Diversifying microRNA sequence and function

Nature Reviews Molecular Cell Biology | AOP, published online 26 June 2013; doi:10.1038/nrm3611

REVIEWS Diversifying microRNA sequence and function Stefan L. Ameres1 and Phillip D. Zamore2

Abstract | MicroRNAs (miRNAs) regulate the expression of most genes in animals, but we are only now beginning to understand how they are generated, assembled into functional complexes and destroyed. Various mechanisms have now been identified that regulate miRNA stability and that diversify miRNA sequences to create distinct isoforms. The production of different isoforms of individual miRNAs in specific cells and tissues may have broader implications for miRNA-mediated gene expression control. Rigorously testing the many discrepant models for how miRNAs function using quantitative biochemical measurements made in vivo and in vitro remains a major challenge for the future. Argonaute (AGO). Proteins that are guided to mRNA targets by small silencing RNAs. AGO proteins can also serve as scaffolds to bind secondary silencing factors such as the GW‑repeatcontaining protein GW182.

Primary miRNAs (pri-miRNAs). Polyadenylated, 7‑methylguanosine-capped RNA polymerase II transcripts containing a stem–loop structure that serves as a substrate for the RNase III enzyme Drosha in animals and Dicer-like 1 (DCL1) in plants. They are processed to liberate a precursor miRNA and two unstable, single-stranded by‑products.

Institute of Molecular Biotechnology of the Austrian Academy of Sciences (IMBA), Dr. Bohr-Gasse 3, 1030 Vienna, Austria. 2 Howard Hughes Medical Institute and University of Massachusetts Medical School, 55 Lake Avenue North, Worcester, Massachusetts 01605, USA. e‑mails: stefan.ameres@ imba.oeaw.ac.at; phillip. [email protected] doi:10.1038/nrm3611 Published online 26 June 2013 1

For more than a decade, miRNAs have enthralled the biological community. Almost everything about microRNAs (miRNAs) is small. At ~22 nucleotides long, they are among the shortest functional eukaryotic RNAs. They were discovered in a diminutive organism, Caenorhabditis elegans, and their physiological relevance in regulating plant and animal gene expression was underappreciated for nearly a decade1,2. Finally, miRNAs repress most of the genes they regulate by just a tiny amount. Yet, collectively miRNAs affect nearly all cellular pathways, from development to oncogenesis. The primary repository for miRNA sequences and annotations, miRBase, debuted with just 218 miRNA loci3. Since then, the application of high-throughput sequencing to miRNA discovery in >193 different plant and animal species has swelled that number to >21,264 loci that produce >25,141 mature miRNAs. The human genome alone comprises >1,500 hairpin structures that produce detectable small RNAs, although the authenticity and functional importance of many of these miRNA candidates remain to be established4. Selective pressure during evolution seems to have maintained the pairing between miRNAs and more than half of all human protein-coding genes5, suggesting that these tiny riboregulators control the expression of most human proteins6. The absence of miRNAs in fungi and the distinct sequences, precursor structures and mechanisms of production for plant and animal miRNAs suggest that the miRNA pathway evolved at least twice7. Small silencing RNAs, including miRNAs, are present in the earliest diverging extant lineage of animal life, the sponge Amphimedon queenslandica, suggesting that the emergence of miRNAs herald the dawn of multicellular

animals8. In the plant kingdom, the miRNA pathway presumably evolved before multicellularity, because the unicellular alga Chlamydomonas reinhardtii produces miRNAs that are similar to those of higher plants9. For several model animals and plants, as well as humans, we now know the majority of miRNAs and these are inferred to be functional on the basis of their evolutionary conservation. But we are only beginning to understand the divers mechanisms that contribute to miRNA production and assemble them into complexes with proteins. For example, although it is established that miRNAs direct Argonaute (AGO) proteins to repress mRNAs, the mechanism of repression remains intensely debated. Here, we review the mechanisms that diversify miRNA sequence and function. We briefly outline the pathways that generate miRNAs and regulate their production, activity and longevity. We describe the mechanisms by which miRNA sequences and functions can be altered, as well as the processes that destroy rogue and old miRNAs. Finally, we consider the evidence for different models of how miRNAs function and present a quantitative framework for evaluating these models.

Making miRNAs Most miRNAs derive from longer, intramolecularly double-stranded RNAs (dsRNAs; termed primary miRNAs (pri-miRNAs)) that are sequentially cleaved into shorter intermediates by specialized ribonuclease III (RNase III) enzymes that partner with specific dsRNA‑binding proteins 10,11 (FIG. 1) . Both transcriptional and posttranscriptional mechanisms regulate miRNA biogenesis (for a detailed discussion, see REFS 12,13). A few miRNAs are produced by alternative pathways that replace

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REVIEWS Standard pathway

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Figure 1 | MicroRNA biogenesis. In the standard microRNA (miRNA) biogenesis Nature Reviews | Molecular Cell Biology pathway, primary miRNA (pri-miRNA) transcripts are processed by Drosha in the nucleus and by Dicer in the cytoplasm. The pri-miRNA, which is transcribed by RNA polymerase II (Pol II), begins with a 7‑methylguanosine cap (m7Gppp) and ends with a 3ʹ poly(A) tail. The pri-miRNA contains a stem–loop structure that is cleaved in the nucleus by the endonuclease Drosha together with its double-stranded RNA (dsRNA)-binding protein parner DGCR8 (in mammals) or Pasha (in flies). The resulting precursor miRNA (pre-miRNA) is exported from the nucleus by exportin 5 and then further cleaved by the endonuclease Dicer together with its dsRNA-binding partner TRBP (transactivation-response RNA-binding protein; in mammals) or Loquacious (Loqs; in flies) to liberate a miRNA–miRNA* duplex. Supported by the HSC70–HSP90 chaperone machinery, this duplex is loaded into an Argonaute (AGO) protein as a dsRNA. Subsequent maturation steps expel the miRNA*, producing a mature RNA-induced silencing complex (RISC). Alternative pathways (shown on the right) typically replace individual steps of miRNA precursor processing. Pri-miRNA cropping can be replaced with nucleases from other cellular pathways, including the general RNA degradation machinery or pre-mRNA splicing factors. In such instances, the pri-miRNA is generated from a branched mirtron structure that undergoes lariat debranching. In the specific case of pre-miR‑451, the pre-miRNA escapes Dicer processing after nuclear export and is instead directly loaded into the AGO2 protein, which triggers its maturation into a single-stranded miRNA. 2ʹ OH, 2ʹ hydroxyl group; HSP, heat shock protein; ORF, open reading frame.

standard biogenesis steps with RNA processing control from pre-mRNA splicing or RNA degradation pathways (for further details, see REF. 14). miRNAs are transcribed by RNA polymerase II. RNA polymerase II (Pol II) produces long pri-miRNAs from independent genomic transcription units or from the introns of protein-coding genes15–22. For miRNAs that arise from introns, splicing is not required for their production23, and the co‑transcriptional processing of primiRNA into precursor miRNAs (pre-miRNAs) does not affect the splicing of the host pre-mRNA24,25. Drosha crops pri-miRNAs into shorter pre-miRNAs. In animals, the RNase III enzyme Drosha converts primiRNAs into pre-miRNAs, which are ~60 nucleotide stem–loop structures. Drosha excises one or more premiRNAs from the pri-miRNA26–29. Thus, pri-miRNAs can produce several pre-miRNAs. Polycistronic miRNAs allow a single promoter to drive co‑expression of multiple miRNAs and, in some cases, mediate the coordinated expression of target factors in a single pathway 10. Drosha acts as a component of a larger complex, the microprocessor. This nuclear complex seems to exist in at least two forms, a ~600 kDa complex of unknown function and a smaller heterodimer comprising Drosha and its dsRNA-binding protein, named DGCR8 in mammals and Pasha in other animals28–31. The collaboration of Pasha with Drosha defines a general paradigm in small RNA biogenesis: dsRNA-binding proteins partner with RNase III enzymes to restrict the substrates they process, increase their affinity for a substrate or improve cleavage site accuracy. The subcellular distribution of miRNA-producing enzymes and their substrates implies that animal pri-miRNAs are cropped in the nucleus, whereas pre-miRNAs are processed in the cytoplasm26,32–36. The nuclear transport receptor exportin 5 (known as Ranbp21 (accession number CG12234) in flies) recognizes the ends and the stem of the premiRNA37 and exports the pre-miRNA from the nucleus to the cytoplasm via the nuclear pore38–41. Dicing makes miRNA–miRNA* duplexes. In the cytoplasm, a second RNase III enzyme, Dicer, liberates a ~22 nucleotide miRNA–miRNA* duplex from the pre-miRNA42–45. In flies, Dicer‑1 cleaves pre-miRNAs, whereas Dicer‑2 generates siRNAs46. Like Drosha, Dicer‑1 recognizes defined RNA structures and then cleaves at a fixed distance away from the base of the pre-miRNA stem, cutting off the loop to produce ~22 nucleotide mature miRNA– miRNA* duplexes47,48. Like cropping of pri‑miRNAs, ‘dicing’ of pre-miRNAs can also be bypassed. For example, the hairpin of pre-miR-451 in zebrafish and mice is too short to be recognized by Dicer; instead, it is directly loaded into the RNA-induced silencing complex (RISC) for further processing by AGO2 into a mature miRNA49–51. Partner proteins shape Dicer function. Continuing the theme of RNase III enzymes that partner with dsRNAbinding proteins, Drosophila melanogaster Dicer‑1 partners with two isoforms of the dsRNA-binding protein

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REVIEWS Flies and mammals

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Figure 2 | Small RNA turnover by tailing and trimming. In the tailing-and-trimming pathway, the presence of a highly complementary target RNA triggers both tailing and trimming of microRNAs (miRNAs), theirBiology Natureultimately Reviews | decreasing Molecular Cell abundance in flies and mammals (left panel). The identity of the exonuclease is not known (denoted by the question mark in the figure). In flies, Argonaute 2 (AGO2)‑bound siRNAs are protected from tailing and trimming because the methyltransferase Hen1 (Hua enhancer 1) methylates siRNAs at the 2ʹ position of the 3ʹ terminal nucleotide. In the absence of Hen1, siRNAs are subjected to target RNA-directed tailing and trimming (middle panel). Unlike fly Hen1, plant HEN1 methylates both siRNA and miRNA duplexes before they are loaded into AGO. As in flies, loss of HEN1 in plants prompts small RNA tailing, which is mediated by the plant terminal nucleotidyltransferase (TNTase) HEN1 SUPPRESSOR 1 (HESO1), ultimately resulting in 3ʹ-to‑5ʹ exonucleolytic decay (right panel). The normal triggers for small RNA decay in plants are unknown. Ribonuclease III (RNAse III). Double-stranded RNA-specific endoribonuclease that generates products with two nucleotide 3ʹ overhangs, a 5ʹ phosphate and a 3ʹ hydroxyl group.

dsRNA-binding protein Proteins containing double-stranded RNA-binding domains, which are ~70 amino acid motifs that bind to RNA helices via their 2ʹ hydroxyl group and phosphate backbone.

Precursor miRNAs (pre-miRNAs). Stem–loop RNAs comprising a single-stranded loop that connects two partially complementary sequences. These sequences pair to form a predominantly double-stranded stem. Pre-miRNAs typically have a 5ʹ phosphate and a two nucleotide 3ʹ overhang, allowing them to serve as substrates for Dicer.

Drosha The nuclear RNase III endonuclease in animals that cleaves the base of a stem–loop structure contained in primary microRNAs (pri-miRNAs) to produce a precursor miRNA. Collaborates with mammalian DGCR8 (or Pasha in other animals), which is its double-stranded RNA-binding protein partner.

Loquacious (Loqs), termed Loqs‑PA and Loqs‑PB. These are required for efficient miRNA processing in flies because they increase the affinity of Dicer‑1 for premiRNAs52–55. Conversely, the Dicer‑2 partner protein R2D2 prevents Dicer‑2 from processing pre-miRNAs, restricting it to long dsRNA substrates56. Alternative partner proteins refine the substrate specificity of individual Dicer proteins, and this may explain how organisms such as worms and mammals produce miRNAs and siRNAs with only a single enzyme. Consistent with this, mammals have two Dicer partners, transactivation-response RNA-binding protein (TRBP) and protein kinase R‑activating protein (PACT)57–59. In addition to their role in defining Dicer substrate specificity, Dicer partner proteins can alter the position of Dicer-mediated cleavage within a pre-miRNA55,60. For example, TRBP, but not PACT, changes the site at which mammalian Dicer cleaves a few pre-miRNAs, such as pre-miR‑132. In flies, Loqs‑PB changes the length and seed sequence of the miRNAs generated by Dicer‑1 from a small subset of pre-miRNAs. A seed for target recognition. After the sequential processing of miRNA precursors, one of the two strands of the miRNA duplex guides AGO proteins to complementary mRNA sequences to repress their expression (see REFS 61,62 for detailed reviews of AGO loading). The major determinant for AGO binding to its target mRNA is a 6–8 nucleotide domain at the 5ʹ end of the miRNA. AGO associates with this region to create the ‘seed’ (for details, see REF. 63). Sequences that are complementary to the seed (‘seed matches’) suffice to trigger a modest

but detectable decrease in the expression of an mRNA. Seed matches can occur in any region of an mRNA but are more likely to decrease mRNA expression when they are in the 3ʹ untranslated region (3ʹ UTR)64–67. Because the region used to create the seed is so short, more than half of all protein-coding genes in mammals are regulated by miRNAs, and thousands of other mRNAs seem to have experienced negative selection to avoid seed matches with miRNAs that are present in the same cells5,68–70.

Target RNAs alter miRNA stability Binding of the AGO–miRNA complex to an mRNA not only regulates gene expression (discussed in detail below) but also alters the stability of the miRNA itself. For example, binding of complementary target RNAs in flies and mammals can direct miRNA destruction. miRNA decay elicited by target binding was first reported for antagomirs, which are chemically modified, synthetic oligonucleotides used to inhibit miRNA function in cells and in vivo71. We now know that antagomirs, as well as highly expressed RNAs containing sites with extensive complementarity to a miRNA, trigger miRNA degradation (FIG. 2). This pathway involves both the addition of adenosine or uracil to the miRNA (‘tailing’) and the 3ʹ-to‑5ʹ exonucleolytic resection of the miRNA 3ʹ end (‘trimming’)72–74. Target RNA-directed tailing and trimming of miRNAs require greater complementarity than is typically found between miRNAs and their targets72,74. This may be because the structural rearrangements associated with target binding provide the tailing and trimming enzymes access to the miRNA 3ʹ end, which would otherwise reside in the AGO PAZ domain75–79. Alternatively, all miRNAs may be susceptible to tailing



REVIEWS Dicer An RNase III endonuclease that is predominantly found in the cytoplasm of animal cells and the nucleus of plant and some fungal cells. Dicer proteins liberate mature microRNA– microRNA* duplexes from pre-microRNAs and siRNA duplexes from long double-stranded RNAs.

siRNAs The ~21 nucleotide small RNAs that mediate RNA interference in plants, animals and some fungi. Typically produced by Dicer processing of long double-stranded RNA (dsRNA) precursors encoded in the genome (endo-siRNAs) or from exogenous dsRNA sources (exo-siRNAs) such as viruses.

RNA-induced silencing complex (RISC). A ribonucleoprotein complex that consists of a small RNA guide strand bound to an Argonaute protein. RISC mediates all RNA silencing pathways, and it can also include auxiliary proteins that extend or modify its function.

Seed sequence A nucleotide motif in the 5ʹ domain of all small silencing RNAs, which is organized by Argonaute to determine target-RNA recognition.

PIWI-interacting RNAs (piRNAs). Small silencing RNAs, 25–35 nucleotides long, that bind PIWI clade Argonaute proteins in animals and silence germline transposons. They are thought to derive from single-stranded RNA precursors and do not require RNase III enzymes for their maturation.

Exoribonuclease Enzymes that successively remove nucleotides from either the 3ʹ (3ʹ-to‑5ʹ exoribonuclease) or the 5ʹ end (5ʹ-to‑3ʹ exoribonuclease) of RNA. They catalyse phosphodiester bond cleavage using water (releasing nucleotide monophosphates) or inorganic phosphate (releasing nucleotide diphosphates) as a nucleophile.

Isomirs microRNA variants containing sequences that deviate from miRBase-annotated or most frequently observed species.

and trimming when bound to target RNAs, but the slower dissociation of miRNAs from high-affinity sites makes highly complementary RNAs more efficient at triggering modification and degradation of miRNAs. The endogenous functions of target RNA-directed tailing and trimming of miRNAs in animals remain elusive, and the proteins that mediate this pathway are unknown. However, tailing and trimming can be triggered by transcribed miRNA inhibitors, and use of these inhibitors in adult mice phenocopies miRNA loss‑of‑function mutations, suggesting that this pathway is functionally relevant for miRNA-mediated regulation of gene expression in vivo74,80,81. Moreover, some viruses use a similar strategy to interfere with host miRNA function during infection (for details, see REFS 82,83). To what extent target RNA-directed miRNA degradation contributes to the regulation of miRNA function in vivo remains to be determined. But it is tempting to speculate that the tailing-and-trimming pathway has contributed to the evolution of target sites in mRNAs in animals, as mRNAs rarely possess sufficient complementarity to miRNAs to trigger miRNA destruction68,69. In D. melanogaster, target RNA-directed small RNA decay is restricted to the miRNA pathway. siRNAs in flies and PIWI-interacting RNAs (piRNAs) in all animals are protected from tailing and trimming by 2ʹ-O‑methylation of their 3ʹ ends, a modification that is introduced during AGO loading by the methyltransferase HEN1 (Hua enhancer 1)72,84–86. Flies use these protected siRNAs to target AGO2–RISC complexes for the repression of extensively complementary targets. In the absence of Hen1, these targets promote the tailing, trimming and depletion of complementary endo genous siRNAs86. HEN1 was first discovered in plants, in which it methylates both miRNAs and siRNAs87,88. Loss of HEN1 in plants provokes uridylation and 3ʹ-to‑5ʹ exonucleolytic decay of miRNAs and siRNAs88–91. Plant and animal HEN1 enzymes catalyse the same chemical reaction92, but plant small RNAs are methylated as duplexes before their loading into AGO88,92,93, whereas animals methylate only a subset of single-stranded small RNAs, typically piRNAs. In flies, and presumably in arthropods more generally, HEN1 methylates siRNAs as the last step in AGO2–RISC assembly 85, but mammalian siRNAs are thought not to be HEN1 substrates. Tailing and trimming distinguish endogenous from foreign small silencing RNAs. In flies, highly complementary target RNAs have not been found for miRNAs bound to AGO1 (the miRNA-binding AGO protein in flies). Animal miRNAs generally lack a protective 2ʹ-O‑methyl modification (FIG. 2). Introduction of RNAs with extensive complementarity to one of these miRNAs triggers tailing and trimming of the miRNA. Thus, in flies, Hen1 and the tailing-and-trimming pathway might collaborate to distinguish self from non-self small RNAs. For example, anti-viral siRNAs bound to AGO2 (the siRNAbinding AGO protein in flies) bind and destroy viral RNA without risk of tailing and trimming, because they have a 2ʹ-O‑methyl-modified 3ʹ end. During viral infection, non-self-directed siRNAs could be so abundant

that they inappropriately accumulate in AGO1 complexes. As a consequence, they would not acquire a protective 2ʹ-O‑methyl modification. Their binding to fully complementary viral RNAs will instead mark them as foreign sequences that have inappropriately entered the miRNA (self) pathway. The tailing and trimming pathway would then initiate their elimination. Target RNAs can stabilize miRNAs. In C. elegans, the 5ʹ-to‑3ʹ exoribonuclease XRN‑2 limits miRNA abundance. However, miRNAs in RISC are protected from XRN‑2 when bound to a partially complementary target RNA94. Similarly, persistence of siRNAs in cell culture is enhanced by the presence of a fully complementary target mRNA95. So, perhaps target binding recruits miRNA protective factors or translocates miRNAs to a subcellular location that lacks small RNA-degrading nucleases such as XRN‑2. Alternatively, target RNAs may protect miRNAs from a release factor that would expel them from RISC and expose them to subsequent destruction94. Supporting this idea, a partially complementary target RNA also protects worm miRNAs from destruction by XRN‑1, another 5ʹ-to‑3ʹ exonuclease96. XRN1 may also degrade miRNAs in animals97.

Diverse strategies to generate isomirs Although miRBase catalogues miRNAs as single sequences, the availability of small RNA sequence data from various organisms, tissues and cell types shows that miRNAs comprise multiple isoforms (also known as isomirs ) 4,98–104. Such sequence heterogeneity may arise from imprecise precursor cropping or dicing, terminal trimming or the addition of non-templated nucleotides98,103,104 (FIG. 3). Trimming produces 5ʹ miRNA isoforms. Because the 5ʹ end of a miRNA defines its seed sequence, a single nucleotide shift at this site will radically alter its target repertoire. The 5ʹ ends of miRNAs are further constrained because AGO loading typically selects for miRNAs with distinct 5ʹ nucleotides99,102,105–107. Changes in the 5ʹ end of a miRNA can therefore alter the efficiency of AGO loading. miRNA 5ʹ isoforms comprise less than 10% of all miRNA reads in humans, flies and worms4,99,108. Nonetheless, some miRNAs produce two or more 5ʹ isomirs that repress distinct target mRNAs4,55. One of the most striking examples is D. melanogaster miR‑210, which exists as two nearly equally abundant 5ʹ isoforms98. miRNA 5ʹ isoforms can also arise from differential processing of paralogous pre-miRNAs, each of which produces a single miRNA sequence, such as D. melanogaster pre-miR‑2‑1 and pre-miR‑2‑2 (REF. 99) (FIG.3a). Other paralogous hairpins, such as human premiR‑133a‑1 and pre-miR‑133a‑2, generate two predominant isoforms 4. Some extreme examples of 5ʹ heterogeneity were discovered in mammals and chickens for a set of mirtrons that contain a long 5ʹ extension connecting the hairpin to the 5ʹ splice site4,109,110. After splicing, the 5ʹ extension is trimmed ‘sloppily’ to produce several mature miRNAs.



REVIEWS a 5′ miRNA isoforms miR-2-1

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Figure 3 | Mechanisms and consequences of microRNA isoforms. Precursor microRNAs (pre-miRNAs) and mature Nature Reviews | Molecular Cell Biology miRNAs can be modified by the addition, editing or subtraction of nucleotides, resulting in distinct miRNA isoforms. Such isomirs can be classified as 5ʹ, internal and 3ʹ isoforms. a | 5ʹ miRNA isoforms can be produced by differential processing of paralogous miRNAs, as exemplified by the miR‑2 family in Drosophila melanogaster or the exonucleolytic processing of 5ʹ tailed mirtrons. This results in miRNAs with different seed sequences, which are the primary sequence determinants for target RNA binding. Consequently, different 5ʹ miRNA isoforms regulate different sets of target RNAs. The identity of the exonuclease that targets 5ʹ tailed mirtrons is not known (indicated by the question mark). b | Internal miRNA isoforms are rare and almost exclusively generated by editing of adenosine to inosine by the enzyme ADAR (adenosine deaminase acting on RNA). c | The 3ʹ end of a miRNA shows the greatest heterogeneity, which is a result of both exonucleolytic trimming and the addition of nucleotides (typically adenine or uridine) to the 3ʹ end (tailing). Exonucleolytic trimming is required for the efficient processing of 3ʹ tailed mirtrons, as well as ~25% of all miRNAs in flies. Dicer-1 initially makes these miRNAs as ~24 nucleotide miRNA duplexes. Upon loading into Argonaute 1 (AGO1), such ‘long’ miRNAs are trimmed by the 3ʹ-to‑5ʹ exoribonuclease Nibbler to produce mature ~22 nt miRNAs. In mammals, monouridylation of selected pre-miRNAs by ZCCHC11, GLD-2 and/or ZCCHC6 enhances substrate recognition and processing by Dicer. In the case of miRNAs derived from the 3ʹ arm of such pre-miRNAs, monouridylation is propagated into the mature miRNAs, contributing to its 3ʹ heterogeneity. In the presence of the pre-miRNA-binding protein LIN28, ZCCHC11 is a processive enzyme, oligouridylating pre-miRNAs that contain a LIN28‑binding motif. Unlike monouridylation, oligouridylation inhibits Dicer-mediated processing and enhances pre-miRNA decay by the 3ʹ-to‑5ʹ exoribonuclease DIS3L2. Loqs, Loquacious; TRBP, transactivation-response RNA-binding protein.

Mirtrons Intron-derived precursor microRNAs excised from primary miRNAs by the splicing machinery and a lariat-debranching enzyme instead of Drosha.

Internal miRNA isoforms. Modifications that change the internal sequence of miRNAs are rare and occur at frequencies that are similar to sequencing errors4,98,111. However, for a few individual miRNA species, as much as 80% of all sequence reads for the miRNA contain at least one base change from the corresponding genomic sequence4,98,112–114. Such editing events nearly always

correspond to deamination of adenosine to inosine by ADAR (adenosine deaminase acting on RNA) 115 (FIG. 3b). ADAR enzymes also edit long dsRNAs, promoting their unwinding and preventing them from triggering RNAi116–118, as well as local double-stranded structures. Indeed, these local structures may have evolved to facilitate ADAR-mediated repair of mRNAs



REVIEWS that lack the appropriate protein-coding sequence. Because ADAR enzymes are preferentially expressed in neural tissues of flies and mammals, miRNAs in the brain are often edited. The biological significance of miRNA editing outside the seed sequence, however, remains to be established. Trimming can produce 3ʹ miRNA isoforms. miRNA isoforms often differ at their 3ʹ ends99,100,102,103,108. miRNAs with a common 5ʹ end but different 3ʹ ends are presumed to target the same mRNAs, but they may differ in the extent of target repression or they might have different half-lifes. Recent studies implicate 3ʹ-to‑5ʹ trimming as one major source of miRNA 3ʹ heterogeneity. In D. melanogaster, ~40% of AGO1‑bound miRNAs are trimmed at their 3ʹ termini, irrespective of whether they are produced via the canonical or the mirtron pathway 104. In extreme cases, such as miR‑34 or miR‑317, four to six variant isoforms accumulate that differ at their 3ʹ ends104,119,120. This 3ʹ trimming requires the 3ʹ-to‑5ʹ exoribonuclease Nibbler 119,120 (FIG. 3c). Nibbler mediates trimming of more than a quarter of all fly miRNAs. It acts predominantly on miRNAs that arise from pre-miRNAs containing bulges or internal loops that force Dicer or Drosha to generate an atypically long product. After the miRNA is loaded into AGO1, Nibbler trims these long miRNAs into shorter isoforms, perhaps to ensure that their 3ʹ ends nestle in the AGO1 PAZ domain. Human miRNAs also seem to be trimmed by a Nibbler-like enzyme120. The C. elegans homologue of Nibbler, MUT‑7, is required for transposon silencing, RNAi and co-suppression121–124. But a role for MUT‑7 in the miRNA pathway has not been reported.

Terminal nucleotidyl transferases (TNTases). Templateindependent polymerases that add nucleotides to the 3ʹ ends of nucleic acids.

Tailing: a special form of 3ʹ heterogeneity. The addition of non-templated nucleotides by terminal nucleotidyl transferases (TNTases) (tailing) gives rise to a special form of 3ʹ heterogeneity in miRNAs and other types of RNA. All TNTases are members of the DNA polymerase‑β superfamily that includes poly(A) polymerase, which adds the poly(A) tail to mRNAs. TNTases typically add adenosine or uridine in vivo125. The consequences of tailing can vary. In bacteria, the poly(A) tail triggers mRNA decay, whereas in eukaryotes it enhances mRNA stability and translation126. Moreover, uridylation has been implicated in the maturation and turnover of coding and non‑coding RNAs127. Several studies have recently implicated TNTases in the regulation of miRNAs, through uridylation of their 3ʹ ends (FIG. 3c). These include ZCCHC6 (also known as TUT7) and ZCCHC11 (also known as TUT4), which are both zinc-finger- and CCHC domain-containing proteins, and GLD2 (also known as TUT2 and PAPD4), which is a poly(A) polymerase domain-containingprotein. For example, the TNTase ZCCHC11 oligo uridylates pre-let‑7 in mouse embryonic stem cells128,129. Oligouridylation requires the formation of a ternary complex, consisting of ZCCHC11, pre-let‑7 and the pluripotency factor LIN28, which directly binds to a sequence motif in the loop of pre-let‑7 (REFS 130,131). Tailing masks the two nucleotide 3ʹ overhang of the

pre-miRNA and prevents efficient substrate recognition by Dicer, whereas the oligo(U) tail serves as a decay signal for the Perlman syndrome exonuclease DIS3L2 (REF. 132). Surprisingly, in the absence of LIN28, ZCCHC11 and its close relative ZCCHC6 monouridylate a selected class of pre-miRNAs, including several members of the pre-let‑7 family (class II pre-miRNAs). Monouridylation enhances dicing, because it restores the two nucleotide 3ʹ overhang of pre-miRNAs that were imprecisely cropped by Drosha133. Mature miRNAs can also serve as substrates for TNTases, but the triggers and biological consequence of this type of modification are less well-established. For example, the pre-miRNA tailing enzyme ZCCHC11 apparently also adds uridine to mature mammalian miRNAs such as miR‑26, and this somehow facilitates cytokine expression134. GLD2 adds one adenosine to miR‑122, which is the most abundant miRNA in the liver 135. miR‑122 derives from the 5ʹ arm of pre-miR‑122, so GLD2 must act on the mature miRNA after dicing. In mice lacking GLD2, both miR‑122 adenylation and miR‑122 abundance decrease, suggesting that adenylation enhances miR‑122 stability. GLD2 adds tails to several other miRNAs in mammals, although this does not always increase their stability 136. A protective function for miRNA adenylation has also been reported in plants137. In the absence of the methyltransferase HEN1, uridylation of mature plant miRNAs by the TNTase HEN1 SUPPRESSOR 1 (HESO1) triggers miRNA decay 88,89,138,139. Loss of HESO1 partially restores small RNA function and suppresses developmental phenotypes in HEN1 mutant plants138,139 (FIG. 2). Because HEN1 methylates miRNAs before they bind AGO proteins, the substrates for tailing are inferred to be duplexes. In the green alga C. reinhardtii, loss of the TNTase MUT68 reduces small RNA uridylation and increases miRNA and siRNA abundance140. In vitro, MUT68 tails synthetic small RNAs, and this promotes their degradation by the catalytic exosome subunit RRP6 (ribo somal RNA-processing protein 6). Like loss of MUT68, loss of RRP6 stabilizes small RNAs in vivo. MUT68 is also implicated in the degradation of the 5ʹ target RNA fragments produced by siRNA- or miRNA-directed RISC cleavage141, which have previously been reported to serve as substrates for tailing in Arabidopsis thaliana and mammals142. Perhaps enzymes that can tail either miRNAs or their targets are both recruited when AGO proteins bind an RNA.

AGO creates the seed AGO proteins create a seed in any small RNA sequence they bind. Mutations in the seed block target cleavage by plant miRNAs in extracts143 and in vivo144. In addition, such mutations relieve the repression of highly complementary mRNAs by miRNAs and siRNAs in animals145,146, almost certainly because a seed match is required for high-affinity binding between a miRNA and its target 147. Structural analyses of archael, bacterial, yeast and human AGO proteins reveal how AGO creates



REVIEWS the seed79,148–154. Upon AGO binding, the seed bases are exposed to the solvent and pre-ordered in a quasihelical structure that ‘pre-pays’ the entropic penalty that comes with nucleic acid pairing 151. Structural and kinetic data suggest that the seed nucleates binding to complementary sequences in the target RNA (the seed match) and that pairing of the target with sequences 3ʹ to the seed can only occur subsequently, after a structural rearrangement of the AGO protein79,147,150,152,155. This nucleation theory was recently modified after the discovery that a set of miR‑124 targets in the mouse brain seem to lack full seed complementarity. Instead, they contain one ‘bulged’ nucleotide across from positions 5 and 6 of the miRNA seed156 (FIG. 4). In these cases, an initial seed–target interaction across five nucleotides has been proposed to rearrange into a six nucleotide helix containing one additional unpaired (that is, flipped out or bulged) target base. Such sites constitute more than 15% of all AGO-bound sequences in the mouse brain, are evolutionary conserved and mediate target mRNA repression156. These studies are consistent with in vitro analyses of AGO in the eubacterium Thermus thermophilus, which tolerates a flipped out base in the seed-matching sequences of its target 150, as well as thermodynamic studies on the recognition of let‑7 targets in worms157. Pairing between a target and miRNA bases that are 3ʹ to the seed — that is, the ‘supplementary’ base pairs from miRNA position 13 to 16 — can decrease the off rate (koff) of RISC from a target RNA, which may explain why such additional pairing can enhance target repression64,147 (FIG. 4e). Moreover, features other than base pairing can have a substantial effect on target regulation and will contribute to the quality of a miRNA-binding site. These include: an adenosine across from the first nucleotide of the miRNA70,158; the accessibility of a target site64,159–161; and the position of a target site within a 3ʹ UTR64. mRNAbinding proteins add an additional layer of complexity, as they can compete with and perhaps even enhance miRNA binding 162–169. Alternative modes of miRNA target recognition have been reported, but these seem to have a modest impact on gene regulation: fewer than 10% of all target sites are predicted to include base pairs that compensate for an incomplete seed match, and even fewer sites lack seed matches altogether 63,170 (FIG. 4e). miRNAs can also contain sequence motifs that regulate their stability or intracellular localization (BOX 1). Nonetheless, the association of a seed sequence with the target mRNA remains the predominant mechanism for target recognition.

The great miRNA function debate Despite a plethora of high-quality studies examining the biochemistry, biology and genomics of miRNA-directed mRNA regulation, how miRNAs repress or activate gene expression in animals remains controversial. There is general agreement that miRNAs regulate gene expression post-transcriptionally, but several mechanisms have been proposed to explain how, ranging from repression of translational initiation or elongation to acceleration of general mRNA decay processes (FIG. 4c,d).

How do miRNAs regulate mRNA expression? Some AGO proteins cleave highly complementary target RNAs, a process that is catalysed by their RNase H‑like PIWI domain, which positions a pair of Mg 2+ ions at the scissile phosphate. This endonucleolytic mechanism derives from the ancestral RNAi pathway, in which target ‘slicing’ provides an anti-viral and anti-transposon defence. In plants, highly complementary miRNA target sites trigger mRNA cleavage143,171–176, but at least some miRNAs can block translation177,178 (FIG. 4). By contrast, just a few miRNA–target pairs in mammals have sufficient complementarity to direct AGO2 to cleave the target 170,179–181. Even for these atypical examples, the contribution of miRNA-directed target cleavage to the overall decrease in target mRNA abundance is unknown. Moreover, three of the four human AGO clade proteins are catalytically inactive; only a miRNA bound to AGO2 can cleave a highly complementary target RNA. This suggests that AGO proteins more often mediate target gene expression control through a mechanism that is independent of RNA endonucleolytic cleavage. The overwhelming majority of miRNAs recognize their targets solely through their seed sequence, and such partial complementarity does not permit target cleavage even when the miRNA is bound to AGO2. Nonetheless, animal miRNAs typically trigger destruction of the mRNAs they bind182–190. Quantitative high-throughput sequencing and proteomics methods for measuring mRNA and protein abundance as well as new methods that measure ribosome occupancy of mRNAs have only fuelled the debate on how miRNAs regulate their targets. Ribosome profiling experiments in mice suggest that mRNA destruction by a process distinct from endonucleolytic cleavage, rather than translational repression, explains the overwhelming majority (>84%) of miRNA-directed repression in mammals188. Ribosome profiling quantitatively determines the positions of ribosomes on cellular mRNAs with codonlevel resolution for the entire transcriptome191. Such data have reinforced the view that most of the decrease in steady-state protein abundance that is observed during miRNA-mediated repression reflects mRNA decay 188,189,192. Nonetheless, a small but significant fraction (11–16%) of miRNA repression seems to derive from a block in translation190. Although mRNA destruction is necessarily mutually exclusive with translational repression, studies in zebrafish suggest that translational repression can precede mRNA degradation. Zebrafish miR‑430 is the predominant miRNA expressed at the onset of zygotic transcription. miR‑430 clears maternal mRNAs from the embryo, a process that is prevented in mutants lacking both maternal and zygotic Dicer 185,193. Ribosome profiling of wildtype and Dicer mutant fish shows that miR‑430 initially reduces ribosome occupancy of target mRNAs — which represses their translation — and then, two hours later, triggers mRNA destruction193. The initial decrease primarily reflects a change in the rate of translational initiation, a mechanism of miRNA action previously observed for fly AGO1 in vitro 194–196 and in cell extracts197,198.



REVIEWS a Endonucleolytic cleavage

b

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Figure 4 | microRNA function in plants and animals. a | Most plant microRNAs (miRNAs) and a few animal miRNAs direct endonucleolytic cleavage (slicing) of their mRNA targets. The 5ʹ‑to‑3ʹ exoribonuclease XRN4 in plants and Nature Reviews | Molecular CellXRN1 Biology in animals, together with the major cellular 3ʹ‑to‑5ʹ exonucleolytic complex, the exosome, subsequently degrade the sliced mRNA fragments. The 3ʹ end of the 5ʹ cleavage product is frequently uridylated, perhaps to mark these fragments for decay. b | miRNA-directed endonucleolytic cleavage of mRNAs requires extensive complementarity between the miRNA and its target site (representative examples in plants and mammals are shown). Endonucleolytic cleavage occurs at the phosphodiester bonds across from nucleotides 10 and 11 of the miRNA, counted from the miRNA 5ʹ end. Although slicing seems to be the dominant mode of action for miRNAs in plants, highly complementary sites in the transcriptome of animals are rare. c | In animals, miRNAs were originally proposed to repress translation of an open reading frame (ORF). Biochemical studies have suggested that miRNAs have a role in blocking translational initiation, in poly(A) tail shortening or in the recruitment of protein cofactors that can interfere with translation. d | In many cells and tissues, miRNA-directed translational repression is indistinguishable from mRNA destruction via decapping and 5ʹ‑to‑3ʹ decay. This has led to the suggestion that miRNAs directly target mRNAs for decay. Another possibility is that the inhibition of translation by miRNAs (part c) triggers subsequent mRNA decay, and the temporal delay between these two effects can vary depending on the surveillance mechanisms in place in particular cellular contexts (depicted by blue arrow on the right). e | Like animal miRNAs, plant miRNAs may regulate target mRNAs via mechanisms other than endonucleolytic cleavage. But the underlying miRNA–target RNA interaction (for example, miR‑172–SCL6) is indistinguishable from sites that trigger cleavage (part b). In contrast to plants, animal miRNAs are almost always imperfectly complementary to the target mRNAs they regulate. The seed sequence (miRNA nucleotides 2 to 8; sometimes interrupted by a specific G‑bulge) is the major determinant for target binding and often suffices to trigger mRNA repression. Additional pairing of miRNA nucleotides 12 to 16 can sometimes bolster seed binding (3ʹ supplementary sites). Only in rare cases can an imperfectly matching seed sequence be compensated for by extensive complementarity between the miRNA 3ʹ region and the target site (3ʹ compensatory sites). Binding of animal miRNAs to partially complementary target sites generally results in translational inhibition and/or mRNA decay via the mechanisms shown in part c and part d. HoxB8, homeobox B8; UTR, untranslated region.



REVIEWS Box 1 | microRNA sequence motifs Sequence motifs within a microRNA (miRNA) can direct its localization to particular sites in a cell or enhance its turnover. For example, a hexamer motif in miR‑29b triggers its nuclear import241. Nuclear miR‑29b may regulate gene expression in the nucleus or, alternatively, the nuclear import of miR‑29b may alter target gene expression in the cytosol by sequestering the miRNA and effectively decreasing its concentration in the cytoplasm. This hexamer motif is necessary and sufficient for nuclear import, as insertion of this sequence into another miRNA directs its nuclear import. Several sequence motifs have been linked to miRNA turnover. In the absence of such motifs, miRNAs are long-lived, with half-lives as long as 5 days242 or even 2 weeks243. For example, a seven nucleotide motif at the 3ʹ end of miR‑382 triggers its decay in HEK293 cells97. Furthermore, destruction of miR‑16 family miRNAs upon re‑entry into the cell cycle after serum starvation relies on both the seed sequence and a sequence at the 3ʹ end of the miRNAs244. Uridylation of transiently transfected miR‑29b and miR‑29c at positions 9–11 elicits their rapid turnover in HeLa cells, a phenomenon that seems to be conserved in the fly miRNA bantam245. For miR‑29b and miR‑29c, the miRNA–miRNA* duplex seems to be targeted for degradation rather than the mature Argonaute (AGO)-bound miRNA. The proteins that recognize the sequence motifs controlling miRNA localization or stability and whether such recognition occurs while the miRNA is still bound to AGO are not yet known.

The timing of mRNA poly(A) tail shortening, which was previously proposed to explain miR‑430‑directed translational repression in the same model system185, is similar to the effects on mRNA decay but not to the initial decrease in translational initiation193. Together with previous biochemical studies198–201 and more recent kinetic analysis in a D. melanogaster cell line202, these studies point to a testable, unifying hypothesis: miRNAs initially repress mRNA translation and later trigger mRNA decay, perhaps in response to the block to translation rather than any specific property of miRNAs or AGO proteins. The timing of decay might vary depending on the context: in cells with robust surveillance mechanisms, mRNAs bound by miRNAs would be rapidly degraded, whereas in early embryos, translationally repressed mRNAs would be stable. Mutations in the mRNA surveillance or degradation machinery should prove useful in testing these ideas. We note that all forms of this ‘translational-repressiontriggers-mRNA-decay’ hypothesis imply biochemical coupling between the rate of translation and the stability of an mRNA. Proof of such coupling will require rigorous, quantitative in vivo measurements of the rates of translation and decay for mRNAs targeted by miRNAs in the presence and absence of miRNA function. miRNAs turn me on? miRNAs generally repress gene expression, but they have also been linked to gene activation. Often, the mechanism of activation is indirect, with repression of a repressor leading to increased expression of specific transcripts. One such case occurs during neuronal differentiation in frogs, chickens and mammals203, in which miR‑128 in the brain targets components of the nonsense-mediated decay machinery and thereby increases the abundance of mRNAs involved in neuronal differentiation and brain function. A second example may be the activation of trans cription by miR‑373 (REF. 204) . In other contexts, repression of non-coding transcripts that cross the promoters of overlapping genes have been reported to increase the abundance of the sense gene transcript205,206.

Although the precise mechanism by which overlapping non-coding transcripts decrease gene expression remains unknown, miR‑373 may similarly destabilize or block the function of non-coding transcripts that cross the promoter regions of protein-coding genes. The requirement for miR‑122 in hepatitis C virus replication is a particularly intriguing example of activation by a miRNA: binding of miR‑122 to complementary sites in the 5ʹ end of the virus genome is required for efficient viral replication and enhances internal ribosome entry (IRES) site-directed translation207. The mechanism of this activation is not known, nor is it clear whether miRNAs have similar effects on viral gene expression as they do on mammalian mRNAs. Nonetheless, the finding that a readily inhibited host factor is required for viral replication makes miR‑122 one of the most promising drug targets for treating hepatitis C208,209. A more unusual and direct form of miRNA activation of gene expression has been described following cell cycle arrest. Upon serum starvation, miR‑369‑3 binds an AU‑rich element in the 3ʹ UTR of the gene encoding tumour necrosis factor (TNF). miR‑369‑3 normally directs repression but, in these specific conditions, it seems to promote translation210,211. A similar phenomenon has also been observed for the translational activation of ribosomal protein genes, although miR‑10a was proposed to bind non-seed-matching sites in the 5ʹ UTR of mRNAs212. Apostasy or prophecy? Not only are the mechanisms by which miRNAs repress gene expression debated but also who represses whom. The crux of the issue is the seemingly modest effect that miRNAs have on the vast majority of mRNAs they target. A generally accepted hypothesis posits that many miRNAs ‘tune’ the expression of most targets, with only a small number of targets experiencing a large change in mRNA or protein abundance63. The biological importance of such tuning is thought to emerge in specific environmental conditions that are not typically encountered by animals reared in a laboratory. Supporting this view, the role of miR‑7 in ensuring correct patterning of the D. melanogaster eye is only apparent in sensitized genetic backgrounds or when the fly experiences ‘wild swings’ in temperature during eye development 213. Similarly, the loss of individual miRNAs or miRNA families in C. elegans rarely produces an obvious phenotypic defect214. An alternative view proposes that most ‘tuning targets’ are actually ‘titrating targets’ that moderate the effective concentration of the miRNA available to bind a smaller set of highly regulated targets215. The concept of titrating sites is supported by the finding that artificial, long, highly abundant RNAs containing miRNA-binding sites can act as competitive inhibitors of specific miRNAs. Indeed, the first specific miRNA inhibitors were synthetic anti-miRNA oligonucleotides216,217. Similarly, transgenic or transfected genes containing multiple miRNA-binding sites can inhibit the function of miRNAs in animals218–220. Such miRNA ‘sponges’ typically contain non-natural miRNA-binding sites that are designed with central mismatches to block target cleavage.



REVIEWS Biologically irrelevant

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Figure 5 | A quantitative framework for microRNA function. In this model, the relative abundance of microRNAs (miRNAs) and their targets inside the cell dictates the regulatory outcome. At concentrations below the affinity of Biology Nature Reviews | Molecular Cell Argonaute (AGO)-bound miRNAs for their targets (Kd), occupancy of miRNAs at mRNA targets is negligible and, thus, the biological function of these miRNAs is irrelevant. If present at concentrations above the Kd (this is typically true at least for the most abundant miRNAs in a given cell type or tissue), virtually every target site will be bound by a miRNA. In this case, the cumulative abundance of all target sites in the cell, relative to the abundance of the miRNA, will determine the extent to which each individual target mRNA is regulated. Fewer total target sites than the corresponding miRNA will prompt a larger extent of regulation compared with miRNAs that are less abundant than their target sites.

miRNA sponges occur naturally in plants and act to competitively inhibit miRNA repression of other targets in a phenomenon termed target mimicry 221. For example, phosphate starvation in A. thaliana induces the expression of both miR‑399 and the non-coding RNA INDUCED BY PHOSPHATE STARVATION 1 (IPS1). IPS1 binds miR‑399 through a site that contains central mismatches that protect IPS1 from AGO-catalysed cleavage. IPS1 thereby reduces the amount of miR‑399 available to repress its target mRNA PHOSPHATE 2. Transgenic non-cleavable miRNA sponges modelled after IPS1 inhibit individual miRNAs in plants221,222. Notably, the titrating targets model leaves unexplained the apparent conservation of miRNA-binding sites in titrating targets: how would so many sites, each of which contributes just a little to reducing the free pool of the miRNA, be under positive evolutionary selection? Should such sites not come and go over evolutionary time, with only their number and not their locations within any specific mRNA being conserved? A more speculative and thermodynamically implausible version of this idea proposes that individual target RNAs — not the collective abundance of miRNA seed matches — can titrate the amount of miRNA that is available to repress their targets223. For example, protein-coding and non-coding transcripts have been predicted to regulate the dosagesensitive tumour suppressor gene PTEN by competing with the PTEN transcript for binding to a similar set of miRNAs224. A decrease or loss of competing transcripts, for example derived from the pseudogene PTENP1 or the protein-coding gene zinc-finger E‑box-binding homeobox 2 (ZEB2), would increase the amount of miRNAs available to repress PTEN and hence accelerate tumorigenesis225–228. Among the objections to this idea are the findings that many miRNAs are more abundant than the sum of their individual mRNA targets and that the concentration of miRNAs in vivo are probably far greater than

their dissociation constant (Kd) values for seed match sites 147. Nematode and human miRNAs can reach 50,000 molecules per cell, with some being as abundant as the spliceosomal U6 small nuclear RNA (snRNA)229,230. By contrast, the 5,000 most abundant mRNA species in nematodes are thought to be present at just ~100 molecules each per cell229. Many miRNAs therefore outnumber any single mRNA target by as much as 500‑fold. However, each animal miRNA has the potential to regulate hundreds of mRNAs, some of which contain more than one miRNA‑binding site. Consequently, the number of some miRNAs may be similar to the number of miRNA‑binding sites, suggesting that a model in which a miRNA is regulated by numerous titrating targets is biochemically more feasible than one in which individual target mRNAs are key. Moreover, computer simulations using experimentally measured Kd and koff values suggest that only miRNAs that are least likely to provide meaningful target repression in a given cell type could be subject to competition by highly abundant, individual non-coding RNAs or mRNAs bearing miRNA-binding sites147. A quantitative framework for miRNA and target RNA function. The biochemical and biophysical properties of AGO-bound miRNAs provide a quantitative framework for understanding the reciprocal function of miRNAs and their targets according to their relative abundance inside the cell (FIG. 5). When a miRNA is dilute, that is, at concentrations lower than the affinity of AGO-bound miRNAs for their targets, the overall impact of the miRNA on gene expression is expected to be negligible147. This prediction is supported by miRNA sensor assays in cultured monocytes, which revealed that only the most abundant miRNAs are able to change the expression of a reporter construct containing a miRNA-binding site231. Highly abundant miRNAs (present at intracellular concentrations that vastly exceed the Kd for miRNA binding to a typical binding site on an mRNA target) are



REVIEWS predicted to be bound to target mRNA. The extent of regulation, that is, the number of sites on each target mRNA occupied by a miRNA, is then determined by the overall abundance of target sites in the transcriptome. In this case, significant regulation of miRNA activity through competitive transcripts is in principle possible, but only if the overall abundance of the competitor is much higher than the total number of miRNAs. Experimentally, this can be achieved by either expressing artificially high levels of sponge transcripts220,232–234 or by vastly increasing the number of miRNA-binding sites in each transcript219. For endogenous mRNAs or non-coding transcripts, it is unlikely that sufficiently high concentrations can be achieved to allow competitive inhibition of abundant miRNAs, particularly because miRNA binding to the competitor transcripts should decrease their steady-state abundance via one of the mechanisms highlighted above. Nevertheless, miRNA decay induced by viral infection suggests that mechanisms exist that efficiently inhibit miRNA function by triggering their decay upon inter action with target RNAs235–237, perhaps because enzymatic degradation and not simple competition renders this process efficient72–74,236. Because viruses only contribute the target RNA signal for miRNA decay, and the molecular machinery seems to be provided by the host cell itself, it is tempting to speculate that this process may be used to regulate miRNA function through cellular transcripts in uninfected cells, but biological examples of this have not yet been identified. Squaring the circle. An unexpected, natural solution to the competitive miRNA inhibitor paradox seems to have now been identified in the form of previously overlooked circular RNAs (circRNAs)238–240. circRNAs are generated by head‑to‑tail splicing of exons. The human circRNA, CDR1as (antisense to the cerebellar degeneration-related protein 1), contains 63 conserved sites with imperfect Lee, R. C., Feinbaum, R. L. & Ambros, V. The C. elegans heterochronic gene lin‑4 encodes small RNAs with antisense complementarity to lin‑14. Cell 75, 843–854 (1993). 2. Wightman, B., Ha, I. & Ruvkun, G. Posttranscriptional regulation of the heterochronic gene lin‑14 by lin‑4 mediates temporal pattern formation in C. elegans. Cell 75, 855–862 (1993). References 1 and 2 report the discovery of the first miRNA, lin‑4, and propose that it regulates mRNA expression post-transcriptionally. 3. Griffiths-Jones, S., Grocock, R. J., van Dongen, S., Bateman, A. & Enright, A. J. miRBase: microRNA sequences, targets and gene nomenclature. Nucleic Acids Res. 34, D140–D144 (2006). 4. Chiang, H. R. et al. Mammalian microRNAs: experimental evaluation of novel and previously annotated genes. Genes Dev. 24, 992–1009 (2010). 5. Friedman, R. C., Farh, K. K., Burge, C. B. & Bartel, D. P. Most mammalian mRNAs are conserved targets of microRNAs. Genome Res. 19, 92–105 (2009). Shows that more than half of all protein-coding genes in mammals have been evolutionarily selected to maintain pairing with miRNAs, indicating that most of the protein-coding transcriptome is regulated by miRNAs. 6. Bartel, D. P. MicroRNAs: genomics, biogenesis, mechanism, and function. Cell 116, 281–297 (2004). 7. Jones-Rhoades, M. W., Bartel, D. P. & Bartel, B. MicroRNAs and their regulatory roles in plants. Annu. Rev. Plant Biol. 57, 19–53 (2006). 1.

complementarity to miR‑7 and is proposed to function as a ‘super-sponge’ to regulate miR‑7 function in the neuronal tissues in which both the miRNA and the circRNA are co‑expressed. Although the biological function and the general scope of this type of regulation are unclear, ectopic expression of CDR1as in zebrafish, which naturally produce miR‑7 but not CDR1as, impairs midbrain development, recapitulating the loss‑of‑function phenotype of miR‑7 (REF. 240). The high abundance and strikingly high numbers of miRNA-binding sites in CDR1as are consistent with its proposed function in miRNA inhibition. More intriguingly, its circular architecture might prevent its degradation by miRNAs, particularly if such miRNA-directed destruction requires ribosome binding or recruitment of exonucleases.

Conclusions and perspectives This year marks the twentieth anniversary of the discovery of miRNAs1,2. In 1993, it would have been hard to imagine the extent and complexity of miRNA-mediated regulation of gene expression. Those studying these tiny riboregulators can deservedly be proud of our current catalogue of miRNAs, their mRNA targets and mechanisms of biogenesis and sequence diversification. Although many details surely remain to be discovered, what we most lack is a deeper understanding of the biological functions of miRNAs. Are unifying principles and deeper explanations for the role that miRNAs have in coordinating gene expression during development and environmental experience to be found? Or does much of what we now view as miRNA-mediated regulation or seemingly purposeful mechanisms to diversify miRNA sequences simply correspond to neutral experiments in evolution, the results of which we will be unable to observe? We believe that the answers to these and other questions about miRNAs lie in rigorous computational, biochemical and genetic tests of quantitative models.

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Acknowledgements

The authors thank the members of the Zamore and Ameres laboratories for helpful discussions and comments. The Ameres laboratory is funded by the Austrian Academy of Sciences and the Austrian Federal Ministry of Economy, Family and Youth (BMFWJ).

Competing interests statement

The authors declare no competing financial interests.

FURTHER INFORMATION Phillip D. Zamore’s homepage: http://www.umassmed.edu/zamore/index.aspx Stefan L. Ameres’ homepage: http://www.imba.oeaw.ac.at/research/stefan-ameres The microRNA database: http://www.mirbase.org ALL LINKS ARE ACTIVE IN THE ONLINE PDF


Diversifying microRNA sequence and function

Diversifying microRNA sequence and function

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