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NATURE|Vol 457|22 January 2009|doi:10.1038/nature07754

On the road to reading theRNA-interference code
Haruhiko Siomi1 & Mikiko C. Siomi1,2
The finding that sequence-specific gene silencing occurs in response to the presence of double-stranded
RNAs has had an enormous impact on biology, uncovering an unsuspected level of regulation of gene
expression. This process, known as RNA interference (RNAi) or RNA silencing, involves small non-coding
RNAs, which associate with nuclease-containing regulatory complexes and then pair with complementary
messenger RNA targets, thereby preventing the expression of these mRNAs. Remarkable progress has been
made towards understanding the underlying mechanisms of RNAi, raising the prospect of deciphering the‘RNAi code’ that, like transcription factors, allows the fine-tuning and networking of complex suites of geneactivity, thereby specifying cellular physiology and development.
The discovery of RNA interference (RNAi)1 heralded a revolution in
RNA biology. Researchers uncovered ‘hidden’ layers of regulation ofgene expression, in which many previously unidentified families of small
RNAs (consisting of ~20–30 nucleotides) mediate gene silencing. A
diverse set of gene-regulatory mechanisms were found to use key steps in
the RNAi process, including mechanisms that silence endogenous genes
and mechanisms that restrain the expression of parasitic and pathogenic
invaders such as transposons and viruses2–5.
The basic RNAi process can be divided into three steps6,7. First, a long
double-stranded RNA (dsRNA) that is expressed in, or introduced into,
the cell (for example, as a result of the base-pairing of sense and antisensetranscripts or the formation of stem–loop structures) is processed into
small RNA duplexes by a ribonuclease III (RNaseIII) enzyme known
as Dicer. Second, these duplexes are unwound, and one strand ispreferentially loaded into a protein complex known as the RNA-induced
silencing complex (RISC). Third, this complex effectively searches thetranscriptome and finds potential target RNAs. The loaded single-stranded
RNA (ssRNA), called the guide strand, then directs an endonuclease that ispresent in the RISC (sometimes called the ‘slicer’ and now known to be an
Argonaute protein8–11) to cleave messenger RNAs that contain sequencehomologous to the ssRNA, over many rounds. In this way, the guide strand
determines the sequence specificity of the RNAi response.
In different organisms, the RNAi pathways comprise different
proteins and mechanisms, but they operate by strikingly convergent
strategies. In all organisms that have been studied, RNAi involves two
main components: small RNAs, which determine the specificity of
the response; and Argonaute proteins, which carry out the repression.
Depending on both the nature of the Argonaute in the RISC and thedegree of complementarity between the small RNA and the target
sequence in the mRNA, the association of the RISC with target mRNAs
has been shown to have different outcomes: it can control protein
synthesis and mRNA stability, maintain genome integrity or produce a
specific set of small RNAs8,12. Analyses of the biogenesis of small RNAs
and their targeting mechanisms have benefited from the advent of highthroughput sequencing technologies and sophisticated bioinformatics13.
The picture emerging from these studies is that RNAi systems in different
organisms have been refined in many ways, and such modifications

include built-in molecular ‘rulers’ that define the size of small RNAs,
structures that determine which strand of a small RNA is selected,
mechanisms that direct further rounds of small RNA amplification, or
safeguards against off-target (unrestricted and unrelated) silencing.
Another emerging finding in the field is that the activity of RNAi
pathways is subject to intense regulation at various levels, from the levelof biogenesis of small RNAs to the silencing mode of the RISC. In thisReview, we describe the biogenesis of the guide strand of small RNAs
and the formation and actions of the RISC, and we discuss the current
understanding of the molecular mechanisms of RNAi in the light of recent
insights into how silencing pathways are specified and regulated.

Biogenesis of small RNAs
A hallmark of RNAi is that short (~20–30 nucleotide) dsRNAs — known
as small RNAs — are generated by the activity of RNaseIII enzymes(either Dicer alone or Drosha and Dicer). Two main categories of small
RNAs have been defined on the basis of their precursors. The cleavageof exogenous long dsRNA precursors in response to viral infection orafter artificial introduction generates short interfering RNAs (siRNAs),
whereas the processing of genome-encoded stem–loop structuresgenerates microRNAs (miRNAs). Using high-throughput sequencing
technology, several new classes of endogenous small RNA species haverecently been uncovered, and these include PIWI-interacting RNAs
(piRNAs) and endogenous siRNAs (endo-siRNAs or esiRNAs).
A common feature of all of these small RNAs is that they are loaded
onto Argonaute proteins to effect their targeting function (discussed
further in the section ‘Loading and sorting of small RNAs by the RISC’).
An overview of the generation of small RNAs is presented in Fig. 1.
siRNA biogenesis
Dicer (Table 1) processes long RNA duplexes and generates siRNAs.
These small RNAs are ~21–25-nucleotide duplexes with a phosphategroup at both 5ʹ ends, and hydroxyl groups and two-nucleotide overhangs
at both 3ʹ ends, all hallmarks of RNaseIII-mediated cleavage. The Dicer
protein contains a PAZ domain, which binds to the 3ʹ end of an siRNA,and two RNaseIII domains, which have the catalytic activity. It functions
as a monomer14, but the RNaseIII domains associate with each other to1
Department of Molecular Biology, Keio University School of Medicine, 35 Shinanomachi, Shinjuku-ku, Tokyo 160-8582, Japan. 2Japan Science and Technology Agency (JST), Core Research for
Evolutional Science and Technology (CREST), 4-1-8 Hon-chou, Kawaguchi, Saitama 332-0012, Japan.

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form an ‘internal dimer’ (see page 405). The distance between the PAZ
domain and the two RNaseIII domains is the length spanned by 25 basepairs (bp) of RNA15. Thus, Dicer itself is a molecular ruler.
miRNA biogenesis
Similarly, miRNAs are short (~21–25-nucleotide) RNA molecules16;
however, their biogenesis differs markedly from that of siRNAs. The
primary precursors of miRNAs (pri-miRNAs) are encoded in the
genome, and the relevant genomic regions are mostly transcribed by RNA
polymerase II (ref. 17). The pri-miRNAs contain stem–loop structuresa5′

dsRNA precursors

b
3′

5′

that harbour the miRNA in the 5ʹ or 3ʹ half of the stem. During miRNA
production in plants, one type of RNaseIII, Dicer-like protein 1 (DCL1),
generates the miRNA–miRNA* duplex in the nucleus (miRNA* being
the sequence in the stem–loop that pairs with the miRNA, equivalent to
the passenger strand of siRNA duplexes; discussed later). By contrast,
in animals, miRNAs are derived in a two-step process, in which thenuclear-localized RNaseIII Drosha defines one end of the miRNA–
miRNA* duplex and releases a precursor miRNA (pre-miRNA) of~65–70 nucleotides. The pre-miRNA hairpin is then exported to thecytoplasm, where Dicer completes the processing.
TranscriptcssRNA precursor

3′
3′

m7G

Synthesis of dsRNA
by RdRP

5′
Primary processing
pathway(RNaseIII independent)

dsRNA processing(RNaseIII dependent)
Small-RNA-duplex
unwinding(Slicer dependent)

Unwindase?

(Slicer independent)

Loading onto
Argonaute

Loading onto
Argonaute

Loading onto
Argonaute
Argonaute

Target recognitionand silencing

Secondary processing
pathway

CleavageTransfer to another
Argonaute protein

Figure 1 | Small RNA production and RNA silencing. a, Natural transcriptsthat form dsRNAs and hairpin-shaped structures can be sources ofsmall RNAs. These precursors are processed by an RNaseIII enzyme(such as Drosha or Dicer), yielding small RNA duplexes. Duplexes witha perfect match (left pathway) are further processed by an enzyme withslicer activity (an Argonaute protein) into single-stranded small RNAs.
By contrast, small RNA duplexes with a mismatch or bulge in the centre(right pathway) are not substrates for the slicer and thus become singlestranded in a cleavage-independent manner. The identity of the proteinthat carries out this unwinding is unknown. Single-stranded small RNAs
are then loaded onto Argonaute proteins. The particular strand thatis selected (sense or antisense) depends on thermodynamic stability.
The loaded Argonaute proteins are guided to target mRNAs containing
complementary sequence, and the expression of the corresponding
genes is silenced. The mode of action of this silencing — whether themRNA is cleaved or whether translation is just repressed — largely
depends on the degree of complementarity between the target mRNAs
and the Argonaute-associated small RNAs. b, Some small RNAs foundin Caenorhabditis elegans and plants are known to be produced in an
RNA-dependent RNA polymerase (RdRP)-dependent manner. Naturaltranscripts (often aberrant RNAs) can be substrates for this type of small
RNA synthesis. This does not occur in organisms that lack RdRP activity,
such as mammals and Drosophila melanogaster. Single-stranded small
RNAs generated in this way can then be loaded onto Argonaute proteinsand silence gene expression. c, The PIWI subfamily of Argonaute proteins,which are germline specific, are loaded with piRNAs. These complexes
function to silence transposons. Single-stranded precursors give rise topiRNAs, through a mechanism called the primary processing pathway.
The proteins required for this pathway are unknown. The silencing oftransposons by PIWI proteins simultaneously amplifies piRNAs in germ
cells. This pathway is known as the secondary processing pathway (or the
ping-pong amplification loop) and is conserved in a variety of organisms,
including mice and zebrafish. In this pathway, the slicer activity of the
PIWI proteins reciprocally forms the 5ʹ ends of piRNAs by cleavingtransposon transcripts (piRNA precursors). Proteins required to form
the 3ʹ end of piRNAs remain unidentified.

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Table 1 | Key proteins in RNA silencing in various organisms
Protein

Yeast(Schizosaccharomyces
pombe)

Plant(Arabidopsis thaliana)

Nematode(Caenorhabditis
elegans)

Fruitfly(Drosophilamelanogaster)

Mammal
Mouse

Human

RNaseIII

Dcr1

DCL1

DCR-1

DCR-1

DICER1

DICER1DCL2

DRSH-1

DCR-2

DROSHADROSHADROSHA

DCL3
DCL4
Argonaute: AGO subfamily

Ago1

AGO1

ALG-1

AGO1

AGO1

AGO1

AGO2

ALG-2

AGO2

AGO2

AGO2

AGO4

3 others

AGO3

AGO3

AGO5

AGO4

AGO4

AGO6

AGO5 (possiblya pseudogene)

AGO7 (ZIPPY)AGO10 (ZLL, PNH)
3 others
Argonaute: PIWI subfamily

None

None

ERGO-1AGO3MILI (PIWIL2)

HILI (PIWIL2)

PRG-1

PIWI

MIWI (PIWIL1)

HIWI (PIWIL1)

PRG-2

AUB

MIWI2 (PIWIL4)

HIWI2 (PIWIL4)

RDE-1

None

None

None

PASH-1

PASHA

DGCR8

DGCR8

RDE-4

R2D2

TRBP (TARBP2)

TRBP (TARBP2)

LOQS

PACT (PRKRA)PACT (PRKRA)

None

None

None

PIWIL3 (HIWI3)
Argonaute: WAGO subfamily

None

None

SAGO-1
SAGO-2
PPW-1
PPW-2
CSR-1
NRDE-311 others
Double-stranded-RNA-binding
domain (dsRBD)-containing
cofactor of RNaseIII

None

RNA-dependent RNA
polymerase (RdRP)

Rdp1

HYL1

RDR1

EGO-1

RDR2 (SMD1)

RRF-1

RDR6 (SDE1, SGS2)

RRF-3

3 others

1 otherMolecules that belong to these categories but have unknown functions are not listed but are indicated as ‘others’. Common synonyms are indicated in parentheses. Data were taken from refs 8, 12, 29, 50 and 98.

Drosha is present in a large complex, known as the microprocessor
complex, which functions like a molecular ruler to determine the cleavage
site in the pri-miRNAs18,19. In this complex, Drosha interacts with its
cofactor, known as DGCR8 or Pasha (depending on the species), which
also binds to dsRNA (through its dsRNA-binding domain; dsRBD)20,21.
A typical metazoan pri-miRNA consists of a 33-bp stem, a terminal
loop and ssRNA flanking segments. The flanking segments are crucial
for binding to DGCR8, and the 33-bp stem is also required for efficientbinding. Drosha can interact transiently with the stem of this ‘pre-cleavage’
complex, and the processing centre of the enzyme, located at ~11 bp fromthe ssRNA–dsRNA junction, makes a staggered pair of breaks in the
RNA to create the ~65–70-nucleotide pre-miRNA. Thus, DGCR8 might
function as the molecular anchor that measures the distance from thessRNA–dsRNA junction. It is possible that the microprocessor complex
could recognize the terminal loop as ssRNA and bind to the stem–loop
structure in the opposite orientation. In this case, abortive cleavage canoccur at an alternative site ~11 bp from the terminal loop. However, mostpri-miRNAs contain internal bulges or weakly paired bases ~11 bp fromthe terminal loop that mitigate processing from this direction18.
Although many of the sequences encoding miRNAs are locatedwithin introns, clusters encoding miRNAs that are processed directlyby the spliceosome, instead of Drosha, were recently identified22,23. The
3ʹ end of the stem–loop precursor of these intronic miRNAs (known as
mirtrons) coincides with the 3ʹ splice site of a small annotated intron
and is cleaved in the same splicing pathway as pre-mRNA in the nucleus
instead of by Drosha. Subsequently, the mirtron precursors, which arereleased by the spliceosome in the shape of a lariat (lasso), are linearized
398

by a de-branching enzyme. They then enter the miRNA-processing
pathway directly (by mimicking the structural features of pre-miRNAhairpins) and are therefore exported to the cytoplasm and processed bya Dicer protein, bypassing Drosha-mediated cleavage.
The imprecision of Drosha or Dicer cleavage could result in the
production of a set of miRNA–miRNA* duplexes with a variety of5ʹ and 3ʹ ends. Most miRNAs in animals form imperfect hybrids withsequences in the target mRNA, with most of the pairing specificity being
provided by the 5ʹ-proximal region of the miRNA (that is, positions 2–8;
also known as the seed region)24,25. Imprecise cleavage either alters theseed sequence or inverts the relative stabilities of the 5ʹ and 3ʹ ends ofthe duplex (see the section ‘Loading and sorting of small RNAs by the
RISC’). The results of recent deep-sequencing studies of small RNAs,
however, indicate that human cells might take advantage of such
imprecise cleavage, because the generation of a diverse set of miRNAs
from a single precursor could be a way of broadening the network offactors and processes that are regulated by miRNAs26–28.
RNaseIII-independent pathways of small RNA biogenesis
In some systems, small RNAs do not seem to be produced in responseto dsRNA, but silencing signals are still amplified. Because these small
RNAs do not arise from dsRNA precursors, RNaseIII enzymes cannot
be involved in their generation. These findings therefore call into
question the definition of RNAi. In this subsection, we describe theknown RNaseIII-independent pathways of small RNA production,
including those that generate piRNAs, 21U-RNAs, and secondarysiRNAs in Caenorhabditis elegans.

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The small RNAs known as piRNAs were named for their ability tobind to a group of Argonaute proteins known as PIWI proteins. As noted
earlier, members of the Argonaute family bind directly to small guide
RNAs and lie at the core of all known RISCs8. Argonaute proteins consistof a variable amino-terminal domain and three conserved domains (thePAZ, middle (MID) and PIWI domains)8,29,30. The 3ʹ end of a small RNA
interacts with the PAZ domain, whereas the phosphate group at the5ʹ end of small RNAs binds to a cleft bridging the MID domain and
the PIWI domain29,30 (see page 405). The PIWI domain has an RNaseH-likefolded structure10 and slicer activity (although some Argonaute proteins
seem to have no slicer activity). There are three phylogenetic groupsof Argonaute proteins29: the AGO subfamily (or AGO clade), named
after the founding member Arabidopsis thaliana ARGONAUTE 1(AGO1); the PIWI subfamily, named after D. melanogaster PIWI
(P-element-induced wimpy testis); and the WAGO (worm-specific
Argonaute) subfamily of C. elegans-specific proteins. PIWI-subfamily
proteins bind to piRNAs31–37 (Table 1). These small RNAs havebeen found only in germ cells, and they are important for germline
development and suppress transposon activity in the germline cells of
mammals, fish and D. melanogaster. They are ~24–31 nucleotides (slightly longer than miRNAs), usually have a uridine at the 5ʹ end and carrya 5ʹ monophosphate. Unlike mammalian miRNAs, but similarly to plant
miRNAs, piRNAs have a 2ʹ-O-methyl (2ʹ-O-Me) modification on the
nucleotide at the 3ʹ end, a modification that is carried out by a HEN1like methyltransferase38–42. If Dicer is mutated, the production of piRNAs
is not affected, indicating that their biogenesis is distinct from that ofmiRNAs and siRNAs and does not involve dsRNA precursors31,42.
The sequencing of small RNAs associated with D. melanogaster PIWIsubfamily proteins (PIWI, Aubergine (AUB) and AGO3)43,44 showed thatpiRNAs associated with AUB and PIWI are derived mainly from theantisense strand of retrotransposons, whereas AGO3-associated piRNAs
arise mainly from the sense strand. AUB- and PIWI-associated piRNAs
show a strong preference for uridine at their 5ʹ ends, whereas AGO3associated piRNAs show a preference for adenosine at nucleotide 10. The
first ten nucleotides of AUB-associated piRNAs can be complementary
to the first ten nucleotides of AGO3-associated piRNAs. In addition,
PIWI-subfamily proteins have slicer activity that allows them to cleave
an RNA substrate opposite position 10 of their bound piRNA32,44. Theseobservations suggest that piRNAs have a self-amplifying loop (Fig. 1),
in which sense piRNAs associated with AGO3 cleave long antisensetranscripts and guide the formation of the 5ʹ end of antisense piRNAs
bound to AUB or PIWI, and vice versa. Thus, in this amplification loop,
which is called the ping-pong cycle43, transposons are both a source ofpiRNAs and a target of piRNA-mediated silencing. After the resultant
cleavage products have been loaded onto another member of the PIWI
subfamily, further (as yet unidentified) nuclease activity generates the3ʹ end of the piRNA, with the specific size of the piRNA determined by
the footprint of the PIWI-subfamily protein on the RNA, a step that seems
to precede 2ʹ-O-Me modification38. In each PIWI-subfamily protein, thePAZ domain might be positioned at a distance from the MID domainthat corresponds to the length of each piRNA. Thus, the PAZ domain
might function as part of a molecular ruler for processing piRNAs of adefined size. Signatures of this amplification cycle are also apparent inzebrafish (Danio rerio) germ cells and in mammalian germ cells before
the pachytene stage of meiosis during spermatogenesis42,45.
PIWI-subfamily proteins and, presumably, their associated piRNAs
are loaded into embryos from the ova8, implying that the piRNAs that
initiate an amplification cycle of piRNA biogenesis (which generates
secondary piRNAs) could be supplied by germline transmission. But
several findings indicate that there must be mechanisms of piRNAbiogenesis other than amplification induced by maternal piRNAs.
First, the amplification cycle in D. melanogaster engages mainly
AGO3 and AUB43,44, but piRNAs are still loaded onto PIWI, which is
spatially separated from these proteins at the subcellular and cell-type
levels32,43,44. Second, piRNAs derived from a particular piRNA clusterin the genome (the flamenco locus) associate almost exclusively withPIWI43. These findings indicate that flamenco-derived piRNAs areproduced by a pathway independent of the amplification loop. Whether
such a piRNA-biogenesis pathway exists remains to be determined.
What at first seemed to be another type of small RNA, 21U-RNA,is found in C. elegans. These small RNAs are precisely 21 nucleotides
and have a bias towards uridine at the 5ʹ end (but not in the remaining
20 nucleotides), and the genetic regions that encode them contain acharacteristic sequence motif ~42 bp upstream of the first nucleotideof the small RNA46. It is possible that these RNAs are derived fromthousands of separate, autonomously expressed, loci that are broadly
scattered in two large regions of one chromosome. They are expressedsolely in the germ line and interact with the PIWI-subfamily protein
PRG-1 (refs 47, 48); therefore, 21U-RNAs are the C. elegans equivalent ofpiRNAs by definition. Like piRNAs, they depend on PRG-1 activity for
their accumulation and are independent of DCR-1 (the C. elegans Dicer
protein) for their production. C. elegans with mutations in prg-1 havea smaller brood and a temperature-sensitive sterile phenotype, whichis consistent with the idea that PIWI-subfamily proteins are involvedin germline maintenance. Like the piRNAs found in mammalian germ
cells in pachytene33,34, 21U-RNAs have remarkable sequence diversity
but lack obvious targets.
Small RNAs with a similar role to piRNAs have also been found in theciliate Tetrahymena thermophila. These scan RNAs (scnRNAs) direct
the elimination of transposon-like DNA sequences and associate witha PIWI-subfamily protein, TWI1 (ref. 8) but, in contrast to piRNAs and
21U-RNAs, are produced by a Dicer-dependent pathway49.
These three examples (piRNAs, 21U-RNAs and scnRNAs) indicatethat the core PIWI and piRNA machinery might have evolved to produce
small RNAs and silence targets by different strategies.
RNA silencing pathways include mechanisms that downregulate
endogenous genes and restrain the expression of selfish or exogenous
genetic material, and these pathways often share common components
such as Dicer. Therefore, there should be competition between different
silencing pathways for particular components. Ways to overcome suchcompetition should also exist; for example, by amplifying a weak silencing
signal. In C. elegans, distinct Argonaute proteins operate at different stagesof RNAi, directing gene silencing in a sequential manner50 — the second
stage of which involves RNaseIII-independent biogenesis of small RNAs.
First, a primary Argonaute protein (such as RDE-1 for exogenous siRNAs(exo-siRNAs) and ERGO-1 for endo-siRNAs) is guided by ‘primary’siRNAs (that is, a first round of siRNAs), which have been generated
from long dsRNAs by DCR-1. Second, the silencing signal is amplified
by the production of ‘secondary’ siRNAs by the action of RNA-dependent
RNA polymerases (RdRPs) (Fig. 1). These secondary siRNAs then binddifferentially to secondary Argonaute proteins (SAGOs, members of
the WAGO subfamily), which mediate downstream silencing. In plants,
RNAs with aberrant features, including lack of a poly(A) tail and lack ofa 5ʹ cap, are copied into double-stranded forms by RdRPs and become
substrates for Dicer, which converts them into siRNA duplexes12. Bycontrast, the C. elegans somatic RdRP mostly produces 21-nucleotide,
single-stranded, 5ʹ-triphosphorylated small RNAs directly from thetarget mRNA in a primer-independent manner without the need for
Dicer-mediated cleavage of dsRNA51–53. Such recruitment of an RdRP
directly to the target mRNA allows dsRNA synthesis without consuming
the siRNAs generated in response to the original trigger, although it isunclear how the 3ʹ end of these secondary siRNAs is formed and what
the molecular ruler is that determines their size.

Blurring of the boundaries between small RNA types
As described above, the three main classes of small RNA — siRNAs,miRNAs and piRNAs — are distinct in their biogenesis and cellularroles. However, recent findings blur these distinctions and show that
there are even more-complex interactions between factors involved insmall RNA biogenesis. Deep sequencing of small RNAs from somatic
tissues and cultured somatic cells in D. melanogaster has uncovered
another class of small RNA, consisting of 3ʹ-methylated, 21-nucleotide
RNAs derived from the D. melanogaster genome. These endogenous
RNAs are derived from transposons and from several loci, including© 2009 Macmillan Publishers Limited. All rights reserved

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NATURE|Vol 457|22 January 2009loci that encode cis-natural antisense transcript pairs, and long stem–
loop structures containing many mismatched pairs in their stems54–57.
In D. melanogaster, distinct Dicer-containing complexes produce
exo-siRNAs and miRNAs58,59. DCR-1 generates miRNAs, acting withits dsRNA-binding protein partner, Loquacious (LOQS)60,61, and themiRNAs are loaded onto AGO1. By contrast, DCR-2, together withits dsRNA-binding protein partner, R2D2 (ref. 62), generates exosiRNAs, which are loaded onto AGO2. Like exo-siRNAs, the recently
discovered endogenous small RNAs are produced by the DCR-2dependent pathway and are loaded onto AGO2, and they are therefore
called endo-siRNAs. However, the generation of many endo-siRNAs
requires LOQS54,56, the dsRBD-containing partner of DCR-1 in themiRNA pathway60,61, but not R2D2, the partner of DCR-2 (ref. 62).
In D. melanogaster deficient in DCR-2 or AGO2, the expression oftransposons increases, so endo-siRNAs might be the main mechanism
for silencing ‘selfish’ genetic elements in somatic cells, which lackthe piRNA pathway. Therefore, endo-siRNAs and piRNAs arefundamentally similar in that they defend organisms against nucleicacid-based ‘parasites’. This finding also shows that D. melanogasterhas two RNAi pathways that repress transposon expression. Mouse
oocytes have also been shown to contain endo-siRNAs. These RNAs
are derived from various sources, including transposons63,64; however,some are processed from overlapping regions of functional genes andtheir cognate pseudogenes. This finding suggests that pseudogenes,
which have been thought to be non-functional protein ‘fossils’, might
regulate the expression of their founder genes.
Although siRNAs and miRNAs are categorized in terms of their origin
rather than their size or function7,12, the discovery of endo-siRNAs makes
it difficult to distinguish between siRNAs and miRNAs. This blurring of
the boundaries between the different types of small RNA has interesting
evolutionary implications. The long stem–loop structures that areprocessed to form endo-siRNAs are reminiscent of the pre-miRNAs inplants. One hypothesis for the evolutionary origin of plant miRNAs isthat new plant miRNA loci might evolve from the inverted duplicationof founder loci, which when transcribed would result in hairpin RNAs12.
These hairpin RNAs would have almost perfect self-complementarityand might be processed by Dicer-like enzymes other than DCL1, themain miRNA-processing enzyme in plants, because DCL1 has limited
activity against such substrates. Subsequent acquisition of mutations
as a result of genetic drift would produce a hairpin with imperfectcomplementarity, which could then be processed by DCL1. Thus, thestem–loop structures from which endo-siRNAs are derived could be
evolutionary intermediates that are gradually transformed into miRNA
precursors. It is possible that such an adaptive switch could also occur
during the evolution of miRNA-encoding genes in D. melanogaster, inwhich DCR-1 would then generate miRNAs instead of endo-siRNAs
being generated by DCR-2.

that thermodynamic asymmetry along small RNA duplex determines
which RNA strand is retained and which is discarded. More specifically,
the strand that has its 5ʹ end at the thermodynamically less stable end of
the small RNA duplex is preferentially loaded into the RISC as the guidestrand, a phenomenon referred to as the asymmetry rule65,66.
For siRNAs, the known interactions between Dicer and the Argonaute
proteins8 indicate that the production of the small RNA and theassembly of the RISC might be physically coupled. For example, in
D. melanogaster, DCR-2 does not simply transfer siRNAs to a distinct
RISC but, instead, forms part of the RISC together with the siRNAs,
indicating that the role of DCR-2 extends beyond the initiation phase.
The loading of siRNA duplexes onto AGO2 is facilitated by the RISCloading complex, which contains DCR-2 and its dsRBD-containing
partner, R2D2 (refs 62, 67). The particular strand of the siRNA duplex
that is loaded onto AGO2 seems to be determined by the orientationof the DCR-2–R2D2 heterodimer on the siRNA duplex68. R2D2 isthought to sense the thermodynamic stability of the siRNA duplexes andbind to the more stable end of the siRNA, whereas DCR-2 is recruited to
the less stable end. The heterodimer probably recruits AGO2 throughan interaction between DCR-2 and AGO2. Previous models haveproposed that the transition from a double-stranded silencing trigger
to a single-stranded one is mediated by an unidentified ATP-dependent
RNA helicase. However, the unwinding of the siRNA duplex and theloading of a single strand into the RISC are facilitated by the slicing of
the unincorporated (passenger) strand by AGO2, a process that does
not require ATP69–71 (Fig. 1). Cleavage in the middle of the passenger
strand, as though the passenger strand were an mRNA target, would be
expected to reduce the annealing temperature and the free energy of
duplex formation, which in turn facilitates the separation of the siRNA
strands. These data support a model in which siRNAs are initially loaded
as duplexes onto an AGO2-containing pre-RISC (Fig. 2).
By contrast, in humans, pre-miRNAs are known to bind to apreformed trimeric complex of AGO2, DICER1 and DICER1’s dsRBDcontaining partner, TRBP72. This complex can cleave target RNAs using
pre-miRNA and can distinguish miRNA from miRNA*, in the absenceof ATP hydrolysis72,73, suggesting that DICER1-mediated cleavage
and sensing of thermodynamic stability occur in series in the AGO2–DICER1–TRBP complex.
This process by which a pre-RISC is converted to a holo-RISC can also
occur by a slicer-independent mechanism. Three of the four Argonaute
proteins in humans (AGO1, AGO3 and AGO4) lack slicer activity but arenonetheless loaded with single-stranded guide siRNAs9,11,28. Similarly,
single-stranded miRNAs are found associated with AGO2 in humans,
despite the expectation that mismatches in the unwound pre-miRNAshould block the passenger-strand cleavage activity of AGO2. Thus,a cleavage-independent (bypass) mechanism for RISC assembly must
exist. RNA helicase A has been identified as a candidate for unwinding
the duplex in this process74.

Loading and sorting of small RNAs by the RISC
In gene silencing pathways initiated by dsRNA precursors, Dicer-mediated cleavage yields small dsRNA intermediates (small RNA duplexes).
These small RNA duplexes must be dissociated into ‘competent’ single
strands in order to function as guides for RISCs. For each small RNA
duplex, only one strand, the guide strand, is loaded onto a specific Argonaute protein and assembled into the active RISC; the other strand, thepassenger strand, is destroyed. Many eukaryotes express more thanone Argonaute protein, and these proteins bind to small RNAs in asequence-independent manner. So how are small RNAs sorted andloaded onto a specific Argonaute protein?
Loading
A small RNA generated from dsRNA precursors is converted from aduplex into a single-stranded form as it is loaded into the RISC. The key
steps in converting the RISC from its precursor form (the pre-RISC),
which contains the small RNA duplex, to its mature form (the holoRISC), which contains the guide strand, are small RNA strand unwinding
and preferential strand selection. The prevalent view of RISC loading is
400

Sorting
Once assembled, RISCs mediate a range of the effector steps in all RNA
silencing mechanisms, from repressing translation to maintaining
genome stability. The specialized functions of RISCs are likely toresult from the particular proteins that associate with each Argonaute
protein. In other words, the different RISC variants are distinguishedby their constituent Argonaute protein. Thus, it is crucial that a specific
set of small guide RNAs is directed to a specific Argonaute protein.
Analyses of how different types of small RNA are channelled to different
Argonaute proteins show that there are multiple mechanisms: thedeterminants for small RNA sorting vary from the structure of the small
RNA duplex to the identity of the 5ʹ nucleotide and the presence andextent of modifications to this nucleotide.
In D. melanogaster, pre-miRNAs are processed by DCR-1, whereas
exo-siRNA duplexes are produced by DCR-2 from long dsRNAs58(Fig. 2a). Small RNAs then seem to be loaded onto either AGO1 or
AGO2, depending on the structure of a small intermediate RNA duplex75.
If the duplex has a bulge in the middle (frequently observed in miRNA© 2009 Macmillan Publishers Limited. All rights reserved

REVIEW INSIGHT

NATURE|Vol 457|22 January 2009asiRNA duplex
5′
3′

miRNA–miRNA*
5′
3′(Perfect match)(Bulge)

DCR-2

DCR-1

R2D2

Figure 2 | Sorting of small RNAs onto distinct Argonaute
proteins. Small RNAs are sorted onto specific Argonaute
proteins, and this process occurs by several mechanisms. a, In
Drosophila melanogaster, small RNAs originating from a duplex
are loaded onto one of two Argonaute proteins (AGO1 or
AGO2), on the basis of the structure of the small RNA duplex. If
the duplex has a mismatch or a bulge in the centre (as miRNAsdo), then the RNA is routed to AGO1. If the duplex is perfectly
matched (as siRNAs are), then the small RNA is routed to AGO2.
This selectivity occurs because the small RNAs are loaded onto
Argonaute proteins from a Dicer-containing complex, and thetwo forms of Dicer, DCR-1 and DCR-2, associate with different
RNA structures. DCR-2 pairs with R2D2, and this heterodimerbinds to highly paired small RNA duplexes but recognizes small
RNA duplexes with a central mismatch only poorly. AGO2
favours binding to DCR-2–R2D2 over binding to the other Dicercontaining complex, DCR-1–LOQS, which binds to small RNAs
with bulges. Further processing into single-stranded
small RNAs is described in Fig. 1. b, Arabidopsis thaliana
miRNAs and trans-acting siRNAs (ta-siRNAs) have a 5ʹ uridine
and preferentially associate with AGO1. By contrast, AGO2 andAGO5 show preferences for small RNAs containing 5ʹ adenosinesand 5ʹ cytidines, respectively. However, it is unlikely that the5ʹ nucleotide is the sole determinant of selective loading in
A. thaliana. c, Secondary endo-siRNAs in Caenorhabditis
elegans, as well in Schizosaccharomyces pombe, have a striking
strand bias in which only the antisense siRNA is loaded onto
Argonaute proteins. These siRNAs correspond to the RNA strand
synthesized by RdRP. In C. elegans, RdRP produces small RNAs
directly from the target mRNA in a primer-independent manner.
Thus, these secondary small RNAs show negative polarity, andthis mechanism reinforces the silencing carried out by
the primary small RNAs.

LOQS

AGO2

AGO1

Unwindase?
U

b
AGO1

AGO2
U-5′

c

ssRNA template
5′

SAGO
3′
5′

AGO5
A-5′

C-5′

RdRP
3′

SAGO
3′
5′

precursors), the small RNA is routed to AGO1. If the duplex is perfectly
matched, the small RNA is channelled to AGO2. This is because the
DCR-2–R2D2 heterodimer, which recruits AGO2 to form the pre-RISC,
binds well to highly paired small RNA duplexes but poorly to duplexes
with central mismatches. Thus, the DCR-2–R2D2 heterodimer not only
determines the polarity of siRNA loading on the basis of thermodynamic
stability rules but also functions as a gatekeeper for AGO2-containing
RISC assembly, promoting the incorporation of siRNAs over miRNAs.
These observations suggest that each siRNA duplex dissociates from theactive site of the Dicer protein after it is produced and is subsequently
recaptured by the DCR-2–R2D2 heterodimer. However, although AGO1
favours binding to small RNA duplexes with central mismatches, a large
proportion of miRNA–miRNA* duplexes with a base-paired central
region still enter into AGO1-containing RISCs55, suggesting that the
AGO1-loading pathway is selective and not a default pathway for small
RNAs rejected by the AGO2 pathway.
The identity of the nucleotide at the 5ʹ end and the extent to which
this nucleotide is phosphorylated also influence which Argonaute

protein the small RNA associates with. In contrast to what is observedin D. melanogaster, processing by Dicer may be uncoupled fromassociation with Argonaute proteins in A. thaliana because, in thisspecies, the miRNAs are all generated by one particular Dicer protein,
DCL1, but are still sorted and loaded onto different Argonaute proteins.
In A. thaliana, miRNAs and trans-acting siRNAs (ta-siRNAs), a classof small RNAs that regulate plant development12, generally have a5ʹ uridine and preferentially associate with AGO1 (ref. 76) (Fig. 2b). By
contrast, AGO2 associates preferentially with small RNAs containing
5ʹ adenosines, and AGO5 prefers 5ʹ cytidines. Interestingly, if the
opposite strand of a miRNA (that is, miRNA*) has a 5ʹ adenosine or a5ʹ cytidine, it is bound to AGO2 or AGO5, respectively. These findings
have led to the hypothesis that the binding affinity of Argonaute proteinsfor small RNAs is determined by the nucleotide at the 5ʹ end. Althoughthese 5ʹ-nucleotide preferences generally hold true for these Argonaute
proteins in plants, exceptions have been observed: the A. thaliana
miRNA known as miR-172 has a 5ʹ adenosine but preferentiallyassociates with AGO1 (ref. 77); and AGO7 preferentially associates with© 2009 Macmillan Publishers Limited. All rights reserved

401

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NATURE|Vol 457|22 January 2009

miR-390, which has a 5ʹ adenosine77. Therefore, the 5ʹ nucleotide does
not seem to be the sole determinant of Argonaute association.
Another mechanism might operate for secondary siRNAs in C. elegans.
These small RNAs are specifically loaded onto SAGOs50. Secondary siRNAs
carry a 5ʹ-triphosphate modification51,52, the hallmark of RdRP products,
which might function as a recognition element for SAGO binding while
excluding binding by a primary Argonaute, such as RDE-1.
Endo-siRNAs in C. elegans (including the secondary siRNAs justmentioned) and Schizosaccharomyces pombe (fission yeast) havea striking strand bias in which only the antisense siRNA strand,corresponding to the RNA strand synthesized by RdRP, is loaded into
Argonaute-containing complexes. Because C. elegans RdRPs produce
small RNAs directly from the target mRNA, in a primer-independent
manner (Fig. 2c), all secondary siRNAs have a negative polarity andfunction to reinforce the silencing of the target mRNA50–52. In S. pombe,
the strand bias is probably the result of a different mechanism. The
physical association of Dicer with an RdRP-containing complex known
as RDRC and an Argonaute-containing complex known as the RNAinduced transcriptional silencing complex (RITS) (see page 413) may
facilitate the loading of siRNAs onto Argonaute proteins in a directionalmanner as Dicer moves along and cleaves the dsRNA products of RdRP,
giving rise to an antisense strand bias. This suggests that the polarity ofDicer processing defines the polarity of the siRNA strand loaded onto
the Argonaute protein.
Argonaute proteins have diversified over evolutionary timescales,
evolving a range of functions8,12. These findings about small RNA sorting
imply that the diversification of the Argonaute proteins is a consequenceof which small RNA they recruit. It is possible that the conformation of
the Argonaute protein dictates which small RNAs it partners, but thestructures of eukaryotic Argonaute proteins will need to be determined
before this can be assessed.

Safeguards in silencing pathways
During RNA silencing, a single non-sequence-specific RNA-binding
protein (Argonaute) is loaded with small guide RNAs with a variety ofsequences, resulting in effector complexes (RISCs). Thus, this system
requires gatekeepers to ensure that Argonaute can bind to small guide
RNAs but not to degraded small RNAs, thereby avoiding ‘off-target’silencing. Such gatekeeper systems seem to depend mainly on structuralfeatures specific for small guide RNAs.
As described earlier, Dicer helps to load siRNAs into the RISC,
preventing siRNAs from diffusing freely in the cytoplasm after
their production. This function of Dicer probably also aids in thediscrimination of genuine siRNAs from various RNA-degradation
products in the cell. Processing by RNaseIII enzymes (such as
Dicer) characteristically yields small RNAs with 5ʹ monophosphates
and 3ʹ two-nucleotide overhangs. The PAZ domain of Argonaute
proteins might, as a first step, distinguish degraded RNAs (derived
from unrelated pathways) from these small RNAs by binding to thecharacteristic 3ʹ overhangs of the small RNAs8,12. In addition, to become
incorporated into the RISC and mediate cleavage of the target mRNA,
the guide strand of an siRNA must have a phosphate group at the5ʹ end78. In humans, the 5ʹ end of siRNAs is phosphorylated by the
enzyme CLP1 (ref. 79), which also has roles in splicing transfer RNAsand forming the 3ʹ ends of mRNAs. Interestingly, both tRNA splicing
and mRNA 3ʹ-end formation occur in the nucleus80,81, suggesting thatsiRNA duplexes with a 5ʹ hydroxyl group are transported to, or diffuse
into, the nucleus and, after phosphorylation by CLP1, are exported tothe cytoplasm and assembled into the RISC.
Amplification of the silencing signal needs to be balanced against
the dangers of amplifying off-target silencing. For example, the slicermediated ping-pong mechanism for piRNA production does notlead to ‘transitive’ RNA silencing (in which RdRPs synthesize siRNAs
complementary to sequences upstream or downstream of the initial
trigger region in the target mRNA). Instead, it leads to conservative
amplification of functional primary piRNA sequences (those inheritedby germline transmission). However, it is conceivable that any off-target
402

events mediated by RdRPs could lead to a chain reaction or transitiveeffect of silencing with deleterious consequences. Thus, there must be
safeguards to prevent the pervasive use of RdRPs. A striking aspect of
RdRP-based trigger amplification is that amplification occurs only
when a target has been engaged, so amplification of the silencing signal
is limited to cases in which there is a real target51,52. In C. elegans, theprocessing of the trigger dsRNA and the loading of primary siRNAs
into the RDE-1-containing complex seem to be inherently inefficient,
limiting the first round of target recognition by RDE-1-containing
complexes and minimizing the risk of amplifying off-target silencing
reactions50. In addition, each secondary siRNA seems to be generated
by non-processive self-termination by RdRP, thereby restricting
transitive effects51–53. Furthermore, secondary siRNAs associate with
SAGOs, which lack catalytic residues for cleaving mRNAs, suggestingthat these complexes cannot generate cleaved substrates for further
amplification, which in turn would prevent them from inducing the
exponential generation of secondary siRNAs50 (but see also ref. 53 fora conflicting viewpoint). SAGOs are also present in limited supply
and thus have a restricted capacity to support multiple simultaneous
silencing reactions.
Another factor is that in C. elegans and S. pombe the RNAi machineryis negatively regulated by a conserved siRNA nuclease called enhanced
RNAi (ERI-1 and Eri1, respectively)82,83. In S. pombe, transgene silencing
is linked to a protein complex resembling the TRAMP complex of
Saccharomyces cerevisiae (budding yeast), which carries out surveillancein the nucleus, targeting aberrant transcripts for degradation by the
exosome84. Thus, RNAi in S. pombe is actively restricted from exerting
its effects throughout the genome and seems to be subject to competition
from RNA quality-control machinery.

Target-sensing modes and effector modes of the RISC
When the RISC is loaded with the guide strand of a small RNA, howdoes it find its target mRNA? Most of the binding energy that tethersa RISC to a target mRNA is from nucleotides in the seed region of thesmall RNA85. It seems that the accessibility of the target site can besensed by the intrinsic, nonspecific affinity of RISC for ssRNA, which
follows the initial specific association between the RISC and the target(through the 5ʹ seed region of the small RNA)86. But the accessibilityof the target site correlates directly with the efficiency of cleavage,
indicating that the RISC cannot unfold structured RNA.
Target mRNAs are present in the cell in complex with ribonucleoproteins (RNPs)87, so target accessibility is also controlled by several
RNA-binding proteins that either mask the target binding site or
facilitate unfolding of the target. Therefore, the function of a RISC seems
to be context-dependent, with its effector mode influenced not only by
the structures of the small-RNA-binding sites on the target but also by theparticular proteins associated with each Argonaute protein. For example,
animal miRNAs silence gene expression by at least three independent
mechanisms through binding sites that are mostly in the 3ʹ untranslated
region of target mRNAs: by cleaving mRNAs, by repressing theirtranslation and/or by promoting mRNA degradation88,89. However, thecontribution of translational repression or mRNA degradation to genesilencing seems to differ for each miRNA–mRNA pair. Thus, the final
outcome of miRNA regulation is probably affected by other proteinsinteracting with the targeted mRNA or RISC and counteracting theeffects of the miRNA, resulting in differential regulation depending on
the proteins present in each tissue90.

Regulation of silencing pathways
So far, the pictures of RNA silencing pathways that we have built up(shown in Figs 1 and 2) are static. To gain further insight into silencing
processes, it is important to incorporate information about how thesepathways are regulated. It is already clear that competition between
different silencing pathways (for example, competition between endosiRNAs and miRNAs for LOQS in D. melanogaster) is a key step inhow each stage of the RNAi mechanism is regulated. Many plant andanimal viruses are known to encode suppressor proteins that block© 2009 Macmillan Publishers Limited. All rights reserved

REVIEW INSIGHT

NATURE|Vol 457|22 January 2009

host RNAi, and therefore silencing, at various stages91. Cellular proteinscan also regulate RNAi. For example, processing to form the humanmiRNA let-7, which is a tumour suppressor and cell-cycle regulator, ispost-transcriptionally inhibited in embryonic cells by the pluripotency
factor LIN28, which seems to block the microprocessor-complexmediated cleavage of pri-let-7 and the Dicer-mediated processing of
pre-let-7 in series92,93. By contrast, in humans, signalling mediated by
the transforming growth factor-β (TGF-β) and bone morphogenetic
protein (BMP) family of growth factors rapidly increases the productionof mature miR-21 (which is oncogenic), by promoting the processing ofpri-miR-21 into pre-miR-21 by DROSHA94. More specifically, TGF-βand BMP-specific signal transducers of the SMAD family are recruited
to pri-miR-21 in complex with the RNA helicase p68, a component of the
microprocessor complex, facilitating the accumulation of pre-miRNA.
In addition, heterogeneous nuclear RNP A1 (hnRNP A1), a well-known
regulator of precursor mRNA splicing, also assists DROSHA to cropand release pre-miR-18 efficiently, perhaps by refolding the hairpin
or by creating a cleavage site for DROSHA through direct binding to
the pri-miRNA95. This implies that some hairpins within pri-miRNAs
might form and be processed only after the binding of a protein withRNA chaperone activity.
The activity of the RISC can also be regulated. In A. thaliana,
the non-protein-coding gene IPS1 (INDUCED BY PHOSPHATESTARVATION 1) contains a motif with sequence complementarity tothe phosphate-starvation-induced miRNA miR-399, but the pairing isinterrupted by a mismatched loop at the expected miRNA cleavage site96.
IPS1 mRNA is not cleaved but, instead, sequesters miR-399. Thus, IPS1
overexpression results in increased accumulation of the target of miR-399,
PHO2 mRNA. The idea of target mimicry introduces unanticipatedcomplexity into the network of RNA-regulatory interactions and raises
the possibility that a large number of mRNA-like non-coding RNAs
recently identified in humans97 could be attenuators of the regulationof small-RNA–Argonaute complexes.

Perspective

1.
2.
3.
4.
5.
6.
7.
8.
9.
10.
11.
12.
13.
14.
15.
16.
17.
18.
19.
20.
21.
22.
23.
24.

Recent studies hint that human cells contain a large number of small
RNAs similar to miRNAs or siRNAs, with the potential to regulate theexpression of almost all human genes. The future challenges in thisfield are clear. Many questions remain to be answered. How many typesof small RNA are there? How are these small RNAs generated? What
are their biological functions? How are these pathways regulated? One
potential problem is that because many types of small RNA are modified
at their 5ʹ and 3ʹ ends98, it is unclear whether the current sequencing
technologies are sampling the entire range of small RNAs present incells. But next-generation sequencing technologies13 should soon help
to uncover the full range of small RNA molecules.
One major challenge will be to identify how specific RNA-binding
proteins affect the final outcome of gene regulation by small RNAs,
given that RNAs in a cell are usually associated with multiple proteinsthat regulate many aspects of gene expression. For example, genomewide in vivo approaches using a combination of immunoprecipitationand high-throughput sequencing will be required to establish protein–mRNA interactions or RNP complex occupancy at certain regions ofmRNA, where expression is suppressed.
Finally, changes in the activity and specificity of silencing pathways could
create quantitative and qualitative genetic variation in gene expression,
thereby generating new gene-expression networks. Such changes might
have contributed to many processes, including human evolution16. Given
that all vertebrates have almost exactly the same number of protein-coding
genes and therefore cannot readily be distinguished in this way, it might
be prophetic that the first small guide RNA to be identified, the C. elegans
miRNA lin-4, has been found to regulate a gene involved in the timing ofdevelopment99,100. In humans, unlike other mammals, the brain tissue of
newborns continues to grow at a similar rate to that of the fetus. This isa good example of a change in developmental timing, and there is much
speculation about whether changes in this rate contributed to the evolutionof humans as a new species.
■

25.
26.
27.
28.
29.
30.
31.

32.
33.
34.
35.
36.

37.
38.
39.
40.
41.
42.
43.

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Acknowledgements We apologize to colleagues whose relevant primary
publications were not cited because of space constraints. We thank Y. Tomari,K. Aoki, Y. Watanabe and all the members of the Siomi laboratory for theircomments and critical reading of the manuscript. Work in our laboratory is
supported by grants from the Ministry of Education, Culture, Sports, Scienceand Technology (MEXT, Japan) and the New Energy and Industrial Technology
Development Organization (Japan). M.C.S. is associate professor of the Global
Centre of Excellence for Human Metabolomics Systems Biology (MEXT).
Author Information Reprints and permissions information is available atwww.nature.com/reprints. The authors declare no competing financial interests.
Correspondence should be addressed to the authors (awa403@sc.itc.keio.ac.jp;
siomim@sc.itc.keio.ac.jp).

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