Biochimie 89 (2007) 1211e1220 www.elsevier.com/locate/biochi

MicroRNAs in Drosophila: The magic wand to enter the Chamber of Secrets? Ste´phanie Jaubert a,*, Agne`s Mereau b, Christophe Antoniewski c, Denis Tagu a a

INRA, Agrocampus Rennes, UMR 1099 BiO3P (Biology of Organisms and Populations applied to Plant Protection), BP 35327, F-35653 Le Rheu, France b UMR 6061 CNRS, Universite´ de Rennes 1, IFR 140 Ge´ne´tique Fonctionnelle, Agronomie et Sante´, Faculte´ de Me´decine, 2 avenue du Pr. Le´on Bernard, 35043 Rennes Cedex, France c Laboratory of Drosophila Genetics and Epigenetics, Department of Developmental Biology, CNRS URA 2578, Institut Pasteur, 25 rue du Docteur Roux, 75724 Paris Cedex 15, France Received 8 January 2007; accepted 31 May 2007 Available online 7 June 2007

Abstract MicroRNAs are small non-coding RNAs that are now recognised as key regulators of gene expression in eukaryotes. Over the past few years, hundreds of miRNAs have been identified from various organisms including vertebrates, nematodes, insects and plants. A high level of conservation of some miRNAs from animals to plants underlines their crucial role in eukaryotes. Although biogenesis and mode of action of miRNAs are now quite well understood, their numerous and specific regulatory functions remain to be unravelled. In this review, we summarise the current knowledge on miRNAs in insects, which was mainly acquired through the study of the fruit fly, Drosophila melanogaster. Ó 2007 Elsevier Masson SAS. All rights reserved. Keywords: miRNA; Gene silencing; Insect

1. What are microRNAs? In the 1990s, mutations affecting Caenorhabditis elegans lin-4 and let-7 genes were shown to be responsible for disorder in timing of larva development [1]. Detailed analysis of both these genes revealed that they encode a small non-coding RNA of about 21e24 nucleotides (nt) long. Similar small non-coding RNAs were described in plants as well as in animals [2e6], but not in fungi, in which they were found to act as post-transcriptional regulators of gene expression. They were called microRNAs (miRNAs) Abbreviations: miRNA, microRNA; nt, nucleotide; pri-miRNA, primary nuclear transcript; dsRNA, double-stranded RNAs; siRNA, small interfering RNA; miRISC, miRNA containing RNA-induced silencing complex; EGFR, epidermal growth factor receptor; SOP, sensory organ precursors; GSC, germ stem cells. * Corresponding author. Tel.: þ33 2 23 48 51 65; fax: þ33 2 23 48 51 50. E-mail address: [email protected] (S. Jaubert). 0300-9084/$ - see front matter Ó 2007 Elsevier Masson SAS. All rights reserved. doi:10.1016/j.biochi.2007.05.012

now defined as 21e24 nt long non-coding small RNAs processed from host genome encoded transcript precursors [7]. In animals, miRNAs differ from small interfering RNAs (siRNAs) which are mainly produced by the processing of exogenous double-stranded RNAs (dsRNAs). However, there is striking similarities between miRNA and siRNA structure, biogenesis and mechanisms of action, reflecting a strong overlap between the miRNA and siRNA core machineries. miRNAs are estimated to represent more than 1% of a genome, which thus represent one of the largest gene families in higher eukaryotes [8]. A class of miRNAs is highly conserved throughout evolution, but recent works reported the existence of taxa-specific miRNAs [9e11], such as iab-4, only described in insect species and involved in wing formation [12]. Most miRNAs are located in regions of the genome corresponding to non-coding sequences and are transcribed by RNA

1212

S. Jaubert et al. / Biochimie 89 (2007) 1211e1220

polymerase II as long primary nuclear miRNAs (pri-miRNAs), which range from hundreds to thousands of nucleotides in length [13]. One pri-miRNA typically contains a single or several miRNA precursors (pre-miRNAs) as stem-loop, hairpin structures flanked by unstructured, single stranded RNA sequences [14]. Mature miRNA sequences reside either in the 50 arm or the 30 arm of the stem. About half of the miRNA genes in Drosophila melanogaster are clustered and transcribed from a single polycystronic pri-miRNA [2,15]. An example of such a cluster is provided by the mir-17e18e 19ae20e19b-1 Drosophila gene [2]. Recent analysis of mammalian genomes showed that approximately half of the known mammalian miRNAs are within the introns of protein-coding genes, or within the introns or exons of non-coding RNAs, rather than in their own unique transcription units [16,17]. Similarly, around 50% of the miRNAs in Drosophila melanogaster are located within introns of protein-coding genes (Kim, personal communication). It is important to note that intronic miRNAs are co-ordinately expressed with the pre-mRNA in which they reside. This

correlation has been recently evidenced by microarray analysis in human tissues [18].

2. Biogenesis and action of miRNAs The multiple steps of miRNA processing and mode of action are described in Fig. 1. Pri-miRNAs undergo an initial processing by a ‘‘microprocessor’’ multiprotein complex’’ which contains the nuclear RNAse III Drosha and a doublestranded RNA binding protein named Pasha or DGCR8 [19,20]. Upon the action of Drosha, the 60e70 nt stem-loop structure of the pre-miRNA is cropped from the pri-miRNA [21] and exported from the nucleus by the Ran-GTP dependent transporter exportin-5 [22]. The pre-miRNA loop is then cutout in the cytoplasm by the RNase III enzyme Dicer. Two distinct Dicer enzymes, Dicer-1 and Dicer-2, have been reported in D. melanogaster. Both play a key role in the processing of small non-coding RNAs but only Dicer-1 is required for the pre-miRNA processing, while Dicer-2 is involved in siRNA

Fig. 1. Biosynthesis and action of miRNAs. miRNA genes are transcribed by RNA polymerase II (Pol II) to generate pri-miRNA. Processing is initiated by the Drosha-Pasha microprocessor complex and results in pre-miRNAs of around 60e70 nt. Pre-miRNA are transferred from nucleus to cytoplasm by the Ran-GTP dependant Exportin-5. Cleavage of pre-miRNAs by Dicer in concert with Loquacious occurs in the cytoplasm and produces miRNA duplexes. The duplex is separated and one strand is selected as the mature miRNA whereas the other strand is degraded. Mature miRNA is integrated in the RISC complex composed of the Argonaute-2 protein and Dicer. The gene silencing strategy is defined by the sequence complementarity between miRNA and targeted mRNA. Incomplete base pairing will led to the inhibition of translation whereas perfect base pairing will target mRNA to be degraded by the RISC complex.

S. Jaubert et al. / Biochimie 89 (2007) 1211e1220

biogenesis [23]. Dicer-1 is associated with Loquaciousda dsRNA binding proteindin a multiprotein complex that cleaves pre-miRNAs into miRNA:miRNA duplexes of 21 to 24 nt [24]. MiRNAs duplexes are in turn incorporated into a miRNA containing RNA-induced silencing complex (miRISC) whose core component is the miRNA binding protein Argonaute 1 [25], a member of the PAZ Piwi domain protein family [26]. Within the miRISC complex, one strand of the miRNA duplex, the ‘‘passenger strand’’, is rapidly unwound and degraded [27,28], while the remaining strand serves as a guide to direct through base pairing miRISC to the 30 UTR of target mRNAs. In animals, with very few exceptions, miRNA: mRNA complementarity is imperfect. Despite this partial sequence complementarity, animal miRNAs can regulate gene expression either by translation inhibition or mRNA decay (reviewed in [29]). The contribution of these mechanisms to miRNA gene silencing seems to vary for each miRNA: mRNA pair. mRNA decay induced by miRNAs has been evidenced in Drosophila. This process involves the RISC complex, the Argonaute protein 1 and the GW182 protein and requires deadenylation and decapping of the target mRNAs [30]. However, since the translational status of an mRNA can affect its stability, it is possible that mRNA decay induced by miRNA is a consequence of translational repression rather than a primary effect of miRNA. The exact mechanism of translational repression is still under discussion. Recent works suggested that miRNA gene silencing requires a polyA tail and occurs early in the translation initiation, probably by involving the recognition of m7G cap [29,30]. However, other studies performed on C. elegans and mammals have suggested that miRNA action occurs after translation initiation since miRNA and targets were correctly loaded in ribosomes [29,30]. The localisation of Argonaute proteins, miRNA and miRNA targets in discrete cytoplasmic domains called P-bodies suggested that these domains are the main sites for mRNA degradation/ translational repression [31,32]. Nevertheless, it is unclear whether the localisation in the P-bodies is the cause or the consequence of translational repression and the role of P-bodies in miRNA gene silencing requires further investigation. MiRNAs are key regulators of gene expression in animals and plants. Since the beginning of 2000, the amount of work to characterise their functions in the cellular regulatory networks has exploded and the main goals now are (i) to identify the miRNA coding region in a genome, (ii) to study their expression, (iii) to seek for their mRNA targets and (iv) to characterise their roles in the biological pathways. Drosophila provides a model species that can be studied through powerful genetic and genomic approaches. The first description of the function for a D. melanogaster miRNA was published in 2003 [33], but most papers assigning precise biological roles to miRNAs have only been published over the last two years and have started to open the chamber of secrets of these regulating RNAs. Below, we review the current knowledge on Drosophila miRNAs and emphasise the different functions of miRNA in Drosophila. The structural and evolutional aspects of insect miRNA genes are not developed here since it has recently been reviewed [34].

1213

3. Identification of miRNAs in Drosophila 3.1. Molecular cloning In the early 2000s, high throughput cloning and sequencing approaches led to identification of small non-coding RNAs and, among them, miRNAs [35]. Small RNAs can be sizefractioned by electrophoresis and ligated to RNA/DNA hybrid adapters using dedicated protocols. The ligation products are amplified by RTePCR, concatemerised, cloned in bacterial vectors and sequenced. Through this approach, a significant fraction of the collected sequences is not related to miRNAs, but instead corresponds to other non-coding RNAs such as siRNAs, rasiRNAs or degradation products from abundant cellular RNAs. Thus, identification of miRNAs by cloning requires further validations [36] such as the identification of hairpin secondary structures in the corresponding precursors of candidate sequences. Lagos-Quintana et al. [2] were among the first to describe the cloning and validation of D. melanogaster miRNAs. They obtained the sequences of 17 different miRNAs and analysed their expression profiles in different stages of D. melanogaster development (embryo, larva, pupa, adult, cultured cells). In a similar study, Aravin et al. [37] identified several miRNAs by sequencing cDNA libraries of small non-coding RNAs from embryonic, larval, pupal and adult stages. A total of 382 miRNA sequences were retrieved, corresponding to 60 different miRNA genes. 3.2. Computational approaches Not all miRNAs may be amenable to direct cloning because of low level or restricted expression profiles. A landmark paper was published by Lai and collaborators in 2003 [15] to propose a computational approach for the identification miRNA genes at the Drosophila genome level. At that time, only a few (24) D. melanogaster miRNAs had been cloned. The strategy developed by Lai et al. used an annotation pipeline called miRseeker to scan and seek for miRNAs in insect genomes. To develop this tool, they first had to define how miRNA genes could be specifically recognised. A first analysis of the primary and secondary structures of genomic precursors of the already cloned Drosophila miRNAs was used to edit a set of rules and parameters that define the miRseeker screen. First, the D. melanogaster genome was scanned to mask repeats and eliminate coding exons where no miRNA genes were expected, as well as genes encoding other small RNAs (snRNAs, snoRNAs, rRNAs, tRNAs). Then, 436,000 conserved nucleotidic regions were extracted and kept for further analyses. The second step of miRseeker was based on conserved secondary structures between D. melanogaster and Drosophila pseudoobscura miRNA genes, according to the already known miRNAs. The conserved regions were folded and candidate structures were selected by taking into account several parameters such as the length of the arms and their free energy folding. After ranking, only the best 25% of folded structures were retained. Finally, miRseeker evaluated the divergence patterns of the conserved stem-loops, since miRNAs

1214

S. Jaubert et al. / Biochimie 89 (2007) 1211e1220

can be strongly conserved in evolution [38]. This analysis identified 204 putative miRNA genes. After experimental verification, the authors estimated that around 100 miRNA genes are present in the D. melanogaster genome (which is obviously underestimatedd10 times more in the most recent predictions, also consistent with recent cloning approaches that identified completely new miRNAs; Carthew, personal communication). This indicates that miRNA genes correspond to about 1% of the predicted genes in Drosophila species, as it was estimated for vertebrates [39]. 3.3. miRNA classification Very early in the process of miRNA discovery, a nomenclature was proposed to allow a consistent gene naming scheme. This takes into account several rules to define whether or not the putative newly identify miRNAs differ from other small RNAs such as siRNAs. A set of criteria define a true new miRNA, which includes experimental proof (e.g. by Northern blot) of their expression or prediction of hairpin structure in the precursor or phylogenetic conservation [40]. The miRBase database [41] is dedicated to the collection of all miRNA sequences from any organism, as well as for navigating within miRNA catalogues and checking for putative genomic targets (see below). About 235 insect miRNAs (spring 2007) have already been stored in miRBase (http://microrna.sanger.ac.uk/ sequences/): 78 and 73 for D. melanogaster and D. pseudoobscura, as well as 38, 25, and 21 for Anopheles gambiae, Apis mellifera and Bombyx mori, respectively. 4. Identification of miRNAs targets Functional characterisation of miRNA functions requires the identification of their mRNA target(s). First targets for miRNAs in Drosophila were found by looking at gainof-function mutants in genes involved in the Notch pathway, which contain conserved motifs in their 30 -UTR [42,43]. This study showed that in the fly, the 50 -end of the miRNAs was complementary of the 30 -end of the target mRNA, within a stretch of 7 perfectly matched nucleotides, with no G:U pairs [44]. As miRNAs recognise their target(s) by base complementarities, it became rapidly evident that targets could be identified in silico. However, because of the small size of miRNAs and the imperfect match to their targets, identification of mRNA targets is not possible by standard methods of base pair similarities and requires the development of specific bioinformatics procedures. Stark et al. [45] presented a screen for miRNAs to identify new targets in Drosophila. They first constructed a database of conserved 30 -UTR sequences of mRNAs by comparing D. melanogaster and D. pseudoobscura transcripts: 22% of the D. melanogaster 30 -UTR were conserved in D. pseudoobscura. Within this set, they searched for sequence complementarity with D. melanogaster miRNAs identified by Lai [42]. This work was combined with a RNA folding algorithm which evaluates and ranks the quality based on

structure and energy of the RNA duplex formed between the miRNA and its putative target(s) [46]. In this way, several new miRNA-targets were identified: for example genes belonging to the Notch pathway (e.g. hairy, HLHm3) as targets for miR-7; proapoptotic genes (e.g. reaper, grim, sickle) as target for the miR-2 and enzymes from a metabolic pathway (valine, leucine, isoleucine degradation) as target for miR-277. Several experimental validations were performed as well as a cross-genome comparison using a third insect: A. gambiae. Similar studies were published by Enright et al. [47] and Rajewsky and Socci [48] who identified new potential target genes for D. melanogaster miRNAs. No experimental validations were performed. The miRanda algorithm developed by Enright et al. [47] can be used online at the miRBase website (see above). The search for miRNA targets in Drosophila species have been extended to seven drosophila genomes, using the PicTar algorithm [49]. This algorithm is based on the notion of a ‘‘seed’’ which is a sequence of 7 nucleotides perfectly matched to the target site [50,51]. In conclusion, new computational approaches are regularly described to improve miRNA target identification, such as the work of Miranda et al. [52] who developed the rna22 algorithm. Brennecke et al. [53] developed a functional assay by using transgenic flies expressing green fluorescent genes modified in their 30 -UTR to be targeted by different miRNAs. Based on these experiments, they defined with greater details the minimal miRNA target site required for repression of expression. Their conclusions confirmed that complementarity of the 50 end of miRNAs was sufficient to confer regulation, but weaker complementarity at the 50 -end could be compensated by pairing at the 30 -end. Altogether, these computational studies indicate that a given miRNA can have several target genes and that the expression of more than 10% of the D. melanogaster genes could be regulated by miRNAs, as it was reported for humans [53e55]. Drosophila mutants devoid of key enzymes for miRNA processing have recently been used as new tools to identify the global range of miRNA-regulated genes. Loss of such enzymes in these mutants led to the complete elimination of miRNAs. Since homozygous mutation of dicer-1 is lethal, alternative methods, such as tissue specific or inducible depletion of dicer-1 activity, had to be developed to analyse the effect of the loss of miRNA processing. The proteome of oocytes mutated for dicer-1 and therefore depleted for miRNAs were compared to wild-type oocyte proteome. 41 proteins, mainly involved in protein metabolism, were identified as down regulated by miRNA during oocyte maturation [56]. Moreover, Rehwinkel et al. [57] compared the mRNA transcriptome of wild-type D. melanogaster S2 cells and S2 cells depleted from Ago-1 or Drosha by RNAi. These wild-genome mRNA profiling experiments showed that miRNAs regulate the expression level of approximately 20% of Drosophila S2 cell transcriptome, supporting the idea that miRNAs action is not restricted to translational repression in animals and can effectively reduce the amount of mRNA targets.

S. Jaubert et al. / Biochimie 89 (2007) 1211e1220

5. Functions of miRNAs in Drosophila The majority of miRNAs appears now to be involved in the fine-tuning biological processes by modulating a precise dosage of regulatory proteins such as transcription factors [58]. 5.1. A single process targeted by multiple miRNAs Several methods are used to investigate biological functions of miRNAs. Studies of the general role and importance of the whole miRNA complement have been developed by analysing mutants for genes encoding key enzymes of miRNA biogenesis such as Dicer-1. The loss of dicer-1 leads to the complete and specific elimination of miRNAs processing and has profound effects on Drosophila development within both somatic and germ-lineages [23]. In order to bypass embryonic lethality generated by dicer-1 homozygous mutation, such analyses have been carried out using S2 cultured cells Dicer-depleted by RNAi, or by inducing mitotic Dicer mutant cell clones in flies [23]. Mutation of dicer-1 in D. melanogaster germline stem cells was used to investigate the role of miRNAs in cell division. It was observed that in S2 cells depleted of Dicer-1 by RNAi, the G1 to S phase transition of the cell cycle was delayed, indicating that miRNAs are required to go beyond the normal G1/S cell cycle checkpoint [59]. However, the miRNA(s) involved in this process remain to be characterised. Recent analyses of Drosophila ovaries that are mutant for loquacious or dicer-1 provided interesting results concerning

1215

the role of miRNA in Drosophila female germ stem cells (GSC) control. By analysing ovaries from drosophila mutated for dicer-1, Jin and Xie [60] showed that dicer-1 mutation induced a loss of GSC suggesting that dicer-1 is required for the control of GSC self-renewal. Moreover, they demonstrated that dicer-1 play a general role in ovaries development: in GSC maintenance, growth and development, in somatic stem cell development and in the control of follicle cell proliferation. Since dicer-1 is specific of miRNA synthesis, the authors proposed that miRNAs processed are responsible for these processes. These results have been partly confirmed by the analysis of Drosophila ovaries mutated for loquacious [61]. As it was demonstrated for dicer-1 mutants, a crucial role for Loquacious B was demonstrated in the self renewal of ovaries GSC. However, whereas dicer-1 appears to control GSC maintenance through a bam-independent pathway; loquacious regulation seems to occur in a bam-dependant manner. In the following paragraphs, we describe the role of specific miRNAs described in D. melanogaster. The classification of the role of miRNAs in biological processes is still arbitrary as one given miRNA is often involved in several different cellular functions (Table 1). 5.2. Cell proliferation, apoptosis and organ growth A group of D. melanogaster miRNAs including bantam, mir-278, mir-14 and the mir-2 family, regulate cell proliferation and apoptosis. In 2002, Hipfner et al. reported a new locus named bantam involved in D. melanogaster tissue growth [62].

Table 1 Drosophila miRNAs, for which biological functions have been identified Mechanism

miRNA

Function

Regulators

Target genes

References

Tissue growth

bantam mir-278 mir-14 mir-2

Anti-apoptosis/cell proliferation Anti-apoptosis/homeostasis Anti-apoptosis/fat storage Anti-apoptosis

Hippo pathway Unknown Unknown Unknown

hid Unknown Caspase? grim/reaper/hid family

[33,62e65] [66] [67] [55,68]

Neurogenesis and neurodegeneration

bantam mir-7 mir-9

Anti-apoptosis/protection Photoreceptor differentiation Determination of precise SOP numbers/formation of wings/ cellularisation

Unknown EGF pathway

Unknown Yan senseless

[69] [70] [71]

Muscle differentiation

mir-1

Differentiation and maintenance of muscle cells

twist

Delta (Notch pathway)

[73e75]

mef2 Homeotic transformation

iab-4

Induction of haltere-to-wing transformation

Unknown

Ubithorax

[76]

Energy homeostasis

Mir-278

Regulation of level of circulating sugar Fat storage/mobilisation

Unknown

Unknown

[77]

Metamorphosis

Let-7 Mir-100 Mir-125 Mir-34

Unknown Unknown Unknown Unknown

Unknown Unknown Unknown Unknown

Lin-41 Unknown

[78,81,82]

1216

S. Jaubert et al. / Biochimie 89 (2007) 1211e1220

Loss-of-function mutation of the bantam locus is lethal at early pupal stage whereas hypomorphic combinations of bantam mutant alleles give rise to adult flies, which are smaller that wild-type adults and have some fertility defects. Conversely, over-expression of bantam induces tissue overgrowth due to an increase in the number of cells. Brennecke et al. [33] identified bantam as a 21 nt RNA involved in the control of cell proliferation and apoptosis in D. melanogaster embryos, mostly in developing larval brains and wing disks. Moreover, restoration of bantam activity was sufficient to direct non proliferating cells to enter S phase. The authors also demonstrated an anti-apoptotic activity for bantam, as bantam over-expression suppressed apoptosis induced by the E2F oncogene. The anti-apoptotic activity of bantam was shown to act through translational inhibition of the proapoptotic hid genes. The mechanism of the control of cell proliferation involving bantam is still under investigation. Bantam was shown to be a critical target of the hippo pathway [63,64], which is a new tumour suppressor pathway that coordinates cell proliferation and apoptosis via the control of the transcriptional factor Yki [65]. In D. melanogaster, the expression of bantam appears to be controlled by Yki in order to control organ growth during development. Mir-278, also named mirvana, has been shown to control cell division necessary for eye growth [66] by affecting cell number to increase the size of adult eye via an anti-apoptotic activity. However, whereas complete loss of bantam is lethal, mirvana function is not crucial for normal growth of the whole organism. Mir-14 was reported to suppress cell death in D. melanogaster [67]. Finally, systematic depletion of miR functions by injection of miRNA antisense nucleic acids [68] identified the mir-2 family as controlling apoptosis during D. melanogaster embryonic development. Inhibition of these miRNAs in embryo resulted in an increase of the number in apoptotic cells. Mir-2 action occurs via the inhibition of the translation of proapoptotic factors of the grim/reaper/hid family. These different reports highlight the importance of the regulation of cell division, apoptosis and cell growth by miRNAs. 5.3. Neurogenesis, neurodegeneration and development Polyglutamine (PolyQ) diseases result in a progressive neuronal degeneration due to the expansion of CAG repeat-encoding glutamine within the open reading frame of specific genes. Analysis of flies mutated for dicer-1, loquacious or dicer-2 in the eye indicated that miRNAs modulate PolyQ toxicity as well as toxicity of the Tau protein associated with Alzheimer disease in human. Such a role was demonstrated for bantam that modulates neurodegeneration induced by the PolyQ Ataxin-3 form, as well as neurodegeneration induced by Tau. Bantam acts downstream of toxic protein accumulation or cellular responses to this accumulation, and appears to directly modulate cell survival independently of its anti-apoptotic function involving hid genes [69]. These data point out a novel role of miRNAs in protection against cellular degeneration and open new perspectives for therapy of neurodegenerative human diseases.

Li and Carthew [70] localised mir-7 in the early photoreceptors during embryonic eye development and demonstrated that this miRNA stimulates photoreceptor differentiation. A reciprocal regulation was evidenced between mir-7 and yan, a gene encoding a transcription factor involved in the differentiation of retinal progenitor cells. In undifferentiated retinal cells, Yan protein inhibits mir-7 transcription by direct interaction with cis-regulatory elements upstream of the mir-7 gene. However, in differentiating cells, mir-7 is expressed and inhibits Yan protein synthesis. The switch from undifferentiated to differentiating stage seems to be triggered by the epidermal growth factor receptor (EGFR) signalling pathway. Both mir-7 and yan genes are regulated by the EGFR signalling pathway. Whereas it activates the rapid turnover of Yan protein, EGFR signalling pathway activates mir-7 transcription. This complex mechanism constitutes a negative feedback loop between a miRNA and a transcription factor to control cell fate decision for retinal cells. Mir-9 is involved in the control of the precise numbers of sensory organs in D. melanogaster embryos and adults [71]. Sensory organ precursors (SOP) produce neurons of external sensory organs as well as multidendritic neurons. In nonSOP cells, mir-9 down-regulates the expression of senseless gene encoding a transcription factor involved in the selection process of SOP cells from ectoderm. This inhibition leads to a differential expression of Senseless in SOPs and adjacent epithelial cells which is essential for the production of a precise number of SOP during development. 5.4. Muscle differentiation MiR-1 is an evolutionary conserved miRNA involved in the control of muscle cell identity [72] and cardiomyocyte proliferation of in mammals [73]. Sokol and Ambros [74] reported that D. melanogaster miR-1 is specifically expressed in the mesoderm during early embryogenesis and in myogenic precursors and muscle cells in late embryos. The expression of miR-1 is regulated by different factors during embryogenesis: whereas the expression of miR-1 in mesoderm is directly controlled by the Twist transcription factor, miR-1 transcription is regulated by the Mef2 protein in late embryos. By analysing D. melanogaster mutants devoid of miR-1, Kwon et al. [75] assessed the essential role of miR-1 for muscle differentiation. They showed that miR-1 regulates the determination of specific cardiac and somatic muscle lineages from pluripotent progenitor cells in early embryogenesis. The Delta protein, a ligand for the Notch signalling pathway, was identified as a miR-1 target in cardiac progenitor cells. 5.5. Homeotic transformation The acquisition of segmental identity is controlled during development by Hox genes, whose loci are clustered in few genomic regions in insects as well as in mammals. Hox gene clusters contain conserved miRNAs such as miR-10 or iab-4 [2,37]. In vivo experiments demonstrated that iab-4 selectively reduces the effect of the Ultrabithorax (Ubx) Hox

S. Jaubert et al. / Biochimie 89 (2007) 1211e1220

protein that is involved in controlling wing formation. In haltere imaginal disc, Ubx proteins impose haltere maintenance by inhibiting gene expression that otherwise would direct wing development [76]. The repression of the anterior Hox gene Ubx by iab-4 in haltere discs results in the induction of halter-to-wing homeotic transformation [12]. In silico analyses identified other targets for iab-4 such as Homothorax that works in parallel with Hox cofactors to control legs and antennae patterning [49]. MiRNAs could therefore contribute to the acquisition of segmental identity by regulating not only the expression of hox genes themselves but also the expression of their downstream targets. 5.6. Energy homeostasis A role for miR-278 has been reported in energy homeostasis [77]. MiR-278 mutants present elevated level of circulating sugar and in parallel an overexpression of the insulin-like peptides that control fat storage and mobilisation. However, in spite of high insulin levels, miR-278 mutants remain hyperglycaemic indicating that miR-278 mutants are insulin resistant. This resistance was shown to occur through the regulation of expanded transcripts, a gene encoding a membrane-associated protein. 5.7. Regulation of metamorphosis (ecdysone and juvenile hormones) In D. melanogaster, moulting and metamorphosis are controlled by juvenile hormones and ecdysone. The involvement of miRNAs in Drosophila metamorphosis has been investigated by analysing the expression profile of 24 miRNAs during normal development or following a treatment with the moult hormones ecdysone or juvenile hormones [78]. Twelve of these miRNAs were regulated during normal development, of which let-7, miR-100, miR-125 were up-regulated after ecdysone treatment whereas miR-34 was down-regulated. Juvenile hormone treatment abolished the up-regulation induced by ecdysone and stimulated the expression of miR-34. Let-7 is a 21nt miRNA highly conserved and present in organism such as insects, human and nematodes [79,80]. In C. elegans, let-7 regulates late developmental events through the down regulation of a RING, B-box, coiled-coil protein encoded by the lin-41 gene. A lin-41 orthologue has been identified in Drosophila. As for all bilateral animals tested so far, the expression of D. melanogaster let-7 is correlated with the initiation of adult differentiation. In D. melanogaster, let-7 transcription starts just before metamorphosis and last throughout adulthood [79]. The regulatory link between let-7 and ecdysone is unclear as conflicting results have been reported. On one hand, Sempere et al. [81] indicated that let-7 transcription requires both a pulse ecdysone and the activity of the Broad complex transcription factor. On the other hand, Bashirullah et al. [82] argued against a role for ecdysone signalling in controlling the timing of let-7 and miR-125 transcription. Characterisation of let-7 knock-out mutations

1217

(Ambros, personal communication) should help to clarify the role of miRNAs in endocrine regulation in D. melanogaster. 5.8. A single miRNA for several biological processes Current experimental and bio-informatic data argue that one miRNA can regulate hundred of targets simultaneously [72]. In many cases, a single miRNA affects multiple biological processes. For instance, beside its role in cell death, miR-14 is involved in fat metabolism by regulating the level of triacylglycerols and diacylglycerols. Targets of miR-14 as a regulator of fat metabolism are probably distinct from those that mediate its role as a cell death inhibitor. MiR-9dpreviously described as playing a role in neurogenesisdis also involved in wing formation since miR-9a mutants exhibit a severe wing margin defects. Moreover, a role for miR-9 in cellularisation has been demonstrated. Injection of wild type embryos with an antisense miR-9 probe induced severe defects in nuclear division and migration, pole cell formation, cellularisation, and in the basal movement of yolk droplets [68]. Finally, a role for miR-278 has been reported in energy homeostasis [77]. 6. Conclusions The discovery of miRNAs added an important layer in the sophisticated mechanisms controlling gene expression in eukaryotes and further emphasised the key role of post-transcriptional regulations during the development of metazoans. miRNAs in Drosophila are delivering their secrets, but still much remains to be done before completely characterising their diverse roles. A single miRNA type can regulate several target mRNAs while a single mRNA can be regulated by several miRNAs [83]. Thus, the complexity in this regulatory network is reminiscent to the complexity of gene promoters/ transcription factors interactions. MiRNA are not the only small non-coding regulatory endogenous RNAs. Thanks to high throughput pyrosequencing methods [84], the identification of new small non-coding RNAs is expanding. Earlier work by Aravin et al. [37] allowed the cloning in D. melanogaster of repeat-associated small interfering RNAs (rasiRNAs) which map to repetitive sequence, including retrotransposons and satellite DNA. Dramatic insights on the role of rasiRNAs have been recently provided [85e88]. They appear to function as guides to silence selfish element transcripts, through their interaction with the Argonaute proteins Piwi, Aubergin and Ago3. Likewise, rasiRNA corresponding to the repetitive Su(Ste) sequences have been shown to be partially responsible for the repression of testisexpressed Stellates genes at the post-transcriptional level [88e90]. Finally, repetitive sequences and retrotransposable elements are highly enriched in heterochromatic regions of chromosomes that mainly map to pericentromeres and telomeres. An increasing number of reports strongly suggest that Drosophila rasiRNAs, as in plants and Schizosaccharomyces pombe [91], might be also involved in the control of heterochromatin formation, thereby potentially mediating silencing at the transcriptional level [92e94]. Altogether, these data

1218

S. Jaubert et al. / Biochimie 89 (2007) 1211e1220

reinforces the role of small non-coding RNAs in regulating multiple cellular functions.

Acknowledgements The authors would like to thank Howard-Beverly Osborne (UMR CNRS 6061, Rennes) for careful reading of the manuscript. This work was supported by the INRA (Action Inte´gre´e Programme´e Se´quenc¸age 2005e2006) and by a grant from the ANR (AKROSS programme) to C.A.

References [1] R.C. Lee, R.L. Feinbaum, V. Ambros, The C. elegans heterochronic gene lin-4 encodes small RNAs with antisense complementarity to lin-14, Cell 75 (1993) 843e854. [2] M. Lagos-Quintana, R. Rauhut, W. Lendeckel, T. Tuschl, Identification of novel genes coding for small expressed RNAs, Science 294 (2001) 853e858. [3] M. Lagos-Quintana, R. Rauhut, J. Meyer, A. Borkhardt, T. Tuschl, New microRNAs from mouse and human, RNA 9 (2003) 175e179. [4] N.C. Lau, L.P. Lim, E.G. Weinstein, D.P. Bartel, An abundant class of tiny RNAs with probable regulatory roles in Caenorhabditis elegans, Science 294 (2001) 858e862. [5] R.C. Lee, V. Ambros, An extensive class of small RNAs in Caenorhabditis elegans, Science 294 (2001) 862e864. [6] B.J. Reinhart, E.G. Weinstein, M.W. Rhoades, B. Bartel, D.P. Bartel, MicroRNAs in plants, Genes Dev. 16 (2002) 1616e1626. [7] V. Ambros, B. Bartel, D.P. Bartel, C.B. Burge, J.C. Carrington, X. Chen, G. Dreyfuss, S.R. Eddy, S. Griffiths-Jones, M. Marshall, M. Matzke, G. Ruvkun, T. Tuschl, A uniform system for microRNA annotation, RNA 9 (2003) 277e279. [8] D.P. Bartel, MicroRNAs: genomics, biogenesis, mechanism, and function, Cell 116 (2004) 281e297. [9] J. Hertel, M. Lindemeyer, K. Missal, C. Fried, A. Tanzer, C. Flamm, I.L. Hofacker, P.F. Stadler, Students of Bioinformatics Computer Labs 2004 and 2005, The expansion of the metazoan microRNA repertoire, BMC Genomics 15 (7) (2006) 25. [10] I. Bentwich, A. Avniel, Y. Karov, R. Aharonov, S. Gilad, O. Barad, A. Barzilai, P. Einat, U. Einav, E. Meiri, E. Sharon, Y. Spector, Z. Bentwich, Identification of hundreds of conserved and nonconserved human microRNAs, Nat. Genet. 37 (2005) 766e770. [11] E. Berezikov, V. Guryev, J. van de Belt, E. Wienholds, R.H. Plasterk, E. Cuppen, Phylogenetic shadowing and computational identification of human microRNA genes, Cell 120 (2005) 21e24. [12] M. Ronshaugen, F. Biemar, J. Piel, M. Levine, E.C. Lai, The Drosophila microRNA iab-4 causes a dominant homeotic transformation of halteres to wings, Genes Dev. 19 (2005) 2947e2952. [13] Y. Lee, Y.K. Jeon, J.T. Lee, S. Kim, V.N. Kim, MicroRNA maturation: stepwise processing and subcellular localization, EMBO J 21 (2002) 4663e4670. [14] T.M. Rana, Illuminating the silence: understanding the structure and function of small RNAs, Nat. Mol. Cell Biol. 8 (2007) 23e36. [15] E.C. Lai, P. Tomancak, R.W. Williams, G.M. Rubin, Computational identification of Drosophila microRNA genes, Genome Biol. 4 (2003) R42. [16] A. Rodriguez, S. Griffiths-Jones, J.L. Ashurst, A. Bradley, Identification of mammalian microRNA host genes and transcription units, Genome Res. 14 (2004) 1902e1010. [17] Y.K. Kim, V.V. Kim, Processing of intronic microRNAs, EMBO J 26 (2007) 775e783. [18] S. Baskerville, D.P. Bartel, Microarray profiling of microRNAs reveals frequent coexpression with neighboring miRNAs and host genes, RNA 11 (2005) 241e247.

[19] M. Landthaler, A. Yalcin, T. Tuschl, The human DiGeorge syndrome critical region gene 8 and Its D. melanogaster homolog are required for miRNA biogenesis, Curr. Biol. 14 (2004) 2162e2167. [20] A.M. Denli, B.B. Tops, R.H. Plasterk, R.F. Ketting, G.J. Hannon, Processing of primary microRNAs by the Microprocessor complex, Nature 432 (2004) 231e235. [21] J. Han, Y. Lee, K.H. Yeom, J.W. Nam, I. Heo, J.K. Rhee, S.Y. Sohn, Y. Cho, B.T. Zhang, V.N. Kim, Molecular basis for the recognition of primary microRNAs by the Drosha-DGCR8 complex, Cell 125 (2006) 887e901. [22] T. Bohnsack, K. Czaplinski, D. Gorlich, Exportin 5 is a RanGTPdependent dsRNA-binding protein that mediates nuclear export of premiRNAs, RNA 10 (2004) 185e191. [23] Y.S. Lee, K. Nakahara, J.W. Pham, K. Kim, Z. He, E.J. Sontheimer, R.W. Carthew, Distinct roles for Drosophila Dicer-1 and Dicer-2 in the siRNA/miRNA silencing pathways, Cell 117 (2004) 69e81. [24] K. Saito, A. Ishizuka, H. Siomi, M.C. Siomi, Processing of pre-microRNAs by the Dicer-1-Loquacious complex in Drosophila cells, PLoS Biol. 3 (2005) e235. [25] K. Okamura, A. Ishizuka, H. Siomi, M.C. Siomi, Distinct roles for Argonaute proteins in small RNA-directed RNA cleavage pathways, Genes Dev. 18 (2004) 1655e1666. [26] M.A. Carmell, Z. Xuan, M.Q. Zhang, G.J. Hannon, The Argonaute family: tentacles that reach into RNAi, developmental control, stem cell maintenance, and tumorigenesis, Genes Dev. 16 (2002) 2733e2742. [27] C. Matranga, Y. Tomari, C. Shin, D.P. Bartel, P.D. Zamore, Passengerstrand cleavage facilitates assembly of siRNA into Ago2-containing RNAi enzyme complexes, Cell 123 (2005) 607e620. [28] K. Miyoshi, H. Tsukumo, T. Nagami, H. Siomi, M.C. Siomi, Slicer function of Drosophila Argonautes and its involvement in RISC formation, Genes Dev. 19 (2005) 2837e2848. [29] R.S. Pillai, S.N. Bhattacharyya, W. Filipowicz, Repression of protein synthesis by miRNAs: how many mechanisms? Trends Cell Biol. 17 (2007) 118e126. [30] I. Behm-Ansmant, J. Rehwinkel, T. Doerks, A. Stark, P. Bork, E. Izaurralde, mRNA degradation by miRNAs and GW182 requires both CCR4:NOT deadenylase and DCP1:DCP2 decapping complexes, Genes Dev. 15 (2006) 1885e1898. [31] J. Liu, M.A. Valencia-Sanchez, G.J. Hannon, R. Parker, MicroRNAdependent localization of targeted mRNAs to mammalian P-bodies, Nat Cell Biol. 7 (2005) 719e723. [32] J. Liu, F.V. Rivas, J. Wohlschlegel, J.R. Yates, R. Parker, G.J. Hannon, A role for the P-body component GW182 in microRNA function, Nat. Cell. Biol. 7 (2005) 1261e1266. [33] J. Brennecke, D.R. Hipfner, A. Stark, R.B. Russell, S.M. Cohen, , bantam encodes a developmentally regulated microRNA that controls cell proliferation and regulates the proapoptotic gene hid in Drosophila, Cell 113 (2003) 25e36. [34] K.S. Behura, Insect microRNAs: Structure, function and evolution, Insect Biochem. Mol. Biol. 37 (2007) 3e9. [35] E. Berezikov, E. Cuppen, R.H. Plasterk, Approaches to microRNA discovery, Nat. Genet. 38 (Suppl) (2006) S2eS7. [36] I. Bentwich, Prediction and validation of microRNAs and their targets, FEBS Lett. 579 (2005) 5904e5910. [37] A.A. Aravin, M. Lagos-Quintana, A. Yalcin, M. Zavolan, D. Marks, B. Snyder, T. Gaasterland, J. Meyer, T. Tuschl, The small RNA profile during Drosophila melanogaster development, Dev. Cell 5 (2003) 337e350. [38] T. Tran, P. Havlak, J. Miller, MicroRNA enrichment among short ’ultraconserved’ sequences in insects, Nucleic Acids Res. 34 (2006) e65. [39] L.P. Lim, M.E. Glasner, S. Yekta, C.B. Burge, D.P. Bartel, Vertebrate microRNA genes, Science 299 (2003) 1540. [40] E. Berezikov, F. Thuemmler, L.W. van Laake, I. Kondova, R. Bontrop, E. Cuppen, R.H. Plasterk, Diversity of microRNAs in human and chimpanzee brain, Nat. Genet. 38 (2006) 1375e1377. [41] S. Griffiths-Jones, R.J. Grocock, S. van Dongen, A. Bateman, A.J. Enright, miRBase: microRNA sequences, targets and gene nomenclature, Nucleic Acids Res. 34 (2006) D140eD144.

S. Jaubert et al. / Biochimie 89 (2007) 1211e1220 [42] E.C. Lai, Micro RNAs are complementary to 30 UTR sequence motifs that mediate negative post-transcriptional regulation, Nat. Genet. 30 (2002) 363e364. [43] E.C. Lai, B. Tam, G.M. Rubin, Pervasive regulation of Drosophila Notch target genes by GY-box-, Brd-box-, and K-box-class microRNAs, Genes Dev. 19 (2005) 1067e1080. [44] E.C. Lai, Predicting and validating microRNA targets, Genome Biol. 5 (2004) 115. [45] A. Stark, J. Brennecke, R.B. Russell, S.M. Cohen, identification of Drosophila MicroRNA targets, PLoS Biol. 1 (2003) E60. [46] M. Rehmsmeier, P. Steffen, M. Hochsmann, R. Giegerich, Fast and effective prediction of microRNA/target duplexes, RNA 10 (2004) 1507e1517. [47] A.J. Enright, B. John, U. Gaul, T. Tuschl, C. Sander, D.S. Marks, MicroRNA targets in Drosophila, Genome Biol. 5 (2003) R1. [48] N. Rajewsky, N.D. Socci, Computational identification of microRNA targets, Dev. Biol. 267 (2004) 529e535. [49] D. Grun, Y.L. Wang, D. Langenberger, K.C. Gunsalus, N. Rajewsky, microRNA target predictions across seven Drosophila species and comparison to mammalian targets, PLoS Comput. Biol. 1 (2005) e13. [50] A. Krek, D. Grun, M.N. Poy, R. Wolf, L. Rosenberg, E.J. Epstein, P. MacMenamin, I. da Piedade, K.C. Gunsalus, M. Stoffel, N. Rajewsky, Combinatorial microRNA target predictions, Nat. Genet. 37 (2005) 495e500. [51] N. Rajewsky, microRNA target predictions in animals, Nat. Genet. 38 (2006) S8eS13. [52] K.C. Miranda, T. Huynh, Y. Tay, Y.S. Ang, W.L. Tam, A.M. Thomson, B. Lim, I. Rigoutsos, A pattern-based method for the identification of MicroRNA binding sites and their corresponding heteroduplexes, Cell 126 (2006) 1203e1217. [53] J. Brennecke, A. Stark, R.B. Russell, S.M. Cohen, Principles of microRNA-target recognition, PLoS Biol. 3 (2005) e85. [54] B.P. Lewis, C.B. Burge, D.P. Bartel, Conserved seed pairing, often flanked by adenosines, indicates that thousands of human genes are microRNA targets, Cell 120 (2005) 15e20. [55] H. Robins, Y. Li, R.W. Padgett, Incorporating structure to predict microRNA targets, Proc. Natl. Acad. Sci. USA 11 (2005) 4006e4009. [56] K. Nakahara, K. Kim, C. Sciulli, S.R. Dowd, J.S. Minden, R.W. Carthew, Targets of microRNA regulation in the Drosophila oocyte proteome, Proc. Natl. Acad. Sci. USA 102 (2005) 12023e12028. [57] J. Rehwinkel, P. Natalin, A. Stark, J. Brennecke, S.M. Cohen, E. Izaurralde, Genome-wide analysis of mRNAs regulated by Drosha and Argonaute proteins in Drosophila melanogaster, Mol. Cell. Biol. 26 (2006) 2965e2975. [58] A. Stark, J. Brennecke, N. Bushati, R.B. Russell, S.M. Cohen, Animal MicroRNAs confer robustness to gene expression and have a significant impact on 30 UTR evolution, Cell 123 (2005) 1133e1146. [59] S.D. Hatfield, H.R. Shcherbata, K.A. Fischer, K. Nakahara, R.W. Carthew, H.J. Ruohola-Baker, Stem cell division is regulated by the microRNA pathway, Nature 435 (2005) 974e978. [60] Z. Jin, T. Xie, Dcr-1 maintains Drosophila ovarian stem cells, Curr Biol. 20 (2007) 539e544. [61] J.K. Park, X. Liu, T.J. Strauss, D.M. McKearin, Q. Liu, The miRNA pathway intrinsically controls self-renewal of Drosophila germline stem cells, Curr. Biol. 17 (2007) 533e538. [62] D.R. Hipfner, K. Weigmann, S.M. Cohen, The bantam gene regulates Drosophila growth, Genetics 161 (2002) 1527e1537. [63] B.J. Thompson, S.M. Cohen, The Hippo pathway regulates the bantam microRNA to control cell proliferation and apoptosis in Drosophila, Cell 126 (2006) 767e774. [64] R. Nolo, C.M. Morrison, C. Tao, X. Zhang, G. Halder, The bantam microRNA is a target of the hippo tumor-suppressor pathway, Curr. Biol. 16 (2006) 1895e1904. [65] B.A. Edgar, From cell structure to transcription: Hippo forges a new path, Cell 124 (2006) 267e273. [66] K. Nairz K, C. Rottig, F. Rintelen, E. Zdobnov, M. Moser, E. Hafen, Overgrowth caused by misexpression of a microRNA with dispensable wild-type function, Dev. Biol. 291 (2006) 314e324.

1219

[67] P. Xu, S.Y. Vernooy, M. Guo, B.A. Hay, The Drosophila microRNA Mir14 suppresses cell death and is required for normal fat metabolism, Curr. Biol. 13 (2003) 790e795. [68] D. Leaman, P.Y. Chen, J. Fak, A. Yalcin, M. Pearce, U. Unnerstall, D.S. Marks, C. Sander, T. Tuschl, U. Gaul, Antisense-mediated depletion reveals essential and specific functions of microRNAs in Drosophila development, Cell 121 (2005) 1097e1108. [69] J. Bilen, N. Liu, B.G. Burnett, R.N. Pittman, N.M. Bonini, MicroRNA pathways modulate polyglutamine-induced neurodegeneration, Mol. Cell 24 (2006) 157e163. [70] X. Li, R.W. Carthew, A microRNA mediates EGF receptor signaling and promotes photoreceptor differentiation in the Drosophila eye, Cell 123 (2005) 1267e1277. [71] Y. Li, F. Wang, J.A. Lee, F.B. Gao, MicroRNA-9a ensures the precise specification of sensory organ precursors in Drosophila, Genes. Dev. 20 (2006) 2793e2805. [72] L.P. Lim, N.C. Lau, P. Garrett-Engele, A. Grimson, J.M. Schelter, J. Castle, D.P. Bartel, P.S. Linsley, J.M. Johnson, Microarray analysis shows that some microRNAs downregulate large numbers of target mRNAs, Nature 433 (2005) 769e773. [73] Y. Zhao, E. Samal, D. Srivastava, Serum response factor regulates a muscle-specific microRNA that targets Hand2 during cardiogenesis, Nature 436 (2005) 214e220. [74] N.S. Sokol, V. Ambros, Mesodermally expressed Drosophila microRNA-1 is regulated by Twist and is required in muscles during larval growth, Genes Dev. 19 (2005) 2343e2354. [75] C. Kwon, Z. Han, E.N. Olson, D. Srivastava, MicroRNA1 influences cardiac differentiation in Drosophila and regulates Notch signalling, Proc. Natl. Acad. Sci. USA 102 (2005) 18986e18991. [76] S.D. Weatherbee, G. Halder, J. Kim, A. Hudson, S. Carroll, Ultrabithorax regulates genes at several levels of the wing-patterning hierarchy to shape the development of the Drosophila haltere, Genes Dev. 12 (1998) 1474e 1482. [77] A.A. Teleman, S. Maitra, S.M. Cohen, Drosophila lacking microRNA miR-278 are defective in energy homeostasis, Genes Dev. 20 (2006) 417e422. [78] L.F. Sempere, N.S. Sokol, E.B. Dubrovsky, E.M. Berger, V. Ambros, Temporal regulation of microRNA expression in Drosophila melanogaster mediated by hormonal signals and broad-Complex gene activity, Dev. Biol. 259 (2003) 9e18. [79] A.E. Pasquinelli, B.J. Reinhart, F. Slack, M.Q. Martindale, M.I. Kuroda, B. Maller, D.C. Hayward, E.E. Ball, B. Degnan, P. Muller, J. Spring, A. Srinivasan, M. Fishman, J. Finnerty, J. Corbo, M. Levine, P. Leahy, E. Davidson, G. Ruvkun, Conservation of the sequence and temporal expression of let-7 heterochronic regulatory RNA, Nature 408 (2000) 86e89. [80] G. Hutvagner, J. McLachlan, A.E. Pasquinelli, E. Balint, T. Tuschl, P.D. Zamore, A cellular function for the RNA-interference enzyme Dicer in the maturation of the let-7 small temporal RNA, Science 293 (2001) 834e838. [81] L.F. Sempere, E.B. Dubrovsky, V.A. Dubrovskaya, E.M. Berger, V. Ambros V, The expression of the let-7 small regulatory RNA is controlled by ecdysone during metamorphosis in Drosophila melanogaster, Dev. Biol. 244 (2002) 170e179. [82] A. Bashirullah, A.E. Pasquinelli, A.A. Kiger, N. Perrimon, G. Ruvkun, C.S. Thummel, Coordinate regulation of small temporal RNAs at the onset of Drosophila metamorphosis, Dev. Biol. 259 (2003) 1e8. [83] K. Chen, N. Rajewsky, The evolution of gene regulation by transcription factors and microRNAs, Nat, Rev. Genet. 8 (2007) 93e103. [84] M. Margulies, M. Egholm, W.E. Altman, S. Attiya, J.S. Bader, L.A. Bemben, J. Berka, M.S. Braverman, Y.J. Chen, Z. Chen, et al., Genome sequencing in microfabricated high-density picolitre reactors, Nature 437 (2005) 376e380. [85] K. Saito, K.M. Nishida, T. Mori, Y. Kawamura, K. Miyoshi, T. Nagami, H. Siomi, M.C. Siomi, Specific association of Piwi with rasiRNAs derived from retrotransposon and heterochromatic regions in the Drosophila genome, Genes Dev. 20 (2006) 2214e2222.

1220

S. Jaubert et al. / Biochimie 89 (2007) 1211e1220

[86] J. Brennecke, A.A. Aravin, A. Stark, M. Dus, M. Kellis, R. Sachidanandam, G.J. Hannon, Discrete small RNA-generating loci as master regulators of transposon activity in Drosophila, Cell 128 (2007) 1089e1103. [87] L.S. Gunawardane, K. Saito, K.M. Nishida, K. Miyoshi, Y. Kawamura, T. Nagami, H. Siomi, M.C. Siomi, A slicer-mediated mechanism for repeat-associated siRNA 50 end formation in Drosophila, Science 315 (2007) 1587e1590. [88] V.V. Vagin, A. Sigova, C. Li, H. Seitz, V. Gvozdev, P.D. Zamore, A distinct small RNA pathway silences selfish genetic elements in the germline, Science 313 (2006) 320e324. [89] A.A. Aravin, N.M. Naumova, A.V. Tulin, V.V. Vagin, Y.M. Rozovsky, V.A. Gvozdev, Double-stranded RNA-mediated silencing of genomic tandem repeats and transposable elements in the D. melanogaster germline, Curr. Biol. 11 (2001) 1017e1027.

[90] A.A. Aravin, M.S. Klenov, V.V. Vagin, F. Bantignies, G. Cavalli, V.A. Gvozdev, Dissection of a natural RNA silencing process in the Drosophila melanogaster germ line, Mol. Cell. Biol. 24 (2004) 6742e6750. [91] M.A. Matzke, J.A. Birchler, RNAi-mediated pathways in the nucleus, Nat. Rev. Genet. 6 (2005) 24e35. [92] K.A. Haynes, A.A. Caudy, L. Collins, S.C. Elgin, Element 1360 and RNAi components contribute to HP1-dependent silencing of a pericentric reporter, Curr. Biol. 16 (2006) 2222e2227. [93] M. Pal-Bhadra, B.A. Leibovitch, S.G. Gandhi, M. Rao, U. Bhadra, J.A. Birchler, S.C. Elgin, Heterochromatic silencing and HP1 localization in Drosophila are dependent on the RNAi machinery, Science 303 (2004) 669e672. [94] J.C. Peng, G.H. Karpen, H3K9 methylation and RNA interference regulate nucleolar organization and repeated DNA stability, Nat. Cell Biol. 9 (2007) 25e35.