Regulation and function of maternal mRNA destabilization during early Drosophila development

r 2007, Copyright the Authors Differentiation (2007) 75:482–506 DOI: 10.1111/j.1432-0436.2007.00178.x Journal compilation r 2007, International Society of Differentiation

RE VIEW

Jennifer L. Semotok . Howard D. Lipshitz

Regulation and function of maternal mRNA destabilization during early Drosophila development

Received December 14, 2006; accepted in revised form February 20, 2007

Abstract Early embryonic development in all animals depends on maternally provided gene products. Posttranscriptional and posttranslational processes control spatial and temporal readout of the maternal information. This review focuses on the control of maternal transcript stability in the early Drosophila embryo and how transcript destabilization is necessary for normal development. The molecular pathways that regulate transcript stability are often intimately linked with other posttranscriptional mechanisms such as mRNA localization and translational regulation. These additional mechanisms are explored here with an emphasis on their relationship to transcript decay. Key words mRNA stability
Drosophila
early embryo
maternal transcripts
posttranscriptional control
RNA localization
translational regulation

. ) Jennifer L. Semotok
Howard D. Lipshitz (* Graduate Department of Molecular and Medical Genetics University of Toronto, 1 King’s College Circle Toronto, ON, Canada M5S 1A8 Tel: 1416 946 5296 Fax: 1416 813 7956 E-mail: [email protected] Howard D. Lipshitz Department of Medical Genetics and Microbiology University of Toronto, 1 King’s College Circle Toronto, ON, Canada M5S 1A8 Jennifer L. Semotok
Howard D. Lipshitz Program in Developmental and Stem Cell Biology Research Institute, The Hospital for Sick Children 101 College Street Toronto, ON, Canada M5G 1L7 This article is dedicated to the memory of Laurence D. Etkin: scientist, colleague, friend.

Oogenesis and early embryogenesis in Drosophila Oogenesis The female reproductive system contains two ovaries, each consisting of about a dozen ovarioles that contain an assembly line of maturing egg chambers (Spradling, 1993a). Each egg chamber contains an oocyte and 15 nurse cells, which derive from the germline and are surrounded by a monolayer of somatically derived follicle cells. The endoreplicated nurse cells undergo massive transcription and are responsible for synthesis of the majority of maternal products, which are deposited into the developing oocyte through cytoplasmic bridges known as ring canals. During its 3-day residence in the egg chamber, the oocyte is arrested at meiotic prophase I. Three hours before it is laid, the oocyte undergoes maturation, during which meiosis progresses to metaphase I and certain dormant maternal mRNAs are translationally activated (reviewed in Tadros and Lipshitz, 2005). Egg activation Egg activation serves to trigger additional molecular and cellular events that prepare the mature oocyte for early embryogenesis. In ascidians, echinoderms, and vertebrates, either sperm–egg contact or sperm–egg fusion initiate egg activation by elevating intracellular Ca21 levels through SRK/PLCg/IP3 signaling (Ciapa and Chiri, 2000; Whitaker, 2006). Subsequent regulation of maturation promoting factor (MPF) and mitogen-activated protein kinase (MAPK) signal transduction then drives completion of meiosis and the embryonic cell cycle. Several additional processes are triggered: dormant maternal mRNAs are translationally activated via the lengthening of the poly(A) tail by cytoplasmic polyadenylation; certain maternal mRNAs, which were stable during maturation, are de-

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stabilized; a subset of maternal proteins is degraded and/or undergoes posttranslational modifications such as phosphorylation; and the egg cortex undergoes cytoskeletal modifications (Richter, 1999; Tadros and Lipshitz, 2005; Roux et al., 2006). Similarly, in Drosophila egg activation triggers the completion of meiosis, cortical microtubule depolymerization, and posttranscriptional changes that include cytoplasmic polyadenylation, translational activation, and mRNA destabilization (Mahowald et al., 1983; Theurkauf et al., 1992; Salles et al., 1994; Lieberfarb et al., 1996; Page and Orr-Weaver, 1996, 1997; Tadros et al., 2003). Additionally, the egg swells upon activation and its vitelline membrane, the innermost component of the egg shell, becomes impermeable (Mahowald et al., 1983; Spradling, 1993b). The precise trigger for Drosophila egg activation has been difficult to study because it occurs very rapidly and within the adult female fly. Although the nature of the trigger is unclear, it is known that mature Drosophila oocytes are activated before and independent of fertilization (Doane, 1960; Mahowald et al., 1983). It has been suggested that egg activation is triggered by mechanical stimulation upon exit from the ovary and/or hydration as it enters the lateral oviduct (Mahowald et al., 1983; Page and OrrWeaver, 1997; Heifetz et al., 2001). Recent analyses of sarah (sra), a gene that encodes an inhibitor of the Ca21-dependent phosphatase, calcineurin, have shown that it is required for meiotic progression and translational activation (Horner et al., 2006; Takeo et al., 2006). Thus, just as in other metazoa, Ca21 may play a role in the signaling events during Drosophila egg activation.

Early embryogenesis Early Drosophila embryogenesis is unusual in that 13 synchronous and rapid nuclear divisions occur without cytokinesis, producing  5,000 nuclei within a common cytoplasm (Foe and Alberts, 1983). The events of early embryogenesis occur in the absence of high-level zygotic transcription and are thus dependent on maternally provided molecules. The first seven nuclear divisions occur within the interior of the embryo. During their eighth and ninth cycles, most of the nuclei migrate toward the periphery. The first nuclei to arrive at the posterior pole bud off to form the pole cells, which give rise to the future germline. For the remaining nuclei, cycles 10–13 occur at the surface of the embryo, the syncytial blastoderm stages. During interphase of the 14th cycle, at about 2.5 hr after fertilization, the plasma membrane invaginates between the nuclei to form the cellular blastoderm. With the exception of a few ‘‘early zygotic genes,’’ it is during interphase of cycle 14 where the majority of zygotic transcription occurs, representing the maternal-to-zygotic transition (MZT) in gene

expression (Edgar and Schubiger, 1986; Edgar and Datar, 1996). Cellularization marks the midblastula transition (MBT) as it is the first developmental event that depends on newly synthesized zygotic products (Merrill et al., 1988; Wieschaus and Sweeton, 1988; Yasuda and Schubiger, 1992).

Mechanisms of maternal mRNA turnover in the early Drosophila embryo Much of our knowledge of eukaryotic mRNA turnover has come from genetic and biochemical analyses in the budding yeast, Saccharomyces cerevisiae, and biochemical studies in mammalian cell culture. In both systems, the major degradation pathways initiate with removal of the poly(A) tail, which is a rate-limiting step in mRNA turnover (see Fig. 1) (Wilson and Treisman, 1988; Shyu et al., 1991; Muhlrad and Parker, 1992; Decker and Parker, 1993). Although the systematic knock-out of all degradation components has been fruitful in yeast for defining the molecular pathways leading to mRNA decay, such rigorous genetic analyses are lacking in Drosophila. This is partly due to a lack of mutants and/or lack of characterization of mutant alleles with respect to effects on mRNA turnover, as well as to the lethality and/or pleiotropy of the mutants. Although only a few transcripts have been assessed for changes in poly(A) tail length and stability in the early embryo, all examples investigated to date indicate that deadenylation-dependent decay is also the prominent degradation pathway. Deadenylation and destabilization have been reported in the early embryo for maternal Heat shock protein 83 (Hsp83), nanos (nos), bicoid (bcd), torso (tor), maternal hunchback (hbmat), and oskar (osk) mRNAs (Salles et al., 1994; Lieberfarb et al., 1996; Wreden et al., 1997; Gamberi et al., 2002; Semotok et al., 2005; Jeske et al., 2006; Zaessinger et al., 2006). Deadenylation-dependent decay Deadenylation: Drosophila possesses genes encoding two of the three deadenylases that have been identified in eukaryotes: the CCR4-NOT deadenylase and the PAN2/PAN3 deadenylase; to date, only the former has been studied in detail. Homologs of the PARN deadenylase are absent. In yeast, the major cytoplasmic deadenylation activity derives from the CCR4-NOT deadenylase, a multisubunit complex that includes CCR4 and POP2/CAF1 (the catalytic subunits), NOT1–5, CAF130, and CAF40. In Drosophila, CCR4 (a.k.a. TWIN) and POP2/CAF1 co-purify with a 3 0 poly(A)-dependent exonuclease activity (Temme et al., 2004). Down-regulation of components of the CCR4NOT complex has been shown to have global effects on

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3‘-5’ decay

5‘-3’ decay

7mGpppN

AAAAAAAAAAAAAA SKI2/SKI3/SKI8 EXOSOME

XRN1

7mGpppN

AAAAAAAAAAAAAA AGO2 IRE1 UPF1/UPF2/UPF3

Deadenylation-independent decay

7mGpppN

AAAAAAAAAAAAAA

Deadenylation-dependent decay 7mGpppN 1. CCR4-NOT 2. PAN2/PAN3

Decapping

3‘-5’ decay 7mGpppN

7mGpp pN DCP1/DCP2 DHH1 PAT1 LSM1-7 EDC1/EDC2

SKI2/SKI3/SKI8 EXOSOME

XRN1

5‘-3’ decay

the poly(A) tail lengths of bulk mRNA in Drosophila S2 cell culture, larvae, and adult flies, as well as on the deadenylation status of specific transcripts such as Hsp70 mRNA during heat shock, cyclin A (cycA) mRNA during early oogenesis, and Hsp83 and nos mRNAs during early embryogenesis, thus again demonstrating a conserved role in regulation of transcript deadenylation in vivo (Temme et al., 2004; Morris et al., 2005; Semotok et al., 2005; Zaessinger et al., 2006). Decapping: Two exoribonucleolytic pathways may attack transcripts following their deadenylation. The first pathway (the major one in yeast) occurs via decapping followed by 5 0 -to-3 0 exonucleolytic degradation of the mRNA body. mRNA decapping in yeast and mammals is accomplished by DCP2, the catalytic subunit, along with its enhancer, DCP1, which together catalyze the hydrolysis of the 5 0 -m7GpppN cap to generate 5 0 -m7GDP and a 5 0 -monophosphate mRNA body (reviewed in Coller and Parker, 2004; Meyer et al., 2004). Additional participants such as DHH1 (an RNA helicase), PAT1, LSM1–7, EDC1, and EDC2 serve as en-

Fig. 1 Cytoplasmic mRNA decay pathways that are known or predicted to function in Drosophila. The targeted 5 0 -capped/3 0 -polyadenylated mRNA entering either mRNA decay pathway is pictured in center (highlighted in green). In the deadenylation-dependent pathway (red arrows), mRNA degradation initiates with deadenylation catalyzed by the two deadenylase activities: the CCR4-NOT and PAN2/PAN3 complexes (red arrows). Following removal of the poly(A) tail, the mRNA may be decapped by DCP1/DCP2 and then degraded in the 5 0 –3 0 direction by XRN1, or the mRNA may be degraded in the 3 0 –5 0 direction by the SKI/exosome complex. Alternatively, an mRNA may be subject to endonucleolytic cleavage via the deadenylation-independent pathway (blue arrows). In siRNA-directed silencing, mRNA surveillance, and the unfolded protein response (UPR), mRNAs are cleaved internally and the 5 0 -fragment is degraded via the SKI/exosome complex, while the 3 0 -fragment is degraded via XRN1. These pathways are based on genetic/biochemical data in Drosophila as well as the functions of homologous components in yeast/mammals.

hancers of the decapping process while PAB1 and eIF4E act as repressors. In Drosophila, recent analyses of dDcp1 mutants have revealed defects in degradation of osk, bcd, and twine (twe) mRNAs in the early embryo suggesting a conserved function in regulation of mRNA destabilization (Lin et al., 2006). Although a direct role in cap hydrolysis has not yet been demonstrated in Drosophila, a transgene with an R57 missense mutation, previously shown to be crucial for decapping activity in both yeast and mammals (Tharun and Parker, 1999; LykkeAndersen, 2002), was unable to rescue the maternal mRNA degradation defects of dDcp1 mutants, providing support for dDCP1’s role in mRNA degradation via decapping (Lin et al., 2006). In contrast to its widely known role in cap hydrolysis and mRNA turnover, an interesting additional role of dDCP1 in Drosophila is during oogenesis in posterior localization of osk mRNA (Lin et al., 2006). Because the catalytic enzyme dDCP2 is not found in these dDCP1-osk mRNA-containing ribonucleoprotein

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(mRNP) complexes at the oocyte posterior, it has been postulated that an active decapping activity is also absent and, thus, association of dDCP1 and osk mRNA during localization prepares the latter for subsequent degradation in the early embryo (Lin et al., 2006). Additional factors in these complexes may act to prevent decapping during oogenesis; these include translational enhancer proteins such as ORB (a Drosophila homolog of Xenopus CPEB, which regulates cytoplasmic polyadenylation), cap-binding eukaryotic initiation factor 4E (eIF4E), poly(A)-binding protein (PABP), and/ or the ribosomal machinery itself. Genetic and biochemical analyses of other components of the decapping machinery in the Drosophila embryo are still in their infancy. The closest DHH1 homolog in Drosophila, ME31B, is encoded by an essential gene that has been shown in genetic experiments to function in translational repression of osk and BicD mRNAs during their active transport to the posterior of the oocyte (Nakamura et al., 2001). This evidence, together with ME31B’s colocalization with maternal mRNAs in cytoplasmic particles in both nurse cells and the oocyte, suggests a mechanism of translational silencing (viz., via regulation of the 5 0 cap), although it has not yet been determined whether ME31B functions directly to stimulate decapping. Interestingly, ME31B colocalizes with dDCP1 foci in the nurse cells, and both proteins co-localize with osk mRNA in the oocyte posterior, perhaps in a pre-decapped state (Lin et al., 2006). However, in the early embryo the patterns of expression—both globally and subcellularly—of these proteins, as well as their biochemical interactions, remain unexplored (subcellular localization of degradation components is discussed in ‘‘Spatial regulation of maternal mRNA turnover’’). 5 0 -to-3 0 exoribonucleolytic degradation: Decapping is typically followed by degradation of the mRNA body in the 5 0 -to-3 0 direction by a highly conserved, processive exoribonuclease, XRN1, which was first characterized in yeast (Hsu and Stevens, 1993; Muhlrad et al., 1994). Although its function in Drosophila development has not yet been reported, transgenes encoding the Drosophila XRN1 homolog, pacman (pcm), can rescue xrn1D yeast strains, suggesting a conserved function in mediating exoribonucleolytic mRNA decay (Till et al., 1998). pcm mRNA is expressed during oogenesis and embryogenesis with levels peaking during the first 8 hr post-fertilization, consistent with a function during early embryogenesis (Till et al., 1998). 3 0 -to-5 0 exoribonucleolytic decay: The second deadenylation-dependent decay pathway requires a large multimeric complex called the exosome. The exosome functions in various RNA-processing events in both the nucleus and the cytoplasm. The cytoplasmic exosome, which is involved in mRNA turnover, includes the core RNase PH-like subunits RRP41/SKI6, RRP42, RRP43, RRP45, RRP46, and MTR3; the S1-domain

containing subunits RRP4, RRP40, and CSL4; and the RNase R-like protein RRP44/DIS3 (reviewed in Raijmakers et al., 2004). In addition to its core subunits, the exosome functions in association with the SKI complex, an assembly of SKI2, SKI3, and SKI8 RNA helicases, which are thought to assist degradation by unwinding higher order RNA structures. Together, these complexes target 3 0 -deadenylated messages and direct degradation in the 3 0 -to-5 0 direction. Analyses of cytoplasmic mRNA turnover in mammalian cell culture have revealed that exosome-mediated mRNA decay is the major mammalian deadenylation-dependent decay pathway (Wang and Kiledjian, 2001; Mukherjee et al., 2002), which has been implicated in the decay of transcripts encoding cytokines, lymphokines, growth factors, and proto-oncogenes. Drosophila homologs have been documented for each of the above-mentioned components except RRP43 (Seago et al., 2001; Andrulis et al., 2002; Cairrao et al., 2005). Their function in cytoplasmic mRNA turnover is yet to be established. Recent work by Graham et al. (2006) has shown that exosome components exhibit differences in their localization in the cytoplasm of S2 cells, suggesting differential regulation of mRNA decay in distinct cytoplasmic compartments. Because these studies were performed in cell culture, their implications for exosome function in the early embryo are unclear. dDis3 (a.k.a. tazman or taz) and dSki2 (a.k.a. twister or tst) transcripts are expressed in dynamic patterns throughout the Drosophila life cycle. dDis3 mRNA and protein are maternally expressed and abundant during early embryogenesis, consistent with a role at this stage (Cairrao et al., 2005). Similarly, tst mRNA is present at high levels during the first 2 hr of embryonic development (Seago et al., 2001).

Deadenylation-independent decay Endoribonucleolytic cleavage: There are a small number of examples of deadenylation-independent cytoplasmic degradation of endogenous mRNAs. Best characterized are several endoribonucleases that mediate deadenylation-independent decay: Xenopus polysomal RNase 1 (PMR-1), which acts to destabilize albumin and vitellogenin liver mRNAs (Pastori et al., 1991; Dompenciel et al., 1995; Chernokalskaya et al., 1998; Cunningham et al., 2000); mammalian erythroidenriched endoribonuclease (ErEN), which specifically targets a-globin mRNA for decay during erythroid differentiation (Wang and Kiledjian, 2000a, 2000b; Rodgers et al., 2002); and mammalian Ras-GTPase activating protein SH3 domain-binding protein (G3BP), which functions in endonucleolytic cleavage of c-myc mRNA within its 3 0 -untranslated region (3 0 UTR) (Gallouzi et al., 1998; Tourriere et al., 2001). While homologs of PMR-1 and ErEN are yet to be

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characterized in Drosophila, the G3BP homolog (RASPUTIN a.k.a. RIN) has been shown to play a role in mediating Ras- and Rho-signaling in the ommatidia (Pazman et al., 2000). Although a role for RIN in the regulation of mRNA stability has not yet been reported, rin mRNA is abundant and ubiquitous in the cytoplasm of the early embryo, which may suggest a potential function in RNA metabolism during this time (Pazman et al., 2000). RNA surveillance and the unfolded protein response (UPR): Degradation of aberrant mRNAs is regulated by the so-called ‘‘mRNA surveillance’’ pathway, which is carried out by a complex of proteins that include UPF1, UPF2, and UPF3, which scans the mRNA for premature stop codons (reviewed in Conti and Izaurralde, 2005; Behm-Ansmant and Izaurralde, 2006). In contrast to the exoribonucleolytic mechanisms of mRNA surveillance in yeast and mammals, Drosophila utilizes an endonucleolytic cleavage event that is poly(A) tail- and cap-independent. Gatfield and Izaurralde (2004) were able to capture decay intermediates in Drosophila S2 cell culture and found that endonucleolytic cleavage occurs near the stop codon of an aberrant, premature stop codon-containing mRNA. Following cleavage, the 5 0 fragment (which lacks a poly(A) tail) was degraded by the exosome, while the 3 0 fragment (which lacks a 5 0 cap) was degraded by PCM, the XRN1 homolog. Related to the mRNA surveillance mechanism, in Drosophila S2 cells both the UPR, which functions during cellular stress to help alleviate an overwhelming of the protein folding capacity of the ER, and RNA interference (to be discussed in detail below), also direct the endonucleolytic cleavage of target mRNAs that are subsequently subject to PCM- and SKI/exosomedependent decay (Orban and Izaurralde, 2005; Hollien and Weissman, 2006). To date, no reports have addressed whether any of the mRNA surveillance- or UPR-related factors play a direct role in mRNA decay during development and/or whether these deadenylation-independent endonucleolytic mechanisms function in maternal mRNA regulation in embryos.

RNA silencing An important mechanism of gene regulation discovered during the last decade is transcriptional and posttranscriptional gene silencing mediated by small RNAs (reviewed in Sen and Blau, 2006). These small RNAs can be categorized according to their origins: microRNAs (miRNAs) are derived by transcription of endogenous miRNA genes that contain stretches of self-complementarity, thus forming a hairpin precursor, while small interfering RNAs (siRNAs) are derived from long double-stranded RNAs (dsRNAs) that are either generated from viruses, overlapping transcripts, or provided ex-

ogenously (Meister and Tuschl, 2004). A third class of small RNAs include repeat-associated small interfering RNAs (rasiRNAs), are expressed in the Drosophila germline and derive predominantly from the antisense strand of repetitive elements including retrotransposon and heterochromatic sequences (Saito et al., 2006; Vagin et al., 2006). With respect to biological function in Drosophila, RNA silencing pathways have been implicated in: (1) silencing transgenes, transposable elements, heterochromatin, and endogenous genes; (2) anti-viral defense; (3) gene regulation during developmental processes that include cell fate specification/differentiation, cell proliferation, apoptosis, stem cell division, and epidermal growth factor receptor and NOTCH-mediated cell signaling; and (4) energy homeostasis and fat metabolism (Pal-Bhadra et al., 2002, 2004; Aravin et al., 2001; Brennecke et al., 2003; Xu et al., 2003; Hatfield et al., 2005; Kwon et al., 2005; Li and Carthew, 2005; Ronshaugen et al., 2005; Sokol and Ambros, 2005; Bilen et al., 2006; Galiana-Arnoux et al., 2006; Li et al., 2006; Nguyen and Frasch, 2006; Teleman et al., 2006; Wang et al., 2006). Collectively, short RNAs have critical roles in both transcriptional and posttranscriptional regulation of gene expression in Drosophila. However, only the posttranscriptional mechanisms of RNA silencing will be discussed here (for a general review of RNAi in Drosophila, see Kavi et al., 2005). In Drosophila, three RNaseIII-type endonucleases, DROSHA, DICER-1 (DCR-1), and DICER-2 (DCR2), and their stabilizing dsRNA-binding domain (dsRBD) partners PASHA, LOQUACIOUS (LOQS), and R2D2, respectively, participate in the production of siRNAs and miRNAs (Liu et al., 2003, 2006; Denli et al., 2004; Tomari et al., 2004b; Forstemann et al., 2005; Jiang et al., 2005; Saito et al., 2005). Typically, long primary miRNA hairpin precursors (pri-miRNAs) are first processed in the nucleus by DROSHA/PASHA into  70 nt pre-miRNAs before their export into the cytoplasm (Lee et al., 2003b; Denli et al., 2004). The DCR-1/LOQS complex subsequently converts the exported pre-miRNAs into 21–23 nt miRNAs, while the DCR-2/R2D2 complex functions in the processing of long cytoplasmic dsRNAs into 21–23 nt short duplexes termed siRNAs. In vivo phenotypic analyses of dcr-1 and dcr-2 mutants have revealed that DCR-1 not only participates in miRNA production in early embryos but is also required for siRNA-directed cleavage of target mRNAs, and that both DCR-1 and DCR-2 participate downstream of siRNA processing in the assembly of active mRNA-degradation machines (Lee et al., 2004). The molecular machinery involved in rasiRNA production is currently uncharacterized and appears to be independent of both DCR-1 and DCR-2 (Vagin et al., 2006). Once processed into short RNA duplexes, both miRNAs and siRNAs are assembled into multimeric protein complexes called RNA interference silencing

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complexes, miRISC and siRISC, respectively (details of RISC assembly can be found in Filipowicz, 2005; Sontheimer, 2005; Tang, 2005). RISC represents the active mRNA silencing machinery, which contains a core protein component, ARGONAUTE (AGO). Once a single strand of the short dsRNA duplex is incorporated into RISC, the active complex utilizes the single-RNA strand to recognize its cognate mRNA in a sequencespecific manner, and subsequently triggers mRNA degradation and/or translational repression of the target mRNA. siRNAs tend to possess perfect complementarity to their target mRNAs and, in Drosophila S2 cell culture, direct deadenylation-independent mRNA turnover via endonucleolytic cleavage followed by PCMmediated decay of the 3 0 -fragment and SKI/exosomedirected degradation of the 5 0 -fragment (Elbashir et al., 2001; Orban and Izaurralde, 2005). In most animals, miRNAs contain base-pair mismatches with respect to their target transcripts, and direct either mRNA degradation or translational repression. In Drosophila S2 cells, miRNAs trigger target mRNAs for degradation via AGO1 interaction with another protein, GW182 (a.k.a. GAWKY or GW), which then triggers CCR4NOT deadenylase- and DCP1/DCP2 decapping-dependent decay (Rehwinkel et al., 2005; Behm-Ansmant et al., 2006). miRNA-directed mRNA deadenylation appears to be a conserved mechanism of decay as it has also been shown to occur in mammalian cells and zebrafish embryos (Giraldez et al., 2006; Wu et al., 2006). In mammalian cells and Caenorhabditis elegans, miRNA-directed translational repression of target mRNAs may occur at translation initiation and/or postinitiation (Olsen and Ambros, 1999; Pillai et al., 2005; Petersen et al., 2006), while the precise mechanisms of miRNA-directed mRNA translational repression in Drosophila remain unclear. Four AGO paralogs have been well characterized in Drosophila, which belong to two subfamilies: AGO1 and AGO2 make up the AGO subfamily; PIWI and AUBERGINE (AUB) are members of the PIWI subfamily. AGO1 and AGO2 have been demonstrated to have ‘‘slicer’’ activity, which is critical for RISC-mediated target mRNA cleavage (Rand et al., 2004; Miyoshi et al., 2005). Recombinant PIWI was recently found to have target mRNA cleavage activity in vitro, suggesting that it too possesses slicer activity (Saito et al., 2006). Whether AUB possesses slicer activity is currently unknown. In Drosophila, AGO1’s primary function appears to be in miRNA-directed regulation of target mRNAs while AGO2 function has been implicated in siRNAdirected mRNA cleavage (Okamura et al., 2004). Recent data have indicated partial redundancy in AGO-family protein function. Meyer et al. (2006) demonstrated both genetic and biochemical interactions between AGO1 and AGO2 while Rehwinkel et al. (2006) identified overlapping sets of target mRNAs for AGO1 and AGO2 in knock-down experiments in Drosophila

S2 cell culture. Additionally, AGO2 co-purifies with miRNAs (Rehwinkel et al., 2006) while AGO1 mutants are defective in siRNA-directed mRNA cleavage (Williams and Rubin, 2002). Thus, a precise division of labor between AGO proteins in Drosophila may not exist. Drosophila AGO proteins are expressed maternally and are present ubiquitously throughout the early embryo (Kataoka et al., 2001; Williams and Rubin, 2002). While initially ubiquitous, mRNA encoding PIWI and AUB becomes restricted to the pole cells by the cellular blastoderm stage; in contrast, AGO1 and AGO2 mRNA remain ubiquitous throughout embryogenesis (Williams and Rubin, 2002). Additional putative components of the siRNA- and miRNA-mediated silencing pathways have been identified in Drosophila S2 cells. Biochemical purification of AGO2-containing siRISC identified siRNAs, a micrococcal nuclease domain-containing protein, TUDORSN, and two RNA-binding proteins (RBPs), VASA INTRONIC GENE (VIG) and FRAGILE X-MENTAL RETARDATION PROTEIN 1 (dFMR1) (Caudy et al., 2002, 2003). While the function of dFMR1 in early embryos has been explored (see ‘‘cis-elements and trans-factors: RNA silencing’’), the functions of TUDOR-SN and VIG in Drosophila development are yet to be established. Interestingly, dFMR1 not only associates with components of siRISC but also physically associates with miRISC components, DCR-1 and miRNAs, as well as with ribosomal proteins L5 and L11, and DMP68, an RNA helicase that is essential for siRNA interference in Drosophila (Ishizuka et al., 2002). Such dFMR1 interactions provide further support for an intersection between these pathways. Genetic analyses in Drosophila have uncovered additional members of the RNA silencing pathway. ARMITAGE (ARMI), an RNA helicase homologous to the RNA silencing protein SDE3 in Arabidopsis, has been shown to be required for siRNA-mediated silencing and siRISC assembly in ovarian extracts (Cook et al., 2004; Tomari et al., 2004a), while SPINDLE-E (SPN-E), along with AUB, has been shown to be required for siRNA function in early embryos (Kennerdell et al., 2002).

Cis-elements and trans-factors Cis-acting elements that regulate stability may reside at various locations within a transcript; these include the 5 0 -cap and 3 0 -poly(A) tail, as well as within the UTR and open reading frame (ORF). Either the primary sequence of an UTR/ORF element or its secondary (or even tertiary) structure is recognized by its cognate trans-acting factor. Trans-acting factors can be RBPs or, in the case of RNA silencing, complementary single strands of siRNA or miRNA. These, in turn, recruit

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the mRNA degradation machinery and thus target the mRNA for decay, or act to block the activity of the degradation machinery and thus result in mRNA stabilization. As mRNAs often contain more than one cisacting sequence, the ultimate fate of an mRNA is dependent on integration of the information provided by the cis-elements, the presence or absence of their cognate trans-acting effectors, and the ability of these factors to either stimulate or inhibit mRNA degradation. Changes in the equilibrium of any of these components can result in changes in mRNA stability and, thus, alter the production of a specific protein. The use of transgenic reporter constructs in cis-mapping experiments has been fruitful in unveiling the presence of instability elements. One of the most noted examples is the AU-rich element (ARE), which contains primary sequence enriched in adenosines and uridines and is typically found in the 3 0 -UTR (reviewed in Barreau et al., 2005). Originally identified due to its ability to rapidly destabilize mRNAs encoding several cytokines, growth factors, lymphokines, and proto-oncogenes (including c-fos and c-myc) in mammalian cells, AREs have since been shown to be highly conserved and capable of functioning in the posttranscriptional control of mRNAs in yeast, Drosophila, Xenopus, and in mammals (Grafi and Galili, 1993; Chen and Shyu, 1995; Voeltz and Steitz, 1998; Vasudevan and Peltz, 2001; Jing et al., 2005). Other instability elements have been identified in the 5 0 -UTRs of yeast sdh1, sdh2, suc2, and ppr1 mRNAs, and in the ORFs of mammalian c-fos, c-myc, manganese superoxide dismutase (MnSOD), plasminogen activator inhibitor type 2 (pai-2), and interferon-b (ifn-b) mRNAs, yeast mat1a mRNA, and Drosophila fushi tarazu (ftz) mRNA (Shyu et al., 1989; Wisdom and Lee, 1991; Caponigro et al., 1993; Pierrat et al., 1993; Cereghino et al., 1995; Davis et al., 2001; Ito and Jacobs-Lorena, 2001; Tierney and Medcalf, 2001; de la Cruz et al., 2002; Paste et al., 2003). Several of the transcripts that contain elements in their ORFs also contain instability elements in their 3 0 -UTR (c-fos, c-myc, ifn-b, and pai-2 contain AREs while ftz contains a ftz instability element 3 [FIE3]). Except for the ARE, evolutionary conservation of these elements has not been analyzed. Genetic and biochemical explorations of mRNA instability during development have identified cis-elements and trans-factors that are important for temporal control of maternal mRNA expression. In early Xenopus development, regulation of maternal mRNA stability is mediated by the ARE and its ARE-binding proteins (ARE-BPs), as well as by the embryo deadenylation element (EDEN) and EDEN-binding protein (EDEN-BP) (reviewed in Paillard and Osborne, 2003; Osborne et al., 2005). These instability elements and RBPs have been shown to function during Xenopus oocyte maturation and fertilization, acting to silence maternal mRNAs in early development via deadenylat-

ion. With respect to Drosophila embryos, ARE-mediated transcript regulation has not yet been demonstrated. However, the ability of AREs to elicit transcript decay in an miRNA-dependent manner in Drosophila S2 cells raises the possibility that this mechanism also functions during development (Jing et al., 2005). The Drosophila homolog of EDEN-BP, BRUNO-3 (BRU-3), has been shown to be maternally expressed and capable of binding EDEN sequences via gel mobility shift assays (Delaunay et al., 2004), however, its endogenous targets are yet to be identified. Despite the lack of knowledge of ARE-dependent and EDEN-dependent maternal mRNA degradation in the early Drosophila embryo, there has been significant progress in the identification of instability elements and trans-acting factors that represent major players in the regulation of maternal mRNA stability. Maternal degradation elements and their RBPs SMG response elements (SREs) and SMG: SMAUG’s (SMG) RNA-binding activity maps to its sterile a motif (SAM) domain (Dahanukar et al., 1999; Smibert et al., 1999; Aviv et al., 2003). SAM domains typically mediate protein–protein interactions; thus, the ability of SMG’s SAM domain to bind RNA defines a novel class of RBPs that includes its homologs in yeast (VTS1), C. elegans (ZC190.4), mouse (mSMG1 and mSMG2), and humans (hSMG1 and hSMG2) (Aviv et al., 2003). Drosophila SMG was first identified as a translational repressor of unlocalized nos mRNA in the bulk cytoplasm of the syncytial embryo (Smibert et al., 1996, 1999; Dahanukar et al., 1999). SMG inhibits nos mRNA translation through its physical interaction with CUP, an eIF4E-binding protein that blocks the recruitment of the translation initiation complex (Fig. 2A) (Nelson et al., 2004). SMG triggers maternal mRNA decay in the early Drosophila embryo. Embryos from smg mutant mothers were shown to be defective in elimination of maternal Hsp83 mRNA from the bulk cytoplasm (Fig. 2B) (Semotok et al., 2005). In wild-type early embryos, Hsp83 mRNA is deadenylated and destabilized; both these processes are blocked in smg mutants (Semotok et al., 2005). SMG triggers Hsp83 mRNA decay by recruiting the CCR4-NOT deadenylase to the transcript; this mechanism is supported by both biochemical evidence (purification of SMG’s binding partners identified components of the CCR4-NOT deadenylase complex) and genetic analysis (smg and ccr4 mutants exhibit dominant genetic interactions) (Semotok et al., 2005). As yeast VTS1 also regulates mRNA stability in a CCR4-dependent manner, SMG’s role as an effector of mRNA degradation may be highly conserved (Aviv et al., 2003). Recent in vitro studies using early embryo extracts have suggested that an additional component,

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whose role is to hydrolyze ATP, functions in SMGmediated deadenylation (Jeske et al., 2006). SMG interacts with nos mRNA via direct binding of its SAM domain to two stem-loop structures in the nos 3 0 -UTR, termed SREs (only one is shown in Fig. 2A for simplicity) (Smibert et al., 1996, 1999; Dahanukar et al., 1999). Functional studies of yeast VTS1 and the use of GFP reporter constructs carrying either three wild-type SREs or three point-mutated SREs revealed that VTS1 regulates mRNA stability in an SRE-dependent manner (Aviv et al., 2003). The decay profiles of reporter mRNA constructs bearing SREs revealed that SREs also function in Drosophila to mediate deadenylation in an SMG-dependent manner, suggesting a conserved role for these cis-elements in mediating posttranscriptional control (Semotok et al., 2005). However, although SREs are capable of eliciting SMG-dependent mRNA deadenylation and decay, they do not appear to function in Hsp83 mRNA degradation (Smibert et al., 1999; Semotok et al., 2005). Identification and subsequent mutation of four SRE-like elements within its ORF revealed that Hsp83 mRNA degradation is SRE-independent (Semotok et al., 2005). Thus, SMG functions in both SRE-dependent and SRE-independent mRNA destabilization pathways. The consensus sequence for SREs has recently been refined (Aviv et al., 2006; Johnson and Donaldson, 2006; Oberstrass et al., 2006); mutation of any additional SRE-like sequences in Hsp83 will be required to determine whether they function in Hsp83 mRNA decay. While smg mutations completely stabilize maternal Hsp83 transcripts, they only partially stabilize nos transcripts (Fig. 3), whose elimination must therefore depend on additional factors (Semotok et al., 2005). Recent experiments have suggested a role for CCR4-dependent deadenylation in nos transcript degradation (Zaessinger et al., 2006); it remains to be determined whether CCR4 functions in both the SMG-dependent and SMG-independent aspects of nos transcript elimination. NOS response elements (NREs) and PUM: PUMILIO (PUM) is a highly conserved RBP that belongs to the PUF family of proteins (named after its two founding members, PUM in Drosophila and FBF in C. elegans). Characteristic of PUF proteins, PUM contains eight repeats of the 40 amino acid ‘‘PUF’’ domain in its Cterminus (Wickens et al., 2002). It is through these domains that PUM specifically recognizes a 16 nt bipartite sequence in the 3 0 -UTR of Drosophila hbmat mRNA, termed the NRE (Zamore et al., 1997). Through the two NREs present within hbmat mRNA, PUM binds to the mRNA, forms a complex with the NOS, BRAIN TUMOR (BRAT), and 4EHP proteins, thus controlling the fate of hbmat mRNA via translational repression and/or transcript deadenylation and decay in the posterior half of the early embryo (Wharton and Struhl, 1991; Murata and Wharton, 1995; Wreden et al., 1997; Wharton et al., 1998; Sonoda and Wharton, 1999, 2001; Cho

et al., 2006). PUM has also been implicated in the deadenylation/degradation of bcd mRNA via NREs; this appears to be a NOS-independent mechanism as NOS is not present in the anterior half of the embryo where bcd mRNA exclusively resides (Wharton and Struhl, 1991; Gamberi et al., 2002). Maternal cyclin B (cycB) mRNA is also regulated in a PUM/NOS-dependent manner via an NRE-like sequence in its 3 0 -UTR (Dalby and Glover, 1993; Asaoka-Taguchi et al., 1999). PUM and NOS function together in the early embryo to prevent precocious germ cell division by repressing cycB mRNA translation in the pole cells where cycB transcripts are present at high levels and are stable. Although a mechanistic link to the deadenylation/ degradation machinery has not been made for Drosophila PUM, one of the budding yeast PUM homologs, PUF5/MPT5, has recently been shown to bind to POP2/CAF1 and recruit the CCR4-NOT deadenylase complex to its target mRNAs (Goldstrohm et al., 2006, 2007). Considering the high degree of sequence conservation in the PUF family of RBPs and the similarity of their mRNA target sequences (i.e., the UGUA tetranucleotide), it is reasonable to speculate that Drosophila PUM regulates its target mRNAs in the early embryo via the CCR4-NOT deadenylase complex. In addition to PUM, bcd mRNA degradation depends on dDCP1, thus it is possible that the decapping machinery is then recruited to initiate PUM-dependent mRNA destabilization (Lin et al., 2006). While the molecular details of PUM-mediated mRNA deadenylation/decay for specific transcripts such as hb and bcd are as yet ill-defined, a recent report has identified additional potential maternal mRNA targets that are highly enriched for NRE-like sequences, supporting a role for this primary sequence in posttranscriptional regulation of transcripts (Gerber et al., 2006).

bcd instability element (BIE) and FIEs In addition to the identification of SREs and NREs as cis-elements that direct maternal mRNA degradation, the mapping of instability elements in bcd and ftz mRNAs using transgenic reporter constructs has revealed additional elements that control mRNA instability in early embryos. A BIE maps to the first 43 nt of the bcd-3 0 -UTR and is distinct from the NRE, while ftz mRNA contains three instability elements: FIE5–1, FIE5–2, and FIE3 (Riedl and Jacobs-Lorena, 1996; Surdej and Jacobs-Lorena, 1998; Fontes et al., 2001; Ito and Jacobs-Lorena, 2001). Both BCD and FTZ proteins are critical for the proper patterning of the embryo and thus their precise temporal and spatial expression warrants tight regulation, presumably via multiple regulatory elements. With respect to the identification of the BIE, it is interesting to note that although PUM has been implicated in the degradation of bcd mRNA via

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NREs, mutation of the NREs or mutation of PUM does not completely abolish bcd mRNA degradation but, instead, results in a delay in transcript deadenylation and decay (Gamberi et al., 2002). This supports the hypothesis that partially redundant pathways mediate bcd mRNA decay. FIE3 is a 201 nt element that resides in the ftz-3 0 UTR and is sufficient to mediate degradation of a stable, maternally synthesized mRNA, rpA1, during the first few hours of embryonic development (Riedl and Jacobs-Lorena, 1996; Fontes et al., 2001). FIE5–1 (63 nt) and FIE5–2 (69 nt) are located within the ORF; the ability of these elements to mediate maternal mRNA degradation depends upon their location within an ORF (Ito and Jacobs-Lorena, 2001). While their cognate RBPs or miRNAs/siRNAs remain to be identified for these elements, the ORF-dependence of their function is reminiscent of c-fos and c-myc mRNA degradation where the coding region instability elements require ribosome transit to trigger destabilization.

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Fig. 3 Maternal mRNA stabilization in smg mutant embryos. Mutations in smg completely stabilize Hsp83 transcripts, but only partially stabilize nos transcripts. (A) Northern blots of RNA purified from embryos produced by wild-type (left) or smg (right) mutant mothers at 0–1, 1–2, or 2–3 hr after egg lay. rpA1 transcript levels are unchanged over this time course and thus served as loading controls. (B) Quantitation of Northern blots in (A) with normalization to rpA1 mRNA levels. Reprinted with modifications from Semotok et al. (2005) with permission from Elsevier.

The poly(A) tail—particularly its length—has proven to be a major cis-element controlling mRNA stability (reviewed in Meyer et al., 2004). PABPs bind to adenosine-rich sequences at the 3 0 -end of the message and dramatically influence the fate of mRNAs. PABP can physically interact with components of the translation initiation machinery, such as the cap-associated protein eIF4G, bringing the 5 0 - and 3 0 -ends of the mRNA into close proximity (and thus forming a closed-loop configuration), which stimulates translation and promotes stability (Munroe and Jacobson, 1990; Sachs, 1990; Gallie, 1991, 1998; Tarun and Sachs, 1995; Wells et al., 1998).

PABPs can regulate mRNA stability via interactions with other RBPs. It has been proposed for untranslated c-fos mRNA that a cold-shock domain-containing protein, upstream of N-ras (UNR), and PABP interact with each other and with their cis-acting elements—the

Fig. 2 Models for maternal mRNA degradation and protection in the Drosophila embryo. Representative RNA in situ hybridizations for nos (A) and Hsp83 (B) transcripts. (A) nos mRNA is stable in mature oocytes from which SMG is absent; nos is translationally repressed in the bulk cytoplasm by GLORUND (GLO) via interaction with stem-loop III of the nos translational control element (TCE) in the nos-3 0 -UTR (red arrows). NOS protein is detected in late-stage oocytes, presumably due to OSK-dependent translational activation and/or relief of GLO-mediated repression (green arrows). Upon egg activation, SMG is synthesized ubiquitously and associates with nos mRNA in the bulk cytoplasm via direct interaction with two SMG response elements (SREs) found within the TCE (red arrows, 1). For simplicity, only one of the two SREs is shown. SMG assumes control over nos mRNA translational repression via interaction with CUP, which in turn binds eIF4E and thus interferes with the assembly of the translation initiation complex. SMG–SRE binding also permits the recruitment of the CCR4-NOT deadenylase. An additional as yet unidentified nos degradation factor (NDF) also plays a role in SMG-independent nos mRNA decay. Recruitment of the deadenylase results in removal of the poly(A) tail (2), and subsequent destabilization from the bulk cytoplasm (3). At the posterior pole (green arrows), SMG protein is present but nos mRNA is translated in an OSK-dependent manner (1). SMG-dependent recruitment of the CCR4-NOT deadenylase and/or the CUP-eIF4E translation repression complex may be

blocked via the actions of OSK. For example, OSK could inhibit the SMG-nos mRNA and/or the NDF-nos mRNA association by competing for mRNA binding (i), or by directly interacting with SMG and/or NDF, thus sequestering these proteins (ii). These mechanisms are not necessarily mutually exclusive and would result in stabilization and translation of nos mRNA at the posterior pole (2). (B) Hsp83 mRNA is stable and translated in mature oocytes where SMG is absent. Egg activation triggers ubiquitous SMG protein synthesis. In the bulk cytoplasm (red arrows), SMG associates with Hsp83 mRNA via an as yet unidentified decay element and mediates recruitment of the CCR4-NOT deadenylase complex (1). CCR4-NOT recruitment results in rapid removal of the poly(A) tail (2) and subsequent elimination of the transcript from the bulk cytoplasm (3). At the posterior pole (green arrows), SMG protein is present (1) but SMG-dependent recruitment of the CCR4-NOT deadenylase is blocked via the action of an Hsp83 protection element (HPE) and its hypothetical trans-acting factor, the Hsp83 protection factor (HPF). The HPF could act to prevent deadenylation by inhibiting SMG-Hsp83 mRNA association (i), inhibiting SMG’s ability to recruit the CCR4-NOT complex (ii), or inhibiting the deadenylase or subsequent events (iii). These mechanisms, which are not mutually exclusive, result in Hsp83 mRNA being protected in the posterior (2). Hsp83 mRNA is actively translated in unfertilized eggs and early embryos and is independent of SMG function.

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mRNA coding region determinant (mCRD) and the poly(A) tail, respectively—to form an mRNP that protects against deadenylation (Chang et al., 2004). Upon ribosome transit during translation, this UNR-PABP mRNP is remodeled, thus granting the CCR4-NOT deadenylase access to the mRNA. In addition, PABP appears to protect a-globin mRNA from ErEN-mediated endonucleolytic attack by stabilizing the a-complex (an mRNP complex consisting of the poly(C)-binding proteins, aCP1 and/or aCP2) to the cytosine-rich element (CRE) in the 3 0 -UTR. The a-complex itself also enhances PABP-poly(A) tail interactions and thus further promotes mRNA stability (Wang and Kiledjian, 2000b).

Examination of maternal factors that influence poly(A) tail status in the early Drosophila embryo have revealed that PABP2 is an important regulator of osk and cyc B transcripts (Benoit et al., 2005). Analysis of embryos lacking PABP2 showed that osk and cycB mRNA deadenylation is perturbed. Genetically, PABP2-mediated deadenylation occurs through the CCR4NOT deadenylase pathway (Benoit et al., 2005). As the osk and cycB mRNAs are regulated posttranscriptionally by dDCP1 and PUM, respectively, it will be interesting to determine whether these factors and the PABP2/CCR4-deadenylation pathway act together or independently to regulate these maternal transcripts.

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RNA silencing The role of siRNA- or miRNA-mediated regulation of maternal mRNAs in the early Drosophila embryo is just beginning to be unveiled. It has been demonstrated through the injection of long dsRNA, siRNA, pre-miRNA, or miRNA into early embryos, as well as their incubation with syncytial embryo lysates, that both siRNA- and miRNA-directed processing and RISC activities are present (Tuschl et al., 1999; Lee et al., 2004; Okamura et al., 2004). The presence of these activities, however, does not necessarily implicate them in developmental regulation of maternal mRNAs. Instead, analyses of mutants in the RNA silencing machinery have highlighted their important role in early Drosophila development. Embryos derived from mothers deficient for DCR-1, AGO2, PIWI, or dFMR1 proteins have defective syncytial nuclear cycles while those produced by mothers deficient for DCR-1, AGO2, PIWI, dFMR1, or AUB proteins lack proper pole plasm assembly and/or pole cell formation (Galewsky and Schulz, 1992; Wilson et al., 1996; Harris and Macdonald, 2001; Deshpande et al., 2005, 2006; Megosh et al., 2006; Meyer et al., 2006). There does not appear to be a functional requirement for DCR-2 in the early embryo as dcr-2 mutant mothers give rise to embryos with normal nuclear divisions and pole cells form normally (Megosh et al., 2006). As DCR-2 is essential for siRNA-directed mRNA degradation, these data indicate either that siRNA function is not important for early embryonic development or, more likely, that DCR-2 function is compensated for by DCR-1, DROSHA, or a yet to be characterized DICER-like protein. The role of AGO1 in early development has not yet been explored. However, the fact that removal of one copy of the gene encoding AGO1 enhances the severity of the phenotype of AGO2-deficient embryos indicates functional redundancy between these proteins in the early embryo (Meyer et al., 2006). Identification of the target mRNAs of the miRNAand siRNA-mediated silencing pathways will be essential to fully understand how and why maternal mRNAs are regulated by these posttranscriptional mechanisms. Analysis of the expression patterns of the 78 experimentally validated Drosophila miRNA genes revealed that at least 46 are expressed during the first half of embryogenesis (Aravin et al., 2003; Lai et al., 2003; Leaman et al., 2005). To understand the function of these miRNA genes in embryonic development, Leaman et al. (2005) blocked the specific activity of each of these 46 miRNAs via the microinjection of antisense 2 0 O-methyl oligoribonucleotides into early embryos. Specific developmental phenotypes were identified for 25 of the 46 embryonic miRNAs tested. Interestingly, inhibition of miR-9 function in the early embryo resulted in defective nuclear division, pole cell formation, lipid

droplet formation, and, ultimately, failure to undergo the MBT (i.e., blastoderm cellularization). These results strongly suggest that miR-9 plays a major role in mediating developmental events during the first few hours of embryogenesis (Leaman et al., 2005). Strikingly, the miR-9 phenotype is similar to that seen in dcr-1-, piwi-, dfmr1-, and ago2-deficient embryos. In particular, ago2 mutants have almost identical defects—including the perturbations in lipid droplet formation—suggesting that miR-9 may function through AGO2 in the early embryo (Galewsky and Schulz, 1992; Deshpande et al., 2005; Meyer et al., 2006).

Temporal control of maternal mRNA decay Certain maternal mRNAs—including Hsp83, nos, bcd, osk, tor, cycB, and polar granule component (pgc), and string (stg)—are stable in stage 14 oocytes but are destabilized in the bulk cytoplasm of early embryos (Salles et al., 1994; Edgar and Datar, 1996; Lieberfarb et al., 1996; Surdej and Jacobs-Lorena, 1998; Bashirullah et al., 1999; Gamberi et al., 2002; Tadros et al., 2003). The degradation profile of several of these messages in activated but unfertilized eggs compared with fertilized, developing embryos revealed that there are two degradation pathways that mediate transcript decay in early development (Fig. 4) (Surdej and Jacobs-Lorena, 1998; Bashirullah et al., 1999). The first pathway, the ‘‘maternal’’ degradation pathway, is activated upon egg activation and requires solely maternally provided degradation factors (there is no transcription in activated eggs). This pathway acts on specific mRNA targets such as Hsp83, pgc, nos, and stg while transcripts such as rpA1, rp49, and aTub84B remain stable throughout a 6-hr time course after deposition of the unfertilized eggs (J. L. S and H. D. L., unpublished data, Surdej and Jacobs-Lorena, 1998; Bashirullah et al., 1999; Tadros et al., 2007). The second, ‘‘zygotic’’ pathway exerts its effects starting at about 2 hr into embryonic development (i.e., after pole cell formation but before the MBT). While rpA1, rp49, and aTub84B remain stable (Matthews et al., 1989; Myers et al., 1995; Surdej and Jacobs-Lorena, 1998; Bashirullah et al., 1999; Tadros et al., 2007), this pathway serves to increase the efficiency of elimination of transcripts already destabilized by the maternal pathway (e.g., the half-life of Hsp83 mRNA decreases from 75 to 25 min, Bashirullah et al., 2001). It is also likely to trigger the degradation of mRNAs that are stable in the absence of zygotic transcription (Surdej and Jacobs-Lorena, 1998; Bashirullah et al., 1999; Tadros et al., 2007). Thus, the dual action of the ‘‘maternal’’ and ‘‘zygotic’’ pathways ensures that a subset of maternal transcripts is eliminated from the bulk cytoplasm before the MBT (Bashirullah et al., 1999).

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Fig. 4 Maternal mRNA degradation and the midblastula transition (MBT). Maternal mRNAs are targeted for degradation via dual activities: the ‘‘maternal’’ degradation activity (blue) and the ‘‘zygotic’’ degradation activity (green). The maternal degradation pathway initiates at egg activation and targets transcripts such as Hsp83 and stg (orange), as well as nos and pgc (purple). Because of their lower initial abundance, the maternal degradation pathway alone is sufficient to eliminate nos and pgc mRNAs before initiation of the zygotic pathway. Trans-acting factors that function in the maternal degradation pathway include SMG and the CCR4-NOT deadenylase. The zygotic degradation activity initiates about 2 hr after egg lay and requires de novo transcription. Transcripts such as bcd and hbmat (dark blue) are destabilized in embryos, but not unfertilized eggs and so may be targeted solely via the zygotic degradation activity. This activity also serves to accelerate the decay of

abundant transcripts (such as Hsp83 and stg) previously targeted via the maternal degradation activity. The onset of the zygotic activity is indicated by the vertical black dashed line; the dashed green line indicates that components of the activity must be synthesized before the initiation of mRNA destabilization. PUM and DCP1 are candidate components of the zygotic degradation activity due to their roles in bcd (PUM and DCP1) and hbmat (PUM) mRNA decay. The hypothesized role of miRNAs in the zygotic degradation activity stems from the role of zygotically synthesized miR-430 in destabilizing maternal mRNAs at the maternal-to-zygotic transition (MZT) in zebrafish. Transcripts such as rpA1, rp49, and aTub84B are resistant to both degradation activities and are thus stable throughout early embryogenesis (red). The timing of the syncytial nuclear cycles and cellularization with respect to the time after egg deposition is given at the bottom.

Trigger for maternal mRNA turnover

(GNU) also result in failure to destabilize maternal Hsp83, pgc, and nos transcripts (Tadros et al., 2003). These proteins are components of a novel protein kinase complex that is required for the S-to-M transition during the nuclear divisions of the early syncytial embryo (Shamanski and Orr-Weaver, 1991; Elfring et al., 1997; Fenger et al., 2000; Lee et al., 2003a). Mutations in any of these components result in a failure to enter the M phase; thus the nuclei remain in the S phase and endoreplicate. Several lines of evidence led to the conclusion that the PNG/PLU/GNU complex independently controls the S-to-M transition and maternal transcript degradation in the early embryo (Tadros et al., 2003). The molecular basis for regulation of both the early cell cycle and transcript degradation by PNG, PLU, and GNU has recently been elucidated (Tadros et al., 2007;

Maternal effect mutant screens as well as candidate gene-based approaches identified a number of genes required for maternal mRNA destabilization. cortex (cort) and grauzone (grau) were found to be defective in destabilization of maternal Hsp83 mRNA (Bashirullah et al., 1999). These proteins also function in cytoplasmic polyadenylation and translational activation of bcd and Toll (Tl) mRNAs, as well as the completion of meiosis upon egg activation (Lieberfarb et al., 1996; Page and Orr-Weaver, 1996). This suggests a quite general effect of these, and several other RNA decay-defective mutants, in egg activation (Tadros et al., 2003). Mutations in the genes encoding the PAN GU (PNG), PLUTONIUM (PLU), and GIANT NUCLEI

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Vardy and Orr-Weaver, 2007): smg and cycB mRNAs are not translated in mutants of the kinase complex, respectively explaining the RNA decay and cell-cycle defects. Cytoplasmic polyadenylation has been linked to the translational activation of dormant maternal mRNAs in Xenopus: increasing the poly(A) tail length promotes the binding of PABP, which in turn stabilizes a closedloop configuration via PABP’s interaction with the translation initiation complex (reviewed in Sachs et al., 1997; Richter, 1999). The PNG-kinase complex is required for cytoplasmic polyadenylation of smg and cycB mRNAs (Tadros et al., 2007; Vardy and OrrWeaver, 2007). Surprisingly, however, rescue of the polyadenylation defect in png mutants by the overexpression of poly(A)-polymerase (PAP) does not result in translation of SMG and CYCB proteins (Tadros et al., 2007). Thus, the PNG-kinase regulates translation of these mRNAs via a poly(A)-independent mechanism. Translation of both transcripts is probably due to relief of translational repression; the PUM repressor is a strong contender as pum mutations result in elevated levels of SMG and CYCB protein in the early embryo (Tadros et al., 2007; Vardy and Orr-Weaver, 2007). An interesting observation with respect to the trigger for maternal mRNA decay is that components of the core degradation machinery such as CCR4 and POP2/ CAF1 (implicated in the decay of Hsp83 and nos mRNA), as well as dDCP1 and dDCP2 (implicated in osk, bcd, and twe mRNA decay), are present during both oogenesis and early embryogenesis (Temme et al., 2004; Morris et al., 2005; Lin et al., 2006). PABP2, which regulates deadenylation of cycB and osk mRNAs in the early embryo, is also present and functional in both mature oocytes and early embryos (Benoit et al., 2005). In contrast, SMG protein synthesis occurs only upon egg activation (Smibert et al., 1999; Tadros et al., 2007). The coincidence of the onset of SMG expression and the onset of maternal mRNA decay has led to the hypothesis that SMG is a prime candidate for being the RBP that links the mRNA degradation machinery to specific decay targets (Semotok et al., 2005; Tadros et al., 2007). In SMG’s absence, such as in stage 14 mature oocytes, the degradation machinery is not recruited to these maternal mRNAs, while in SMG’s presence, the degradation machinery is effectively targeted to these mRNAs to initiate destabilization. Unlike SMG, PUM protein is present in late-stage oocytes when maternal transcripts are stable; thus its presence is not sufficient to trigger maternal mRNA decay. This suggests that additional factors present in the early embryo act in conjunction with PUM to direct maternal mRNA decay. As two of PUM’s known direct mRNA targets, bcd and hbmat, are stable in unfertilized eggs and yet unstable in embryos (Surdej and JacobsLorena, 1998; Tadros et al., 2007), it is possible that the PUM-associated factor is zygotically rather than maternally encoded.

Magnitude of maternal mRNA turnover Two RBPs, SMG and PUM, were initially shown to govern the stability of a small number of specific maternal mRNAs in the early embryo. Recent genomewide surveys have permitted the systematic discovery of potential mRNA targets at a global level (Gerber et al., 2006; Tadros et al., 2007). Using cDNA microarrays representing two-thirds of the genome to examine maternal transcript destabilization in wild-type and smg mutant unfertilized eggs, Tadros et al. (2007) were able to show that at least 55% of the protein-coding genome is maternally expressed; at least 20% of the maternally expressed mRNAs are destabilized upon egg activation (  1,600 mRNAs; representing those whose degradation is targeted via the ‘‘maternal’’ degradation pathway); and that 76% of these destabilized mRNAs are eliminated in an SMGdependent manner (Fig. 5). These results highlight the major contribution of SMG in mediating transcript destabilization. Considering that SMG also acts as a translational repressor of nos mRNA through its interaction with the eIF4E-binding protein CUP (Smibert et al., 1996, 1999; Dahanukar et al., 1999; Nelson et al., 2004), this is likely to be an underestimate of the true number of maternal mRNA targets under SMG regulation as it is conceivable that not all SMG-dependent translationally repressed mRNAs are destabilized. Although it is not clear how many of the potential SMG targets are directly regulated by SMG, these data make SMG a frontrunner to be the major posttranscriptional regulator of maternal mRNAs. Interestingly, however, this genome-wide analysis of SMG’s role in transcript decay demonstrated that at least one other, as yet unidentified, degradation factor is required to destabilize the remaining 33% of unstable mRNAs in activated eggs. While SMG-binding sites (i.e., SREs) are significantly enriched in the unstable transcript class relative to the stable class, there is no enrichment of these stem-loop structures in the SMG-dependent versus the SMG-independent class (Tadros et al., 2007). This strongly suggests that SMG-dependent unstable transcripts contain an alternative cis-element(s). This conclusion is consistent with the fact that SMG-dependent destabilization of maternal Hsp83 mRNA appears to occur via an SRE-independent mechanism (Semotok et al., 2005). Furthermore, 48 of the 79 putative mRNA targets of SMG’s S. cerevisiae homolog, VTS1, which were identified via copurification with VTS1 followed by microarray analysis, also did not possess any consensus pentanucleotide-loop SREs (Aviv et al., 2006). Further delineation of the in vivo role of the new consensus SRE stem loop (Aviv et al., 2006) as well as identification of mRNAs that are specifically associated with SMG in early embryos will provide clues as to SMG’s postulated alternative binding-site as well as identify directly bound target mRNAs.

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0-2

2-4

0-2

4-6

2-4

smg 4-6

0-2

wild-type

C 0-2

2-4

2-4

4-6

smg 4-6

0-2

2-4

4-6

712 SMG-dependent transcripts 1 0 –1 –2 –3

357 SMG-independent transcripts 1 0 >2 fold higher than reference sample –1 1:1 >2 fold lower than reference sample

–2 –3

Fig. 5 Genome-scale analysis of the maternal degradation pathway in Drosophila. Microarray-based gene expression profiling of maternal transcript stability in activated unfertilized eggs from wildtype and smg mutant females. Unfertilized eggs were collected 0–2, 2–4, and 4–6 hr after laying. (A) A total of 5,097 maternal mRNAs sorted according to instability at 4–6 hr; each is represented by a horizontal bar with black indicating no change, green a decrease and red an increase in transcript abundance relative to stage 14. Transcripts above the dashed yellow line (ratio’s log base 2 of

 0.59 5 1.5-fold decrease) are significantly destabilized. (B) Left: the 1,069 transcripts that are significantly destabilized in wild-type. Right: the same transcripts in eggs from smg mutant females, showing that many of these are stabilized. (C) Unstable transcripts can be subdivided into two classes. Upper, 712 SMG-dependent transcripts; Lower, 357 SMG-independent transcripts; Y-axis, ratio’s log base 2; dashed yellow line as in (A). Reprinted from Tadros et al. (2007) with permission from Elsevier.

The above-mentioned microarray and bioinformatic analyses solely addressed the degradation of maternal mRNAs in in vivo-activated, unfertilized eggs, in which only the ‘‘maternal’’ degradation activity is active (Bashirullah et al., 1999; Tadros et al., 2003, 2007). To fully understand the global regulation of maternal mRNAs in early embryonic development, where both

the ‘‘maternal’’ and ‘‘zygotic’’ degradation activities function, similar analyses need to be conducted on embryos. A previous study calculated that approximately 27% of maternal mRNAs are degraded within the first 6 hr of embryonic development (Arbeitman et al., 2002); their methods were designed to identify strictly maternally expressed mRNAs, thus leading to an underesti-

496

mate of the number of maternal mRNAs that are degraded in the early embryo. Experiments on embryos using the same reference sample and methods as in Tadros et al. (2007) will be required to identify all maternally loaded transcripts that are targeted for degradation via the maternal pathway, the zygotic pathway, or both. PUM had also been examined previously for its role in posttranscriptional control of maternal mRNAs in Drosophila, and several target transcripts were identified, including hbmat, bcd, cycB, and eIF4E mRNAs (Wharton and Struhl, 1991; Asaoka-Taguchi et al., 1999; Sonoda and Wharton, 1999, 2001; Gamberi et al., 2002; Menon et al., 2004). Additional candidate targets were recently identified by the co-purification of TAPtagged RNA-binding domain of PUM with its associated mRNAs (Gerber et al., 2006). Bioinformatic analysis of the 3 0 -UTRs of 113 of the 150 most highly enriched maternal mRNAs revealed a 16-nucleotide consensus PUM-binding sequence that contains a highly conserved core of eight nucleotides UGUA(A/U/C)AUA (Gerber et al., 2006). This core sequence contains the UGUA tetranucleotide sequence also found among yeast PUF protein target mRNAs, thus demonstrating conserved PUM RNA-binding across evolution.

‘‘Maternal’’ versus ‘‘zygotic’’ decay pathways Bashirullah et al. (1999) first identified the two degradation pathways that promote the proper degradation of maternal mRNAs coincident with the MBT (Fig. 4). From the genome-scale analyses described above, it is clear that SMG represents the major mediator of mRNA decay via the ‘‘maternal’’ degradation pathway. At present, the trans-acting factors involved in the ‘‘zygotic’’ degradation pathway are unknown. This pathway requires Pol II-dependent zygotic transcription as a-amanitin blocks the degradation of reporter transcripts targeted by the zygotic pathway (Cooperstock, 2002; Semotok et al., 2005). The accelerated decay in embryos of transcripts destabilized by the ‘‘maternal’’ pathway could be caused by an increase in abundance of the same degradation machinery, when zygotic transcription begins. Alternatively, new factors might be synthesized that either enhance the activity of the same machinery and/or produce a different machinery. Although little is known about maternal mRNA decay driven by the zygotic pathway in Drosophila, recent work from Giraldez et al. (2006) has shown that an miRNA-directed pathway acts to destabilize maternal mRNAs before the MBT in zebrafish. The miRNA implicated in this process, miR-430, is not expressed maternally but is instead synthesized at the onset of zygotic gene expression in the embryo, at which time it regulates 4750 maternal mRNAs (Giraldez et al., 2006).

These mRNAs show a four- to 10-fold enrichment for miR-430 target sites. Thus, miR-430 is a zygotically synthesized degradation factor responsible for the rapid and specific degradation of maternal mRNAs. While Drosophila does not appear to have a miR-430 homolog, bioinformatic analyses have revealed a significant enrichment for miRNA target sites in maternally expressed Drosophila mRNAs; in particular, target sites for 11 miRNA families are significantly enriched (Tadros et al., 2007). Most of these miRNAs are highly expressed in the early embryo (Aravin et al., 2003; Leaman et al., 2005). As, however, there is no enrichment of miRNA target sites in unstable mRNAs eliminated by the maternal degradation pathway in in vivo-activated, unfertilized eggs, it was hypothesized that miRNA-directed degradation might contribute to the ‘‘zygotic’’ but not to the ‘‘maternal’’ decay pathway (Tadros et al., 2007).

Biological role of maternal mRNA turnover During the 2.5 hr between fertilization and cellularization, the Drosophila embryo rapidly cycles through 13 synchronous syncytial nuclear divisions while simultaneously establishing axial pattern and germline-versussomatic fate. Thus, many of the maternal products deposited into the developing oocyte during oogenesis are devoted to the cell-cycle and cell-fate specification. The MZT occurs in all animals before the MBT, and represents a shift from maternally provided products to newly synthesized zygotic ones (Schultz, 1993). Regulation of the mRNA pool during the MZT has three components: first, elimination of a subset of maternal mRNAs; second, de novo expression of zygotic messages not present in the maternal pool; third, the ongoing expression of mRNAs already present in the mature oocyte (Schultz, 2002). During Drosophila embryogenesis, the MZT is roughly coincident with the MBT, the first developmental event that is absolutely dependent on de novo zygotic mRNA synthesis (i.e., cellularization of the syncytial blastoderm). Elimination of maternal mRNAs during the MZT has been hypothesized to be a pre-requisite for the newly synthesized zygotic messages to take control of development (Fig. 4). Edgar and Datar (1996) provided the first clues regarding the biological role of maternal products in timing the MBT in Drosophila. The two Drosophila Cdc25 phosphatase paralogs, TWE and STG, which are encoded maternally, function in the early embryonic nuclear divisions. If the presence of these maternal products was extended beyond the MZT (by increasing the gene dosage), an additional nuclear division occurred before cellularization (i.e., the MBT was delayed, occurring during interphase of cycle 15 instead of 14). Reciprocally, if the dosage of the twe and stg genes was halved, as a result of which their maternal

5

Repressed Repressed Active 5

Stable Stable Stable 5

5

NA NA Active 5

Stable Stable Stable 5

5

Active Active Active3 rpA1 Stable rp49 NA aTub84B NA

5

Ubiquitous, posterior enriched

Unstable Unstable Unstable Unstable None Anterior

NA Active NA Protein present2 Unstable Stable Stable Stable NA Repressed Repressed Protein present2 Stable Stable NA NA stg bcd hbmat cycB

Ubiquitous, posterior enriched Ubiquitous Anterior Ubiquitous Ubiquitous, posterior enriched Stable pgc

(continued )

(17, 20–23) (10, 20, 22, 24) (23, 25–28) 5

5

(1, 5, 7–10) (11–19) (24, 30–38) (17, 33, 39–50) None Anterior Anterior Posterior pole, perinuclear in bulk cytoplasm Protein present Active Active Protein present2, repressed in pole cells

Posterior pole (1, 5, 6)

2

Unstable NA

Unstable Active Posterior pole (1–5) Unstable Repressed in bulk Posterior pole (1, 5–16) cytoplasm, active in posterior pole plasm

Unstable Active Posterior pole Unstable 6Repressed in bulk Posterior pole cytoplasm, protein present2 in posterior pole plasm Unstable NA Posterior pole Ubiquitous Ubiquitous, posterior enriched

Protein present Repressed in bulk cytoplasm, protein present2 in posterior pole plasm NA Stable Stable

Localization

2

Hsp83 nos

Localization Translation Stability Stability Translation

Localization

Early embryo Unfertilized egg

4

Translation and mRNA stability are often coordinately regulated, enhancing the ability of a cell to precisely and rapidly control protein expression. For example, reduction of protein expression can be achieved by a combination of elimination of the mRNA and translational repression of residual transcripts. There are several cases in the early Drosophila embryo where transcripts such as nos and hbmat are coordinately translationally repressed and destabilized (Table 1; and see ‘‘NOS response elements (NREs) and PUM’’). In both of these cases, the complete absence of detectable protein before mRNA elimination suggests that transcript destabilization is a secondary consequence of translational inhibition (Wreden et al., 1997; Smibert et al., 1999). Conversely, the translational activation of an mRNA along with its stabilization can lead to the rapid production and the ongoing expression of a protein. For example, maternal aTub84B mRNA is translationally active and stable during early embryogenesis (Matthews et al., 1989; Patel and Jacobs-Lorena, 1992).

Maternal Stage 14 oocyte transcript Stability Translation

Maternal mRNA degradation and translational regulation

Table 1 Regulation of maternal mRNAs via stability, translation, and localization in early development

mRNA disappeared earlier, cellularization took place during cycle 13. Thus, the timing of turnover of maternal products may be essential in coordinating the MZT and the MBT. Although direct evidence for the above hypothesis is lacking, both developmental and molecular analyses of the mutant phenotypes of regulators of maternal mRNA instability are beginning to provide support. The smg mutant phenotype in embryos includes defective nuclear divisions after cycle 10 followed by failure to cellularize (Dahanukar et al., 1999). As mentioned earlier, SMG destabilizes 20% of maternal mRNAs in the absence of fertilization ( 1,600 mRNAs). Gene ontology (GO) term analysis has shown that transcripts encoding proteins predicted to function in the cell cycle are significantly enriched in the class of mRNAs that degrade in an SMG-dependent manner (Tadros et al., 2007). Failure of the MBT thus correlates with the persistence of SMG-dependent cell-cycle-related mRNAs, suggesting that elimination of these mRNAs is essential for the MBT. Phenotypic analyses of embryos produced by mother’s mutant for dcr-1, ago2, dfmr1, piwi, and gw has further highlighted the biological importance of posttranscriptional control of maternal mRNAs. All of these mutants exhibit defects in nuclear migration, mitosis, and cellularization (Lee et al., 2004; Deshpande et al., 2005, 2006; Megosh et al., 2006; Meyer et al., 2006; Schneider et al., 2006). The phenotypes of these mutants may, as for smg, be a direct result of failure to eliminate specific maternal transcripts during the MZT.

References

497

NA NA NA NA

twe Tl tor smg

NA Protein present2 Repressed Repressed Ubiquitous Ubiquitous Ubiquitous

5

Repressed in bulk Ubiquitous, cytoplasm, present2 posterior enriched in posterior pole plasm

Localization4 Localization

Unstable 6Repressed in bulk Posterior pole6 cytoplasm, protein present2 in posterior pole plasm 5 Stable NA 5 Stable NA 5 Stable NA Stable NA NA

Stability Translation

Unfertilized egg Translation

Unstable Unstable1 Unstable Unstable

NA Active Active Active

Unstable Present2

Stability

Early embryo

None None None Adjacent to pole cells

(7, 9, 24, 32, 34, 35) (14, 16, 24, 36–39) (14, 16, 24, 40, 41) (24, 42, 43)

Posterior pole (13, 16, 24, 29–33)

Localization

References

Tl mRNA is both maternally and zygotically expressed and thus Northern analyses suggest a stable maternal message while in situ hybridization analyses in Tl zygotic mutants revealed that maternal Tl mRNA disappears throughout the embryo. 2 Where only steady state protein levels were examined, the protein is designated as present. 3 aTub84B mRNA is polysome-associated in ovaries. 4 In some cases, localization is inferred from very early embryos or unfertilized eggs. 5 Transcripts that are known to be expressed during the developmental stage (via genetic or Northern blot analyses) but their spatial information is lacking, are denoted. 6 Inferred from localization in oocytes and embryos. NA: not assayed. References 1. Bashirullah, A., et al. (1999) EMBO J 18:2610. 2. Ding, D., Parkhurst, S.M., Halsell, S.R. and Lipshitz, H.D. (1993) Mol Cell Biol 13:3773. 3. Graziosi, G., Micali, F., Marzari, R., De Cristini, F. and Savoini, A. (1980) J Exp Zool 214:141. 4. Semotok, J.L., et al. (2005) Curr Biol 15:284. 5. Tadros, W., et al. (2003) Genetics 164:989. 6. Nakamura A., Amikura R., Mukai M., Kobayashi S. and Lasko P.F. (1996) Science 274:2075. 7. Edgar B.A. and Datar S.A. (1996) Genes Dev 10:1966. 8. Edgar, B. A. and O’Farrell P.H. (1989) Cell 57:177. 9. Jimenez, J., Alphey, L., Nurse, P., Glover and D.M. (1990) EMBO J 9:3565. 10. Myers, F.A., Francis-Lang, H., Newbury and S.F. (1995) Mech Dev 51:217. 11. Berleth, T. et al. (1988) EMBO J 7:1749. 12. Driever, W. and Nusslein-Volhard, C. (1988) Cell, 54:83. 13. Gamberi, C., Peterson, D.S., He, L. and Gottlieb, E. (2002) Development 129:2699. 14. Lieberfarb, M.E. et al.(1996) Development 122:579. 15. Macdonald, P.M. and Struhl, G. (1988) Nature 336595. 16. Salles, F.J., Lieberfarb, M. E., Wreden, C., Gergen, J.P. and Strickland, S. (1994) Science 266:1996. 17. Surdej, P. and Jacobs-Lorena, M. (1998) Mol Cell Biol 18:2892. 18. Weil, T.T., Forrest, K.M. and Gavis, E.R. (2006) Dev Cell 11:251. 19. Tautz, D., et al. (1987) Nature, 327:383. 20. Al-Atia, G.R., Fruscoloni, P. and Jacobs-Lorena, M. (1985) Biochemistry 24:5798. 21. Fruscoloni, P., Al-Atia, G.R. and Jacobs-Lorena, M. (1983) Proc Natl Acad Sci USA 80:3359. 22. Kay, M.A. and Jacobs-Lorena, M. (1985) Mol Cell Biol 5:3583. 23. Patel, R.C. and Jacobs-Lorena, M. (1992) J Biol Chem 267:1159. 24. Tadros, W. et al. (2007) Dev Cell, 12. 25. Kalfayan, L. and Wensink, P.C. (1982) Cell 29:91. 26. Matthews, K.A., Miller, D.F. and Kaufman, T.C. (1989) Dev Biol 132:45. 27. Mischke, D. and Pardue, M.L. (1982) J Mol Biol 156:449. 28. Natzle, J.E. and McCarthy, B.J. (1984) Dev Biol 104:187. 29. Bergsten S.E. and Gavis, E.R. (1999) Development 126:659. 30. Ephrussi, A., Dickinson, L.K. and Lehmann, R. (1991) Cell 66:37. 31. Kim-Ha, J., Smith, J.L. and Macdonald, P.M. (1991) Cell 66:23. 32. Lin, M.D., Fan, S.J. , Hsu, W.S. and Chou, T.B. (2006) Dev Cell 10:601. 33. Megosh, H.B., Cox, D.N., Campbell, C. and Lin, H. (2006) Curr Biol 16:1884. 34. Alphey, L., et al., (1992) Cell, 69:977. 35. Courtot, C., Fankhauser, C., Simanis, V. and Lehner, C.F. (1992) Development 116:405. 36. Gay, N.J. and Keith, F.J. (1992) Biochim Biophys Acta 1132:290. 37. Gerttula, S., Jin, Y.S. and Anderson, K.V. (1988) Genetics 119:123. 38. Hashimoto, C., Gerttula, S. and Anderson, K.V. (1991) Development 111:1021. 39. Schisa, J. A. and Strickland, S. (1998) Development 125:2995. 40. Casanova, J. and Struhl, G. (1989) Genes Dev 3:2025. 41. Sprenger, F., Stevens, L.M. and Nusslein-Volhard, C. (1989) Nature 338:478. 42. Dahanukar, A., Walker, J. A. and Wharton, R.P. (1999) Mol Cell 4:209. 43. Smibert, C.A., Lie, Y.S., Shillinglaw, W., Henzel, W.J. and Macdonald, P.M. (1999) RNA 5:1535.

1

Maternal mRNAs are designated as stable/unstable with respect to mRNA decay time course analyses. With respect to translational status, maternal mRNAs are labeled as repressed if no protein is detectable in the presence of the mRNA. Translation of maternal mRNAs is labeled active if either de novo protein synthesis experiments were performed or protein expression increased following a developmental stage where it was undetectable or only low levels were found. For mRNA localization transcripts are annotated based on in situ hybridization experiments.

NA

osk

Maternal Stage 14 oocyte transcript Stability Translation

Table 1 Continued

498

499

There are several counter examples to the abovementioned correlations. Active translation of mRNAs such c-fos and c-myc in mammalian cells (Herrick and Ross, 1994; Grosset et al., 2000; Lemm and Ross, 2002; Chang et al., 2004), and mata1 in yeast (Caponigro and Parker, 1996; Hennigan and Jacobson, 1996), is essential to trigger their degradation via deadenylation. Furthermore, in Drosophila, maternal Hsp83 mRNA is actively translated in the early embryo but is degraded (Semotok et al., 2005). In contrast, mRNAs such as those encoding the ribosomal proteins, RPA1 and RP49, are translationally repressed in the early Drosophila embryo and yet are completely stable (Al-Atia et al., 1985; Patel and Jacobs-Lorena, 1992; Bashirullah et al., 1999; Tadros et al., 2007). Thus, knowledge of the translational status of an mRNA is not a means to predict its stability and vice versa. Cis-acting elements such as SREs and NREs regulate both translation and stability in Drosophila (Fig. 2 and ‘‘SMG response elements (SREs) and SMG’’ and ‘‘NOS response elements (NREs) and PUM’’ above). The two SREs within the nos-3 0 -UTR bind SMG, which recruits the eIF4E-binding protein CUP, and blocks translation initiation (Smibert et al., 1999; Nelson et al., 2004). SREs also confer rapid transcript deadenylation and instability to reporter mRNAs as well as to endogenous nos mRNA (Smibert et al., 1996; Semotok et al., 2005; Jeske et al., 2006). The NREs in the hbmat-3 0 -UTR mediate transcript destabilization and translational repression. To accomplish the latter, the NREs, PUM, NOS, and BRAT form a quarternary complex that silences translation of the maternal mRNA in the posterior half of the embryo. The mechanism is similar to that exerted by CUP: BRAT interacts with d4EHP, an eIF4E-like cap-binding protein, which, in turn, blocks eIF4E binding to the 5 0 -cap and thus prevents translation initiation (Cho et al., 2006). The precise mechanisms by which SMG independently regulates translational repression and mRNA deadenylation remain to be elucidated. SMG is found in two distinct complexes in embryos: a SMG/CCR4-NOT deadenylase complex and a SMG/CUP/eIF4E translational repression complex (Semotok et al., 2005). SMG’s physical association with an mRNA is not sufficient to target that transcript for translational repression: maternal Hsp83 mRNA is translated in a SMG-independent manner in early embryos but its deadenylation and degradation is SMG-mediated (Semotok et al., 2005). As Hsp83 mRNA degradation appears to be SREindependent, this suggests that an alternative SMGdependent element mediates deadenylation. How PUM mediates both deadenylation and translational repression is even less clear. A PUM/deadenylase complex is yet to be identified in Drosophila but direct association of PUF5/MPT5 and the CCR4-NOT deadenylase has been demonstrated in S. cerevisiae (Goldstrohm et al., 2006, 2007).

Spatial regulation of maternal mRNA turnover Subcellular localization of the degradation machinery In the last several years, it has been found that at least a subset of posttranscriptional regulation occurs in specialized compartments within a cell’s cytoplasm. mRNP complexes known as processing (P) bodies, which were first identified in yeast, form large cytoplasmic foci that contain decapping components (DHH1, DCP1, DCP2, LSM1–7, PAT1) and the 5 0 –3 0 -exoribonuclease, XRN1 (Sheth and Parker, 2003). In mammalian cells, mRNAprocessing particles termed GW bodies contain the marker GW182, which is required for complex integrity, the CCR4 deadenylase, the cap-binding protein, eIF4E, and its transporter, eIF4-T (homologous to Drosophila CUP), in addition to the above-mentioned P-body components (Cougot et al., 2004; Andrei et al., 2005; Ferraiuolo et al., 2005). P/GW-bodies are dynamic and appear to act as a decision-making crossroad where the fates of specific mRNAs are determined. Within the P-/ GW-body, an mRNA can be translationally silenced and then enter the mRNA degradation pathway or await return to the pool of actively translating mRNAs in the cytoplasm (Sheth and Parker, 2003; Brengues et al., 2005). The Drosophila GW182 homolog, GW, has recently been identified and found to reside in large cytoplasmic foci within the cytoplasm of syncytial-stage embryos (Schneider et al., 2006). A link between P/GW-bodies and the miRNA pathway has been made recently in both mammalian and Drosophila cell culture. Components of the miRNA-directed machinery such as the AGO proteins, miRNAs, and their mRNA targets have been found to be localized to P/GW-bodies (Liu et al., 2005; Rehwinkel et al., 2005; Behm-Ansmant et al., 2006). In Drosophila S2 cells, GW localizes to foci in an RNA-dependent manner and interacts with AGO1, AGO2, PCM, and LSM4 (Rehwinkel et al., 2005; Behm-Ansmant et al., 2006; Schneider et al., 2006). Thus the miRNA/RISC-guided pathway may use cytoplasmic mRNA-processing sites to regulate both the translational status and stability of its target mRNAs. SMG has been shown previously to exist in large particulate foci throughout the bulk cytoplasm of the syncytial embryo, with larger aggregates found at the posterior adjacent to the budded pole cells (Smibert et al., 1999). It is not known whether these SMG-containing particles represent P/GW-bodies, or a distinct RNP where mRNAs are sequestered and silenced via eIF4E-CUP and/or CCR4-NOT complexes. The human SMG homolog, hSMG1, is present in cytoplasmic foci in cultured fibroblasts where it colocalizes with the RNA-binding factors PABP, STAUFEN (STAU), Tcell intracellular antigen 1 (TIA-1), TIA-1-related protein (TIAR), and HuR but not GW182 (Baez and Boccaccio, 2005). Thus hSMG1 particles may not be

500

GW-bodies but, instead, may be stress granules, mRNA triage sites generated during cellular stress. Future analysis of the biochemical composition of SMG particles and their component RNAs and proteins will shed light on their composition and function.

Role of degradation in subcellular transcript localization In Drosophila, the cytoplasm at the posterior pole (the ‘‘posterior pole plasm’’) of the oocyte and early embryo contains polar granules, electron-dense nonmembranebound organelles (reviewed in Mahowald, 2001). The polar granules are essential for germ cell (‘‘pole cell’’) formation and specification in the early embryo. A number of the proteins resident in the granules have been implicated in posttranscriptional control, including translational repression, transport and/or anchoring of transcripts, splicing and RNA silencing; these proteins include OSK, STAU, AUB, PIWI, VASA (VAS), TUDOR (TUD), VALOIS (VLS), FAT FACETS (FAF), and GERMCELL-LESS (GCL). Certain transcripts have also been shown to reside within polar granules, including osk, nos, and pgc mRNAs. Certain maternal transcripts are localized to the polar granules/pole plasm and, subsequently, to the pole cells of the early embryo by a mechanism that combines generalized degradation in the bulk cytoplasm with protection from the degradation machinery in the pole plasm (Fig. 2). The first of these to be identified, Hsp83 mRNA, is ubiquitously distributed throughout the mature oocyte (Ding et al., 1993). Upon egg activation, Hsp83 transcripts are destabilized in the bulk cytoplasm via SMG-dependent recruitment of the CCR4 deadenylase (Bashirullah et al., 1999; Semotok et al., 2005; Tadros et al., 2007). Additional maternal transcripts that are localized to the pole plasm and pole cells by degradation-protection include osk, nos, and pgc (Bashirullah et al., 1999; Bergsten and Gavis, 1999). Unlike Hsp83, these are already localized to the posterior pole plasm during oogenesis (see, e.g., Forrest and Gavis, 2003). This localization is inefficient and thus a large proportion of the mRNA remains in the bulk cytoplasm (e.g., 96% of nos mRNA is unlocalized, see Bergsten and Gavis, 1999). Upon egg activation, the transcripts in the bulk cytoplasm are eliminated (Bashirullah et al., 1999). How are transcripts protected from degradation in the pole plasm? It is known that the pole plasm itself is both necessary and sufficient for protection: mutants that fail to assemble polar granules and the associated pole plasm—such as osk, vas, vls, tud, and stau—fail to protect Hsp83 transcripts, while ectopic assembly of polar granules and pole plasm at the anterior (using an osk-bcd3 0 UTR transgene) results in ectopic protection of Hsp83 transcripts (Ding et al., 1993). Interestingly,

Bashirullah et al. (1999) demonstrated using reporter mRNAs that a transcript degradation activity is present in the posterior pole plasm. The presence of trans-acting factors such as SMG in this subcellular domain (Smibert et al., 1999) supports this conclusion and suggests that transcript protection may occur by blocking the activity of the degradation machinery. One model for maternal mRNA protection is that ‘‘protective’’ factors compete for binding to cis-element(s) that mediate mRNA decay, thus preventing binding/activity of degradation factors (Fig. 2A). Recent studies of nos mRNA provide support for this model: OSK protein may block the association of SMG with the nos-3 0 -UTR, allowing translation and promoting transcript decay (Dahanukar et al., 1999; Zaessinger et al., 2006). Alternatively, cis-acting protection element(s) may be distinct from cis-acting instability element(s) (Fig. 2B). In this case, a bound protection factor could block degradation by establishing an mRNP that is unable to recruit the degradation factor(s) or by acting ‘‘downstream’’ of the degradation element(s) to prevent recruitment of the deadenylase complex or the function of exo- or endonucleases subsequent to deadenylation. Mapping of distinct ‘‘degradation’’ and ‘‘protection’’ elements in transgenic Hsp83 reporter mRNAs (Bashirullah et al., 1999) is consistent with this model. An alternative model that can account for persistence of certain maternal mRNAs in the posterior pole plasm is that transcript-specific degradation components are excluded from the pole plasm. Detailed examination of the subcellular localization of degradation factors will be required to assess this possibility. Degradation-protection-based maternal transcript localization is not restricted to Drosophila. In zebrafish embryos, maternal vasa and nanos mRNAs are restricted to germ cells via this mechanism (Koprunner et al., 2001; Wolke et al., 2002), which also appears to operate in C. elegans (Seydoux and Fire, 1994).

Conclusions Significant progress has been made in understanding of the mechanisms of maternal mRNA degradation in early embryos and its relationship to other posttranscriptional processes, including translation and localization. The identification of the SMG and PUM RBPs as key players in transcript destabilization in Drosophila as well as the discovery of a key role for miRNAs in maternal mRNA destabilization in zebrafish have provided entre´es into both mechanism and function. Future genome-scale analyses in selected mutants, as well as computational identification of conserved cis-regulatory elements (including miRNA target sites) in the different classes of post-transcriptionally regulated maternal mRNAs, will be essential for further progress.

501

Given the high level of evolutionary conservation of cisacting elements and trans-acting factors, as well as of temporal and spatial regulation of stability (as in degradation-protection based localization), studies in Drosophila are likely to continue to lead the way in elucidating both biochemical mechanisms and developmental functions of maternal mRNA destabilization in metazoa. Acknowledgments We thank Dr. Craig Smibert for critical comments on the manuscript. J. L. S. has been supported in part by a Natural Sciences and Engineering Research Council of Canada Graduate Scholarship, a Canada Graduate Scholarship, and a studentship from the Ontario Student Opportunity Trust-Hospital for Sick Children Foundation Student Scholarship Program. H. D. L. is Canada research Chair (CRC) in Developmental Biology at the University of Toronto. Our research on posttranscriptional regulation and RNA localization is supported by funds from the CRC Program; a CIHR Team Grant in mRNP Systems Biology (to H. D. L. and others); and an operating grant from the Canadian Institutes for Health Research (CIHR) to H. D. L.

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