Oogenesis and the Oocyte

The process of oogenesis

Development in the fly begins well before the moment of conception. Hundreds of genes participate in oogenesis, the building of the egg. The egg is a highly structured chamber. Its RNA, DNA and proteins do not roam freely, but rather are directed and controlled by the egg's dorsoventral polarity and anterior-posterior axis. Neither is the egg cell an independently created entity. It receives input from two other cell sources: nurse cells and follicle cells.

Follicle cells envelop the maturing, pre-fertilized egg. Information is exchanged between oocyte and follicle cells, conditioning and preparing all parties for the rapid changes that will ensue after fertilization (See Gurken). Nurse cells provide nutrients the embryo will require, storing them in the yolk, as though filling pantry shelves in anticipation of future need.

Given this simplified picture of oocyte development in a pre-fertilized state, one may ask not only what happens next, but working backward, where does the egg comes from in the first place? The fly needs to build not only eggs, but an organ system "factory" for their production and delivery. Soon after fertilization, pole cells bud from the posterior end of the blastula [Images], the earliest cellular phase of embryonic development. Budding from this terminal or polar region gives them their name. They are the germ line for the next generation, and also referred to as such. These germ line stem cells, that will eventually locate to the adult ovary, are the precursors of the eggs (or in males, the sperm) that will produce the next generation.

Pole cells are carried dorsally, enter the posterior midgut rudiment, move between the endodermal cells and split into two groups, one for each ovary-to-be. The pole/germ line stem cells are then enveloped by somatic mesodermal cells. In effect, the ovary structure is built around these germ line stem cells. Oviducts, accessory glands, uterus, vagina and external genitalia are all of ectodermal and mesodermal (non germ cell) lineage, derived from the genital imaginal disc. The separation of the pole cells from the rest of the developing embryo is one of the first events in morphogenesis. They will be held in reserve, locate to the adult gonads that will be assembled during the third instar larva stage. The maturation of the oocyte with which this discussion began takes place inside a germarium surrounded by follicle cells.

The germarium is an assembly line for new egg chambers. Each ovary contains more than a dozen germaria. At the start of the assembly line in any single germarium are two germ line stem cells. At the end of the line, new single stage-1 egg chambers roll off, like kits for building model flies; packed, boxed, and ready to go. One can imagine the various work stations along the way: the stem cells alternate in producing one cystoblast at a time, (the precursor cell of the egg); next, the cystoblast passes through three successive stages of cysts in formation; further along, 6 cysts, each composed of 16 cells, mature sequentially, each yielding a single stage-1 egg chamber that will exit the factory seven days from the time its originating cystoblast was formed.

A more detailed look at stem cell division reveals the two resulting cells are asymmetric in both function and size. What had been a single stem cell is now two new cells: a cystoblast and a daughter stem cell (from which a new daughter stem cell and a new cystoblast will soon arise). The cystoblast continues along the line, undergoing four cycles of cell division to form a 16 cell cyst. During this process, a single cystocyte develops a microtubule organizing center (MTOC) and forms a polarized microtubule network that extends into all 16 cells. Ring canals maintain the connection among cells in the dividing cyst. The single cell of cyst that contains the MTOC develops into the oocyte. Only the oocyte and will continue through meiosis, while the remainder will differentiate to become nurse cells for the presumptive oocyte. The nurse cell genome becomes polytene, a process in which DNA is replicated without cell division. Nurse cells grow and accumulate yolk material in a process termed vitellogenesis (oogenesis stages 8 and 9).

To understand oocyte determination, one needs to ask how a particular cystocyte is singled out to acquire an active microtubule organizing center. Germ cells contain a spherical cytoplasmic structure called the spectrosome, which contains components of the sub-membrane cytoskeleton. At the first cystoblast division, the spectrosome is inherited by only one of the two daughter cells. During the following divisions, the spectrosome grows from this cystocyte into the other cells to form a branched structure called the fusome, which interacts with one spindle pole at each division, before disappearing after the 4th division. These observations have led to the suggestion that the asymmetric inheritance of the spectrosome determines which of the two daughter cells will give rise to the cell that becomes the oocyte. How this happens is unclear, but one possibility is that the position of the spectrosome polarizes the fusome, which in turn directs the formation of an active MTOC in a single cell.

During stage 9, follicle cells, not of germ cell origin, migrate over the surface of the egg chamber, the majority drawn to the oocyte, leaving only a few follicle cells over the nurse cells. A small group of border cells, similarly non-germ cell in origin, that have lagged behind at the anterior end of the developing cystoblast-becoming-oocyte, pass through three nurse cell junctions to arrive at the anterior end of the oocyte. The border cells are implicated in the terminal system, which creates and maintains anterior-posterior polarity in the egg. Follicle cells at the posterior end of the oocyte develop a posterior cell fate. This induction requires gurken function in the oocyte and Epidermal growth factor receptor (Torpedo) in the follicle cell layer. The posterior follicle cells send an unidentified signal back to the oocyte to repolarize the the anterior-posterior axis. Protein kinase A is involved in signalling that results in a disassembly and reassembly of the MTOC.

By stage 10, the follicle cells have matured and begun secretion of a vitelline membrane around the oocyte. The mature nurse cells cytoplasmic contents flow in bulk into the oocyte by means of a cytoskeletal-based mechanism, and having completed their task, the nurse cells are then broken down (stage 11). The follicle cells that earlier produced the vitelline membrane, now add a shell over that membrane. Membrane and shell are both external to the egg, and covered by follicle cells. Specialized follicle cells also make the micropyle, the egg's terminal structure through which sperm pass into the egg.

Mature eggs ovulate one at a time and pass into the uterus. Once mated, females are able to release stored sperm from their seminal receptacles. Sperm enter the micropyle while the egg is still in the uterus. The first meiotic division has begun and is still in progress when fertilization and ovulation occur. Males contribute more than just DNA to the zygote. The zygotic centrosomes, essential for cell division, are derived from the male.

For information about the establishment of oocyte axes, messenger RNA localization in the oocyte, and the basis of polarity in the developing embryo, see posterior group genes, anterior group genes, dorsal group genes and terminal genes

Populations of follicle cells

The Drosophila egg is an intricately patterned structure with distinct specializations and polarities. These features are critical to subsequent embryonic development because the polarities of the egg are transmitted to the embryo, establishing the initial pattern in a developing zygote. The pattern of the mature egg is established by complex cellular interactions among and between both somatic follicle cells and germline cells. Each egg begins as a 16-cell germline cyst, from which one cell will become the oocyte and the remainder will become the supporting nurse cells. In the germarium, the anterior structure in which oogenesis is initiated, the germline cyst, is surrounded by a monolayer of somatic follicle cell precursors. As the encapsulated cyst exits from the germarium, approximately 10-14 of the somatic cells cease proliferation and differentiate. This group of cells forms two distinct populations: two polar cells at the anterior and posterior poles of each chamber and approximately seven stalk cells that form a bridge between the consecutive cysts. As the cyst exits the germarium, the other somatic cells covering each chamber, the epithelial follicle cells, remain undifferentiated (Xi, 2003 and references therein).

After pinching off from the germarium, each germline cyst grows, while the epithelial follicle cells proliferate. During this time, the anterior-posterior polarity that will ultimately determine all of the epithelial follicular fates is established. Elegant experiments have shown that the underlying prepattern of the follicular epithelium displays mirror image symmetry at the termini in the anterior-posterior (A/P) axis. Cells adopt one of three anterior terminal fates [border, stretched, and centripetal cells (terminal to central)], depending on proximity to the poles. In the intervening region between the terminal domains, cells will adopt a default 'main body' identity, and the posterior terminal cells form nearest the posterior pole. The symmetry of the A/P pattern is broken by EGFR signaling at the posterior. Secreted Grk from the posteriorly localized oocyte activates EGFR on the overlying follicle cells, establishing posterior terminal fate. In the absence of EGFR signaling, the anterior pattern is repeated at the posterior (Xi, 2003 and references therein).

By stage 7, the epithelial follicle cells cease proliferation and enter an endocycle. Afterward, these cells begin to show morphological and molecular signs of differentiation into the five epithelial fates: border, stretched, centripetal, posterior, and main body cells. Each of these subpopulations of follicle cells has a specific function with respect to the production of a mature egg, such that the correct number and position of each type is critical to ultimate egg morphology. These functions inluence the production of structures that are essential to the egg, such as the dorsal respiratory appendages and the micropyle. These functions are also critical for proper anterior-posterior organization of the oocyte and, therefore, also for the resulting embryo (Xi, 2003 and references therein).

Gene amplification in follicle cells: Isolation of developmental amplicons

Gene amplification is known to be critical for upregulating gene expression in a few cases, but the extent to which amplification is utilized in the development of diverse organisms remains unknown. By quantifying genomic DNA hybridization to microarrays to assay gene copy number, two additional developmental amplicons, termed DAFC (Drosophila Amplicon in Follicle Cells)-30B and -62D were identified in the follicle cells of the Drosophila ovary. Both amplicons contain genes which, following their amplification, are expressed in the follicle cells, and the expression of three of these genes becomes restricted to specialized follicle cells late in differentiation. Genetic analysis establishes that at least one of these genes, yellow-g, is critical for follicle cell function, because mutations in yellow-g disrupt eggshell integrity. Thus, during follicle cell differentiation the entire genome is overreplicated as the cells become polyploid, and subsequently specific genomic intervals are overreplicated to facilitate gene expression (Claycomb, 2004).

The maximally amplified genes in DAFC-62D, yellow-g and yellow-g2, are members of the yellow gene family that are predicted to encode secreted proteins. The family shares homology with the Major Royal Jelly Protein Family in honeybees (Apis mellifera), involved in the specification of the queen bee. The founding member of the Yellow family, Yellow-y, is known to play a role in mating behavior and in the melanization and hardening of the adult cuticle. Other Yellow family members have been shown to act as dopachrome-conversion enzymes that catalyze a key reaction in the melanization process. Interestingly, a similar process is used in the hardening of the egg chorion in mosquitoes and suggests that Yellow-g and Yellow-g2 may play a catalytic role in the crosslinking of the chorion and/or underlying vitelline membrane proteins in Drosophila (Claycomb, 2004).

A second group of genes encodes proteins with chitin binding motifs that could function in egg production. Genes of this type are present in both amplicons, with DAFC-62D containing two such genes and DAFC-30B containing one. Chitin binding domains serve an antimicrobial function in a variety of plants and marine invertebrates. Homologs of marine invertebrate proteins, such as tachycitin, could provide the egg with protection against microbes. Alternatively, chitin, a structural polysaccharide found in many organisms, could also be a component of the eggshell, and interaction with the chitin binding proteins might contribute to eggshell integrity (Claycomb, 2004).

In both DAFC-30B and 62D, there are also a number of genes whose role in follicle cells is not yet clear. These include both genes encoding proteins without known sequence motifs and genes whose products are predicted to have the enzymatic activities of adenylate cyclases, membrane transporters, calcium-transporting ATPases, GTP dissociation inhibitors, and others (Claycomb, 2004).

The yellow-g gene is essential for a rigid eggshell, and the predicted gene products of the yellow-g and yellow-g2 genes suggest a molecular explanation for these mutant defects. The eggshell is composed of several layers, including the outermost exochorion, the endochorion, the inner chorion layer, and the vitelline membrane, which is the innermost structure that also contacts the oocyte. The collapsed embryos and disrupted vitelline membranes that result from mutation of yellow-g indicate that yellow-g is necessary for the structural integrity of the eggshell. At the level of the light microscope, the exochorion of embryos laid by mutant mothers appears normal. The collapsed embryos are reminiscent of vitelline membrane defects, leading to the hypothesis that yellow-g is necessary for proper vitelline membrane formation (Claycomb, 2004).

It is proposed that Yellow-g and Yellow-g2 act to crosslink the vitelline membrane, or perhaps the inner chorion layer. The Yellow family members, Yellow-f and Yellow-f2, are capable of catalyzing the conversion of dopachrome to dihydroxyindole, a limiting step in the melanization pathway, during larval, pupal, and adult stages. The enzymatic events leading to the crosslinking of the vitelline membrane are not well understood, but seem to involve one phase of disulfide bond formation and a subsequent disulfide bond-independent phase. Additionally, the α methyl dopa resistant (amd) gene product, which acts in the conversion of dopamine during the polymerization of the adult cuticle, is required in the follicle cells for proper vitelline membrane crosslinking. This suggests that a similar set of dopamine conversion reactions catalyzed by Yellow-g and Yellow-g2 may be necessary for the crosslinking of the vitelline membrane just prior to egg laying. Consistent with this hypothesis, it is observed that eggs laid by homozygous yellow-g mutant females are highly sensitive to sodium hypochlorite (bleach), and the majority of these embryos burst upon brief exposure. Of the remaining, intact embryos, 100% were permeable to the dye neutral red, which has been used to assay vitelline membrane defects. These results are indicative of a failure to crosslink the vitelline membrane and further implicate yellow-g in the crosslinking process. However, this hypothesis does not explain the specific expression of the yellow-g and yellow-g2 genes in the follicle cells producing the micropyle late in egg chamber development. It is possible that crosslinking of the vitelline membrane or inner chorion layer within this specialized structure requires distinct regulation or timing. A more detailed analysis of the eggshell defect and biochemical studies of Yellow-g and Yellow-g2 will help gain a better understanding of the steps necessary for vitelline membrane crosslinking and will uncover any specialized micropyle functions (Claycomb, 2004).

DAFC-30B and DAFC-62D provide insights into the use of amplification as a developmental strategy. All of the previously characterized amplified genes play a purely structural role in eggshell formation; no enzymes necessary for proper eggshell formation have been examined. None of the genes of DAFC-30B and DAFC-62D encode known structural components of the eggshell. However, several of the amplified genes that are highly expressed in follicle cells, including CG18419 and the yellow-g genes, encode products predicted to possess enzymatic, signal transduction, or transporting activities. Furthermore, at least yellow-g is essential for proper egg formation, thus revealing an additional function of amplification: to increase the levels of enzymes needed to catalyze developmentally important reactions. Thus the identification of additional amplicons highlights genes likely to be crucial in developmental events and opens the possibility that other tissues employ amplification to maximize gene expression during differentiation. It is surprising that a 4- to 6-fold increase in gene copy number would affect gene product levels in a developmentally significant manner. It is possible, however, that copy number increases are considerably higher in subsets of follicle cells, or that the replication process itself facilitates transcription (Claycomb, 2004).

The follicle cell amplicons serve as superb model metazoan replicons, permitting delineation of cis-regulatory elements, identification of replication proteins, and clarifying the developmental control of the initiation and elongation. Developmental distinctions between DAFC-62D and the previously studied DAFCs provide clues into how origin firing can be linked to developmental signals. It has been shown by real-time PCR that replication initiates at DAFC-66D and -7F, coupled with replication fork movement, during egg chamber stages 10B and 11. Subsequently (stages 12 and 13), origins cease firing and only existing replication forks move bidirectionally to produce a gradient of copy number that extends over 100 kb. Furthermore, the replication initiation factor ORC2 localizes to amplification origins only during the initiation phase and dissociates at the onset of the elongation phase. Replication factors involved in multiple steps of DNA replication, such as MCM2-7 and PCNA, colocalize with BrdU throughout amplification (Claycomb, 2004).

DAFC-62D behaves differently from these amplicons and from DAFC-30B. There is a final increase in copy number at a very precise region of the amplicon, about 1.5 kb downstream of yellow-g2, during stage 13. As it is the peak of amplification, this region is likely to possess a replication origin. Understanding how DAFC-62D can undergo a final initiation hours after ORC is no longer detectable at origins by immunofluorescence will provide insights into the control of replication initiation. The additional replication in stage 13 may occur in only subsets of follicle cells, and ORC could persist specifically at DAFC-62D in these cells. For example, additional gene copies could permit optimal levels of expression of the yellow-g genes in the follicle cells building the micropyle (Claycomb, 2004).

These studies were initiated to devise a systematic approach for finding developmental amplicons. The microarray assay is sensitive and can detect low levels of gene amplification, and amplification levels as low as 4-fold can be developmentally important. Thus, this approach will be invaluable in surveying for gene amplification in a number of tissues and in a variety of organisms where amplification has not been detected. Not only has the microarray strategy identified additional amplicons, but when coupled with the power of a genetic organism, it has proven to be a functional genomics approach for highlighting genes involved in specific developmental pathways (Claycomb, 2004).

Development of the dorsal appendages

Dorsal appendage morphogenesis in Drosophila oogenesis has been used as a model system for studying the relationship between patterning and morphogenesis. Each of the two dorsal respiratory appendages of the Drosophila egg chamber is formed by secretion of eggshell proteins into a tube of follicle cells. This tube is generated by cell shape changes and rearrangements within an epithelial sheet. Dorsal appendage formation is therefore similar to more complicated examples of organogenesis. In addition, the study of dorsal appendage formation provides several advantages that make it an excellent system for investigating the regulation of epithelial morphogenesis. For example, the signaling events that determine two populations of dorsal follicle cells are well understood. This understanding facilitates an ability to uncouple effects on patterning from morphogenesis. Further, powerful genetic tools, including mutations that disrupt dorsal appendage formation, have allowed for an unraveling of the genetic circuitry underlying the regulation of epithelial morphogenesis (French, 2003).

The Drosophila egg chamber contains 16 interconnected germline cells, consisting of 1 oocyte nourished by 15 highly polyploid nurse cells; these germline cells are surrounded by a monolayer of ~1000 somatic follicle cells. The follicle cells secrete the chorion that makes up the three layers of the eggshell: the vitelline envelope, the endochorion, and the exochorion. A subset of these follicle cells undergoes morphogenesis to generate the dorsal appendages, specialized structures that facilitate gas exchange in the developing embryo (French, 2003).

At stage 10 of oogenesis, the oocyte occupies the posterior half of the egg chamber, the nurse cells the anterior half, and the oocyte nucleus is positioned at the dorsal anterior corner of the oocyte. The majority of follicle cells forms a columnar layer over the oocyte, while a few follicle cells are stretched out over the nurse cells. During stage 10B, those follicle cells closest to the nurse cell/oocyte boundary begin to migrate centripetally, between nurse cells and oocyte. The centripetal cells secrete the operculum (a thin layer of chorion that functions as an escape hatch for the larva), the collar (a hinge on which the operculum swings), and the micropyle, a coneshaped structure through which the sperm enters (French, 2003).

Shortly after centripetal migration (stage 10B), the nurse cells rapidly transfer their contents into the oocyte (stage 11) then begin to degenerate and undergo apoptosis (stages 12-14). At the same time, two groups of approximately 65-80 anterior, dorsal follicle cells, one on each side of the dorsal midline of the egg chamber, migrate over the nurse cells, laying down the chorion of the two dorsal appendages. Extensive studies have defined the signaling events that determine two populations of dorsal follicle cells. Dorsal follicle-cell fate determination begins when transcripts encoding the TGFalpha-like signaling molecule Gurken (Grk) become localized in a cap above the oocyte nucleus. Grk signals via the epidermal growth factor receptor homolog (Egfr) to the follicle cells, activating a signal transduction cascade involving the Ras/Raf/MAPK pathway. This initial signaling event defines a set of dorsal anterior follicle cells and induces a second signaling cascade involving three additional Egfr ligands. This second cascade amplifies and refines the initial Grk signal, leading to the definition of two separate populations of dorsal follicle cells. These events are required for the production of two separate dorsal appendages. Disruptions of this process result in dorsalization or ventralization of the follicular epithelium and the eggshell. Partial ventralization generally results in failure to determine two separate populations of cells, leading to the production of a single dorsal appendage at the dorsal midline. Complete ventralization results in the absence of dorsal cell fates and the concomitant loss of dorsal appendages (French, 2003).

Information along the anterior-posterior axis also contributes to cell-fate determination within the dorsal appendage primordia. The BMP2/4 homolog encoded by dpp is expressed in the stretch cells and a single row of centripetally migrating cells. This morphogen radiates posteriorly and alters columnar cell fates. High levels of Dpp repress dorsal identities and specify operculum; moderate levels synergize with Grk to define dorsal, while low levels of Dpp are insufficient to allow cells to respond to Egfr signaling. Thus, loss-of-function mutations generate short, often paddleless appendages, while overexpression either expands the operculum at the expense of appendage material or creates multiple, often antler-shaped dorsal structures. The subsequent events underlying dorsal appendage morphogenesis are only beginning to be understood. Analyses of cultured wild-type egg chambers have revealed several phases of dorsal appendage morphogenesis. From stages 10B to 12, two groups of dorsal anterior follicle cells move out from the follicular epithelium to form short tubes. Each tube extends forward over the nurse cells, secreting chorion proteins that make up the cylindrical stalk of the dorsal appendage. Cells at the anterior end of the tube change shape to produce the flattened paddle of the distal dorsal appendage. Finally, upon oviposition, the entire follicular epithelium sloughs off, leaving behind the chorionic structures (French, 2003).

mRNA localization and translational control during oogenesis

RNA localization is often combined with local translational control. This allows simultaneous spatial and temporal control of protein synthesis within a particular region of the cell. In the case of neurons it is speculated that localized mRNAs may be regulated translationally in response to synaptic activity. The combination of translational control with RNA localization can also serve to restrict protein activities to defined regions in the cytoplasm, thereby preventing deleterious interactions from occurring elsewhere in the cell. This seems to be the case for myelin basic protein, which causes membranes to compact, and for developmental determinants whose activities specify the basic body axes and early differentiation of the embryo (Gunkel, 1998 and references).

Most of the current knowledge of how localized mRNAs are controlled translationally has come from studies in Drosophila of the determinants of embryonic polarity encoded by bicoid (bcd), nanos (nos), and oskar (osk). In all three examples, the mRNA is made in nurse cells, transported into the adjacent oocyte, and subsequently localized within the cell. Translation of these mRNAs is silenced transiently during their transport and until the protein is required. The importance of controlling translation during mRNA transport is underlined by the fact that premature or ectopic translation leads to severe developmental defects (Gunkel, 1998 and references). All three of these proteins have additional roles in localization of egg components or in regulation of mRNA translation. Besides acting as a transcriptional regulator, Bicoid functions as a translational regulator of Caudal mRNA. Nanos is an RNA binding protein that binds to and inhibits translation of Bicoid and Hunchback mRNAs. Oscar is a novel protein that functions in the assembly of the germ plasm (Gunkel, 1998 and references).

In a wide range of organisms, including Drosophila, Xenopus, and mouse, many mRNAs appear to be silenced by underadenylation, and their translation is activated by the cytoplasmic elongation of their poly(A) tails. In Drosophila, the poly(A) tail of anteriorly localized bcd mRNA increases from ~50 nucleotides in oocytes, where it is translationally silent, to 150 nucleotides in early embryos, coincident with its activation. Experiments with injected Bcd mRNAs show that a poly(A) tail of 150 nucleotides rescues the bcd phenotype of embryos, whereas a mere 50 nucleotides, as present in the oocyte, do not suffice (Gunkel, 1998 and references).

For NOS, OSK, and very likely a number of other localized mRNAs, translational regulation does not involve modulation of the length of the poly(A) tail. In contrast to Bcd, localization of OSK and NOS mRNAs is required for their translation. Upon fertilization, NOS mRNA is activated only if it resides at the posterior pole. Similarly, OSK mRNA remains repressed translationally in mutants that prevent OSK RNA localization to the posterior pole. Localization and translational repression of NOS and OSK transcripts require regulatory sequences in the 3' UTR. RNA-binding proteins that mediate repression have been identified. A 130-kD protein named Smaug is thought to prevent translation of nos transcripts that have failed to become localized. Smaug repression is mediated by multiple sites, namely the SREs or TCE (Smaug response elements or translation control element) within the Nos 3' UTR. The TCE mediates localization and activation of Nos mRNA, indicating that these aspects of Nos translational regulation are tightly linked and perhaps interdependent. In the case of OSK, premature translation is prevented by Bruno, a 68-kD protein encoded by the arrest (aret) locus. Bruno recognizes a repeated conserved sequence (BRE, for Bruno response element) in the OSK 3' UTR, and colocalizes with OSK mRNA to the posterior pole. The aret mutant phenotype and the colocalization of the protein with other mRNAs in the oocyte suggest that Bruno-mediated repression is not limited to OSK mRNA (Gunkel, 1998 and references).

Translation of localized OSK mRNA is activated specifically through a discrete element situated at the 5' end of the transcript. This element is only active at the posterior pole and is only required when the transcript is repressed through the BRE, suggesting that it functions as a derepressor rather than as a simple enhancer of translation. There is a direct correlation between translational derepression and the binding of a 50-kD (p50) and a 68-kD protein (p68) to this element. One of the 5' binding proteins, p50, also interacts with the BRE in the 3' UTR, and this binding appears to be required for full translation repression. It is concluded that translational activation of localized OSK mRNA is caused not by the local inactivation of repressor, but rather by an active and specific derepression event mediated by a prelocalized machinery. These findings add the notion of "derepressor element", in addition to repressor removal and poly(A) tail lengthening, as means to achieve translational activation (Gunkel, 1998 and references).

Additional information about mRNA localization in Drosophila eggs can be found at the following Interactive Fly sites: Gurken, Homeless, Hunchback, Orb, Pumillo, Staufen, and Vasa.

REFERENCES

Claycomb, J. M., et al. (2004). Gene amplification as a developmental strategy: Isolation of two developmental amplicons in Drosophila. Dev. Cell 6: 145-155. 14723854
French, R. L., Cosand, K. A. and Berg, C. A. (2003). The Drosophila female sterile mutation twin peaks is a novel allele of tramtrack and reveals a requirement for Ttk69 in epithelial morphogenesis. Dev. Bio. 253: 18-35. 12490195

Grünert, S. and St. Johnston, D. (1996). RNA localization and the development of asymmetry during Drosophila oogenesis. Curr. Opinion Gen. Dev. 6: 395-402

Gunkel, N., Yano, T., Markussen, F.-H., Olsen, L. C. and Ephrussi, A. (1998). Localization-dependent translation requires a functional interaction between the 5' and 3' ends of oskar mRNA. Genes Dev. 12: 1652-1664

Spradling, A.C. (1993). Developmental genetics of oogenesis, pp 1-70. In: The Development of Drosophila melanogaster. eds. Bate, M. and Martinez Arias, A. Cold Spring Harbor Press: Long Island, NY.

Xi, R., McGregor, J. R. and Harrison, D. A. (2003). A gradient of JAK pathway activity patterns the anterior-posterior axis of the follicular epithelium. Dev. Cell 4: 167-177. 12586061

date revised:  25 May 2003

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