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Logo of nihpaAbout Author manuscriptsSubmit a manuscriptNIH Public Access; Author Manuscript; Accepted for publication in peer reviewed journal;
Semin Cell Dev Biol. Author manuscript; available in PMC Jun 1, 2009.
Published in final edited form as:
PMCID: PMC2430523
NIHMSID: NIHMS49237

Drosophila follicle cells: morphogenesis in an eggshell

Abstract

Epithelial morphogenesis is important for organogenesis and pivotal for carcinogenesis, but mechanisms that control it are poorly understood. The Drosophila follicular epithelium is a genetically tractable model to understand these mechanisms in vivo. This epithelium of follicle cells encases germline cells to create an egg. In this review, we summarize progress toward understanding mechanisms that maintain the epithelium or permit migrations essential for oogenesis. Cell-cell communication is important, but the same signals are used repeatedly to control distinct events. Understanding intrinsic mechanisms that alter responses to developmental signals will be important to understand regulation of cell shape and organization.

Keywords: review, oogenesis, migration, epithelial-mesenchymal transition, apical-basal polarity

Introduction

Studies in cultured cells have driven our understanding of mechanisms that regulate cell shape and migration. The application of this mechanistic information to living tissues remains a challenge. Genetic studies in the model invertebrate Drososphila melanogaster have been instrumental for illuminating how cell biological mechanisms are orchestrated during tissue morphogenesis. An important tissue for such studies is the follicular epithelium that surrounds the maturing fly oocyte. Elegant developmental and genetic studies have revealed that this epithelium develops coordinately with the oocyte and establishes the body axes of the resultant embryo. Cell-cell communication coordinates the morphogenesis of the follicular epithelium with the associated germ cells, in much the same way as epithelial-mesenchymal interactions drive tissue formation in vertebrates.

Once formed, the follicular epithelium is subdivided to yield distinct regions of epithelial organization, small groups of migratory follicle cells, and ultimately a secretory epithelium that progressively deposits the layers of the eggshell. The follicular epithelium has been an important mode for studies of receptor tyrosine kinase signal transduction (recently reviewed by [1-3]), epithelial cell polarity (recently reviewed by [4, 5]), and stem cell/niche biology (recently reviewed by [6, 7]). Here we highlight recent progress towards understanding the mechanisms of morphogenesis.

Overview of oogenesis

Each adult ovary has 14-16 ovarioles, which contain strings of developing oocytes of progressive ages (Fig. 1). At the end of each ovariole are 2-3 germline stem cells (GSCs) in a structure called the germarium. Differentiating daughters of GSCs, the cystoblasts, undergo four mitotic divisions to form a 16-cell syncytium, or cyst. One germ cell becomes an oocyte; the remaining 15 become nurse cells. Nearby, follicle cell stem cells (FCSCs) give rise to precursor follicle cells [8]. About 16 precursor follicle cells invade between cysts, cease division and become pre-polar cells, which ultimately become polar cells and stalk cells [9-11]. Other precursor cells form a simple epithelium around the cyst, creating an egg chamber. Differentiation of the 5-8 stalk cells separates the newly formed egg chamber from the germarium. The oocyte grows substantially, due to uptake of yolk protein synthesized in the follicle cells and fat, and occupies almost half the egg chamber by stage 10A [12-14].

Figure 1
Cell outlines visualized by phalloidin staining of the actin cytoskeleton (white). Stages of egg chambers are indicated by the numbers; and G is for germarium. Anterior is to the left. Follicle cells form an epithelium to cover the underlying germline ...

The follicle cells (FCs) remain in a cuboidal epithelium through stage 8. Beginning in stage 9, they reorganize in a series of migrations. The 6-10 anterior-most FCs become the border cells, which migrate through the nurse cells to the oocyte. Another fifty anterior FCs form a squamous epithelium overlying the nurse cells. Most FCs become a columnar epithelium over the oocyte. During stage 10B, FCs move inward to cover the anterior end of the oocyte.

During stages 10B to stage 14, nurse cells transfer their cytoplasmic contents to the oocyte [15]. The FCs deposit the vitelline membrane and eggshell over the oocyte. The oocyte completes maturation; nurse cells and FCs undergo apoptosis. The anterior end of a mature egg has a pair of dorsal appendages for embryonic respiration, an operculum for larval exit, and a micropyle for sperm entry (Fig. 2) [12, 13, 16].

Figure 2
Scanning electron micrograph of an eggshell (dorso-lateral view). Image adapted from Ref. [5] copyright 2000 Wiley-Liss.

The germarium: a complex niche with multiple stem cell types

The adult ovary develops from a larval primordium, which contains mitotically active populations of mesodermally-derived somatic cells and primordial germ cells (PGCs). A central mass of mesenchymal and primordial germ cells forms the ovarioles. During larval development, signaling between PGCs and intermingled somatic cells balances their numbers [17]. At the larval to pupal transition, posterior PGCs differentiate into cystoblasts, which become the first eggs laid by the adult female.

Formation of the niche and stem cells

Ovariole formation starts in third instar, when somatic cell migrations form ovarian structures, the niche for GSCs, and the FCSCs [13]. The central mesenchymal cells form a niche that maintains anterior PGCs in an undifferentiated state [20]; these PGCs become the GSCs [13, 14]. The terminal filaments, which sit at the anterior end of each germarium (Fig. 3), differentiate first, forming an array of 8-9 cell stacks. The Bric-a-brac (Bab) transcription factor is expressed in niche-forming somatic cells, and required for terminal filament formation [18, 19], which can occur in the absence of PGCs [13]. Terminal filament cells form clusters by cell intercalation and cell recruitment [18, 19]. The organization of terminal filament cells requires the transcription factor Engrailed, but subsequent cell flattening does not [21].

Figure 3
A. Diagram of a Drosophila germarium. This illustration is shows the location of germline stem cells (GSCs; red) and follicle cell stem cells (FCSCs; yellow) and their niche cells. Two GSCs cells are located close to their niche composed of cap cell (bright ...

The mature germarium

Niche cells, GSCs and mitotically dividing cystoblasts lie in region 1 of the germarium (Fig. 3). The newly formed 16-cell cyst moves into region 2a as oocyte differentiation progresses [reviewed in 23]. FCSCs are located at the 2a/2b boundary [8]. In region 2b, the cyst reorganizes with the oocyte located in its center. Here, precursor FCs envelope the cyst, replacing the escort cells. A nascent egg chamber comprises region 3 [12-14, 24].

In adults, a terminal filament and cap cells form a niche that maintains GSCs in every germarium [6, 7, 17, 22, 25-27]. At the base of the terminal filament, a cluster of cap cells contact the GSCs. GSCs are identified by the location of their fusomes, a large membrane cytoskeleton complex [38, 39]. As differentiating daughters of GSCs move away from the cap cells, somatic escort cells wrap around them and support cystoblast differentiation [22]. The terminal filament and cap cells promote GSC self-renewal and prevent their differentiation. Somatic cells regulate germ cell proliferation and differentiation in other well-studied animal models. Mammalian Sertoli and Leydig cells regulate spermatogonia [28-31]; C. elegans distal tip cells maintain the mitotic population of germ cells [32-35]. Even in the mammalian ovary, which lacks GSCs after birth, cumulus cells regulate the oocyte cell cycle [29]. The Drosophila germarium has easily identifiable, actively dividing stem cells and an architecturally defined niche, providing a powerful system to study niche function.

Contacts between cap cells and GSCs [36] are necessary to maintain GSCs in an undifferentiated state. Adherens junctions, composed of DE-cadherin (Shotgun) and β-catenin (Armadillo), between cap and germ cells are required for GSC maintenance; indeed cap cells are identified by their high Armadillo levels [37]. Endosomal trafficking is important for this association; Rab11 null GSCs detach from the cap cells, undergo abnormal divisions, and arrest cyst development [40]. Gap junctions are present between germ cells and somatic cells, and may be important for maintenance of GSCs and their progeny [41].

Cap cells also regulate GSCs via secreted signals, notably Decapentaplegic (Dpp), the Drosophila homolog of Bone Morphogenetic Proteins (BMP) 2/4 [42]. Dpp is necessary to maintain GSCs in an undifferentiated state [43-48]. The niche regulates numbers of both GSCs and escort cell progenitors through the Jak/Stat (Janus kinase/signal transducer and activators of transcription) pathway. This pathway is activated by Unpaired ligands (Upd, and related genes, Upd2 and Upd3), acting through the transmembrane receptor Domeless (reviewed by [57, 58]). Conversely, GSCs regulate cap cell number and function via Delta signaling to its receptor Notch [49, 50]. Ectopic expression of Notch or Delta leads to ectopic cap cells that are sufficient to maintain GSCs through local Dpp signaling [50]. Thus, a GSC signal stimulates the somatic niche to produce factors for stem cell maintenance [49].

Follicle cell stem cells

Production of both germ cells and FCs is coordinately regulated by the same niche cells. FCs are born from 2-3 somatic stem cells, which we call follicle cell stem cells (FCSCs) as in [21]. FCSCs are 2-5 cell diameters away from the cap cells and terminal filament, which maintain FCSCs by producing Hh, Wingless (Wg), and Dpp [52, 59-61]. Like GSCs, FCSCs are short-lived if they cannot respond to BMP signals [59]. Hh regulates the number of FCSCs [52, 61]; Wg regulates their proliferation and differentiation [60].

A few inner germarial sheath cells lie between the cap cells and the FCSCs. DE-cadherin and Armadillo are strongly localized at the interface between FCSCs and inner germarial sheath cells, and are required to maintain the FCSCs long term [62]. The role of adherens junctions for either GSCs or FCSCs remains unclear: they may mediate a structural or tensile signal, or they may hold the stem cells near the source of other niche signals.

Disruption of the coordinated proliferation of GSCs and FCSCs leads to defective egg chamber development [52]. When niche cells are mutant for Yb, FCSCs are fewer and GSCs are lost, whereas overexpression increases the numbers of both populations [53, 54]. In addition to developmental regulation, the proliferation rate of both GSCs and FCSCs responds to nutrition of the female over at least a four-fold range [63]. The mechanisms that mediate this response are unknown, but may occur through Drosophila insulin-like peptides produced by the nervous system [64].

Precursor follicle cells and egg chamber formation

Inward migration of precursor FCs separates individual germline cysts into discrete egg chambers. This process involves the specification of three FC types: polar cells, stalk cells, and epithelial FCs.

Prepolar cells and separation of cysts

A polyclonal group of precursor FCs migrates between cysts, stops dividing and becomes prepolar cells, founding the polar/stalk cell lineage [65]. This migration requires Hedgehog (Hh) signaling through a Fused-dependent response pathway [66]. β-integrin is required in precursor FCs to prevent them from migrating inside cysts [21]; additional mechanisms are reviewed elsewhere [5]. Prepolar cell differentiation into polar and stalk cells maintains egg chamber separation.

Selection of the polar cell fate requires a Delta signal from the germline to stimulate Notch activity in the prepolar cells. The Notch receptor has two membrane-bound ligands, Delta and Serrate; the Fringe glycosyltransferase biases Notch to bind Delta preferentially [67][68-70]. Fringe is expressed in prepolar cells as they intercalate between cysts and determines the balance between polar and stalk cells [71]. No polar cells form without Fringe; whereas elevated Fringe increases their numbers. Importantly, egg chambers mosiac for Notch mutant FCs develop normally whenever the polar cells are wild type. Absence of Delta in the germline is associated with absence of polar cells and large, multicyst egg chambers [11, 72-74]. Thus, polar cells are essential to separate cysts into individual egg chambers.

An early function for polar cells is to specify the stalk cells. Prepolar cells between stage 1 and stage 2 egg chambers are induced to form stalk cells by Delta ligand from anterior polar cells of the stage 2 egg chamber [75]. Different levels of Notch activation distinguish the polar and stalk cell fates; high levels induce polar cells. In polar cells, the metalloprotease Kuzbanian-like attenuates the Delta signal to induce stalk cells [76]. Overexpression of either Notch or Delta in the FCs leads to abnormally long stalks between egg chambers [77].

Additional signals modulate polar and stalk cell differentiation. Upd/Jak-Stat signaling is necessary for stalk cells; when FCs cannot respond to Upd, there are excess polar cells [78, 79]. The stacked organization of stalk cells requires Hh signaling through Fused; stalk cells without Fused differentiate in a cluster [66]. Conversely, Fused-independent Hh signaling biases prepolar cells become polar cells [80]. Even within the epithelial FCs, ectopic Hh activity can induce ectopic polar cells [52], by repressing an epithelial FC-determining gene [81]. We will discuss the maturation of polar cells and epithelial FCs below.

Formation of the follicular epithelium

The initial encapsulation of germline cysts involves a mesenchymal to epithelial transition. Mesenchymal precursor FCs divide a few more times, then encapsulate the egg chamber in the follicular epithelium [8, 65]. Differential adhesion, in which tissues reorganize by altering adhesion to their neighbors (reviewed by [82]), drives many of the FC reorganizations during oogenesis. Homotypic associate of DE-cadherin mediates adhesion of precursor FCs to germline cells during epithelium formation. Prepolar cells maintain the oocyte at the posterior of the egg chamber via higher levels of DE-cadherin [83-85].

Glycosphingolipids are required for FCs to encapsulate the egg chamber. egghead and brainiac encode proteins that add sugar residues to glycosphingolipids [86]; both are required in germ cells to separate cysts into individual egg chambers [87, 88]. Similarities in mutant phenotypes between egghead, brainiac, and Notch suggested that they might be involved in a common signaling pathway; however, egghead and brainiac are not required for polar and stalk cell specification. brainiac may also modulate Gurken-EGFR signaling, as simultaneous reduction in brainiac and gurken produces multicyst follicles with discontinuities in the follicular epithelium [88, 89]. It is unlikely that all signals needed for this transition have been identified; indeed, the Upd-Domeless/Jak-Stat pathway is also involved [90].

Some Polycomb group (PcG) genes promote differentiation of both the prepolar and epithelial lineages during egg chamber formation [99]. Because PcG proteins mediate gene silencing and chromatin remodeling (recently reviewed by [98]), it is tempting to speculate that these two lineages are irreversibly determined in the germarium. However, forced expression of certain signals, such as Hh [52, 81] or Delta [76], can convert early epithelial FCs to polar cell or intermediate fates, respectively. Thus, a gradual differentiation of polar cells and epithelial FCs is a more likely scenario.

Maturation of polar cells

Polar cells reside at the anterior and posterior termini of egg chambers (Fig. 3) [14, 94] and organize FC fates throughout oogenesis. They provide cues for axial patterning of the embryo, create a porthole for sperm entry, and organize the specialized eggshell domains thought to enhance embryonic survival [61, 65, 80, 91-93]. Initially, they are distinguished from epithelial FCs by their expression of a reporter in neuralized [11]. During stages 1 and 2, both polar cells and epithelial FCs have high levels of Fasciclin III (Fas3), a homophilic adhesion molecule [95], on their lateral membranes. Polar cells maintain high Fas3 levels as oogenesis proceeds (Fig. 3).

Initially, clusters of about 3-5 polar cells join each end of an egg chamber [80]. During stages 2-5, excess polar cells undergo apoptosis, leaving only two in every cluster. During stages 6-8, polar cells become rounded, reorganize adherens junction proteins, undergo transient apical capping of basement membrane proteins, and detach from the basement membrane [11, 96, 97]. Thus, polar cell maturation involves a progressive loss of epithelial cell features.

Maturation of epithelial follicle cells

One of the earliest markers to distinguish epithelial FCs is Eyes absent (Eya) [81], a DNA binding protein phosphatase [100, 101]. Eya represses polar/stalk cell fates; epithelial FCs that lack Eya form ectopic polar cells; whereas overexpression of Eya in this lineage prevents or delays their differentiation. The Hh signal transducer Cubitus interruptis (Ci) may reinforce the decision between the polar cell and epithelial FC fates.

Extra machrochaetae (Emc), is required in the epithelial FCs for Eya expression, and for other aspects of their maturation [102]. Emc inhibits the transcriptional activity of basic helix-loop-helix proteins, like its mammalian homologs, Id proteins [103, 104]. Emc is detected in precursor FCs, but is not detected in polar cells or stalk cells beginning at stage 2. In the absence of Emc, epithelial FCs continue to express high levels of Fas3, and do not stop proliferating.

Epithelial FCs stop dividing at the end of stage 6, and enter a modified cell cycle where DNA is duplicated without cell division [105, 106]. Hh signaling through activated Ci promotes FC proliferation [52, 61]. Delta in the germline and Notch in the FCs are required for the switch from mitotic cycles to endocycles. Notch activates this switch by inducing expression of the transcription factor Hindsight, which down-regulates expression of Cut, Eya, and Ci [109]. The Salvador-Warts-Hippo pathway also limits FC proliferation by regulating expression of Hindsight and Cut [107, 108].

This change in cell cycle is sometimes called the onset of differentiation, or the switch from immature to differentiated FCs (e.g. [105, 109, 110]). The notion that Notch triggers differentiation is a tempting parallel to Notch function in some mammalian tissues, such as epidermis [113]. Many observations seem to support it, for Notch is required in all epithelial FCs to stop proliferation and to express differentiation markers [11, 71, 72, 105, 110]. However, epithelial FCs begin differentiating at stage 2, with progressive down-regulation of Fas3. Furthermore, studies that manipulate Notch activity late in oogenesis have demonstrated requirements for Notch signaling during FC migrations [114-116], when epithelial FCs reactivate expression of early, ‘immature’ markers such as Fas3 [111], and Emc [112]. Thus, a nuanced model for dynamic modulations in Notch activity throughout FC morphogenesis is more appropriate.

During stages 7-9, FCs complete three endocycles [117]; the resultant polyploidy is necessary for their final size [105, 106]. Drosophila Myc is required for endocycles and resultant growth [118]. As in other tissues, Akt kinase is required for normal FC size and PTEN limits the size increase [119-121]. Some isoforms of the transcription factor Bunched are required for FCs to achieve normal size during these stages (Wu et al, submitted); however, these isoforms are dispensible for the endocycles. The mechanisms that link cellular growth and DNA replication in the FCs are unknown.

Subdivision of epithelial follicle cell types

At about stage 5, FCs at the ends of the egg chamber, terminal FCs, become distinct from those in the center, mainbody FCs. Polar cells induce the terminal FCs [74]. Genetic ablation of polar cells at the anterior results in loss of the border, stretched, and centripetal fates; at the posterior, it results in failure of FCs to polarize the oocyte axis. Moreover, ectopic polar cells recruit neighboring FCs to become border cells or posterior terminal cells, depending on their location [52, 93, 122, 123].

The terminal domains are symmetrically patterned in concentric rings, leading to the model that polar cells produce a morphogen that specifies different cell types at different concentrations [74]. Consistent with this model, Upd is produced from polar cells [78, 79], and one study supports a morphogen function [92]. However, conflicting data exist; the model is reviewed critically elsewhere [124]. Overall, it is clear that the Upd/Jak-Stat pathway subdivides terminal and mainbody FCs, with mainbody FCs marked by expression of the homeodomain gene mirror [92].

The terminal domains have the default, anterior fate, until stage 6, when the TGFα-like ligand Gurken (Grk) activates sufficient levels of epidermal growth factor receptor (EGFR) activity in posterior FCs [9, 10, 74]. Strikingly, both the Jak-Stat and EGFR/Ras pathways must be activated to induce expression of posterior markers in mainbody cells [92]. Posterior FCs signal back to polarize the oocyte cytoskeleton, which localizes determinants to the anterior and posterior poles of the oocyte. This polarization also directs the oocyte nucleus to migrate anteriorly. Beginning in stage 9, the newly positioned oocyte nucleus specifies the dorsal FCs, which define the dorsal-ventral axis of the embryo (reviewed in [125]).

Reorganization of the epithelium

Starting in stage 9, FCs undergo extensive cell shape changes and morphogenetic movements. Different groups of cells undergo different kinds of reorganization and migration (Fig. 4), discussed separately here.

Figure 4
A-D. Follicle cell morphology visualized by α-spectrin staining. Anterior is to the left. A. At stage 8, epithelial follicle cells are morphologically uniform and have a cuboidal shape. B. At stage 9, follicle cells change their shapes as they ...

Border cell migration

At the beginning of stage 9, anterior polar cells recruit 4-6 adjacent FCs to become border cells, which delaminate from the epithelium, surround the polar cells, and migrate posteriorly between nurse cells. The centrally localized polar cells are not migratory; they are carried by the border cells. Elegant studies have identified signaling systems important for border cell migration, including Jak-Stat [90, 91, 126, 127], Notch [128, 129], PDGF/VEGF receptor (PVR) [130], and EGFR pathways [131].

Notch must be activated in border cells for delamination [128, 129]. Border cells then form an extension several cell diameters in length; similar extensions may generally initiate invasive migration [132]. DE-cadherin-mediated adhesion is necessary for movement of the border cell cluster between nurse cells [85, 133, 134]. Since loss of apical/basal cell polarity is associated with invasion and metastasis in cancer [135], one might expect that border cells would lose polarity. Surprisingly, this is not the case; polarity is required for both reorganization and migration [136].

Border cell migration is guided by chemotaxis. Two growth factor receptors, PVR and EGFR, have partially redundant roles in guiding migration towards the oocyte, which expresses their ligands, PDGF/VEGF factor 1 (PVF1) and Grk, respectively [130, 137]. Only EGFR is required for subsequent migration along the oocyte to appose the dorsal nucleus [131]. Initially, border cells migrate toward a localized signal detected by a few cells [138]. Later, differences in signal strength detected by different cells within the cluster guides migration [139].

Transcription factors are the link between extracellular signals and expression of genes required for adhesion and motility. In border cells, Upd from anterior polar cells induces expression of the basic leucine zipper transcription factor, Slow border cells (Slbo), the fly CCAAT/enhancer binding protein (C/EBP) [91, 126]. Slbo is necessary and sufficient for border cell migration [140], and regulates expression of genes such as FAK [141, 142], DE-cadherin [133, 134], the FGF receptor Breathless [143], and genes encoding other nuclear factors, such as jing and yan [144, 145]. Microarray studies have identified a large set of genes differentially expressed in border cells, many of which are induced by Slbo [146, 147].

Formation of squamous and columnar domains

Concurrent with border cell migration, mainbody FCs migrate posteriorly to cover the oocyte. This reorganization involves the formation of a squamous epithelium from the anterior terminal FCs, and a columnar epithelium of the mainbody and posterior terminal FCs. At the beginning of stage 10, the boundary for squamous and columnar FCs is aligned with the nurse cell-oocyte interface. Columnar FCs are a secretory epithelium, which sequentially deposits eggshell components.

About 50 anterior terminal FCs stretch flat to cover the nurse cells. These stretched cells are also called nurse cell FCs or squamous FCs. The name “stretched” implies that flattening is a passive event. However, stretched cells require Notch activation to disassemble their adherens junctions in an ordered fashion [116]. Furthermore, inactivation of Notch signaling in anterior FCs, either by lack of Fringe glycosylase alone, or both Delta and Serrate ligands, impedes posterior migration, so that clusters of mainbody cells linger over the nurse cells. It is likely that anterior cells must interact with each other and the underlying nurse cells to flatten.

Terminal FCs will flatten unless the posterior identity is induced by EGFR activity [9, 10]. However, absence of EGFR activity is not sufficient for flattening; posterior FCs lacking EGFR activation become columnar while expressing anterior markers [74]. Constriction of the apical surface is important for cuboidal FCs to become columnar; this requires karst, the gene for apical βHeavy-spectrin (βH) [148]. Mainbody FCs that lack βH-spectrin do not all migrate to overlie the oocyte; some lag over the nurse cells. This suggests that posterior migration is driven in part by compaction as cells become columnar.

The oocyte is important for the change to columnar shape. When the oocyte is mislocalized at the center of the egg chamber, the columnar cells form at this position [149]. When the germline is mutant for either egghead or brainiac, posterior migration is accelerated relative to border cell migration [87]. Conversely, posterior migration is blocked when the germline lacks the microtubule-associated protein Toucan [151]. However, contact with the oocyte is not necessary for the cuboidal to columnar transition. When the oocyte is abnormally small, columnar FCs form over nurse cells [150]. Altogether, it appears that the oocyte initiates the transition to columnar FCs.

Centripetal migration

At stage 10B, columnar FCs migrate inward at the interface between the oocyte and nurse cells. These cells are called centripetal migrating FCs, and form the operculum and ventral collar of the eggshell. Centripetal migration is regulated by a now-familiar set of signaling pathways, including Dpp, Notch, Jak-Stat, and EGFR.

At the end of stage 9, Dpp is permissive for centripetal migration by downregulating expression of bunched (bun), which encodes multiple isoforms of a TSC-22/DIP/BUN type leucine zipper transcription factor [153]. At this time, Dpp also stimulates Fas3 expression in a broad domain [111]; the Fas3-positive cells are likely competent for migration. Dpp expression continues in leading edge cells during centripetal migration [152, 153]; perhaps it stimulates migration of the following cells, as in embryonic dorsal closure [156].

Notch is required for centripetal migration [114]. In nearby cells, bun down-regulates Notch receptor, ligand, and target genes, as well as other migration markers such as non-muscle myosin [162]. Thus, Bun maintains the columnar epithelium by preventing recruitment of excess centripetal cells. Slbo is expressed in centripetally migrating cells [154]. Bun and Slbo mutually antagonize each other's expression, maintaining a boundary between stationary and migrating cell fates.

Centripetal migrating FCs elongate apically to move over the oocyte surface. Consistent with remodeling of membrane cytoskeletal domains, lateral α-spectrin levels increase [114], and βH-spectrin mutant egg chambers show a mild defect in centripetal migration [148]. DE-cadherin-mediated adhesion promotes centripetal migration, which is delayed in its absence [85]. The ecdysone receptor (EcR) is required for normal centripetal migration, and up-regulates DE-cadherin levels [155]. EcR activity is negatively regulated by EGFR/Ras activity, whereas bun expression is upregulated by this pathway [153]. Intriguingly, both DE-cadherin and βH-spectrin influence the site of centripetal migration. FCs that lack either protein occasionally migrate between nurse cells, instead of along the anterior oocyte. However, for βH-spectrin mutants, defects in columnar epithelium formation may misposition FCs competent for centripetal migration.

Centripetal migration is fundamentally different from border cell migration, and has superficial similarities to convergent extension [157]. Understanding the commonalities and the differences between these processes will provide important insights into the highly regulated process of normal epithelial to mesenchyme transitions and resultant migrations.

Formation of dorsal appendages

The most complex set of migrations forms the dorsal appendages. Two patches of dorsal appendage-forming cells are specified by combinatorial signaling by BMP, EGF, and Notch pathways during stages 9-10 [2, 3, 115, 158]. Within each patch, two cell types, the floor cells and roof cells [159], cooperate to form a tube. Specialized chorion proteins are secreted into each lumen. Each tube extends anteriorly between the basal lamina and the squamous FCs. By stage 13, a flattened paddle is formed at the anterior tip of each tube, by shortening of cells along their apicobasal axes. These morphogenetic movements are reviewed in detail elsewhere [160].

Maintenance of the epithelium

Border cell and centripetal migration have features reminiscent of epithelial-to-mesenchymal transition (EMT). In both cases, FCs detach from the epithelium, invade the germline tissue, and share mechanisms with cancer metastasis. Jak-Stat signaling is required for border cell migration in flies [91, 126] and promotes cell invasion in certain human cancers [161]. Notch signaling regulates both centripetal and border cell migration [74, 146][162]. For human cancers, Notch can be either a tumor suppressor or an oncogene (reviewed in [163]); it can promote metastasis in primary melanomas [164]. VEGF recruits new blood vessels, which support tumorigenesis (reviewed in [165]). Interestingly, Drosophila VEGF is an attractant for border cell migration [130]. Elevated BMP expression appears to be associated with metastatic state of some human cancers (reviewed in [163]); fly BMP signaling makes epithelial cells competent for migration [166]. Despite mechanistic similarities with carcinogenesis, migrating FCs are well-behaved, and most FCs remain in an epithelial monolayer. Many factors influence maintenance of the monolayer epithelium, including cell junctions, apical/basal cell polarity, and association with the extracellular matrix (ECM).

Loss of adherens junctions, as from armadillo mutations, disrupts the membrane cytoskeleton, resulting in irregular FC morphology and degeneration of follicles [170]. Spectraplakin, a cytoskeleton protein encoded by short stop, is localized at adherens junctions; when absent, the epithelium is double layered [171]. Some mechanisms that are required for initial formation of the epithelium in the germarium are also required to maintain a monolayer epithelium, particularly Notch signaling and sphingolipid production by germline cells [172].

FCs are polarized cells, with differential distribution of membrane proteins along the apical/basal axis (reviewed in [4, 167-169]). Disruption of different apical/basal polarity complexes has different consequences. Early loss of the apical determinant Crumbs leads to multilayering or discontinuity of the epithelium [170]; however, once the epithelium has formed, Crumbs is dispensible. In contrast, the loss of basolateral proteins has profound consequences. discs large (dlg) mutant FCs invade between germline cells and overproliferate in the pole regions [173-175]. FCs lacking either Scribble or Lethal giant larvae (Lgl) show the same phenotypes [174], consistent with their function as an interdependent protein complex. Two basolateral proteins, Fasciclin-2 and Neuroglian, suppress epithelial invasion [176]. The apical protein Bazooka is required for invasion by dlg mutant FCs; implicating the apical domain in aberrant adhesion between FCs and the germ cells [177]. These studies led to identification of Drosophila neoplastic tumor suppressors, such as Lgl, Dlg and Scribble (reviewed in [178]). Interestingly, human basolateral proteins, such as Dlg1, are implicated in tumorigenesis [179].

Protein trafficking is essential for apical/basal polarity by targeting proteins to specific intracellular locations. Two core components of the vesicle trafficking machinery, a syntaxin (Avl) and a Rab protein (Rab5) are required to maintain the epithelium. Loss of either results in accumulation of membrane proteins such as Notch and Crumbs, loss of cell polarity, and overproliferation [180].

Epithelial organization of posterior terminal FCs is particularly vulnerable. The membrane cytoskeleton is necessary here; loss of α-spectrin leads to multilayering and hyperplasia at this location [181]. The proteoglycan Perlecan is required here for monolayer organization and proper localization of polarity proteins [184]. The ECM receptor Dystroglycan is required to maintain apical/basal polarity of posterior FCs [182]. Integrin, another transmembrane receptor that links ECM to the cytoskeleton, maintains monolayer organization by orienting planar FC division independent of apical/basal polarity [183]. The vulnerability of this domain has been attributed to the curvature of the epithelium [181], or to adoption of the migratory border cell fate [74]. A third possibility is the activity of two potentially oncogenic signaling pathways in these cells: the Jak-Stat and EGFR/Ras pathways [92].

Summary

Several themes emerge from studies of FC morphogenesis. Cell-cell communication and adhesion are important for every change in cell shape and organization. However, the same signals are used repeatedly; what mechanisms generate different responses in different locations and at different times? Multiple signals are active in every FC rearrangement; it is unlikely that combinatorial signaling alone confers the different responses. More likely, each round of cell-cell interactions induces cell-intrinsic changes that confer distinct responses to the same signals. One example is the expression of different modulators for Notch signaling at different locations and stages. Transcription factors such as Slbo (C/EBP), Mirror, and Bunched also may change cells' competence to respond to certain signals.

In summary, the follicular epithelium is an excellent model to study EMT and migration as well as mechanisms for normal organogenesis. Each egg chamber, comprised of about 1000 cells, undergoes many of the morphogenetic events of larger organs, and uses most of the same regulatory networks. Importantly, sophisticated genetic methods can manipulate individual cells to separate events in space and time. Thus, FCs are a powerful system both to integrate newly identified effector molecules into known networks, and to identify novel mechanisms for cell shape, organization and migration in vivo.

Acknowledgments

We thank D. Bilder, L. Dobens, S. Goode, and D. Montell for discussions, and Raftery lab members for comments. We apologize to colleagues whose work was inadvertently omitted. The Raftery lab is supported by NIH grant GM60501.

Footnotes

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References

1. van Eeden F, St Johnston D. The polarisation of the anterior-posterior and dorsal-ventral axes during Drosophila oogenesis. Curr Opin Genet Dev. 1999;9(4):396–404. [PubMed]
2. Nilson LA, Schupbach T. EGF receptor signaling in Drosophila oogenesis. Curr Top Dev Biol. 1999;44:203–43. [PubMed]
3. Van Buskirk C, Schupbach T. Versatility in signalling: multiple responses to EGF receptor activation during Drosophila oogenesis. Trends Cell Biol. 1999;9(1):1–4. [PubMed]
4. Horne-Badovinac S, Bilder D. Mass transit: epithelial morphogenesis in the Drosophila egg chamber. Dev Dyn. 2005;232(3):559–74. [PubMed]
5. Dobens L, Raftery L. Integration of epithelial patterning and morphogenesis in the Drosophila ovarian follicle cells. Dev Dyn. 2000;218:80–93. [PubMed]
6. Xie T, et al. Intimate relationships with their neighbors: tales of stem cells in Drosophila reproductive systems. Dev Dyn. 2005;232(3):775–90. [PubMed]
7. Lin H. The stem-cell niche theory: lessons from flies. Nat Rev Genet. 2002;3(12):931–40. [PubMed]
8. Margolis J, Spradling A. Identification and behavior of epithelial stem cells in the Drosophila ovary. Development. 1995;121(11):3797–807. [PubMed]
9. Roth S, et al. cornichon and the EGF receptor signaling process are necessary for both anterior-posterior and dorsal-ventral pattern formation in Drosophila. Cell. 1995;81(6):967–78. [PubMed]
10. Gonzalez-Reyes A, Elliott H, St Johnston D. Polarization of both major body axes in Drosophila by gurken-torpedo signalling. Nature. 1995;375(6533):654–8. [PubMed]
11. Ruohola H, et al. Role of neurogenic genes in establishment of follicle cell fate and oocyte polarity during oogenesis in Drosophila. Cell. 1991;66(3):433–49. [PubMed]
12. Koch EA, King RC. The origin and early differentiation of the egg chamber of Drosophila melanogaster. J Morphol. 1966;119(3):283–303. [PubMed]
13. King RC, Aggarwal SK, Aggarwal U. The development of the female Drosophila reproductive system. J Morphol. 1968;124(2):143–66. [PubMed]
14. Spradling AC. Developmental genetics of oogenesis. In: Martinez-Arias, editor. the development of Drosophila melanogaster. cold spring harbor: cold spring harbor laboratory press; 1993. pp. 1–70.
15. Cooley L, Verheyen E, Ayers K. chickadee encodes a profilin required for intercellular cytoplasm transport during Drosophila oogenesis. Cell. 1992;69(1):173–84. [PubMed]
16. Spradling AC. Germline cysts: communes that work. Cell. 1993;72(5):649–51. [PubMed]
17. Gilboa L, Lehmann R. Soma-germline interactions coordinate homeostasis and growth in the Drosophila gonad. Nature. 2006;443(7107):97–100. [PubMed]
18. Godt D, Laski FA. Mechanisms of cell rearrangement and cell recruitment in Drosophila ovary morphogenesis and the requirement of bric a brac. Development. 1995;121(1):173–87. [PubMed]
19. Sahut-Barnola I, et al. Drosophila ovary morphogenesis: analysis of terminal filament formation and identification of a gene required for this process. Dev Biol. 1995;170(1):127–35. [PubMed]
20. Gilboa L, Lehmann R. Repression of primordial germ cell differentiation parallels germ line stem cell maintenance. Curr Biol. 2004;14(11):981–6. [PubMed]
21. Bolivar J, et al. Genetic dissection of a stem cell niche: the case of the Drosophila ovary. Dev Dyn. 2006;235(11):2969–79. [PubMed]
22. Decotto E, Spradling AC. The Drosophila ovarian and testis stem cell niches: similar somatic stem cells and signals. Dev Cell. 2005;9(4):501–10. [PubMed]
23. Huynh JR, St Johnston D. The origin of asymmetry: early polarisation of the Drosophila germline cyst and oocyte. Curr Biol. 2004;14(11):R438–49. [PubMed]
24. Koch EA, Smith PA, King RC. The division and differentiation of Drosophila cystocytes. J Morphol. 1967;121(1):55–70. [PubMed]
25. Fuller MT, Spradling AC. Male and female Drosophila germline stem cells: two versions of immortality. Science. 2007;316(5823):402–4. [PubMed]
26. Xie T, Li L. Stem cells and their niche: an inseparable relationship. Development. 2007;134(11):2001–6. [PubMed]
27. Fuchs E, Tumbar T, Guasch G. Socializing with the neighbors: stem cells and their niche. Cell. 2004;116(6):769–78. [PubMed]
28. Yoshida S, Sukeno M, Nabeshima Y. A vasculature-associated niche for undifferentiated spermatogonia in the mouse testis. Science. 2007;317(5845):1722–6. [PubMed]
29. Matzuk MM, et al. Intercellular communication in the mammalian ovary: oocytes carry the conversation. Science. 2002;296(5576):2178–80. [PubMed]
30. Meng X, et al. Regulation of cell fate decision of undifferentiated spermatogonia by GDNF. Science. 2000;287(5457):1489–93. [PubMed]
31. Ohta H, et al. Regulation of proliferation and differentiation in spermatogonial stem cells: the role of c-kit and its ligand SCF. Development. 2000;127(10):2125–31. [PubMed]
32. Berry LW, Westlund B, Schedl T. Germ-line tumor formation caused by activation of glp-1, a Caenorhabditis elegans member of the Notch family of receptors. Development. 1997;124(4):925–36. [PubMed]
33. Hall DH, et al. Ultrastructural features of the adult hermaphrodite gonad of Caenorhabditis elegans: relations between the germ line and soma. Dev Biol. 1999;212(1):101–23. [PubMed]
34. Austin J, Kimble J. glp-1 is required in the germ line for regulation of the decision between mitosis and meiosis in C. elegans. Cell. 1987;51(4):589–99. [PubMed]
35. Kadyk LC, Kimble J. Genetic regulation of entry into meiosis in Caenorhabditis elegans. Development. 1998;125(10):1803–13. [PubMed]
36. Lin H, Spradling AC. A novel group of pumilio mutations affects the asymmetric division of germline stem cells in the Drosophila ovary. Development. 1997;124(12):2463–76. [PubMed]
37. Song X, et al. Germline stem cells anchored by adherens junctions in the Drosophila ovary niches. Science. 2002;296(5574):1855–7. [PubMed]First study provided genetic evidence showing requirement of adherens junctions in stem cell maintenance.
38. Lin H, Yue L, Spradling AC. The Drosophila fusome, a germline-specific organelle, contains membrane skeletal proteins and functions in cyst formation. Development. 1994;120(4):947–56. [PubMed]
39. de Cuevas M, Lilly MA, Spradling AC. Germline cyst formation in Drosophila. Annu Rev Genet. 1997;31:405–28. [PubMed]
40. Bogard N, et al. Rab11 maintains connections between germline stem cells and niche cells in the Drosophila ovary. Development. 2007;134(19):3413–8. [PubMed]
41. Tazuke SI, et al. A germline-specific gap junction protein required for survival of differentiating early germ cells. Development. 2002;129(10):2529–39. [PubMed]
42. Xie T, Spradling AC. A niche maintaining germ line stem cells in the Drosophila ovary. Science. 2000;290(5490):328–30. [PubMed]
43. Xie T, Spradling AC. decapentaplegic is essential for the maintenance and division of germline stem cells in the Drosophila ovary. Cell. 1998;94(2):251–60. [PubMed]This is the elegant study depicting the role of dpp in stem cell maintenance.
44. Song X, et al. Bmp signals from niche cells directly repress transcription of a differentiation-promoting gene, bag of marbles, in germline stem cells in the Drosophila ovary. Development. 2004;131(6):1353–64. [PubMed]
45. Chen D, McKearin D. Dpp signaling silences bam transcription directly to establish asymmetric divisions of germline stem cells. Curr Biol. 2003;13(20):1786–91. [PubMed]
46. McKearin DM, Spradling AC. bag-of-marbles: a Drosophila gene required to initiate both male and female gametogenesis. Genes Dev. 1990;4(12B):2242–51. [PubMed]
47. Ohlstein B, McKearin D. Ectopic expression of the Drosophila Bam protein eliminates oogenic germline stem cells. Development. 1997;124(18):3651–62. [PubMed]
48. McKearin D, Ohlstein B. A role for the Drosophila bag-of-marbles protein in the differentiation of cystoblasts from germline stem cells. Development. 1995;121(9):2937–47. [PubMed]
49. Ward EJ, et al. Stem cells signal to the niche through the Notch pathway in the Drosophila ovary. Curr Biol. 2006;16(23):2352–8. [PubMed]
50. Song X, et al. Notch signaling controls germline stem cell niche formation in the Drosophila ovary. Development. 2007;134(6):1071–80. [PubMed]
51. Forbes AJ, et al. The role of segment polarity genes during early oogenesis in Drosophila. Development. 1996;122(10):3283–94. [PubMed]
52. Forbes AJ, et al. hedgehog is required for the proliferation and specification of ovarian somatic cells prior to egg chamber formation in Drosophila. Development. 1996;122(4):1125–35. [PubMed]
53. King FJ, Lin H. Somatic signaling mediated by fs(1)Yb is essential for germline stem cell maintenance during Drosophila oogenesis. Development. 1999;126(9):1833–44. [PubMed]
54. King FJ, et al. Yb modulates the divisions of both germline and somatic stem cells through piwi- and hh-mediated mechanisms in the Drosophila ovary. Mol Cell. 2001;7(3):497–508. [PubMed]
55. Cox DN, Chao A, Lin H. piwi encodes a nucleoplasmic factor whose activity modulates the number and division rate of germline stem cells. Development. 2000;127(3):503–14. [PubMed]
56. Cox DN, et al. A novel class of evolutionarily conserved genes defined by piwi are essential for stem cell self-renewal. Genes Dev. 1998;12(23):3715–27. [PMC free article] [PubMed]
57. Hombria JC, Brown S. The fertile field of Drosophila Jak/STAT signalling. Curr Biol. 2002;12(16):R569–75. [PubMed]
58. Arbouzova NI, Zeidler MP. JAK/STAT signalling in Drosophila: insights into conserved regulatory and cellular functions. Development. 2006;133(14):2605–16. [PubMed]
59. Kirilly D, et al. BMP signaling is required for controlling somatic stem cell self-renewal in the Drosophila ovary. Dev Cell. 2005;9(5):651–62. [PubMed]
60. Song X, Xie T. Wingless signaling regulates the maintenance of ovarian somatic stem cells in Drosophila. Development. 2003;130(14):3259–68. [PubMed]
61. Zhang Y, Kalderon D. Hedgehog acts as a somatic stem cell factor in the Drosophila ovary. Nature. 2001;410(6828):599–604. [PubMed]
62. Song X, Xie T. DE-cadherin-mediated cell adhesion is essential for maintaining somatic stem cells in the Drosophila ovary. Proc Natl Acad Sci U S A. 2002;99(23):14813–8. [PMC free article] [PubMed]
63. Drummond-Barbosa D, Spradling AC. Stem cells and their progeny respond to nutritional changes during Drosophila oogenesis. Dev Biol. 2001;231(1):265–78. [PubMed]
64. LaFever L, Drummond-Barbosa D. Direct control of germline stem cell division and cyst growth by neural insulin in Drosophila. Science. 2005;309(5737):1071–3. [PubMed]This study indicated role of Drosophila neural insulin in controlling germline stem maintenance.
65. Tworoger M, et al. Mosaic analysis in the drosophila ovary reveals a common hedgehog-inducible precursor stage for stalk and polar cells. Genetics. 1999;151(2):739–48. [PMC free article] [PubMed]
66. Besse F, Busson D, Pret AM. Fused-dependent Hedgehog signal transduction is required for somatic cell differentiation during Drosophila egg chamber formation. Development. 2002;129(17):4111–24. [PubMed]
67. Okajima T, Xu A, Irvine KD. Modulation of notch-ligand binding by protein O-fucosyltransferase 1 and fringe. J Biol Chem. 2003;278(43):42340–5. [PubMed]
68. Fleming RJ, Gu Y, Hukriede NA. Serrate-mediated activation of Notch is specifically blocked by the product of the gene fringe in the dorsal compartment of the Drosophila wing imaginal disc. Development. 1997;124(15):2973–81. [PubMed]
69. Panin VM, et al. Fringe modulates Notch-ligand interactions. Nature. 1997;387(6636):908–12. [PubMed]
70. Moloney DJ, et al. Fringe is a glycosyltransferase that modifies Notch. Nature. 2000;406(6794):369–75. [PubMed]
71. Grammont M, Irvine KD. fringe and Notch specify polar cell fate during Drosophila oogenesis. Development. 2001;128(12):2243–53. [PubMed]
72. Larkin MK, et al. Expression of constitutively active Notch arrests follicle cells at a precursor stage during Drosophila oogenesis and disrupts the anterior-posterior axis of the oocyte. Development. 1996;122(11):3639–50. [PubMed]
73. Keller Larkin M, et al. Role of Notch pathway in terminal follicle cell differentiation during Drosophila oogenesis. Dev Genes Evol. 1999;209(5):301–11. [PubMed]
74. Gonzalez-Reyes A, St Johnston D. Patterning of the follicle cell epithelium along the anterior-posterior axis during Drosophila oogenesis. Development. 1998;125(15):2837–46. [PubMed]
75. Torres IL, Lopez-Schier H, St Johnston D. A Notch/Delta-dependent relay mechanism establishes anterior-posterior polarity in Drosophila. Dev Cell. 2003;5(4):547–58. [PubMed]
76. Assa-Kunik E, et al. Drosophila follicle cells are patterned by multiple levels of Notch signaling and antagonism between the Notch and JAK/STAT pathways. Development. 2007;134(6):1161–9. [PubMed]
77. Larkin M, et al. Expression of constitutively active Notch arrests follicle cells at a precursor stage during Drosophila oogenesis and disrupts the anterior-posterior axis of the oocyte. Development. 1996;132:3639–50. [PubMed]
78. McGregor JR, Xi R, Harrison DA. JAK signaling is somatically required for follicle cell differentiation in Drosophila. Development. 2002;129(3):705–17. [PubMed]
79. Baksa K, et al. The Drosophila STAT protein, stat92E, regulates follicle cell differentiation during oogenesis. Dev Biol. 2002;243(1):166–75. [PubMed]
80. Besse F, Pret AM. Apoptosis-mediated cell death within the ovarian polar cell lineage of Drosophila melanogaster. Development. 2003;130(5):1017–27. [PubMed]
81. Bai J, Montell D. Eyes absent, a key repressor of polar cell fate during Drosophila oogenesis. Development. 2002;129(23):5377–88. [PubMed]
82. Steinberg MS. Differential adhesion in morphogenesis: a modern view. Curr Opin Genet Dev. 2007;17(4):281–6. [PubMed]
83. Gonzalez-Reyes A, St Johnston D. The Drosophila AP axis is polarised by the cadherin-mediated positioning of the oocyte. Development. 1998;125(18):3635–44. [PubMed]
84. Godt D, Tepass U. Drosophila oocyte localization is mediated by differential cadherin-based adhesion. Nature. 1998;395(6700):387–91. [PubMed]
85. Oda H, Uemura T, Takeichi M. Phenotypic analysis of null mutants for DE-cadherin and Armadillo in Drosophila ovaries reveals distinct aspects of their functions in cell adhesion and cytoskeletal organization. Genes to Cells. 1996;2:29–40. [PubMed]
86. Wandall HH, et al. Drosophila egghead encodes a beta 1,4-mannosyltransferase predicted to form the immediate precursor glycosphingolipid substrate for brainiac. J Biol Chem. 2003;278(3):1411–4. [PubMed]
87. Goode S, et al. The neurogenic genes egghead and brainiac define a novel signaling pathway essential for epithelial morphogenesis during Drosophila oogenesis. Development. 1996;122(12):3863–79. [PubMed]
88. Goode S, et al. Brainiac encodes a novel, putative secreted protein that cooperates with Grk TGF alpha in the genesis of the follicular epithelium. Dev Biol. 1996;178(1):35–50. [PubMed]
89. Goode S, Wright D, Mahowald AP. The neurogenic locus brainiac cooperates with the Drosophila EGF receptor to establish the ovarian follicle and to determine its dorsal-ventral polarity. Development. 1992;116(1):177–92. [PubMed]
90. Ghiglione C, et al. The Drosophila cytokine receptor Domeless controls border cell migration and epithelial polarization during oogenesis. Development. 2002;129(23):5437–47. [PubMed]
91. Beccari S, Teixeira L, Rorth P. The JAK/STAT pathway is required for border cell migration during Drosophila oogenesis. Mech Dev. 2002;111(12):115–23. [PubMed]
92. Xi R, McGregor JR, Harrison DA. A gradient of JAK pathway activity patterns the anterior-posterior axis of the follicular epithelium. Dev Cell. 2003;4(2):167–77. [PubMed]
93. Grammont M, Irvine KD. Organizer activity of the polar cells during Drosophila oogenesis. Development. 2002;129(22):5131–40. [PubMed]
94. Ruohola-Baker H, et al. Role of neurogenic genes in establishment of follicle cell fate and oocyte polarity during oogenesis in Drosophila. Cell. 1991:433–449. [PubMed]
95. Snow PM, Bieber AJ, Goodman CS. Fasciclin III: a novel homophilic adhesion molecule in Drosophila. Cell. 1989;59(2):313–23. [PubMed]
96. Peifer M, et al. A role for the Drosophila segment polarity gene armadillo in cell adhesion and cytoskeletal integrity during oogenesis. Development. 1993;118:1191–1207. [PubMed]
97. Medioni C, Noselli S. Dynamics of the basement membrane in invasive epithelial clusters in Drosophila. Development. 2005;132(13):3069–77. [PubMed]
98. Muller J, Kassis JA. Polycomb response elements and targeting of Polycomb group proteins in Drosophila. Curr Opin Genet Dev. 2006;16(5):476–84. [PubMed]
99. Narbonne K, et al. polyhomeotic is required for somatic cell proliferation and differentiation during ovarian follicle formation in Drosophila. Development. 2004;131(6):1389–400. [PubMed]
100. Jemc J, Rebay I. The eyes absent family of phosphotyrosine phosphatases: properties and roles in developmental regulation of transcription. Annu Rev Biochem. 2007;76:513–38. [PubMed]
101. Tootle TL, et al. The transcription factor Eyes absent is a protein tyrosine phosphatase. Nature. 2003;426(6964):299–302. [PubMed]
102. Adam JC, Montell DJ. A role for extra macrochaetae downstream of Notch in follicle cell differentiation. Development. 2004;131(23):5971–80. [PubMed]
103. Ellis HM, Spann DR, Posakony JW. extramacrochaetae, a negative regulator of sensory organ development in Drosophila, defines a new class of helix-loop-helix proteins. Cell. 1990;61(1):27–38. [PubMed]
104. Garrell J, Modolell J. The Drosophila extramacrochaetae locus, an antagonist of proneural genes that, like these genes, encodes a helix-loop-helix protein. Cell. 1990;61(1):39–48. [PubMed]
105. Lopez-Schier H, St Johnston D. Delta signaling from the germ line controls the proliferation and differentiation of the somatic follicle cells during Drosophila oogenesis. Genes Dev. 2001;15(11):1393–405. [PMC free article] [PubMed]
106. Deng WM, Althauser C, Ruohola-Baker H. Notch-Delta signaling induces a transition from mitotic cell cycle to endocycle in Drosophila follicle cells. Development. 2001;128(23):4737–46. [PubMed]
107. Polesello C, Tapon N. Salvador-warts-hippo signaling promotes Drosophila posterior follicle cell maturation downstream of notch. Curr Biol. 2007;17(21):1864–70. [PubMed]
108. Meignin C, et al. The salvador-warts-hippo pathway is required for epithelial proliferation and axis specification in Drosophila. Curr Biol. 2007;17(21):1871–8. [PMC free article] [PubMed]
109. Sun J, Deng WM. Hindsight mediates the role of notch in suppressing hedgehog signaling and cell proliferation. Dev Cell. 2007;12(3):431–42. [PMC free article] [PubMed]
110. Roth S. Drosophila oogenesis: coordinating germ line and soma. Curr Biol. 2001;11(19):R779–81. [PubMed]
111. Shravage BV, et al. The role of Dpp and its inhibitors during eggshell patterning in Drosophila. Development. 2007;134(12):2261–71. [PubMed]
112. Papadia S, et al. emc has a role in dorsal appendage fate formation in Drosophila oogenesis. Mech Dev. 2005;122(9):961–74. [PubMed]
113. Lefort K, Dotto GP. Notch signaling in the integrated control of keratinocyte growth/differentiation and tumor suppression. Semin Cancer Biol. 2004;14(5):374–86. [PubMed]
114. Dobens L, et al. bunched sets a boundary of Notch signaling to pattern anterior eggshell structures during Drosophila oogenesis. Dev Biol. 2005 in press. [PubMed]
115. Ward EJ, et al. Border of Notch activity establishes a boundary between the two dorsal appendage tube cell types. Dev Biol. 2006;297(2):461–70. [PubMed]
116. Grammont M. Adherens junction remodeling by the Notch pathway in Drosophila melanogaster oogenesis. J Cell Biol. 2007;177(1):139–50. [PMC free article] [PubMed]
117. Edgar BA, Orr-Weaver TL. Endoreplication cell cycles: more for less. Cell. 2001;105(3):297–306. [PubMed]
118. Maines JZ, et al. Drosophila dMyc is required for ovary cell growth and endoreplication. Development. 2004;131(4):775–86. [PubMed]
119. Stocker H, Hafen E. Genetic control of cell size. Curr Opin Genet Dev. 2000;10(5):529–35. [PubMed]
120. Ruggero D, Sonenberg N. The Akt of translational control. Oncogene. 2005;24(50):7426–34. [PubMed]
121. Cavaliere V, et al. dAkt kinase controls follicle cell size during Drosophila oogenesis. Dev Dyn. 2005;232(3):845–54. [PMC free article] [PubMed]
122. Liu Y, Montell DJ. Identification of mutations that cause cell migration defects in mosaic clones. Development. 1999;126(9):1869–78. [PubMed]
123. Zhang Y, Kalderon D. Regulation of cell proliferation and patterning in Drosophila oogenesis by Hedgehog signaling. Development. 2000;127(10):2165–76. [PubMed]
124. Denef N, Schupbach T. Patterning: JAK-STAT signalling in the Drosophila follicular epithelium. Curr Biol. 2003;13(10):R388–90. [PubMed]
125. Lopez-Schier H. The polarisation of the anteroposterior axis in Drosophila. Bioessays. 2003;25(8):781–91. [PubMed]
126. Silver DL, Montell DJ. Paracrine signaling through the JAK/STAT pathway activates invasive behavior of ovarian epithelial cells in Drosophila. Cell. 2001;107(7):831–41. [PubMed]
127. Silver DL, Geisbrecht ER, Montell DJ. Requirement for JAK/STAT signaling throughout border cell migration in Drosophila. Development. 2005;132(15):3483–92. [PubMed]
128. Wang X, Adam JC, Montell D. Spatially localized Kuzbanian required for specific activation of Notch during border cell migration. Dev Biol. 2007;301(2):532–40. [PubMed]
129. Prasad M, Montell DJ. Cellular and molecular mechanisms of border cell migration analyzed using time-lapse live-cell imaging. Dev Cell. 2007;12(6):997–1005. [PubMed]
130. Duchek P, et al. Guidance of cell migration by the Drosophila PDGF/VEGF receptor. Cell. 2001;107(1):17–26. [PubMed]This paper describes two signal pathways, EGFR and PVR, that guide migration of border cells toward the oocyte during Drosophila oogenesis. The two signals function redundantly, so that loss of either signal only had mild defect, while loss of both signals resulted in severe defect in border cell migration. This is an excellent example where redundant pathways function to control a single process in lower organisms.
131. Duchek P, Rorth P. Guidance of cell migration by EGF receptor signaling during Drosophila oogenesis. Science. 2001;291(5501):131–3. [PubMed]
132. Fulga TA, Rorth P. Invasive cell migration is initiated by guided growth of long cellular extensions. Nat Cell Biol. 2002;4(9):715–9. [PubMed]
133. Niewiadomska P, Godt D, Tepass U. DE-Cadherin is required for intercellular motility during Drosophila oogenesis. J Cell Biol. 1999;144(3):533–47. [PMC free article] [PubMed]
134. Pacquelet A, Rorth P. Regulatory mechanisms required for DE-cadherin function in cell migration and other types of adhesion. J Cell Biol. 2005;170(5):803–12. [PMC free article] [PubMed]
135. Gumbiner BM. Regulation of cadherin-mediated adhesion in morphogenesis. Nat Rev Mol Cell Biol. 2005;6(8):622–34. [PubMed]
136. Pinheiro EM, Montell DJ. Requirement for Par-6 and Bazooka in Drosophila border cell migration. Development. 2004;131(21):5243–51. [PubMed]
137. McDonald JA, Pinheiro EM, Montell DJ. PVF1, a PDGF/VEGF homolog, is sufficient to guide border cells and interacts genetically with Taiman. Development. 2003;130(15):3469–78. [PubMed]
138. Jekely G, et al. Regulators of endocytosis maintain localized receptor tyrosine kinase signaling in guided migration. Dev Cell. 2005;9(2):197–207. [PubMed]
139. Bianco A, et al. Two distinct modes of guidance signalling during collective migration of border cells. Nature. 2007;448(7151):362–5. [PubMed]
140. Montell D, Rørth P, Spradling A. slow border cells, a locus required for a developmentally regulated cell migration during oogenesis, encodes Drosophila C/EBP. Cell. 1992;121:51–62. [PubMed]
141. Bai J, Uehara Y, Montell DJ. Regulation of invasive cell behavior by taiman, a Drosophila protein related to AIB1, a steroid receptor coactivator amplified in breast cancer. Cell. 2000;103(7):1047–58. [PubMed]
142. Fox GL, Rebay I, Hynes RO. Expression of DFak56, a Drosophila homolog of vertebrate focal adhesion kinase, supports a role in cell migration in vivo. Proc Natl Acad Sci U S A. 1999;96(26):14978–83. [PMC free article] [PubMed]
143. Murphy AM, et al. The breathless FGF receptor homolog, a downstream target of Drosophila C/EBP in the developmental control of cell migration. Development. 1995;121(8):2255–63. [PubMed]
144. Liu Y, Montell DJ. Jing: a downstream target of slbo required for developmental control of border cell migration. Development. 2001;128(3):321–30. [PubMed]
145. Schober M, Rebay I, Perrimon N. Function of the ETS transcription factor Yan in border cell migration. Development. 2005;132(15):3493–504. [PubMed]
146. Wang X, et al. Analysis of cell migration using whole-genome expression profiling of migratory cells in the Drosophila ovary. Dev Cell. 2006;10(4):483–95. [PubMed]
147. Borghese L, et al. Systematic analysis of the transcriptional switch inducing migration of border cells. Dev Cell. 2006;10(4):497–508. [PMC free article] [PubMed]
148. Zarnescu DC, Thomas GH. Apical spectrin is essential for epithelial morphogenesis but not apicobasal polarity in Drosophila. J Cell Biol. 1999;146(5):1075–86. [PMC free article] [PubMed]
149. González-Reyes A, St Johnston D. Role of oocyte position in establishment of anterior-posterior polarity in Drosophla. Science. 1994;266:639–642. [PubMed]
150. Swan A, Suter B. Role of Bicaudal-D in patterning the Drosophila egg chamber in mid- oogenesis. Development. 1996;122(11):3577–86. [PubMed]
151. Grammont M, Dastugue B, Couderc JL. The Drosophila toucan (toc) gene is required in germline cells for the somatic cell patterning during oogenesis. Development. 1997;124(24):4917–26. [PubMed]
152. Twombly V, et al. The TGF-beta signaling pathway is essential for Drosophila oogenesis. Development. 1996;122(5):1555–65. [PubMed]
153. Dobens LL, et al. Drosophila bunched integrates opposing DPP and EGF signals to set the operculum boundary. Development. 2000;127:745–754. [PubMed]
154. Levine B, et al. Notch signaling links interactions between the C/EBP homolog slow border cells and the GILZ homolog bunched during cell migration. Dev Biol. 2007;305(1):217–31. [PubMed]
155. Hackney JF, et al. Ras signaling modulates activity of the ecdysone receptor EcR during cell migration in the Drosophila ovary. Dev Dyn. 2007;236(5):1213–26. [PubMed]
156. Riesgo-Escovar JR, Hafen E. Drosophila Jun kinase regulates expression of decapentaplegic via the ETS-domain protein Aop and the AP-1 transcription factor DJun during dorsal closure. Genes Dev. 1997;11(13):1717–27. [PubMed]
157. Keller R. Shaping the vertebrate body plan by polarized embryonic cell movements. Science. 2002;298(5600):1950–4. [PubMed]
158. Dobens LL, Raftery LA. Integration of epithelial patterning and morphogenesis in Drosophila ovarian follicle cells. Dev Dyn. 2000;218(1):80–93. [PubMed]
159. Ward EJ, Berg CA. Juxtaposition between two cell types is necessary for dorsal appendage tube formation. Mech Dev. 2005;122(2):241–55. [PubMed]
160. Berg CA. The Drosophila shell game: patterning genes and morphological change. Trends Genet. 2005;21(6):346–55. [PubMed]
161. Lai SY, et al. Erythropoietin-mediated activation of JAK-STAT signaling contributes to cellular invasion in head and neck squamous cell carcinoma. Oncogene. 2005;24(27):4442–9. [PubMed]
162. Dobens L, et al. Bunched sets a boundary for Notch signaling to pattern anterior eggshell structures during Drosophila oogenesis. Dev Biol. 2005;287(2):425–37. [PubMed]
163. Bailey JM, Singh PK, Hollingsworth MA. Cancer metastasis facilitated by developmental pathways: Sonic hedgehog, Notch, and bone morphogenic proteins. J Cell Biochem. 2007;102(4):829–39. [PubMed]
164. Balint K, et al. Activation of Notch1 signaling is required for beta-catenin-mediated human primary melanoma progression. J Clin Invest. 2005;115(11):3166–76. [PMC free article] [PubMed]
165. Hanahan D, Weinberg RA. The hallmarks of cancer. Cell. 2000;100(1):57–70. [PubMed]
166. Dobens LL, et al. Drosophila bunched integrates opposing DPP and EGF signals to set the operculum boundary. Development. 2000;127(4):745–54. [PubMed]
167. Knust E. Control of epithelial cell shape and polarity. Curr Opin Genet Dev. 2000;10(5):471–5. [PubMed]
168. Knust E. Regulation of epithelial cell shape and polarity by cell-cell adhesion (Review) Mol Membr Biol. 2002;19(2):113–20. [PubMed]
169. Tepass U, et al. Epithelial cell polarity and cell junctions in Drosophila. Annu Rev Genet. 2001;35:747–84. [PubMed]
170. Tanentzapf G, et al. Apical, lateral, and basal polarization cues contribute to the development of the follicular epithelium during Drosophila oogenesis. J Cell Biol. 2000;151(4):891–904. [PMC free article] [PubMed]
171. Roper K, Brown NH. Maintaining epithelial integrity: a function for gigantic spectraplakin isoforms in adherens junctions. J Cell Biol. 2003;162(7):1305–15. [PMC free article] [PubMed]
172. Goode S, et al. The neurogenic genes egghead and brainiac define a novel signaling pathway essential for epithelial morphogensis during Drosophila oogenesis. Development. 1996;122:3863–3879. [PubMed]
173. Goode S, Perrimon N. Inhibition of patterned cell shape change and cell invasion by Discs large during Drosophila oogenesis. Genes Dev. 1997;11(19):2532–44. [PMC free article] [PubMed]
174. Bilder D, Li M, Perrimon N. Cooperative regulation of cell polarity and growth by Drosophila tumor suppressors. Science. 2000;289(5476):113–6. [PubMed]
175. Goode S, Wei J, Kishore S. Novel spatiotemporal patterns of epithelial tumor invasion in Drosophila discs large egg chambers. Dev Dyn. 2005;232(3):855–64. [PubMed]
176. Szafranski P, Goode S. Basolateral junctions are sufficient to suppress epithelial invasion during Drosophila oogenesis. Dev Dyn. 2007;236(2):364–73. [PubMed]
177. Abdelilah-Seyfried S, Cox DN, Jan YN. Bazooka is a permissive factor for the invasive behavior of discs large tumor cells in Drosophila ovarian follicular epithelia. Development. 2003;130(9):1927–35. [PubMed]
178. Bilder D. Epithelial polarity and proliferation control: links from the Drosophila neoplastic tumor suppressors. Genes Dev. 2004;18(16):1909–25. [PubMed]
179. Fuja TJ, et al. Somatic mutations and altered expression of the candidate tumor suppressors CSNK1 epsilon, DLG1, and EDD/hHYD in mammary ductal carcinoma. Cancer Res. 2004;64(3):942–51. [PubMed]
180. Lu H, Bilder D. Endocytic control of epithelial polarity and proliferation in Drosophila. Nat Cell Biol. 2005;7(12):1232–9. [PubMed]This paper demonstrated that endocytosis is involved in both epithelial cell polarity and proliferation control. By analyzing mutant phenotypes in two components of the endocytic pathway, a syntaxin (Avl) and Rab5, the authors showed that failure of endosomal entry and progression towards lysosome degradation results in protein accumulation. Aberrant protein accumulation leads to defects in apical/basal cell polarity and overproliferation.
181. Lee JK, et al. α-spectrin is required for follicle cell monolayer integrity in Drosophila melanogaster. Development. 1997;124:353–362. [PubMed]
182. Deng WM, et al. Dystroglycan is required for polarizing the epithelial cells and the oocyte in Drosophila. Development. 2003;130(1):173–84. [PubMed]
183. Fernandez-Minan A, Martin-Bermudo MD, Gonzalez-Reyes A. Integrin signaling regulates spindle orientation in Drosophila to preserve the follicular-epithelium monolayer. Curr Biol. 2007;17(8):683–8. [PubMed]This paper reported that Integrin is required for maintenance of monolayer epithelium during Drososphila oogenesis. Interestingly, Integrin is not required for apical/basal cell polarity, but for mitotic spindle orientation in follicle cells. This work suggests a possible role that mitosis plays in maintenance of epithelial organization.
184. Schneider M, et al. Perlecan and Dystroglycan act at the basal side of the Drosophila follicular epithelium to maintain epithelial organization. Development. 2006;133(19):3805–15. [PMC free article] [PubMed]
185. Li L, Xie T. Stem cell niche: structure and function. Annu Rev Cell Dev Biol. 2005;21:605–31. [PubMed]
186. Spradling A, Drummond-Barbosa D, Kai T. Stem cells find their niche. Nature. 2001;414(6859):98–104. [PubMed]
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