MEETING REVIEW 3093
Development 138, 3093-3102 (2011) doi:10.1242/dev.065284© 2011. Published by The Company of Biologists Ltd

Olivia S. Rissland1,* and Eric C. Lai2,*
Summary
The tenth annual Keystone Symposium on the Mechanism andBiology of Silencing convened in Monterey, California, in March
2011. Those seeking some West Coast sunshine were,
unfortunately, met with incessant precipitation throughout themeeting. Nevertheless, attendees were brightened byenlightening and vigorous scientific discussions. Here, wesummarize the results presented at the meeting, which inspire
and push this expanding field into new territories.
Key words: microRNA, piRNA, siRNA

Introduction
Central to understanding the process of development is knowledgeof how gene activity can be controlled precisely and robustly. Of
course, transcriptional regulation is crucial for development, and thefunctional complexities of DNA-binding proteins, their cofactors,
and chromatin structure remain actively pursued. However, the past
decade has witnessed a sea change in appreciating the contributionof post-transcriptional gene control by non-coding RNAs. Manyadvances in this field were first discussed during symposia in theannual Keystone series on RNA silencing, which convened mostrecently this spring in Monterey. Although the meeting covered
various aspects of the non-coding RNA world, emphasis was placedon the microRNA (miRNA), small interfering RNA (siRNA) andPiwi-interacting RNA (piRNA) pathways.
We divide this review into five themes. First, we discuss new
mechanistic insights into the biogenesis and post-transcriptional
regulation of miRNAs and siRNAs. Second, we report the latest
views on the still-controversial topic of how animal miRNAs
mediate silencing. Third, we describe new biological functions ofmiRNAs during animal development and physiology. Fourth, we
recount strong progress on elucidating piRNA biogenesis andfunction. Fifth, we consider the involvement of plant and fungal
small RNAs in transcriptional silencing. Together, these talks united
researchers in diverse disciplines and model systems, and theircollective progress has broad implications for post-transcriptional
gene regulation during development and metabolism, its dysfunction
during disease and its possible exploitation for therapy.
miRNAs and siRNAs: biogenesis and posttranscriptional regulation
The biogenesis of miRNAs and siRNAs has been well characterized
over the past decade (Fig. 1) (Kim et al., 2009). Briefly, primarymiRNA transcripts (pri-miRNAs) bearing miRNA hairpins are firstcleaved by the nuclear RNase III enzyme Drosha and its cofactor
Dgcr8. The resulting pre-miRNA hairpins are exported to thecytoplasm and cleaved by the RNase III enzyme Dicer to generate~22 nucleotide (nt) duplexes. One strand (the mature miRNA, or‘guide strand’) is preferentially loaded into an Argonaute (Ago)protein, which associates with Tnrc6 to yield the RNA-induced
silencing complex (RISC). siRNA biogenesis, by contrast, involves
cleavage of double-stranded RNAs (dsRNAs) by Dicer and loadingof mature siRNAs (guide strands) into Ago2 (Eif2c2), which is
specialized for target cleavage (‘slicing’). Many talks highlighted
outstanding questions regarding miRNA and siRNA biogenesis andregulation (Fig. 1).
Dicer processing

Canonical Dicers have a common architecture that includes ahelicase domain, a PAZ domain and two RNase III domains. Studies
of Giardia Dicer revealed that its PAZ domain binds the RNA 3⬘ end
to position the RNase III domains for substrate cleavage (MacRae
et al., 2007; Macrae et al., 2006). However, the role of the Nterminal helicase domain has been less clear. Brenda Bass
(University of Utah, UT, USA) reported that the helicase domain of
Caenorhabditis elegans Dicer (DCR-1 – WormBase) enables it,upon recognition of substrates with blunt termini, to engage Dicer
for processive cleavage (Welker et al., 2011). By contrast, RNAs
with a 3⬘ overhang (such as pre-miRNAs) are not cleaved
processively and do not require the helicase domain; these data help
explain the in vivo importance of the helicase domain in processing
nematode siRNAs, but not miRNAs.
Previous work from Narry Kim (Seoul National University,
Korea) showed that Dicer-mediated cleavage (referred to as‘dicing’) of pre-let-7 was inhibited by 3⬘ oligouridylation (Heo etal., 2009). However, her group recently found that human Dicer(DICER1) is not deterred by short 3⬘additions. These studies led tothe discovery that human Dicer can not only measure cleavage fromthe 3⬘ end, but can also recognize the 5⬘-phosphate to ‘count’ fromthe 5⬘ end (Park et al., 2011). In collaboration with Dinshaw Patel(Memorial Sloan-Kettering Cancer Center, NY, USA) structural
studies identified both a 5⬘-phosphate-binding pocket in the PAZcontaining region of human Dicer, which might serve as an anchor
point for measuring duplexes from the 5⬘ end, and a helical segment
implicated in orienting the duplex RNA.
David Bartel (Whitehead Institute, MA, USA) previously
reported that some budding yeasts, though notably not
Saccharomyces cerevisiae, harbor a functional RNAi pathway(Drinnenberg et al., 2009). However, budding yeast Dicers lack mostdomains found in canonical Dicers, including the PAZ domain.
How, then, does this Dicer accurately generate 23-nt siRNAs? Joint
efforts by the Bartel and Patel groups showed that multiple Dicer
dimers (the functional units of Dicer in budding yeast) bindcooperatively to the interior of the dsRNA substrate, with the
distance between active sites of adjacent dimers determining siRNA
product length (Weinberg et al., 2011). This ‘inside-out’ mechanism,
starting in the middle of dsRNA and working outwards, differs fromthe canonical mechanism observed with all other Dicers, whichstarts at dsRNA ends and works inwards.
Ago loading and activation

1

Whitehead Institute, Massachusetts Institute of Technology, Cambridge, MA 02142,
USA. 2Department of Developmental Biology, Sloan-Kettering Institute, 1275 YorkAve, Box 252, New York, NY 10065, USA.
*Authors for correspondence (rissland@wi.mit.edu; laie@mskcc.org)

Qinghua Liu (UT Southwestern, TX, USA) recently identified an
endoribonuclease that facilitates RISC activation by removing
passenger-strand cleavage products (Liu et al., 2009). Composed oftwo subunits, Translin and Trax, he dubbed this complex ‘C3PO’

DEVELOPMENT

RNA silencing in Monterey

3094 MEETING REVIEW

Development 138 (15)miRNA geneTranscription• How are hairpin structures
recognized by Drosha/Dgcr8?

Microprocessor

Droshapri-miRNA m7G

AAAAAAAA
Dgcr8(Pasha in flies)

Microprocessor
cleavage

Drosha-independent substrates(mirtrons, endo-shRNA, etc.)

pre-miRNA
Guide strand

Passenger strand

Nucleus
Cytoplasm
Dicer• How does Dicer measureits cleavage position?

Fig. 1. Animal microRNA biogenesis. Most microRNA (miRNA) genes
are transcribed by RNA polymerase II, resulting in the production of a
primary miRNA (pri-miRNA). This pri-miRNA is recognized by the‘microprocessor complex’ (composed of the RNase III enzyme Drosha
and its co-factor Dgcr8) and is cleaved (yellows arrows) by Drosha togive rise to an intermediate called the pre-miRNA. After being exported
to the cytoplasm, the pre-miRNA binds to a complex consisting of Dicer(an RNase III enzyme) and Trbp, and is cleaved (yellow arrows) by Dicer
to generate a small RNA duplex. The mature miRNA (the ‘guide’ strand;red) is loaded into Argonaute (Ago) to create a functional miRNAArgonaute effector complex (RNA-induced silencing complex or RISC).
The other strand of the duplex (the ‘passenger’ or ‘star’ strand; gray) is
degraded. RISC is then guided to its target messenger RNAs (mRNAs)
through base-pairing interactions between the guide strand andsequences in the target message, which are found predominantly
within the 3⬘untranslated region (UTR). The accessory protein Tnrc6(GW182 in Drosophila) mediates interactions with additional proteinsto elicit both translational repression and degradation of the target
message. Note that some substrates (e.g. mirtrons) generate premiRNAs independently of Drosha, whereas some rare miRNAs (e.g.
miR-451) are Dicer-independent and mature via Ago2-mediated slicing
reactions. Some open questions in the pathway are highlighted in the
text bubbles; recent advances that address these questions were
presented at the symposium. dsRBD, double-strand RNA binding
domain; endo-shRNA, endogenous short hairpin RNA; m7G, 7-methyl
guanosine; ORF, open reading frame; siRNA, small interfering RNA.

Trbp

DicingLoadingRNA-induced silencing
complex (RISC)

Argonaute(Ago1-4 in mammals,
Ago2 is Slicer)• How does RLC
transition to RISC?
Tnrc6(GW182 in flies)
Dicer-independent, Ago2
substrates (pre-miR-451)

Target binding
and repression

ORF

Key

• How are levels of mature
miRNAs/siRNAs regulated?

AAAAAAAAAA

RNase III enzyme(Drosha, Dicer)

ArgonautedsRBD partner(Dgcr8, Trbp)

Tnrc6(after the chatty Star Wars robot, a nod to their previous designationof the siRNA loading factor R2D2). At this year’s meeting, Liu andPatel independently presented C3PO structures. Liu’s group
crystallized human C3PO and likened it to an elongated Death Star
(Ye et al., 2011). Their studies revealed a hollow octomeric complex
containing six Translin and two Trax subunits. Patel also observed
a central cavity in Drosophila C3PO (Tian et al., 2011), half ofwhich he likened to an opened, de-pitted avocado. Intriguingly, thecatalytic residues were found on the interior surface, raisingquestions of how the passenger strand becomes enveloped by C3PO
for degradation.
Olivier Voinnet (Swiss Federal Institute of Technology,
Switzerland) discussed the transition of Ago from the RNA-loading
complex (RLC) into RISC (Fig. 1). Previously, he and Rich Carthew

described Ago and its effector partner GW182 (Tnrc6) associatingwith endosomes and multivesicular bodies (MVBs) (Gibbings et al.,2009; Lee et al., 2009). Voinnet reported that the prion protein (PrP),a known MVB marker, contains extensive GW-repeats resemblingthe ‘Ago hooks’ found in other Ago-interacting proteins. Voinnet
proposed that PrP directly recruits Ago to the membrane of MVBs.
Upon relocalization of Ago and subsequent Tnrc6 binding, atargeting-competent RISC forms. Future studies of the subcellular
localization of Ago and its associated proteins should provide moreinsight into miRNA biology.
miRNA turnoverWhat determines the fate and longevity of miRNA-Ago
complexes? Although most miRNAs seem stable, there is growinginterest in unstable miRNAs and the enzymes mediating theirdecay (Kai and Pasquinelli, 2010; Siomi and Siomi, 2010). Using
the cell cycle as a system, Bartel and co-workers identified miR503 as an unusually unstable miRNA, the levels of which decreaseby ~90% during the G0-G1 transition. miR-503 belongs to theextended miR-16 family, several members of which similarly
decrease upon this transition. Because this family represses genesthat promote the G1-S transition (Cimmino et al., 2005; Linsley etal., 2007), Bartel proposed that high levels of the miR-16 family
reinforce G0 arrest, whereas the subsequent downregulation of thisfamily, mediated by the unusual instability of some members,
facilitates cell cycle re-entry.
Continuing on this theme, Kim reported that miR-141 is rapidly
degraded upon loss of cell contact, and that this instability requires
a 3-nt sequence in the center of miR-141. By profiling miRNAs
following loss of cell adhesion, her group observed destabilizationof other miRNAs known to be involved in the epithelialmesenchymal transition, suggesting dynamic control of miRNAstability during this process.

DEVELOPMENT

RNA loading complex(RLC)

Development 138 (15)MEETING REVIEW 3095
Translational
repression

A Deadenylation

Decay

Ccr4-Caf1
deadenylase

Xrn1
AAAA

AAAA

AAAA

Pan2-Pan3
deadenylase

B Translational repression

eIF4E

Deadenylation

AAAA

AAAAAAA
AAAA
AAAA
A
AAA

eIF4G

Decay

AAAA

AAAAAAA
AAAA
AAAA
AAA
PABP

Coincident deadenylationand translational repression

C Decircularization

RISC

AAAAAAA
AAAA
AAAA
AAA

Decay

AAAA

AAAA

Coincident Deadenylation
and Translational Repression

D Unknown process

Decay

AAAA

AAAA
A
A

AAAAAAA
AAAA
AAAA
AAA

Key
RNA-induced silencing
complex (RISC)

eIF4E

PABP

Ccr4-Caf1 deadenylase complex

Ribosome

eIF4G

Xrn1

Pan2-Pan3 deadenylase complex

Highly stable miRNA-Ago complexes help to protect miRNAs
from exonucleases. Ian MacRae (Scripps Research Institute, CA,
USA) reported that, surprisingly, in vitro, upon the addition of
fully complementary target RNA, guide RNAs were rapidly
released from human AGO2 (hAGO2). A few 3⬘ mismatches
inhibited such unloading, thus allowing more Ago-mediatedcleavage reactions to be catalyzed in vitro and enhancing silencing
in vivo. These results are similar to those from Brian Brown(Mount Sinai School of Medicine, NY, USA), who showed thatoverexpression of highly complementary targets destabilizes the
cognate miRNA (Baccarini et al., 2011). How such unloadinginterfaces with the trimming-tailing pathway of sRNA degradation(Ameres et al., 2010) remains to be determined.

miRNA-mediated silencing mechanisms in animals
miRNAs act predominantly via target mRNA destabilization, withsmaller contributions from translational repression (Guo et al., 2010;
Hendrickson et al., 2009) (Fig. 2). Given this, monitoring mRNAchanges can identify bulk miRNA targets. Nevertheless, the precise
mechanism of repression still eludes the field, and, of all the topics
covered at this Keystone symposium, this stimulated the mostintense discussions.
Ago cap-binding and allostery

Although it has been established that the Ago MID domain binds tothe 5⬘ end of miRNAs (Hutvagner and Simard, 2008), the hypothesisthat it also binds the 5⬘ cap of target mRNAs has been both

DEVELOPMENT

Fig. 2. Proposed mechanisms of microRNA-mediated repression. The majority of microRNA (miRNA)-mediated regulation is due to mRNAdegradation. However, the precise mechanism(s) underlying this process remain a topic of lively discussion. Initially, the mature, fully transcribedmRNA is thought be in a circular messenger ribonucleoprotein (mRNP) form composed of, at least, the cap-binding protein (eIF4E), the scaffold
protein eIF4G, poly(A) binding protein (PABP), ribosomes and other related proteins not shown here. Once deadenylation, which is mediated by thePan2-Pan3 and Ccr4-Caf1 deadenylase complexes, has occurred, mRNAs are decapped by the decapping enzyme (not shown) and subsequently
degraded by the 5⬘r3⬘ exonuclease Xrn1. Current studies aim to elucidate the order and relative contributions of target deadenylation, capdependent translational repression and target decay. Evidence for several potential scenarios has been reported, including: (A) mRNA deadenylation
leading to translational repression and mRNA decay; (B) translational repression of target mRNAs leading to deadenylation and decay; (C)decircularization of target mRNAs, which then leads to coincident deadenylation and translational repression, and then to decay; and (D) an
unknown/undiscovered mechanism, which occurs coincidently with deadenylation and translational repression.

captivating and contentious (Djuranovic et al., 2010; Eulalio et al.,2008; Kiriakidou et al., 2007). Previous work from Rachel Green’slaboratory (Johns Hopkins University, MD, USA) suggested that
this domain possesses two distinct, and allosteric, sites: one binding
the guide strand, the other binding the 5⬘ cap (Djuranovic et al.,2010). Crystallization of the MID domain of QDE-2 (theNeurospora crassa homolog of Ago) provided structural support fora second ligand-binding site (Boland et al., 2010). However, Filipp
Frank (Nahum Sonenberg and Bhushan Nagar laboratories, McGill
University, Canada) reported that nucleotides and cap analogs
readily interact with the hAGO2 MID domain through the miRNAbinding pocket, but that additional nucleotide interactions outside
this region were not observed (Frank et al., 2011).
Elisa Izaurralde (Max Planck Institute for Developmental
Biology, Germany) presented new work on the MID-PIWI lobe of
QDE-2 (Boland et al., 2011). Siwi also stably bound longer
substrates with 1U preference. Excitingly, addition of appropriate
lysates to these complexes revealed a processive 3⬘ to 5⬘
exonuclease (‘trimming’) activity that resected the 3⬘ ends of long
substrates yielding 27-nt products. Although structurally similar to
Thermus thermophilus Ago (Wang et al., 2008), QDE-2 contains aeukaryotic-specific C-terminal insertion within the PIWI domain.
She proposed that this insertion can sense the functional state of theprotein, i.e. whether or not the guide RNA is bound to target RNA
in an extended duplex. In this structure, the binding pocket for theguide RNA 5⬘ end forms at the MID-PIWI interface, as observed inprokaryotic Ago proteins (Wang et al., 2008). However, thepreviously proposed cap-binding pocket (Boland et al., 2010) is
occluded by C-terminal residues inserting into this interface. Finally,
this structure demonstrated that residues previously implicated inallosteric regulation (Djuranovic et al., 2010) are involved in MIDPIWI interdomain interactions and facilitate guide RNA binding bystabilizing this interface.
Rachel Green noted that Drosophila AGO1-mediated repressionin tethering assays correlates with the binding of miRNAs, m7G-cap
resin and GW182 (Gawky – FlyBase), and she re-iterated a model
whereby the second binding site might predominantly regulate Agofunction allosterically. She presented new work on Ago proteinsbearing PAZ mutations or those lacking a PAZ domain; suchproteins were surprisingly functional in tethering assays. During his
keynote talk, Patel discussed how formation of an extended duplex
within T. thermophilus Ago requires the release of the guide strand
3⬘ end from PAZ (Wang et al., 2009). In agreement with this, Green
showed that dAGO1 PAZ mutants, which are impaired for 3⬘ guidestrand binding, show increased and promiscuous cleavage activity.
Understanding the importance of miRNA-binding, and the role ofallostery, for the ability to mediate repression remains an importantgoal.
miRNA-mediated repression and deadenylation

Many studies have shown that miRNA binding triggers target
deadenylation via Ago and GW182 (Tnrc6) complexes (BehmAnsmant et al., 2006; Giraldez et al., 2006; Lai et al., 1998; Wu etal., 2006). Deadenylation is an initial step in mRNA decay, which is
required for both decapping (and subsequent 5⬘-3⬘ decay) and 3⬘-5⬘
degradation. Deadenylation also inhibits translation, as synergism
between the poly(A) tail and the 5⬘ cap promotes translation
initiation. But what is the mechanistic relationship between target
deadenylation and translational repression (Fig. 2)? Is thisrelationship causal and, if so, does translational repression trigger
deadenylation or vice versa? Alternatively, do these processes occur
coincidently but independently?

Development 138 (15)

Numerous studies have provided evidence for miRNA-mediatedinhibition of canonical translation initiation by various mechanisms(Fabian et al., 2010; Humphreys et al., 2005; Mathonnet et al., 2007;
Pillai et al., 2005). Citing the non-physiological nature of such
experiments, Bartel discussed the repression of endogenous long,non-coding RNAs (lncRNAs). If deadenylation is purely aconsequence of translational repression, then lncRNAs should be
refractory to miRNA-mediated repression. However, Bartel reported
that targeted lncRNAs are effectively degraded, suggesting thattranslational repression is not required for deadenylation. Izaurralde
supported the view that miRNAs can mediate deadenylation
directly, as her group detected interactions between Tnrc6 and thecytoplasmic deadenylase complexes Pan2-Pan3 and Ccr4-Not. Her
studies show that Pan3 bound directly to Tnrc6 and suggest thatTnrc6 and the Pan3-Pan2 complex might form a platform onto
which the processive Ccr4/Not deadenylase is recruited.
Thomas Duchaine (McGill University, Canada) described a C.
elegans extract-based system that recapitulates miRNA-mediateddeadenylation and translational repression. The observation that the3⬘ untranslated region (UTR) influences both the rate and the extent
of deadenylation (Wu et al., 2010) led his group to investigate the‘dialogue’ between UTRs and miRNAs. They find that certain
alternative poly(A) sites (APAs) in C. elegans 3⬘ UTRs can split
cooperative pairs of miRNA binding sites, adding a twist to how
APAs can modulate 3⬘-UTR-mediated repression (Mayr and Bartel,
2009; Sandberg et al., 2008). It will be interesting to integrate these
insights with recent genome-wide surveys of C. elegans poly(A)sites (Jan et al., 2010; Mangone et al., 2010). Nikolaus Rajewsky(Max Delbruck Center for Molecular Medicine, Germany) also
investigated the impact of mRNA-protein (mRNP) complexes on
RNA regulation, using PAR-CLIP (photoactivatable ribonucleosideenhanced cross-linking and immunoprecipitation). This technique
was developed to map RNA binding sites in cultured cells (Hafner
et al., 2010) and Rajewsky’s group has now extended its usage tointact C. elegans animals. They also combined data from published
CLIP studies and from the UCSC genome browser to generate aconvenient portal for integrated database searches(http://dorina.mdc-berlin.de).
The zebrafish embryo is another powerful system forinvestigating mechanisms of miRNA-mediated repression. Here,maternal mRNAs are deadenylated and cleared upon zygotic
expression of miR-430 (Giraldez et al., 2006). Antonio Giraldez(Yale University, CT, USA) profiled ribosome occupancy and total
mRNA levels at various time points post fertilization in wild-type
embryos and in dicer mutants (which lack miR-430). Many of themRNAs that were repressed in wild-type cells contained miR-430
sites: strikingly, these were first translationally repressed andsubsequently degraded. Such results are consistent with Green’s
kinetic data from human cells, in which translational repression was
detected prior to either full deadenylation or reduction in target
messages. Although some deadenylation of selected zebrafish
transcripts was observed, Giraldez proposed that this wasinsufficient to explain the observed translational repression. He thus
favors a model in which deadenylation and translational repression
are coincident processes, perhaps initiated by Tnrc6-mediatedmRNA decircularization (Fig. 2).
Biological functions of animal miRNAs
Although studies of miRNAs are aided by genome-wide target
predictions and convenient methods for transcriptome profiling,understanding miRNA biology in vivo remains challenging. In
contrast to mutants of the founding miRNAs lin-4 and let-7, many

DEVELOPMENT

3096 MEETING REVIEW

other individual miRNA mutants lack obvious phenotypes (Miska
et al., 2007). Instead, mutants of core miRNA pathway components
provide a means to assess miRNA requirements broadly. Eric Miska(University of Cambridge, UK) exploited a temperature-sensitive
allele of C. elegans Pasha (PASH-1 – WormBase), which encodes
an obligate cofactor for Drosha (DRSH-1 – WormBase) (Fig. 1), to
inactivate miRNA biogenesis with temporal control. This study
revealed that miRNA production is continuously required in adult
C. elegans for diverse behaviors and lifespan. This mutant also
provides a unique background in which differential miRNA stability
can be examined in the intact animal.
Victor Ambros (University of Massachusetts Medical School, MA,
USA) noted that many miRNA mutants do exhibit phenotypes ingenetically sensitized or stressed conditions (Brenner et al., 2010). He
found previously that C. elegans miRNAs of the let-7 ‘sister’ family
function redundantly to control the transition between the L2 and L3
larval stages by repressing hunchback-like 1 (hbl-1) (Abbott et al.,2005). Surprisingly, they could bypass the requirement for let-7 sister
miRNAs by forcing animals through dauer (a stage of arrested
development adopted under harsh environmental conditions), a
phenomenon termed ‘post-dauer suppression’ (PDS). A genetic screen
for loci required for PDS revealed NHL-2, which augments let-7
family activity (Hammell et al., 2009), and lin-4, which acts in parallelto let-7 sisters in repressing hbl-1. This latter finding was unexpected
given that lin-4 is not required per se for the L2 to L3 transition duringcontinuous development. Ambros speculated that other miRNA
functions might be revealed under the appropriate sensitization.
The group of Eric Olson (UTSW, TX, USA) deleted ~20 miRNAs
expressed in heart and/or muscle. Although these knockouts are all
homozygous viable, fertile and of normal morphology, they havebeen a treasure trove for elucidating essential roles of miRNAs
under stressed conditions (Small and Olson, 2011). At the meeting,Olson reported how the dysregulation of miR-378 in skeletal muscle
affected glucose uptake: its loss normalized glucose sensitivity in an
obesity model and was cardioprotective under a high fat diet, raising
the possibility of therapeutic manipulation of this miRNA. Olson
also exploited Mir451 knockouts, which he and other groups foundto be relatively normal, except that they were defective inregenerating red blood cells (RBCs) following oxidative stress(Patrick et al., 2010; Rasmussen et al., 2010; Yu et al., 2010). In newwork, Olson reported that inhibition of miR-451 reduced hematocrit
levels in a mouse model of polycythemia vera, a disease involving
excess RBCs. These miRNA inhibitors were well tolerated in wildtype mice supporting their potential therapeutic usage.
Eric Lai (Sloan-Kettering Institute, NY, USA) discussed miRNAs
embedded within the Drosophila Bithorax-complex (BX-C), which
encodes conserved homeodomain (‘Hox’) proteins that execute
segmental identities along the anterior-posterior axis. In bothmammals (Mansfield et al., 2004; Yekta et al., 2004) and Drosophila(Stark et al., 2008; Tyler et al., 2008), Hox miRNAs strongly target
Hox mRNAs, yet changes in segmental identity were not observedin a BX-C miRNA deletion mutant (Bender, 2008). In new work, Lai
reported that multiple BX-C miRNA targets were strongly derepressed in the posterior domains of larval brains that lack BX-C
miRNAs. Importantly, heterozygosity of these miRNA targets could
rescue the sterility of the BX-C miRNA mutant, establishing themas key target genes in the Hox miRNA network.
Elucidating the mysterious piRNA pathway
The Piwi-interacting RNA (piRNA) pathway generates ~24- to 32nt RNAs that load Piwi-class proteins in gonads (Siomi et al., 2011).
Despite a rich collection of piRNA mutants, our knowledge of

MEETING REVIEW 3097

piRNA biogenesis and regulatory mechanisms remains rudimentary.
Several talks highlighted cell biological, genetic and in vitro
biochemical strategies to elucidate the piRNA pathway.
Lysate systems for primary and secondary piRNAbiogenesis

Biochemistry was instrumental in the mechanistic dissection of thesiRNA and miRNA pathways but, until now, the major insightgained from in vitro studies of piRNAs has been that Piwi proteins
exhibit slicer activity, i.e. they can mediate target cleavage. This
activity drives a feed-forward loop, termed ‘piRNA ping pong’, that
amplifies piRNAs, yielding pairs that overlap by 10 nt (Brennecke
et al., 2007; Gunawardane et al., 2007) (Fig. 3).
Yuki Tomari (University of Tokyo, Japan) described an in vitro
system for primary piRNA biogenesis based on Bombyx mori(silkworm) Bm-N4 cells. In lysates, Siwi preferentially loaded 27nt, 5⬘ monophosphorylated RNAs with 5⬘ uridine (1U), mirroring
the known characteristics of primary piRNAs. Although this doesn’t
address how long pre-piRNA substrates are initially fragmented, it
indicates that piRNA biogenesis need not involve a 5⬘ U-specificnuclease: Siwi can simply select 1U RNAs. Siwi also stably boundlonger substrates with 1U preference. Excitingly, addition of
appropriate lysates to these complexes revealed a processive 3⬘ to 5⬘
exonuclease (‘trimming’) activity that resected the 3⬘ ends of long
substrates yielding 27-nt products. These products (but not longer
Siwi-bound species) were methylated at their 3⬘ ends, indicative of
genuine piRNA maturation.
Kuniaki Saito, from the laboratory of Mikiko and Haru Siomi(Keio University, Japan), described a complementary Bm-N4 cellbased assay for secondary piRNA production. Saito reported that
immunopurified Siwi cleaved a complementary target, yielding an
expected 47-nt product that reflected the generation of a secondary
piRNA precursor. Strikingly, when performed in lysates, thiscleavage reaction produced a 28-nt product, which was interpreted
as a mature secondary piRNA generated by piRISC cleavage andsubsequent trimming by unknown factors. With these revelations ofin vitro piRNA biogenesis, the hunt is undoubtedly on to establish
analogous systems from more genetically tractable species, as wellas to purify the nucleases and other factors that mediate piRNAbiogenesis.
Tudor domain proteins and piRNAs

Several talks highlighted the spatial compartmentalization of piRNAcomponents (Fig. 3), many of which reside in ‘nuage’. As Toshie
Kai (Temasek Life Sciences Laboratory, Singapore) reminded us,
this French term for ‘cloud’ describes electron-dense perinuclear
structures that are a key cellular location for piRNA function.
Another relevant cellular structure is the ‘Yb body’, which wasrecently inferred to be a location in which Piwi proteins are loaded(Siomi et al., 2011; Szakmary et al., 2009). Many proteins
containing Tudor domains localize to nuage or Yb bodies and,
importantly, are required for piRNA pathway function (Siomi et al.,2011). Tudor domains contain a conserved structural motif that canbind to symmetrically dimethylated arginines (SDMAs), which arefound on most Piwi proteins. Although the functions of most Tudorcontaining proteins (TDRDs) are not known, a ‘Tudor code’ hasbeen postulated to explain how these domains link Piwi proteins toother functional domains on Tudor proteins or associatedcomplexes.
Julius Brennecke (IMBA Vienna, Austria) screened 18 Drosophila
Tudor genes in the somatic and germline portions of the ovary andfocused on functional analysis of Avocado, an MYND-Tudor protein

DEVELOPMENT

Development 138 (15)

3098 MEETING REVIEW

Development 138 (15)Primary processing

Secondary processing
piRNA clusters

Active transposons

TE antisense strand TE sense strand

TE sense strand

piRNA clusters, mRNA/3⬘UTRs, transposons• What is the role ofnuclear Piwi/piRNAs?

U

Nucleus
Cytoplasm

Armitage
Aub

Piwi
U

U

AGO3

ZucchiniYb

Yb bodyUA

Mitochondria
U• How are primary piRNA transcripts fragmented?
• How are the 3⬘ ends of piRNAs formed?
• What is the relationship between mitochondria and piRNAs?

‘Ping-pong’amplification

AAU• Is there sub-nuage segregation of piRNA biogenesis?
• What are the roles of other germline Tudors and helicases?

Nuage
Key

Piwi proteins(Piwi, Aub, AGO3)

Tudor domain proteins(Yb, Tudor, other Tudor proteins, etc.)

most similar to vertebrate TDRD1. They generated an Avocado
mutant, which was sterile and displayed upregulation of transposable
elements (TEs) in the germline and soma, owing to a defect in primarypiRNA biogenesis. Avocado was required for piRNA biogenesis andloading into Piwi, similar to the RNA helicases Armitage and Yb[fs(1)Yb – FlyBase]. In fact, Avocado binds Armitage directly, andboth are localized to Yb bodies of follicle cells. Epistasis tests showedthat Avocado requires armi and yb to localize to Yb bodies, whereas
loss of the nuclease zucchini causes Armitage and Avocado toaccumulate in super-sized Yb bodies.
Kai reported mutants of several Tudor proteins, focusing on Tejas(which is most similar to vertebrate TDRD5) (Patil and Kai, 2010),
Tapas (which is similar to vertebrate TDRD7) and Kumo (which issimilar to vertebrate ring finger protein 17; RNF17). Whereas Tejas
mutants are sterile and exhibit accumulation of TEs, Tapas mutants
were fertile with lower levels of TEs. Strikingly, the double mutants
exhibited synergistic phenotypes, with nearly complete loss of theovary within 5-7 days, germarium arrested cysts and increased TE
levels. Finally, Kumo might represent one of the most upstream
components of the piRNA pathway; Kumo localization to nuage wasindependent of all known piRNA factors and, reciprocally, thelocalization of all other piRNA factors was defective in Kumo
mutants.
Rene Ketting’s group (Hubrecht Institute, The Netherlands) foundmany Tudor proteins in immunoprecipitates of the zebrafish Piwi
proteins Ziwi (Piwil1 – Zebrafish Information Network) and Zili

DEVELOPMENT

Fig. 3. Biogenesis and function of Piwi-interacting RNAs. The process of Piwi-interacting (piRNA) biogenesis is summarized, and the key
regulatory factors and mechanisms involved are shown; for simplicity, only the Drosophila orthologs are named. In the primary processing pathway,
diverse transcripts, including active transposable elements (TEs), piRNA clusters encoding dead TEs, and 3⬘ UTRs (untranslated regions) of many
mRNAs, are converted into 5⬘U-biased piRNAs and loaded into Piwi (which associates with the RNA helicase Armitage) in Yb bodies, which exist in
proximity to mitochondria. Some Piwi proteins, such as fly Piwi and mouse Piwil4, also accumulate in the nucleus. Primary piRNAs can engage insecondary processing, known as the ‘ping-pong’ amplification pathway, if antisense counterparts (shown in red) exist, as is the case with piRNA
clusters and active transposons. In this pathway, Aub, loaded with primary piRNAs, slices complementary transcripts (as indicated by the yellow
arrow) to generate secondary piRNAs. These have an A10 bias, are loaded in AGO3 and can mediate the regeneration of primary 5⬘U piRNAs.
Many ‘ping-pong’ factors localize to nuage, an electron-dense perinuclear structure that appears to be a key cellular location for piRNA function;
additional components include a host of Tudor domain and helicase proteins, not shown. Some open questions in the pathway are highlighted in
the text bubbles and progress on these topics was presented at the symposium.

Development 138 (15)Genetic screens for new piRNA factors

A strength of model organism genetics is the capacity to perform
unbiased forward screening. Brennecke’s group has embarked on afunctional genome-wide screen for primary piRNA pathway
components using the Vienna Drosophila RNAi Center Collectionof knockdown transgenes (http://stockcenter.vdrc.at), an ambitious
effort poised to yield new factors.
Vasily Vagin (Greg Hannon’s laboratory, Cold Spring Harbor
Laboratory, NY, USA) screened a collection of female sterile
Drosophila lines for signs of TE deregulation, and found that loss ofa mitochondrial glycerol 3-phosphate O-acyltransferase wasdetrimental to piRNA-mediated silencing. Encouraged by Hannon
to ‘make it sexy’, Vagin noticed that this mitochondrial enzyme isinvolved in generating phosphatidic acid (PA). This bears relation
to recent reports that MITOPLD, a mammalian homolog of the
Drosophila piRNA factor Zucchini, localizes to mitochondria, isrequired for piRNA-mediated silencing and male fertility, and also
generates PA (Huang et al., 2011; Watanabe et al., 2011). Vagin
speculated on a potential cooperation between metabolic processes
generating PA and piRNA biogenesis, and the intriguing possibility
that PA might serve as a second messenger that regulates nuage
formation and/or piRNA biogenesis.
Possible non-TE functions of the piRNA pathway

Although the major known functional piRNA targets are repetitiveelements and TEs, the range of piRNA targets might be broader. For
example, the most abundant mammalian piRNAs, expressed inpachytene testis from discrete genomic clusters, do not have obvious
targets (Siomi et al., 2011). Similarly, an abundant class of piRNAs
derives from the 3⬘ UTRs of many protein-coding genes in
Drosophila and vertebrate gonads. These might function as transregulatory RNAs, or might exert a cis-regulatory effect on hostmRNAs (Robine et al., 2009; Saito et al., 2009).
Anastasios Vourekas (laboratory of Zissimos Mourelatos,
University of Pennsylvania, PA, USA) performed high-throughput
sequencing of RNA isolated by crosslinking immunoprecipitation(HITS-CLIP) on piRNAs and longer RNA fragments associatedwith the mouse Piwi homologs Mili (Piwil2 – Mouse Genome
Informatics) and Miwi (Piwil1 – Mouse Genome Informatics) andthe mouse Vasa homolog MVH (Ddx4 – Mouse Genome
Informatics). The larger tags comprised many reads mapping topachytene piRNA clusters, and analysis of Mili large CLIP tags
suggested that the larger fragments form by endonucleolytic
cleavage and 3⬘ trimming, in agreement with the lysate-based
studies of Tomari and Siomi. Many Miwi large CLIP tags lacking
corresponding piRNAs mapped to spermiogenic genes, reminiscentof the hypothesis that Miwi stabilizes these transcripts (Deng andLin, 2002). However, as most target interactions of Ago proteins are
driven by small RNAs, the apparent recruitment of Miwi tospermiogenic transcripts awaits further study.
Martine Simonelig (Institute of Human Genetics, Montpellier,France) reported that piRNA factors are required for timely
deadenylation and degradation of maternal transcripts, including theposterior determinant nanos (Rouget et al., 2010). Now, she
described that, in flies with newly acquired piRNA-mediatedimmunity to the P element transposon, insertion of a P-piRNA site
into the nanos 3⬘ UTR could mediate functional repression of nanos.
Additional work is needed to determine breadth of piRNA-mediatedregulation of mRNAs.
Transcriptional gene silencing in fungi and plants
Although most talks focused on animal systems, RNAi operates
much more broadly, and often in ways that are mechanisticallydistinct between organisms. For example, Kathleen Collins
highlighted the fact that various Tetrahymena Piwi proteins carry an
unexpectedly diverse set of small RNA cargoes (Couvillion et al.,2009), including 3⬘ fragments of tRNAs (Couvillion et al., 2010).
Furthermore, a unique aspect of plant and fungal systems has been
the usage of small RNAs to direct transcriptional gene silencing, andwe highlight several new findings here.
Plant 24-nucleotide siRNAs

David Baulcombe (University of Cambridge, UK) discussed plant
24-nt siRNAs, which target nascent transcripts and elicit
epigenetic modifications of genomic targets. These were
implicated in non-cell-autonomous silencing, although definitive
proof from Arabidopsis grafting experiments has only recently
been obtained (Dunoyer et al., 2010; Molnar et al., 2010).
Baulcombe now described that in tomato hybrids, 24-nt siRNAs
were expressed at much higher levels in hybrids than in either
tomato parent, a phenomenon he termed ‘transgressive
expression’. This expression correlates with target gene silencing
and DNA methylation. Moreover, in subsequent hybrid
generations, transgressive phenotypes were observed, indicating
previously unappreciated functional roles for 24-nt siRNAs inplants.
disiRNAs in Neurospora crassa

Yi Liu (UT Southwestern Medical School, TX, USA) recently
reported that the Neurospora Ago QDE-2 loads small RNAs that arederived from diverse, non-canonical biogenesis pathways (Lee et al.,2010). One such class originated from gene-rich regions thatgenerate siRNAs from both strands, and was unexpectedly
generated independently of Dicer. These were thus termed dicerindependent siRNAs (disiRNAs). Their function is not clear but all
disiRNA loci checked were cytosine methylated, similar totransposon relics and repeats in Neurospora. Surprisingly,
methylation at disiRNA loci was reduced but persisted in mutants ofall known DNA or histone methyltransferases. Therefore,
Neurospora disiRNA loci require an unknown nuclease for theirbiogenesis, and might engage an unknown methyltransferase for
their function.
RNAi in Schizosaccharomyces pombeRNAi-mediated transcriptional gene silencing (TGS) has beenstudied extensively in Schizosaccharomyces pombe (Grewal,2010). Briefly, Dcr1 processes long dsRNA into siRNAs that load
Ago1, a component of the RNA-induced transcriptional silencing(RITS) effector complex. RITS is directed to nascent transcripts,

DEVELOPMENT(Piwil2 – Zebrafish Information Network), and used binding assays
with SDMA (symmetrically dimethylated arginine) peptides from
Piwi proteins to define aspects of the Tudor recognition code.
Analysis of Tdrd1 immunoprecipitates revealed short and long RNA
populations; the former comprised a mix of mature piRNAs carriedin Ziwi and Zili whereas the latter derived from the same genomic
regions as mature piRNAs and exhibited signatures of being piRNAtargets. This led to a model in which Tdrd1 scaffolds a spatial
domain in which Ziwi, Zili and their targets interact. Ketting also
reported that Tdrd6 binds specifically to Ziwi and to mainly primarypiRNAs, and thus might define a spatial domain relevant to theprimary pathway.

MEETING REVIEW 3099

a response that is amplified by the RNA-dependent RNA
polymerase Rdp1. Ultimately, the histone methyltransferase Clr4
is recruited and heterochromatic factors such as Swi6 areassembled at silenced regions, including centromeres, telomeresand mating-type loci.
Nick Proudfoot (University of Oxford, UK) earlier reported thatloci of convergent transcripts are transiently marked asheterochromatin by RNAi, resulting in cell cycle-modulated gene
expression (Gullerova and Proudfoot, 2008). As many RNAi genes
are arranged in convergent pairs, does this cell-cycle loop modulate
activity of the RNAi pathway? Indeed, ‘tandemization’ of ago1
abrogated its cell cycle-dependent heterochromatic marking andregulation, and this induced aberrant mitotic spindles (Gullerova etal., 2011). Proudfoot concluded his talk with unpublished data on
TGS in trans, in which convergently transcribed genes expressedfrom plasmids could silence themselves and endogenous genecopies. Even more tantalizingly, Proudfoot indicated that such asystem might also work in human cells.
Marc Bühler (Friedrich Miescher Institute, Switzerland) mapped
genomic binding of Dcr1, Rdp1 and Swi6 in S. pombe using DNA
adenine methyltransferase identification (DamID) (Woolcock et al.,2011). As expected, Swi6, Dcr1 and Rdp1 bound known
heterochromatin loci similarly. Surprisingly, Dcr1 (and to a lesser
extent, Swi6 and Rdp1) also associated with euchromatic regions.
Moreover, although Clr4 is required for accumulation ofcentromeric siRNAs, the DamID profile of Dcr1 was unaffected in
clr4delta cells. Bühler hypothesized that although Dcr1 is recruited
to chromatin independently of heterochromatin marks, downstream
processes, such as Ago-loading, might require histone methylation.
How are the euchromatic loci described by Proudfoot andBühler, which exhibit transient heterochromatin and subtle RNAi
effects, distinguished from canonical heterochromatic loci? Shiv
Grewal (NCI, National Institutes of Health, MD, USA) addressed
this question through studies of Mlo3, an mRNA export-related
factor partially required for centromeric silencing (Zhang et al.,2011). Mlo3 interacts with both the RITS complex (via Clr4) andthe nuclear RNA surveillance complex. Grewal proposed that
Mlo3 is able to sense aberrant transcripts, which are then either
directed to the RNAi pathway or degraded by the exosome, a steppresumably influenced by the presence of siRNAs. Curiously,
deletion of mlo3 in RNAi mutants restored silencing andheterochromatin formation, suggesting the existence of afundamental heterochromatin pathway in which non-coding RNAs
can target loci independently of RNAi.
Conclusions
This symposium convened in California, a state whose motto(‘Eureka!’) is customarily applied to the discovery of gold, but isequally appropriate for the scientific process. As we highlighted
above, this annual meeting of the silencing field reliably provides
new insights and prompts further questions. The powerful pairing ofgenome-wide techniques with biochemical, molecular, genetic andstructural approaches will undoubtedly maintain progress. The
ongoing discovery of processes and relationships, so many of whichwere unimaginable even last year, highlights the vibrancy of thisfield and underscores how much remains to be uncovered. Meeting
attendees left California, dreaming of the vistas in silencing that are
to come.
Acknowledgements
We thank members of the Bartel and Lai laboratories, R. Green, A. Grimson, V.
N. Kim, N. Lau, I. J. MacRae and C. Norbury for helpful discussions andsuggestions. We are grateful to speakers for allowing their unpublished work

Development 138 (15)to be discussed, and apologize to meeting participants whose work could not
be covered owing to space constraints. O.S.R. was supported by a Ruth L.
Kirschstein National Research Service Award. E.C.L. was supported by the
Burroughs Wellcome Fund, the Starr Cancer Consortium and the NIH.
Competing interests statement
The authors declare no competing financial interests.
References
Abbott, A. L., Alvarez-Saavedra, E., Miska, E. A., Lau, N. C., Bartel, D. P.,
Horvitz, H. R. and Ambros, V. (2005). The let-7 MicroRNA family members
mir-48, mir-84, and mir-241 function together to regulate developmental timing
in Caenorhabditis elegans. Dev. Cell 9, 403-414.
Ameres, S. L., Horwich, M. D., Hung, J. H., Xu, J., Ghildiyal, M., Weng, Z.
and Zamore, P. D. (2010). Target RNA-directed trimming and tailing of small
silencing RNAs. Science 328, 1534-1539.
Baccarini, A., Chauhan, H., Gardner, T. J., Jayaprakash, A. D.,
Sachidanandam, R. and Brown, B. D. (2011). Kinetic analysis reveals the fateof a microRNA following target regulation in mammalian cells. Curr. Biol. 21,
369-376.
Behm-Ansmant, I., Rehwinkel, J., Doerks, T., Stark, A., Bork, P. andIzaurralde, E. (2006). mRNA degradation by miRNAs and GW182 requires both
CCR4:NOT deadenylase and DCP1:DCP2 decapping complexes. Genes Dev. 20,
1885-1898.
Bender, W. (2008). MicroRNAs in the Drosophila bithorax complex. Genes Dev. 22,
14-19.
Boland, A., Tritschler, F., Heimstadt, S., Izaurralde, E. and Weichenrieder, O.
(2010). Crystal structure and ligand binding of the MID domain of a eukaryotic
Argonaute protein. EMBO Rep. 11, 522-527.
Boland, A., Huntzinger, E., Schmidt, S., Izaurralde, E. and Weichenrieder, O.
(2011). Crystal structure of the MID-PIWI lobe of a eukaryotic Argonaute
protein. Proc. Natl. Acad. Sci. USA 108, 10466-10471.
Brennecke, J., Aravin, A. A., Stark, A., Dus, M., Kellis, M., Sachidanandam,
R. and Hannon, G. J. (2007). Discrete small RNA-generating loci as master
regulators of transposon activity in Drosophila. Cell 128, 1089-1103.
Brenner, J. L., Jasiewicz, K. L., Fahley, A. F., Kemp, B. J. and Abbott, A. L.
(2010). Loss of individual microRNAs causes mutant phenotypes in sensitized
genetic backgrounds in C. elegans. Curr. Biol. 20, 1321-1325.
Cimmino, A., Calin, G. A., Fabbri, M., Iorio, M. V., Ferracin, M., Shimizu, M.,
Wojcik, S. E., Aqeilan, R. I., Zupo, S., Dono, M. et al. (2005). miR-15 andmiR-16 induce apoptosis by targeting BCL2. Proc. Natl. Acad. Sci. USA 102,
13944-13949.
Couvillion, M. T., Lee, S. R., Hogstad, B., Malone, C. D., Tonkin, L. A.,
Sachidanandam, R., Hannon, G. J. and Collins, K. (2009). Sequence,biogenesis, and function of diverse small RNA classes bound to the Piwi family
proteins of Tetrahymena thermophila. Genes Dev. 23, 2016-2032.
Couvillion, M. T., Sachidanandam, R. and Collins, K. (2010). A growthessential Tetrahymena Piwi protein carries tRNA fragment cargo. Genes Dev. 24,
2742-2747.
Deng, W. and Lin, H. (2002). miwi, a murine homolog of piwi, encodes acytoplasmic protein essential for spermatogenesis. Dev. Cell 2, 819-830.
Djuranovic, S., Zinchenko, M. K., Hur, J. K., Nahvi, A., Brunelle, J. L., Rogers,
E. J. and Green, R. (2010). Allosteric regulation of Argonaute proteins by
miRNAs. Nat. Struct. Mol. Biol. 17, 144-150.
Drinnenberg, I. A., Weinberg, D. E., Xie, K. T., Mower, J. P., Wolfe, K. H.,
Fink, G. R. and Bartel, D. P. (2009). RNAi in budding yeast. Science 326, 544550.
Dunoyer, P., Brosnan, C. A., Schott, G., Wang, Y., Jay, F., Alioua, A., Himber,
C. and Voinnet, O. (2010). An endogenous, systemic RNAi pathway in plants.
EMBO J. 29, 1699-1712.
Eulalio, A., Huntzinger, E. and Izaurralde, E. (2008). GW182 interaction with
Argonaute is essential for miRNA-mediated translational repression and mRNA
decay. Nat. Struct. Mol. Biol. 15, 346-353.
Fabian, M. R., Sonenberg, N. and Filipowicz, W. (2010). Regulation of mRNAtranslation and stability by microRNAs. Annu. Rev. Biochem. 79, 351-379.
Frank, F., Fabian, M. R., Stepinski, J., Jemielity, J., Darzynkiewicz, E.,
Sonenberg, N. and Nagar, B. (2011). Structural analysis of 5⬘-mRNA-cap
interactions with the human AGO2 MID domain. EMBO Rep. 12, 415-420.
Gibbings, D. J., Ciaudo, C., Erhardt, M. and Voinnet, O. (2009). Multivesicularbodies associate with components of miRNA effector complexes and modulate
miRNA activity. Nat. Cell Biol. 11, 1143-1149.
Giraldez, A. J., Mishima, Y., Rihel, J., Grocock, R. J., Van Dongen, S., Inoue,K., Enright, A. J. and Schier, A. F. (2006). Zebrafish MiR-430 promotes
deadenylation and clearance of maternal mRNAs. Science 312, 75-79.
Grewal, S. I. (2010). RNAi-dependent formation of heterochromatin and its
diverse functions. Curr. Opin. Genet. Dev. 20, 134-141.
Gullerova, M. and Proudfoot, N. J. (2008). Cohesin complex promotes
transcriptional termination between convergent genes in S. pombe. Cell 132,
983-995.

DEVELOPMENT

3100 MEETING REVIEW

Gullerova, M., Moazed, D. and Proudfoot, N. J. (2011). Autoregulation of
convergent RNAi genes in fission yeast. Genes Dev. 25, 556-568.
Gunawardane, L. S., Saito, K., Nishida, K. M., Miyoshi, K., Kawamura, Y.,
Nagami, T., Siomi, H. and Siomi, M. C. (2007). A slicer-mediated mechanism
for repeat-associated siRNA 5⬘ end formation in Drosophila. Science 315, 15871590.
Guo, H., Ingolia, N. T., Weissman, J. S. and Bartel, D. P. (2010). Mammalian
microRNAs predominantly act to decrease target mRNA levels. Nature 466, 835840.
Hafner, M., Landthaler, M., Burger, L., Khorshid, M., Hausser, J., Berninger,
P., Rothballer, A., Ascano, M., Jr, Jungkamp, A. C., Munschauer, M. et al.
(2010). Transcriptome-wide identification of RNA-binding protein and microRNAtarget sites by PAR-CLIP. Cell 141, 129-241.
Hammell, C. M., Lubin, I., Boag, P. R., Blackwell, T. K. and Ambros, V. (2009).
nhl-2 modulates microRNA activity in Caenorhabditis elegans. Cell 136, 926938.
Hendrickson, D. G., Hogan, D. J., McCullough, H. L., Myers, J. W., Herschlag,
D., Ferrell, J. E. and Brown, P. O. (2009). Concordant regulation of translationand mRNA abundance for hundreds of targets of a human microRNA. PLoS Biol.
7, e1000238.
Heo, I., Joo, C., Kim, Y. K., Ha, M., Yoon, M. J., Cho, J., Yeom, K. H., Han, J.
and Kim, V. N. (2009). TUT4 in concert with Lin28 suppresses microRNAbiogenesis through pre-microRNA uridylation. Cell 138, 696-708.
Huang, H., Gao, Q., Peng, X., Choi, S. Y., Sarma, K., Ren, H., Morris, A. J. andFrohman, M. A. (2011). piRNA-associated germline nuage formation andspermatogenesis require MitoPLD profusogenic mitochondrial-surface lipid
signaling. Dev. Cell 20, 376-387.
Humphreys, D. T., Westman, B. J., Martin, D. I. and Preiss, T. (2005).
MicroRNAs control translation initiation by inhibiting eukaryotic initiation factor
4E/cap and poly(A) tail function. Proc. Natl. Acad. Sci. USA 102, 16961-16966.
Hutvagner, G. and Simard, M. J. (2008). Argonaute proteins: key players in RNA
silencing. Nat. Rev. Mol. Cell Biol. 9, 22-32.
Jan, C. H., Friedman, R. C., Ruby, J. G. and Bartel, D. P. (2010). Formation,
regulation and evolution of Caenorhabditis elegans 3⬘UTRs. Nature 469, 97101.
Kai, Z. S. and Pasquinelli, A. E. (2010). MicroRNA assassins: factors that regulate
the disappearance of miRNAs. Nat. Struct. Mol. Biol. 17, 5-10.
Kim, V. N., Han, J. and Siomi, M. C. (2009). Biogenesis of small RNAs in animals.
Nat. Rev. Mol. Cell Biol. 10, 126-139.
Kiriakidou, M., Tan, G. S., Lamprinaki, S., De Planell-Saguer, M., Nelson, P. T.
and Mourelatos, Z. (2007). An mRNA m7G cap binding-like motif withinhuman Ago2 represses translation. Cell 129, 1141-1151.
Lai, E. C., Burks, C. and Posakony, J. W. (1998). The K box, a conserved 3⬘ UTR
sequence motif, negatively regulates accumulation of Enhancer of split Complex
transcripts. Development 125, 4077-4088.
Lee, H. C., Li, L., Gu, W., Xue, Z., Crosthwaite, S. K., Pertsemlidis, A., Lewis,
Z. A., Freitag, M., Selker, E. U., Mello, C. C. et al. (2010). Diverse pathways
generate microRNA-like RNAs and Dicer-independent small interfering RNAs in
fungi. Mol. Cell 38, 803-814.
Lee, Y. S., Pressman, S., Andress, A. P., Kim, K., White, J. L., Cassidy, J. J., Li,
X., Lubell, K., Lim, d. H., Cho, I. S. et al. (2009). Silencing by small RNAs islinked to endosomal trafficking. Nat. Cell Biol. 11, 1150-1156.
Linsley, P. S., Schelter, J., Burchard, J., Kibukawa, M., Martin, M. M., Bartz, S.
R., Johnson, J. M., Cummins, J. M., Raymond, C. K., Dai, H. et al. (2007).
Transcripts targeted by the microRNA-16 family cooperatively regulate cell cycle
progression. Mol. Cell. Biol. 27, 2240-2252.
Liu, Y., Ye, X., Jiang, F., Liang, C., Chen, D., Peng, J., Kinch, L. N., Grishin, N.
V. and Liu, Q. (2009). C3PO, an endoribonuclease that promotes RNAi byfacilitating RISC activation. Science 325, 750-753.
Macrae, I. J., Zhou, K., Li, F., Repic, A., Brooks, A. N., Cande, W. Z., Adams, P.
D. and Doudna, J. A. (2006). Structural basis for double-stranded RNA
processing by Dicer. Science 311, 195-198.
MacRae, I. J., Zhou, K. and Doudna, J. A. (2007). Structural determinants of
RNA recognition and cleavage by Dicer. Nat. Struct. Mol. Biol. 14, 934-940.
Mangone, M., Manoharan, A. P., Thierry-Mieg, D., Thierry-Mieg, J., Han, T.,
Mackowiak, S. D., Mis, E., Zegar, C., Gutwein, M. R., Khivansara, V. et al.
(2010). The landscape of C. elegans 3⬘UTRs. Science 329, 432-435.
Mansfield, J. H., Harfe, B. D., Nissen, R., Obenauer, J., Srineel, J., Chaudhuri,A., Farzan-Kashani, R., Zuker, M., Pasquinelli, A. E., Ruvkun, G. et al.
(2004). MicroRNA-responsive ‘sensor’ transgenes uncover Hox-like and otherdevelopmentally regulated patterns of vertebrate microRNA expression. Nat.
Genet. 36, 1079-1083.
Mathonnet, G., Fabian, M. R., Svitkin, Y. V., Parsyan, A., Huck, L., Murata, T.,
Biffo, S., Merrick, W. C., Darzynkiewicz, E., Pillai, R. S. et al. (2007).
MicroRNA inhibition of translation initiation in vitro by targeting the cap-binding
complex eIF4F. Science 317, 1764-1767.
Mayr, C. and Bartel, D. P. (2009). Widespread shortening of 3⬘UTRs by alternative
cleavage and polyadenylation activates oncogenes in cancer cells. Cell 138, 673684.

MEETING REVIEW 3101
Miska, E. A., Alvarez-Saavedra, E., Abbott, A. L., Lau, N. C., Hellman, A. B.,
McGonagle, S. M., Bartel, D. P., Ambros, V. R. and Horvitz, H. R. (2007).
Most Caenorhabditis elegans microRNAs are individually not essential fordevelopment or viability. PLoS Genet. 3, e215.
Molnar, A., Melnyk, C. W., Bassett, A., Hardcastle, T. J., Dunn, R. andBaulcombe, D. C. (2010). Small silencing RNAs in plants are mobile and direct
epigenetic modification in recipient cells. Science 328, 872-875.
Park, J. E., Heo, I., Tian, Y., Simanshu, D. K., Chang, H., Jee, D., Patel, D. J.
and Kim, V. N. (2011). Dicer recognizes the 59 end of RNA for efficient andaccurate processing. Nature (in press). doi:10.1038/nature10198.
Patil, V. S. and Kai, T. (2010). Repression of retroelements in Drosophila germline
via piRNA pathway by the tudor domain protein tejas. Curr. Biol. 20, 724-730.
Patrick, D. M., Zhang, C. C., Tao, Y., Yao, H., Qi, X., Schwartz, R. J., Jun-Shen
Huang, L. and Olson, E. N. (2010). Defective erythroid differentiation in miR451 mutant mice mediated by 14-3-3zeta. Genes Dev. 24, 1614-1619.
Pillai, R. S., Bhattacharyya, S. N., Artus, C. G., Zoller, T., Cougot, N., Basyuk,
E., Bertrand, E. and Filipowicz, W. (2005). Inhibition of translational initiation
by Let-7 MicroRNA in human cells. Science 309, 1573-1576.
Rasmussen, K. D., Simmini, S., Abreu-Goodger, C., Bartonicek, N., Di
Giacomo, M., Bilbao-Cortes, D., Horos, R., Von Lindern, M., Enright, A. J.
and O’Carroll, D. (2010). The miR-144/451 locus is required for erythroid
homeostasis. J. Exp. Med. 207, 1351-1358.
Robine, N., Lau, N. C., Balla, S., Jin, Z., Okamura, K., Kuramochi-Miyagawa,S., Blower, M. D. and Lai, E. C. (2009). A broadly conserved pathway
generates 3⬘UTR-directed primary piRNAs. Curr. Biol. 19, 2066-2076.
Rouget, C., Papin, C., Boureux, A., Meunier, A. C., Franco, B., Robine, N., Lai,
E. C., Pelisson, A. and Simonelig, M. (2010). Maternal mRNA deadenylationand decay by the piRNA pathway in the early Drosophila embryo. Nature 467,
1128-1132.
Saito, K., Inagaki, S., Mituyama, T., Kawamura, Y., Ono, Y., Sakota, E.,
Kotani, H., Asai, K., Siomi, H. and Siomi, M. C. (2009). A regulatory circuit
for piwi by the large Maf gene traffic jam in Drosophila. Nature 461, 12961299.
Sandberg, R., Neilson, J. R., Sarma, A., Sharp, P. A. and Burge, C. B. (2008).
Proliferating cells express mRNAs with shortened 3⬘ untranslated regions andfewer microRNA target sites. Science 320, 1643-1647.
Siomi, H. and Siomi, M. C. (2010). Posttranscriptional regulation of microRNAbiogenesis in animals. Mol. Cell 38, 323-332.
Siomi, M. C., Sato, K., Pezic, D. and Aravin, A. A. (2011). PIWI-interacting small
RNAs: the vanguard of genome defence. Nat. Rev. Mol. Cell Biol. 12, 246-258.
Small, E. M. and Olson, E. N. (2011). Pervasive roles of microRNAs in
cardiovascular biology. Nature 469, 336-342.
Stark, A., Bushati, N., Jan, C. H., Kheradpour, P., Hodges, E., Brennecke, J.,
Bartel, D. P., Cohen, S. M. and Kellis, M. (2008). A single Hox locus in
Drosophila produces functional microRNAs from opposite DNA strands. Genes
Dev. 22, 8-13.
Szakmary, A., Reedy, M., Qi, H. and Lin, H. (2009). The Yb protein defines anovel organelle and regulates male germline stem cell self-renewal in Drosophilamelanogaster. J. Cell Biol. 185, 613-627.
Tian, Y., Simanshu, D. K., Ascano, M., Diaz-Avalos, R., Park, A. Y., Juranek, S.
A., Rice, W. J., Yin, Q., Robinson, C. V., Tuschl, T. et al. (2011). Multimericassembly and biochemical characterization of the Trax-translin endonuclease
complex. Nat. Struct. Mol. Biol. 18, 658-664.
Tyler, D. M., Okamura, K., Chung, W. J., Hagen, J. W., Berezikov, E., Hannon,
G. J. and Lai, E. C. (2008). Functionally distinct regulatory RNAs generated bybidirectional transcription and processing of microRNA loci. Genes Dev. 22, 2636.
Wang, Y., Sheng, G., Juranek, S., Tuschl, T. and Patel, D. J. (2008). Structure of
the guide-strand-containing argonaute silencing complex. Nature 456, 209-213.
Wang, Y., Juranek, S., Li, H., Sheng, G., Wardle, G. S., Tuschl, T. and Patel, D.
J. (2009). Nucleation, propagation and cleavage of target RNAs in Ago silencing
complexes. Nature 461, 754-761.
Watanabe, T., Chuma, S., Yamamoto, Y., Kuramochi-Miyagawa, S., Totoki,Y., Toyoda, A., Hoki, Y., Fujiyama, A., Shibata, T., Sado, T. et al. (2011).
MITOPLD is a mitochondrial protein essential for nuage formation and piRNAbiogenesis in the mouse germline. Dev. Cell 20, 364-375.
Weinberg, D. E., Nakanishi, K., Patel, D. J. and Bartel, D. P. (2011). The insideout mechanism Dicers from budding yeasts. Cell (in press).
doi:10.1016/j.cell.2011.06.021.
Welker, N. C., Maity, T. S., Ye, X., Aruscavage, P. J., Krauchuk, A. A., Liu, Q.
and Bass, B. L. (2011). Dicer’s helicase domain discriminates dsRNA termini topromote an altered reaction mode. Mol. Cell 41, 589-599.
Woolcock, K. J., Gaidatzis, D., Punga, T. and Buhler, M. (2011). Dicer
associates with chromatin to repress genome activity in Schizosaccharomyces
pombe. Nat. Struct. Mol. Biol. 18, 94-99.
Wu, E., Thivierge, C., Flamand, M., Mathonnet, G., Vashisht, A. A.,
Wohlschlegel, J., Fabian, M. R., Sonenberg, N. and Duchaine, T. F. (2010).
Pervasive and cooperative deadenylation of 3⬘UTRs by embryonic microRNAfamilies. Mol. Cell 40, 558-570.

DEVELOPMENT

Development 138 (15)Wu, L., Fan, J. and Belasco, J. G. (2006). MicroRNAs direct rapid deadenylationof mRNA. Proc. Natl. Acad. Sci. USA 103, 4034-4039.
Ye, X., Huang, N., Liu, Y., Paroo, Z., Huerta, C., Li, P., Chen, S., Liu, Q. andZhang, H. (2011). Structure of C3PO and mechanism of human RISC activation.
Nat. Struct. Mol. Biol. 18, 650-657.
Yekta, S., Shih, I. H. and Bartel, D. P. (2004). MicroRNA-directed cleavage of
HOXB8 mRNA. Science 304, 594-596.

Development 138 (15)Yu, D., dos Santos, C. O., Zhao, G., Jiang, J., Amigo, J. D., Khandros, E., Dore,
L. C., Yao, Y., D’Souza, J., Zhang, Z. et al. (2010). miR-451 protects against
erythroid oxidant stress by repressing 14-3-3zeta. Genes Dev. 24, 1620-1633.
Zhang, K., Fischer, T., Porter, R. L., Dhakshnamoorthy, J., Zofall, M., Zhou,
M., Veenstra, T. and Grewal, S. I. (2011). Clr4/Suv39 and RNA quality control
factors cooperate to trigger RNAi and suppress antisense RNA. Science 331,
1624-1627.

DEVELOPMENT

3102 MEETING REVIEW