Oocyte polarity
appears to be regulated differently, however, as this
early polarisation step is unaffected in germline clones
of null alleles of all these genes (J.-R. H. and Uwe Irion,
unpublished data). These results suggest that the
polarisation of the oocyte may rely not only on intrinsic
mechanisms, but may also require an extrinsic signal.
It is interesting to note that oocyte re-polarisation
occurs exactly when the follicle cells first surround and
contact the oocyte. A signal from the follicle cells
might, therefore, induce the reorganisation of oocyte
polarity in region 3. In support of this hypothesis, Dystroglycan,
which encodes a receptor for multiple extracellular
matrix molecules, has been shown to be
required in the germline for the repolarisation of the
oocyte at this precise stage [101].
In conclusion, the fusome establishes an axis of
polarity as early as in region 1, but this polarity is reorganised
in a Par-dependent manner in region 3.
However, this is not the final polarity of the oocyte, as it
repolarises at stage 7 to define the anterior–posterior
axis of the embryo, raising the question of how these
two polarisation events are related.
The Positioning of the Oocyte Links the Early and
Late Polarisation of the Egg Chamber
The oocyte acquires its final anterior–posterior polarity
in a two step process. First, gurken mRNA is translated
at the posterior of the oocyte and the resulting protein
signals to the adjacent terminal follicle cells to induce
them to adopt a posterior rather than an anterior fate
[102,103]. These posterior follicle cells then send an
unknown signal back to the oocyte at stage 7, which
induces a repolarisation of the microtubule cytoskeleton,
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Figure 4. Similarities between the early
steps of Drosophila and Xenopus oogen-
esis.
In both Drosophila and Xenopus, the
oocyte inherits an anterior–posterior axis of
symmetry from the cyst divisions (for a
review on Xenopus early oogenesis, see
[111–115]). Specific cytoplasmic components
(green) accumulate in a depression
above the nucleus (red) [18,81,111,
113,116]. The oocyte then polarises along
this axis when the somatic follicle cells surround
it. We suggest that a signal coming
from the follicle cells may trigger this polarisation.
This polarisation is clearly seen in
Drosophila with the translocation of specific
cytoplasmic proteins, mRNAs and the
centrosomes to the posterior of the oocyte
(black arrow). The situation is less clear in
Xenopus, as the cell rounds up and seems
to lose any polarity [113,115,117].
However, we propose that the same components
that were located above the
nucleus after the cyst division, are now
part of the Balbiani body on the vegetal
side (green sphere). At the following stage,
these components migrate to the posterior/vegetal
cortex of the oocyte. This has
been clearly demonstrated in Xenopus,
and is here hypothesized for Drosophila
[117–119]. Both oocytes then enter the
vitellogenic stages.
4 rounds of mitosis
with incomplete cytokinesis
Polarity inherited from the
divisions
Follicle cell invasion
(signal?)
Repolarisation
Migration to the cortex
posterior/vegetal
Vitellogenic stages
Cystoblast Oogonium
16-cell cyst 16-cell nest
One cell differentiates
as an oocyte
(region 2b)
All 16 cells become
oocytes
(Stage “0”)
Mid-stage I Mid-stage I
Late stage I/
stage II
Late stage I/
stage II
Mitochondrial
aggregates
vesicles, centrosomes
mitochondria, MT
minus-ends, proteins,
mRNAs
vesicles, centrosomes
mitochondria, MT
minus-ends, proteins,
mRNAs
Early stage I
(oocyte separation)
Early stage I
(cyst encapsulation)
Posterior Vegetal
Drosophila Xenopus
Current Biology
which directs the transport of bicoid mRNA to the anterior
of the oocyte and of oskar mRNA to the posterior.
This chain of polarisation depends on at least two
events taking place in the germarium.
Firstly, gurken mRNA must be localized at the posterior
of the oocyte for Gurken protein to signal to the
adjacent follicle cells (Figure 1). As gurken mRNA is part
of the Balbiani body, its posterior localisation is a consequence
of the Par-dependent repolarisation of the
oocyte in region 3. The posterior localisation of gurken
mRNA, therefore, provides a crucial link between the
early polarisation events taking place in the germarium
and the final anterior–posterior polarity acquired at stage
7. This is best demonstrated by the phenotype of maelstrom
mutants, in which oocyte identity is maintained,
but gurken mRNA fails to switch from the anterior to the
posterior of the oocyte [78,79]. As a consequence, all
the subsequent polarisation steps are disrupted.
Secondly, the oocyte needs to be localized at the
posterior of the egg chamber, in contact with the terminal
follicle cells that are competent to receive the
Gurken signal [104]. Indeed, mutants that disrupt the
movement of the oocyte to the posterior of the cyst
abolish all late anterior–posterior asymmetry, and give
rise to symmetric egg chambers with two sets of anterior
follicle cells, and oocytes that localize bicoid mRNA
to both poles and oskar mRNA to the center. This invariant
localisation of the oocyte is due to an up-regulation
of DE-Cadherin in the oocyte and in somatic cells that
contact the posterior of the oocyte [105,106]. The
oocyte, therefore, out-competes the nurse cells for
adhesion to these posterior follicle cells, and is pulled to
the posterior when the cyst changes shape on entering
region 3. Thus, the AP axis originates from the adhesiveness
of the posterior follicle cells, raising the question
of how these cells are specified and positioned.
The role of the posterior follicle cells in oocyte positioning
raises a paradox. On the one hand, the anterior
and posterior follicle cells are thought to be equivalent
until Gurken signaling breaks the symmetry. On the
other hand, these cells must already be different in the
germarium to up-regulate Cadherin and thus to position
the oocyte. This paradox has recently been resolved
with the discovery that the early difference between the
anterior and posterior follicle cells is only temporary,
and arises from a series of inductive signals that are
transmitted from the anterior follicle cells of the adjacent
older egg chamber to the posterior follicle cells of the
younger, anterior egg chamber [107]. Each cyst signals
through Delta to induce the differentiation of the anterior
and posterior pairs of polar follicle cells at each end
of the egg chamber, but the anterior polar cells differentiate
much earlier than the posterior ones. These cells
turn on Unpaired, the ligand for the JAK/STAT pathway,
which induces the adjacent anterior polar stalk cell precursors
to become stalk cells, and it is thought that
these cells then up-regulate Cadherin to adhere to the
oocyte of the adjacent younger egg chamber. Therefore,
the anterior–posterior axis is established by a relay
mechanism in which each cyst induces the positioning
of the oocyte of the next cyst.
These last results emphasize the importance of the
communication between the germline and the somatic
cells to coordinate the formation of an egg chamber.
Signals from the germline to the somatic cells control
the migration and differentiation of the follicle cells,
whereas signals from the somatic cells to the germline
are required to position and polarize the oocyte.
Conclusion and Perspectives
Although we have focused on Drosophila oogenesis in
this review, the formation of a germline cyst seems
widely conserved throughout the animal kingdom
[108]. In particular, the first steps of Drosophila oogenesis
show a striking similarity to the early steps of
Xenopus oogenesis (Figure 4) [109]. Furthermore, PAR-
6b has recently been found to localize to the animal
pole of unfertilised mouse eggs, suggesting that PAR
proteins may play a conserved role in oocyte polarity
in mammals [110]. Thus, the eggs of many species,
including vertebrates, may be polarised much earlier
than previously thought, and the Drosophila egg
chamber will provide a useful paradigm for understanding
this process.
Acknowledgments
The writing of this review was greatly helped by many
people in the field, who openly discussed ideas and/or
shared results before publication. In particular, we are
very grateful to the following people: Adelaide Carpenter,
David Ish-Horowicz, Scott Hawley, Mary Lilly, Ruth
Lehmann, Anthony Mahowald, Scott Page and Allan
Spradling. J.-R.H. is supported by the EMBO and
D.St.J. by the Wellcome Trust.
Supplemental data
Supplemental data are available at http://www.current-
biology.com/cgi/content/full/14/11/R438/DC1/
References
1. Gerhart, J., and Kirschner, M. (1997). Cells, embryos and evolution
(Blackwell Science).
2. van Eeden, F., and St Johnston, D. (1999). The polarisation of the
anterior–posterior and dorsal–ventral axes during Drosophila oogenesis.
Curr. Opin. Genet. Dev. 9, 396–404.
3. Riechmann, V., and Ephrussi, A. (2001). Axis formation during
Drosophila oogenesis. Curr. Opin. Genet. Dev. 11, 374–383.
4. Telfer, W. (1975). Development and physiology of the oocyte-nurse
cell syncitium. Adv. Insect Physiol. 11, 223–319.
5. Deng, W., and Lin, H. (2001). Asymmetric germ cell division and
oocyte determination during Drosophila oogenesis. Int. Rev. Cytol.
203, 93–138.
6. Theurkauf, W.E. (1994). Microtubules and cytoplasm organisation
during Drosophila oogenesis. Dev. Biol. 165, 352–360.
7. Spradling, A. (1993). Developmental genetics of oogenesis. In The
Development of Drosophila melanogaster, Volume 1, 1 Edition, M.
Bate and A. Martinez-Arias, eds. (New York: Cold Spring Harbor
Laboratory press), pp. 1–70.
8. Spradling, A., Drummond-Barbosa, D., and Kai, T. (2001). Stem cells
find their niche. Nature 414, 98–104.
9. Gonzalez-Reyes, A. (2003). Stem cells, niches and cadherins: a view
from Drosophila. J. Cell Sci. 116, 949–954.
10. Lin, H., Yue, L., and Spradling, A.C. (1994). The Drosophila fusome,
a germline-specific organelle, contains membrane skeletal proteins
and functions in cyst formation. Development 120, 947–956.
11. Buning, J. (1994). The Insect Ovary: Ultrastructure, Previtellogenic
Growth and Evolution (New-York: Chapman and Hall).
12. Suter, B., Romberg, L.M., and Steward, R. (1989). Bicaudal-D, a
Drosophila gene involved in developmental asymmetry: localized
transcript accumulation in ovaries and sequence similarity to
myosin heavy chain tail domains. Genes Dev. 3, 1957–1968.
13. Wharton, R.P., and Struhl, G. (1989). Structure of the Drosophila
Bicaudal-D protein and its role in localizing the posterior determinant
nanos. Cell 59, 881–892.
Review
R446
14. Lantz, V., Chang, J., Horabin, J., Bopp, D., and Schedl, P. (1994).
The Drosophila orb RNA-binding protein is required for the formation
of the egg chamber and establishment of polarity. Genes Dev.
8, 598–613.
15. van Eeden, F.J.M., Palacios, I.M., Petronczki, M., Weston, M.J.D.,
and St Johnston, D. (2001). Barentsz is essential for the posterior
localization of oskar mRNA and colocalizes with it to the posterior.
J. Cell Biol. 154, 511–524.
16. Keyes, L.N., and Spradling, A.C. (1997). The Drosophila gene
fs(2)cup interacts with otu to define a cytoplasmic pathway required
for the structure and function of germ-line chromosomes. Development
124, 1419–1431.
17. Ephrussi, A., Dickinson, L.K., and Lehmann, R. (1991). oskar organizes
the germ plasm and directs localization of the posterior determinant
nanos. Cell 66, 37–50.
18. Cox, R.T., and Spradling, A.C. (2003). A Balbiani body and the
fusome mediate mitochondrial inheritance during Drosophila oogenesis.
Development 130, 1579–1590.
19. Grieder, N.C., de Cuevas, M., and Spradling, A.C. (2000). The
fusome organizes the microtubule network during oocyte differentiation
in Drosophila. Development 127, 4253–4264.
20. Bolivar, J., Huynh, J.R., Lopez-Schier, H., González, C., St Johnston,
D., and González-Reyes, A. (2001). Centrosome migration into
the Drosophila oocyte is independent of BicD, egl and of the organisation
of the microtubule cytoskeleton. Development 128,
1889–1897.
21. Carpenter, A. (1975). Electron microscopy of meiosis in Drosophila
melanogaster females. I Structure, arrengement, and temporal
change of the synaptonemal complex in wild-type. Chromosoma
51, 157–182.
22. Huynh, J.R., and St Johnston, D. (2000). The role of BicD, Egl, Orb
and the microtubules in the restriction of meiosis to the Drosophila
oocyte. Development 127, 2785–2794.
23. Page, S.L., and Hawley, R.S. (2001). c(3)G encodes a Drosophila
synaptonemal complex protein. Genes Dev. 15, 3130–3143.
24. Hong, A., Lee-Kong, S., Iida, T., Sugimura, I., and Lilly, M.A. (2003).
The p27(cip/kip) ortholog dacapo maintains the Drosophila oocyte
in prophase of meiosis I. Development 130, 1235–1242.
25. Dej, K.J., and Spradling, A.C. (1999). The endocycle controls nurse
cell polytene chromosome structure during Drosophila oogenesis.
Development 126, 293–303.
26. Huynh, J.R., Shulman, J.M., Benton, R., and St Johnston, D. (2001).
PAR-1 is required for the maintenance of oocyte fate in Drosophila.
Development 128, 1201–1209.
27. Carpenter, A. (1994). egalitarian and the choice of cell fates in
Drosophila melanogaster oogenesis. In Germline development, J.
Marsh and J. Goode, eds. (Chichester: John Wiley & Sons), pp.
223–246.
28. Lin, H., and Spradling, A.C. (1995). Fusome asymmetry and oocyte
determination in Drosophila. Dev. Genet. 16, 6–12.
29. Storto, P., and King, R. (1989). The role of polyfusomes in generating
branched chains of cystocytes during Drosophila oogenesis.
Dev. Genet. 10, 70–86.
30. Deng, W., and Lin, H. (1997). Spectrosomes and fusomes anchor
mitotic spindles during asymmetric germ cell divisions and facilitate
the formation of a polarized microtubule array for oocyte specification
in Drosophila. Dev. Biol. 189, 79–94.
31. McGrail, M., and Hays, T.S. (1997). The microtubule motor cytoplasmic
dynein is required for spindle orientation during germline
cell divisions and oocyte differentiation in Drosophila. Development
124, 2409–2419.
32. de Cuevas, M., and Spradling, A.C. (1998). Morphogenesis of the
Drosophila fusome and its implications for oocyte specification.
Development 125, 2781–2789.
33. de Cuevas, M., Lee, J.L., and Spradling, A.C. (1996). a-spectrin is
required for germline cell division and differentiation in the
Drosophila ovary. Development 124, 3959–3968.
34. Lambert, J.D., and Nagy, L.M. (2002). Asymmetric inheritance of
centrosomally localized mRNAs during embryonic cleavages.
Nature 420, 682–686.
35. Liu, Z., Xie, T., and Steward, R. (1999). Lis1, the Drosophila homolog
of a human lissencephaly disease gene, is required for germline cell
division and oocyte differentiation. Development 126, 4477–4488.
36. Huynh, J.-R., Petronczki, M., Knoblich, J.A., and St Johnston, D.
(2001). Bazooka and PAR-6 are required with PAR-1 for the maintenance
of oocyte fate in Drosophila. Curr. Biol. 11, 901–906.
37. Yue, L., and Spradling, A. (1992). hu-li tai shao, a gene required for
ring canal formation during Drosophila oogenesis, encodes a
homolog of adducin. Genes Dev. 6, 2443–2454.
38. Theurkauf, W., Alberts, M., Jan, Y., and Jongens, T. (1993). A central
role for microtubules in the differentiation of Drosophila oocytes.
Development 118, 1169–1180.
39. Koch, E., and Spitzer, R. (1983). Multiple effects of colchicine on
oogenesis in Drosophila; Induced sterility and switch of potencial
oocyte to nurse-cell developmental pathway. Cell Tissue Res. 228,
21–32.
40. Roper, K., and Brown, N.H. (2004). A Spectraplakin Is Enriched on
the Fusome and Organizes Microtubules during Oocyte Specification
in Drosophila. Curr. Biol. 14, 99–110.
41. Schüpbach, T., and Wieschaus, E. (1991). Female sterile mutations
on the second chromosome of Drosophila melanogaster. II. Mutations
blocking oogenesis or altering egg morphology. Genetics 129,
1119–1136.
42. Ran, B., Bopp, R., and Suter, B. (1994). Null alleles reveal novel
requirements for Bic-D during Drosophila oogenesis. Development
120, 1233–1242.
43. Mach, J.M., and Lehmann, R. (1997). An Egalitarian-BicaudalD
complex is essential for oocyte specification and axis determination
in Drosophila. Genes Dev. 11, 423–435.
44. Swan, A., and Suter, B. (1996). Role of Bicaudal-D in patterning the
Drosophila egg chamber in mid-oogenesis. Development 122,
3577–3586.
45. Bullock, S.L., and Ish-Horowicz, D. (2001). Conserved signals and
machinery for RNA transport in Drosophila oogenesis and embryogenesis.
Nature 414, 611–616.
46. Wilkie, G.S., and Davis, I. (2001). Drosophila wingless and pair-rule
transcripts localise apically by dynein-mediated transport of RNA
particles. Cell 105, 209–219.
47. Hoogenraad, C.C., Akhmanova, A., Howell, S.A., Dortland, B.R., De
Zeeuw, C.I., Willemsen, R., Visser, P., Grosveld, F., and Galjart, N.
(2001). Mammalian Golgi-associated Bicaudal-D2 functions in the
dynein-dynactin pathway by interacting with these complexes.
EMBO J. 20, 4041–4054.
48. Matanis, T., Akhmanova, A., Wulf, P., Del Nery, E., Weide, T.,
Stepanova, T., Galjart, N., Grosveld, F., Goud, B., De Zeeuw, C.I., et
al. (2002). Bicaudal-D regulates COPI-independent Golgi-ER transport
by recruiting the dynein-dynactin motor complex. Nat. Cell
Biol. 4, 986–992.
49. Hoogenraad, C.C., Wulf, P., Schiefermeier, N., Stepanova, T.,
Galjart, N., Small, J.V., Grosveld, F., de Zeeuw, C.I., and
Akhmanova, A. (2003). Bicaudal D induces selective dynein-mediated
microtubule minus end-directed transport. EMBO J. 22,
6004–6015.
50. Navarro, C., Puthalakath, H., Adams, J.M., Strasser, A., and
Lehmann, R. (2004). Egalitarian binds dynein light chain to establish
oocyte polarity and maintain oocyte fate. Nat. Cell Biol., 6, 427–435.
51. Swan, A., Nguyen, T., and Suter, B. (1999). Drosophila
Lissencephaly-1 functions with Bic-D and dynein in oocyte determination
and nuclear positioning. Nat. Cell Biol. 1, 444–449.
52. Januschke, J., Gervais, L., Dass, S., Kaltschmidt, J.A., LopezSchier,
H., St Johnston, D., Brand, A.H., Roth, S., and Guichet, A.
(2002). Polar transport in the Drosophila oocyte requires Dynein and
Kinesin I cooperation. Curr. Biol. 12, 1971–1981.
53. Hawkins, N.C., Thorpe, J., and Schüpbach, T. (1996). encore, a
gene required for the regulation of germ line mitosis and oocyte differentiation
during Drosophila oogenesis. Development 122,
281–290.
54. Buskirk, C.V., Hawkins, N.C., and Schüpbach, T. (2000). Encore is a
member of a novel family of proteins and affects multiple processes
in Drosophila oogenesis. Development 127, 4753–4762.
55. Van Buskirk, C., and Schupbach, T. (2002). Half pint regulates alternative
splice site selection in Drosophila. Dev. Cell 2, 343–353.
56. Storto, P.D., and King, R.C. (1988). Multiplicity of functions for the
otu gene products during Drosophila oogenesis. Dev. Genet. 9,
91–120.
57. King, R.C., and Storto, P.D. (1988). The role of the otu gene in
Drosophila oogenesis. Bioessays 8, 18–24.
58. Clark, K.A., and McKearin, D.M. (1996). The Drosophila stonewall
gene encodes a putative transcription factor essential for germ cell
development. Development 122, 937–950.
59. Mata, J., Curado, S., Ephrussi, A., and Rorth, P. (2000). Tribbles
coordinates mitosis and morphogenesis in Drosophila by regulating
string/CDC25 proteolysis. Cell 101, 511–522.
60. Lilly, M.A., and Spradling, A.C. (1996). The Drosophila endocycle is
controlled by Cyclin E and lacks a checkpoint ensuring S-phase
completion. Genes Dev. 10, 2514–2526.
61. Tearle, R., and Nüsslein-Volhard, C. (1987). Tübingen mutants and
stocklist. Drosoph. Inf. Serv. 66, 209–269.
62. González-Reyes, A., and St Johnston, D. (1994). Role of oocyte
position in the establishment of anterior–posterior polarity in
Drosophila. Science 266, 639–642.
63. Gillespie, D.E., and Berg, C.E. (1995). homeless is required for RNA
localization in Drosophila oogenesis and encodes a new member of
the DE-H family of RNA-dependent ATPases. Genes Dev. 9,
2495–2508.
Current Biology
R447
64. González-Reyes, A., Elliot, H., and St Johnston, D. (1997). Oocyte
determination and the origin of polarity in Drosophila: the role of the
spindle genes. Development 124, 4927–4937.
65. Findley, S.D., Tamanaha, M., Clegg, N.J., and Ruohola-Baker, H.
(2003). Maelstrom, a Drosophila spindle-class gene, encodes a
protein that colocalizes with Vasa and RDE1/AGO1 homolog,
Aubergine, in nuage. Development 130, 859–871.
66. Ghabrial, A., Ray, R.P., and Schüpbach, T. (1998). okra and spindleB
encode components of the RAD52 DNA repair pathway and affect
meiosis and patterning in Drosophila oogenesis. Genes Dev. 12,
2711–2723.
67. Abdu, U., Gonzalez-Reyes, A., Ghabrial, A., and Schupbach, T.
(2003). The Drosophila spn-D gene encodes a RAD51C-like protein
that is required exclusively during meiosis. Genetics 165, 197–204.
68. Staeva-Vieira, E., Yoo, S., and Lehmann, R. (2003). An essential role
of DmRad51/SpnA in DNA repair and meiotic checkpoint control.
EMBO J. 22, 5863–5874.
69. Ghabrial, A., and Schüpbach, T. (1999). Activation of a meiotic
checkpoint regulates translation of Gurken during Drosophila oogenesis.
Nat. Cell Biol. 1, 354–357.
70. Gonzalez-Reyes, A. (1999). DNA repair and pattern formation come
together. Nat. Cell Biol. 1, E150–E152.
71. Morris, J., and Lehmann, R. (1999). Drosophila oogenesis: versatile
spn doctors. Curr. Biol. 9, R55–R58.
72. Kennerdell, J.R., Yamaguchi, S., and Carthew, R.W. (2002). RNAi is
activated during Drosophila oocyte maturation in a manner dependent
on aubergine and spindle-E. Genes Dev. 16, 1884–1889.
73. Aravin, A.A., Naumova, N.M., Tulin, A.V., Vagin, V.V., Rozovsky,
Y.M., and Gvozdev, V.A. (2001). Double-stranded RNA-mediated
silencing of genomic tandem repeats and transposable elements in
the D. melanogaster germline. Curr. Biol. 11, 1017–1027.
74. Findley, S.D., Tamanaha, M., Clegg, N.J., and Ruohola-Baker, H.
(2003). Maelstrom, a Drosophila spindle-class gene, encodes a
protein that colocalizes with Vasa and RDE1/AGO1 homolog,
Aubergine, in nuage. Development 130, 859–871.
75. Tomari, Y., Du, T., Haley, B., Schwarz, D.S., Bennett, R., Cook, H.A.,
Koppetsch, B.S., Theurkauf, W.E., and Zamore, P.D. (2004). RISC
assembly defects in the Drosophila RNAi mutant armitage. Cell 116,
831–841.
76. Cook, H.A., Koppetsch, B.S., Wu, J., and Theurkauf, W.E. (2004).
The Drosophila SDE3 homolog armitage is required for oskar mRNA
silencing and embryonic axis specification. Cell 116, 817–829.
77. Gillespie, D., and Berg, C. (1995). Homeless is required for RNA
localization in Drosophila oogenesis and encodes a new member of
the DE-H family of RNA-dependent ATPases. Genes Dev. 9,
2495–2508.
78. Clegg, N.J., Findley, S.D., Mahowald, A.P., and Ruohola-Baker, H.
(2001). maelstrom is required to position the MTOC in stage 2–6
Drosophila oocytes. Dev. Genes Evol. 211, 44–48.
79. Clegg, N.J., Frost, D.M., Larkin, M.K., Subrahmanyan, L., Bryant, Z.,
and Ruohola-Baker, H. (1997). maelstrom is required for an early
step in the establishment of Drosophila oocyte polarity: posterior
localization of grk mRNA. Development 124, 4661–4671.
80. Martin, S.G., Leclerc, V., Smith-Litiere, K., and St Johnston, D.
(2003). The identification of novel genes required for Drosophila
anteroposterior axis formation in a germline clone screen using
GFP-Staufen. Development 130, 4201–4215.
81. Paré, C., and Suter, B. (2000). Subcellular localization of Bic-D::GFP
is linked to an asymmetric oocyte nucleus. J. Cell Sci. 113,
2119–2127.
82. Guo, S., and Kemphues, K.J. (1995). par-1, a gene required for
establishing polarity in C. elegans embryos, encodes a putative
Ser/Thr kinase that is asymmetrically distributed. Cell 81, 611–620.
83. Cox, D.N., Lu, B., Sun, T., Williams, L.T., and Jan, Y.N. (2001).
Drosophila par-1 is required for oocyte differentiation and microtubule
organization. Curr. Biol. 11, 75–87.
84. Boyd, L., Guo, S., Levitan, D., Stinchcomb, D.T., and Kemphues,
K.J. (1996). PAR-2 is asymmetrically distributed and promotes
association of P granules and PAR-1 with the cortex in C. elegans
embryos. Development 122, 3075–3084.
85. Etemad-Moghadam, B., Guo, S., and Kemphues, K.J. (1995). Asymmetrically
distributed PAR-3 protein contributes to cell polarity and
spindle alignment in early C. elegans embryos. Cell 83, 743–752.
86. Watts, J.L., Etemad-Moghadam, B., Guo, S., Boyd, L., Draper, B.W.,
Mello, C.C., Priess, J.R., and Kemphues, K.J. (1996). par-6, a gene
involved in the establishment of asymmetry in early C. elegans
embryos, mediates the asymmetric localization of PAR-3. Development
122, 3133–3140.
87. Hung, T.J., and Kemphues, K.J. (1999). PAR-6 is a conserved PDZ
domain-containing protein that colocalizes with PAR-3 in
Caenorhabditis elegans embryos. Development 126, 127–135.
88. Joberty, G., Petersen, C., Gao, L., and Macara, I.G. (2000). The cellpolarity
protein Par6 links Par3 and atypical protein kinase C to
Cdc42. Nat. Cell Biol. 2, 531–539.
89. Lin, D., Edwards, A.S., Fawcett, J.P., Mbamalu, G., Scott, J.D., and
Pawson, T. (2000). A mammalian PAR-3-PAR-6 complex implicated
in Cdc42/Rac1 and aPKC signalling and cell polarity. Nat. Cell Biol.
2, 540–547.
90. Benton, R., Palacios, I.M., and St Johnston, D. (2002). Drosophila
14–3-3/PAR-5 is an essential mediator of PAR-1 function in axis formation.
Dev. Cell 3, 659–671.
91. Martin, S.G., and St Johnston, D. (2003). A role for Drosophila LKB1
in anterior–posterior axis formation and epithelial polarity. Nature
421, 379–384.
92. Lyczak, R., Gomes, J.E., and Bowerman, B. (2002). Heads or tails:
cell polarity and axis formation in the early Caenorhabditis elegans
embryo. Dev. Cell 3, 157–166.
93. Cox, D.N., Seyfried, S.A., Jan, L.Y., and Jan, Y.N. (2001). Bazooka
and atypical protein kinase C are required to regulate oocyte differentiation
in the Drosophila ovary. Proc. Natl. Acad. Sci. USA 98,
14475–14480.
94. Vaccari, T., and Ephrussi, A. (2002). The fusome and microtubules
enrich Par-1 in the oocyte, where it effects polarization in conjunction
with Par-3, BicD, Egl, and dynein. Curr. Biol. 12, 1524–1528.
95. Benton, R., Palacios, I.M., and St Johnston, D. (2002). Drosophila
14–3-3/PAR-5 is an essential mediator of PAR-1 function in axis formation.
Dev. Cell 3, 659–671.
96. Benton, R., and St Johnston, D. (2003). Drosophila PAR-1 and 14–3-
3 inhibits Bazooka/Par-3 to establish complementary cortical
domains in polarized cells. Cell 115, 691–704.
97. Hurov, J.B., Watkins, J.L., and Piwnica-Worms, H. (2004). Atypical
PKC phosphorylates PAR-1 kinases to regulate localization and
activity. Curr. Biol. 14, 736–741.
98. Shulman, J.M., Benton, R., and St. Johnston, D. (2000). The
Drosophila homolog of C. elegans PAR-1 organizes the oocyte
cytoskeleton and directs oskar mRNA localisation to the posterior
pole. Cell 101, 1–20.
99. Bilder, D., Schober, M., and Perrimon, N. (2003). Integrated activity
of PDZ protein complexes regulates epithelial polarity. Nat. Cell
Biol. 5, 53–58.
100. Tanentzapf, G., and Tepass, U. (2003). Interactions between the
crumbs, lethal giant larvae and bazooka pathways in epithelial
polarization. Nat. Cell Biol. 5, 46–52.
101. Deng, W.M., Schneider, M., Frock, R., Castillejo-Lopez, C., Gaman,
E.A., Baumgartner, S., and Ruohola-Baker, H. (2003). Dystroglycan
is required for polarizing the epithelial cells and the oocyte in
Drosophila. Development 130, 173–184.
102. González-Reyes, A., Elliott, H., and St Johnston, D. (1995). Polarization
of both major body axes in Drosophila by gurken-torpedo
signalling. Nature 375, 654–658.
103. Roth, S., Neuman-Silberberg, F.S., Barcelo, G., and Schüpbach, T.
(1995). cornichon and the EGF receptor signaling process are necessary
for both anterior–posterior and dorsal–ventral pattern formation
in Drosophila. Cell 81, 967–978.
104. Gonzalez-Reyes, A., and St Johnston, D. (1994). Role of oocyte
position in establishment of anterior–posterior polarity in
Drosophila. Science 266, 639–642.
105. Godt, D., and Tepass, U. (1998). Drosophila oocyte localisation is
mediated by differential cadherin-based adhesion. Nature 395,
387–391.
106. Gonzalez-Reyes, A., and St Johnston, D. (1998). The Drosophila AP
axis is polarised by the cadherin-mediated positioning of the
oocyte. Development 125, 3635–3644.
107. Torres, I.L., Lopez-Schier, H., and St Johnston, D. (2003). A
Notch/Delta-dependent relay mechanism establishes anterior–posterior
polarity in Drosophila. Dev. Cell 5, 547–558.
108. Pepling, M.E., de Cuevas, M., and Spradling, A.C. (1999). Germline:
a conserved phase of germ cell development. Trends Cell Biol. 9,
257–262.
109. Kloc, M., Bilinski, S., Dougherty, M.T., Brey, E.M., and Etkin, L.D.
(2004). Formation, architecture and polarity of female germline cyst
in Xenopus. Dev. Biol., 266, 46-61.
110. Vinot, S., Le, T., Maro, B., and Louvet-Vallee, S. (2004). Two PAR6
proteins become asymmetrically localized during establishment of
polarity in mouse oocytes. Curr. Biol. 14, 520–525.
111. Coggins, L.W. (1973). An ultrastructural and radioautographic study
of early oogenesis in the toad Xenopus laevis. J. Cell Sci. 12, 71–93.
112. Klymkowsky, M.W., and Karnovsky, A. (1994). Morphogenesis and
the cytoskeleton: studies of the Xenopus embryo. Dev. Biol. 165,
372–384.
113. Gard, D.L. (1995). Axis formation during amphibian oogenesis:
reevaluating the role of the cytoskeleton. Curr. Top. Dev. Biol. 30,
215–252.
Review
R448
114. Gard, D.L., Affleck, D., and Error, B.M. (1995). Microtubule organization,
acetylation, and nucleation in Xenopus laevis oocytes: II. A
developmental transition in microtubule organization during early
diplotene. Dev. Biol. 168, 189–201.
115. Gard, D.L., Cha, B.J., and Schroeder, M.M. (1995). Confocal
immunofluorescence microscopy of microtubules, microtubuleassociated
proteins, and microtubule-organizing centers during
amphibian oogenesis and early development. Curr. Top. Dev. Biol.
31, 383–431.
116. Kloc, M., Larabell, C., Chan, A.P., and Etkin, L.D. (1998). Contribution
of METRO pathway localized molecules to the organization of
the germ cell lineage. Mech. Dev. 75, 81–93.
117. Heasman, J., Quarmby, J., and Wylie, C.C. (1984). The mitochondrial
cloud of Xenopus oocytes: the source of germinal granule material.
Dev. Biol. 105, 458–469.
118. Schnapp, B.J., Arn, E.A., Deshler, J.O., and Highett, M.I. (1997). RNA
localization in Xenopus oocytes. Semin. Cell Dev. Biol. 8, 529–540.
119. King, M.L., Zhou, Y., and Bubunenko, M. (1999). Polarizing genetic
information in the egg: RNA localization in the frog oocyte. Bioessays
21, 546–557.
Current Biology
R449