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RNAi Collection
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TABLE
2
OF
CONTENTS
Introduction: SCIENCE RNAi
Keith Jolliff
3
Introduction: The RNAi World
Guy Riddihough
4
A Species of Small Antisense RNA in
Posttranscriptional Gene Silencing in Plants
Andrew J. Hamilton and David C. Baulcombe
Science 29 October 1999 286: 950–952
8
Argonaute2 Is the Catalytic Engine of Mammalian RNAi
Jidong Liu, Michelle A. Carmell, Fabiola V. Rivas, Carolyn G. Marsden,
J. Michael Thomson, Ji-Joon Song, Scott M. Hammond, Leemor JoshuaTor, Gregory J. Hannon
Science 3 September 2004 305: 1437–1441; published online 29 July
2004
16
Structural Basis for Double-Stranded RNA
Processing by Dicer
Ian J. MacRae, Kaihong Zhou, Fei Li, Adrian Repic, Angela N. Brooks,
W. Zacheus Cande, Paul D. Adams, Jennifer A. Doudna
Science 13 January 2006 311: 195–198
22
RNAi-Mediated Targeting of Heterochromatin by the
RITS Complex
André Verdel, Songtao Jia, Scott Gerber, Tomoyasu Sugiyama, Steven
Gygi, Shiv I. S. Grewal, Danesh Moazed
30 January 2004 303: 672–676; published online 2 January 2004
30
Advertising Supplement: Sigma-Aldrich Corporation
COVER: Leaves from Nicotiana benthamiana showing RNA silencing of gene expression.
Lighter areas are those in which the amount of green fluorescent protein is enhanced as a
result of RNA silencing of a suppressor. [Credit: Shou-Wei Ding] [Image from the cover of
Science 296, 1319 (2002)]
Copyright © 2006 by The American Association for the Advancement of Science. All
rights reserved.
I NTRODUCTION
Science RNAi Collection
Supported by Sigma-Aldrich
RNA interference (RNAi) has emerged as one of the most promising tools for
biological research. It is thought that RNAi may have evolved originally as a defense mechanism against foreign parasitic nucleic acid sequences and functions by
specifically knocking down the activity of target genes. RNAi has become a key
research tool, and is now routinely used by scientists as it offers many advantages
over more traditional knockdown technologies.
The ability of RNAi to target specific genes for silencing has extended its use
into the development of new experimental therapeutic approaches for various diseases, such as cancer, neurodegeneration, and AIDS. Although encouraging results
have been achieved in animal models, the development of new and innovative techniques is crucial to the acceleration of RNAi-based approaches.
Sigma-Aldrich is dedicated to providing the scientific community with the most
advanced and innovative tools in RNAi and functional genomics. This commitment
is exemplified by our collaboration with leading RNAi scientists, including The
RNAi Consortium (TRC), a collaborative group of 11 world-renowned academic
and corporate life science research groups, including the Massachusetts Institute of
Technology, Harvard Medical School, and the Broad Institute.
Our aim is to create a portfolio of comprehensive tools and make them broadly
available to scientists worldwide. Underlying this strategy, our product range in
RNAi and functional genomics has been substantially enhanced by the introduction
of novel technologies, including the first lentiviral MISSION™ shRNA gene family
sets (in collaboration with TRC) and the revolutionary TargeTron™ gene disruption
technology (in partnership with InGex).
The utility of RNAi knockdown approaches is attracting scientists to use and
develop the technology further. The rate of progress is astonishing and the groundbreaking research assembled in this Science RNAi Collection represents the breadth
and depth of progress in RNAi over recent years. We are pleased to have this opportunity to work together with Science to sponsor the Science RNAi Collection.
Keith Jolliff
Global Director of Research Biotech Marketing
Page 2
I NTRODUCTION
The RNAi World
RNA interference (RNAi) is an evolutionary ancient mechanism for silencing
gene expression. It is found in plants and animals and thus was presumably present
in the common ancestor of both. RNAi may have arisen as a host defense mechanism against viruses and other foreign nucleic acids. This role persists: RNAi acts
as an innate and adaptive “immune” response against viruses in plants and animals,
and in turn viruses have evolved counter measures to abrogate the RNAi response.
RNAi forms the mechanistic core of a number of closely related endogenous
RNA silencing pathways: for example, posttranscriptional gene silencing, quelling,
and transcriptional gene silencing. Common to all of them is the presence of short
~21 to 25 nt RNAs generated by the action of members of the Dicer enzyme family.
Those of the RNAi pathway, derived from viral and cellular dsRNA, are known as
small interfering RNAs (siRNAs), whereas a large family of small noncoding RNA
genes found in the genomes of animals (a thousand and counting in humans) and
plants and their viruses are processed into micro-(mi)RNAs.
These various small RNAs are bound by Argonaut (Ago) protein family members, part of the RNA-induced silencing complex (RISC). RISC binds target RNAs
through sequence complementarity with the bound small RNA, and silencing is
effected through degradation (siRNAs) and/or repression of translation of the target
RNA (miRNAs can use both mechanisms). siRNAs can also direct transcriptional gene silencing through the formation of repressive heterochromatin and/or the
methylation of DNA. This mode of silencing provides a form of genome protection, in suppressing the action of transposons, retro-elements and other repeated
sequences.
RNAi-driven resistance to an initial viral infection in a single leaf can spread
throughout the whole plant, and although the nature of the mobile signal is not
yet known, it presumably involves a species of nucleic acid homologous to the
viral target. Similarly, spreading of the RNAi signal beyond the initial site of inoculation is seen in the nematode C. elegans, which can also take up RNA directly
from the environment—these process involving the membrane proteins Sid-1 and
Sid-2. Furthermore, RNAi-induced silencing in C. elegans can be inherited for
many generations, although the inheritance does not itself require genes in the RNAi
pathway.
Tapping into the RNAi pathway has provided the biological research community
with a powerful tool for manipulating gene expression. Genomewide RNAi-based
screens where RNAi-targeted genes are decreased in abundance in tissue culture
cells are now common, with results providing insights into fundamental cellular
processes as well as fueling research into the treatment of human disease (with
specific miRNAs themselves implicated in tumorigenesis). Indeed, siRNAs as drugs
are now moving into clinical trials. Issues of delivery remain a problem, although
various chemical modifications and conjugations can prolong half-lives and enhance
cell up-take. Surprisingly, mucosal tissues are particularly permissive in their ability
to absorb even unmodified RNAs, which retain potent biological activity.
In a few short years RNAi has become a standard technique in the molecular
genetic toolkit and a highly active area of basic research in its own right. It may
also become part of our pharmaceutical armory in the future and will very likely
continue to surprise us with its functions in the cell.
Guy Riddihough
Senior Editor, Science
Page 3
A Species of Small Antisense
RNA in Posttranscriptional
Gene Silencing in Plants
Andrew J. Hamilton and David C. Baulcombe*
Posttranscriptional gene silencing (PTGS) is a nucleotide sequence–
specific defense mechanism that can target both cellular and viral mRNAs.
Here, three types of transgene-induced PTGS and one example of virus-induced PTGS were analyzed in plants. In each case, antisense RNA complementary to the targeted mRNA was detected. These RNA molecules were of a
uniform length, estimated at 25 nucleotides, and their accumulation required
either transgene sense transcription or RNA virus replication. Thus, the 25nucleotide antisense RNA is likely synthesized from an RNA template and may
represent the specificity determinant of PTGS.
Posttranscriptional gene silencing occurs in plants and fungi transformed with foreign
or endogenous DNA and results in the reduced accumulation of RNA molecules with
sequence similarity to the introduced nucleic acid (1, 2). Double-stranded RNA induces
a similar effect in nematodes (3), insects (4), and protozoa (5). PTGS can be suppressed
by several virus-encoded proteins (6) and is closely related to RNA-mediated virus
resistance and cross-protection in plants (7, 8). Therefore, PTGS may represent a natural antiviral defense mechanism and transgenes might be targeted because they, or their
RNA, are perceived as viruses. PTGS could also represent a defense system against
transposable elements and may function in plant development (9–11).
To account for the sequence specificity and posttranscriptional nature of PTGS, it
has been proposed that antisense RNA forms a duplex with the target RNA, thereby
promoting its degradation or interfering with its translation (12). If these hypothetical antisense RNA molecules are of a similar size to typical mRNAs, they would have
been readily detected by routine RNA analyses. However, there have been no reports of
such antisense RNA that is detected exclusively in plants or animals exhibiting PTGS.
Nevertheless, PTGS-specific antisense RNA may exist, but may be too short for easy
detection. We carried out analyses specifically to detect low molecular weight antisense
RNA in four classes of PTGS in plants (13). The first class tested was transgene-induced
PTGS of an endogenous gene (“cosuppression”). We used five tomato lines (T1.1, T1.2,
T5.1, T5.2, and T5.3), each transformed with a tomato 1-aminocyclopropane- 1-carboxylate oxidase (ACO) cDNA sequence placed downstream of the cauliflower mosaic virus 35S promoter (35S). Two lines (T5.2 and T5.3) exhibited PTGS of the
endogenous ACO mRNA (Fig. 1A). Low molecular weight nucleic acids purified from
the five lines were separated by denaturing polyacrylamide gel electrophoresis, blotted,
and hybridized to an ACO sense (antisense-specific) RNA probe (Fig. 1B). A discrete,
ACO antisense RNA (14) of 25 nucleotides (nt) was present in both PTGS lines but
absent from the nonsilencing lines. Twenty-five–nucleotide ACO RNA of sense polarity
and at the same abundance as the 25-nt ACO antisense RNA was also present only in
the PTGS lines (Fig. 1C).
PTGS induced by transgenes can also occur when a transgene does not have
homology to an endogenous gene (1). Therefore, we tested whether this type of PTGS
was also associated with small antisense RNA. We analyzed three tobacco lines carrySainsbury Laboratory, John Innes Centre, Colney Lane, Norwich NR4 7UH, UK.
* To whom correspondence should be addressed. E-mail: david.baulcombe@bbsrc.
ac.uk
Page 4
Fig. 1. Twenty-five–nucleotide ACO antisense and
sense RNA in PTGS lines. (A) Endogenous ACO mRNA
abundance in five tomato lines containing 35S-ACO
transgenes. ACO mRNA was amplified by reverse transcriptase–polymerase chain reaction and detected
by hybridization with labeled ACO cDNA. (B and C)
Low molecular weight RNA from the same five lines and a
30-nt ACO antisense RNA were fractionated, blotted, and
hybridized with either ACO sense RNA (B) or antisense
RNA (C) transcribed from full-length ACO cDNA. The low
hybridization temperature permitted some nonspecific hybridization to tRNA and small ribosomal RNA species,
which constitute most of the RNA mass in these fractions.
The oligonucleotide hybridized only to the antisense-specific probe (B). Twenty-five–nucleotide, PTGS-specific
RNA is indicated.
ing 35S-ßglucuronidase (GUS) transgenes. Two of these lines, T4 (15) and 6b5 (16),
exhibited PTGS of GUS. The third line (6b5×271) tested was produced by crossing
6b5 with line 271 (17), in which there is a transgene suppressor of the 35S promoter in
6b5. There was no PTGS of GUS in 6b5×271 because of the transcriptional suppression of the 35S GUS transgene (18). Hybridization with a GUS-specific probe revealed
that low molecular weight GUS antisense RNA was present in T4 and 6b5 (Fig. 2, lanes
1 and 2) but absent from line 6b5×271 (Fig. 2, lane 3). The amount of antisense RNA
correlated with the extent of PTGS: Line 6b5 has stronger PTGS of GUS than line T4
(18) and had more GUS antisense RNA (Fig. 2). As for PTGS of ACO in tomato, the
GUS antisense RNA was a discrete species of ~25 nt.
In some examples of PTGS, silencing is initiated in a localized region of the
plant. A signal molecule is produced at the site of initiation and mediates systemic
spread of silencing to other tissues of the plant (19, 20). We
investigated whether systemic PTGS of a transgene en- Fig. 2. Twenty-five–nucleotide antisense GUS RNA is dependent
coding the green fluorescent
on transcription from the 35S
protein (GFP) is associated promoter. Twenty-five–nucleotide
with 25-nt GFP antisense GUS antisense RNA was detected
RNA. PTGS was initiated in by hybridization with hydrolyzed
Nicotiana benthamiana ex- GUS sense RNA transcribed from
pressing a GFP transgene by the 3´ 700 base pairs of the
infiltration of a single leaf GUS cDNA.
with Agrobacterium tumefaciens containin GFP sequences in a binary plant transformation vector (19). Two to 3 weeks after this infiltration, the GFP
fluorescence disappeared owing to systemic spread of PTGS as described (11, 20). We detected 25-nt GFP antisense RNA in systemic
tissues exhibiting PTGS of GFP. It was not detected in equivalent leaves of plants that
had not been infiltrated or in nontransformed plants that had been infiltrated with A.
tumefaciens (Fig. 3).
A natural manifestation of PTGS is the RNA-mediated defense induced in virusinfected cells (8). Therefore we investigated whether virus-specific, 25-nt RNA could
be detected in a virus-infected plant. Twenty-five–nucleotide RNA complementary
to the positive strand (genomic) of potato virus X (PVX) was detected 4 days after
inoculation of N. benthamiana and continued to accumulate for at least another 6 days
Page 5
Fig. 3. Twenty-five–nucleotide antisense GFP RNA in systemically silenced tissue. Lower leaves of untransformed N. benthamiana (WT) and N. benthamiana carrying an active 35SGFP transgene (35S-GFP) were infiltrated with A. tumefaciens
containing the same 35S-GFP transgene in a binary vector.
RNA from upper, noninfiltrated leaves of these plants (inf.)
and from equivalent leaves of noninfiltrated plants (–) was
hybridized with GFP sense RNA transcribed from a full-length
GFP cDNA. Only the transgenic N. benthamiana infiltrated with
the A. tumefaciens accumulated 25-nt GFP antisense RNA.
in the inoculated leaf (Fig. 4). Twenty-five–nucleotide PVX RNA accumulated to a
similar extent in systemically infected leaves but was not detected in mock-inoculated
leaves.
Thus, 25-nt antisense RNA, complementary to targeted mRNAs, accumulates in
four types of PTGS. We have detected 25-nt RNA in other examples of PTGS (22),
and never detected 25-nt RNA in the absence of PTGS. This correlation and the properties of 25-nt RNA are consistent with a direct role for 25-nt RNA in PTGS induced
by transgenes or viruses (12). Twenty-five–nucleotide RNA species also serve as molecular markers for PTGS. Their presence could be used to confirm other examples of
transgene- or virus-induced PTGS and perhaps also to identify endogenous genes that
are targeted by PTGS in nontransgenic plants.
The 25-nt antisense RNA species are not degradation products of the target RNA
because they have antisense polarity. A more likely source of these RNAs is the
transcription of an RNA template. This is consistent with the presence of the 25-nt PVX
RNA in PVX-infected cells that do not contain a DNA template (Fig 4, “syst. leaf ”).
The dependency of 25-nt GUS antisense RNA accumulation on sense transcription of a
GUS transgene also supports the RNA template model (Fig. 2). An RNAdependent RNA
Fig. 4. Twenty-five–nucleotide antisense PVX RNA
accumulates during virus replication. RNA was
extracted from inoculated leaves after 2, 4, 6, and
10 days and from systemic (syst.) leaves after 6
and 10 days (d.p.i.: days post inoculation). RNA
was extracted from mock-inoculated leaves after
2 days. Twenty-five–nucleotide PVX antisense RNA
was detected by hybridization with PVX sense RNA
transcribed from a full-length PVX cDNA.
Page 6
polymerase, as required by this model, is required for PTGS in Neurospora crassa (23).
With the present data, we cannot distinguish whether the antisense RNA is made directly as a 25-nt species or as longer molecules that are subsequently processed. The
precise role of 25-nt RNA in PTGS remains to be determined. However, because they
are long enough to convey sequence specificity yet small enough to move through
plasmodesmata, it is possible that they are components of the systemic signal and specificity determinants of PTGS.
1.
2.
3.
4.
5.
6.
7.
8.
9.
10.
11.
12.
13.
14.
15.
References and Notes
H. Vaucheret et al., Plant J. 16, 651 (1998).
C. Cogoni and G. Macino, Trends Plant Sci. 2, 438 (1997).
A. Fire et al., Nature 391, 806 (1998).
J. R. Kennerdell and R. W. Carthew, Cell 95, 1017 (1998).
H. Ngo, C. Tschudi, K. Gull, E. Ullu, Proc. Natl. Acad. Sci. U.S.A. 95, 14687 (1998).
G. Pruss, X. Ge, X. M. Shi, J. C. Carrington, V. B. Vance, Plant Cell 9, 859 (1997); R.
Anandalakshmi et al., Proc. Natl. Acad. Sci. U.S.A. 95, 13079 (1998); K. D. Kasschau
and J. C. Carrington, Cell 95, 461 (1998); G. Brigneti et al., EMBO J. 17, 6739 (1998);
C. Beclin, R. Berthome, J.-C. Palauqui, M. Tepfer, H. Vaucheret, Virology 252, 313
(1998).
J. A. Lindbo, L. Silva-Rosales, W. M. Proebsting, W. G. Dougherty, Plant Cell 5,
1749 (1993).
F. Ratcliff, B. D. Harrison, D. C. Baulcombe, Science 276, 1558 (1997); S. N. Covey, N.
S. Al-Kaff, A. Langara, D. S. Turner, Nature 385, 781 (1997); F. Ratcliff, S. MacFarlane,
D. C. Baulcombe, Plant Cell 11, 1207 (1999).
R. B. Flavell, Proc. Natl. Acad. Sci. U.S.A. 91, 3490 (1994).
R. A. Jorgensen, R. G. Atkinson, R. L. S. Forster, W. J. Lucas, Science 279, 1486
(1998).
O. Voinnet, P. Vain, S. Angell, D. C. Baulcombe, Cell 95, 177 (1998).
D. Grierson, R. G. Fray, A. J. Hamilton, C. J. S. Smith, C. F. Watson, Trends Biotechnol.
9, 122 (1991); W. G. Dougherty and T. D. Parks, Curr. Opin. Cell Biol. 7, 399 (1995);
D. C. Baulcombe and J. J. English, Curr. Opin. Biotechnol. 7, 173 (1996).
Total RNA was extracted from leaves of tomato, tobacco, and N. benthamiana as
described [E. Mueller, J. E. Gilbert, G. Davenport, G. Brigneti, D. C. Baulcombe,
Plant J. 7, 1001 (1995)]. From these preparations, low molecular weight RNA was
enriched by ion-exchange chromatography on Qiagen columns after removal of
high molecular weight species by precipitation with 5% polyethylene glycol 8000–
0.5 M NaCl (for tobacco and N. benthamiana) or by filtration through Centricon
100 concentrators (Amicon) (for tomato). Low molecular weight RNA was separated by electrophoresis through 15% polyacrylamide–7 M urea–0.5× tris-borate
EDTA gels, transferred onto Hybond Nx filters (Amersham), and fixed by ultraviolet
cross-linking. Prehybridization was in 45% formamide, 7% SDS, 0.3 M NaCl, 0.05
M Na2HPO4–NaH2PO4 (pH 7), 1× Denhardt’s solution, and sheared, denatured,
salmon sperm DNA (100 mg/ml) at between 30° and 40°C. Hybridization was in
the same solution with single-stranded RNA probes transcribed with α-32P-labeled
uridine triphosphate. Before addition to the filters in the prehybridization solution,
probes were hydrolyzed to lengths averaging 50 nt. Hybridization was for 16 hours
at 30°C (ACO probes), 35°C (GUS probe), or 40°C (GFP and PVX probes). Sizes of
RNA molecules were estimated by comparison with 33P-phosphorylated DNA oligonucleotides run on the same gels but imaged separately. Additionally, samples
from different types of PTGS, including those shown, were frequently run on the
same gel. Alignment of the filters after hybridization with different specific probes
confirmed that the PTGS-specific signals were identical in size. The probes used are
in each case sequence specific. We have observed no cross-hybridization between
25-nt signals in different PTGS systems using either filter hybridization or RNAase
protection (www.sciencemag.org/feature/data/1042575.shl). We do not have an exact measurement of the amount of 25-nt RNA per cell, but given the short exposure
times routinely used to detect these molecules and taking into account their size,
they are likely to be abundant in cells exhibiting PTGS.
The 25-nt ACO antisense signal was completely abolished by pretreatment with either RNAaseONE (Promega) or NaOH.
S. L. A. Hobbs, T. D. Warkentin, C. M. O. DeLong, Plant Mol. Biol. 21, 17 (1993).
Page 7
16.
17.
18.
19.
20.
21.
22.
23.
24.
T. Elmayan and H. Vaucheret, Plant J. 9, 787 (1996).
H. Vaucheret, C. R. Acad. Sci. Paris 316, 1471 (1993).
J. J. English, G. F. Davenport, T. Elmayan, H. Vaucheret, D. C. Baulcombe, Plant J.
12, 597 (1997).
O. Voinnet and D. C. Baulcombe, Nature 389, 553 (1997).
J.-C. Palauqui and S. Balzergue, Curr. Biol. 9, 59 (1999).
A high-titer, synchronized PVX infection on leaves of untransformed N. benthamiana was initiated by infiltration of single leaves with A. tumefaciens containing a
binary plasmid incorporating a 35S-PVX-GFP sequence. Once transcribed, the PVX
RNA replicon is independent of the 35S-PVX-GFP DNA, replicates to high levels,
and moves systemically through the plant. The A. tumefaciens does not spread
beyond the infiltrated patch and is not present in systemic leaves (20). The GFP reporter in the virus was used to allow visual monitoring of infection progress. We have
obtained similar signals with wild-type PVX inoculated as virions in sap taken from
an infected plant.
The other examples of PTGS tested were in N. benthamiana (spontaneous silencing of
a 35S-GFP transgene), tomato (35S-ACO containing an internal direct and inverted
repeat), petunia (cosuppression of chalcone synthase transgenes and endogenes),
and Arabidopsis thaliana (PTGS of 35S-GFP by a 35S-PVX-GFP transgene).
C. Cogoni and G. Macino, Nature 399, 166 (1999).
We thank D. Grierson, C. DeLong, H. Vaucheret, and R. Hellens for transgenic plants.
We are also grateful to O. Voinnet, D. Bradley, A. Bendahmane, and Ratcliff for helpful
comments and suggestions. This work was carried out under M.A.F.F. licence PHL
24A/2921. Funded by the Biotechnology and Biological Sciences Research Council
and the Gatsby Charitable Foundation.
11 June 1999; accepted 14 September 1999.
Science 286, 950-952 (1999).
Argonaute2 Is the Catalytic Engine of
Mammalian RNAi
Jidong Liu,1* Michelle A. Carmell,1,2* Fabiola V. Rivas,1 Carolyn
G. Marsden,1 J. Michael Thomson,3 Ji-Joon Song1, Scott M.
Hammond,3 Leemor Joshua-Tor,1 Gregory J. Hannon1†
Gene silencing through RNA interference (RNAi) is carried out by RISC,
the RNA-induced silencing complex. RISC contains two signature components, small interfering RNAs (siRNAs) and Argonaute family proteins.
Here, we show that the multiple Argonaute proteins present in mammals
are both biologically and biochemically distinct, with a single mammalian
family member, Argonaute2, being responsible for messenger RNA cleavage activity. This protein is essential for mouse development, and cells
lacking Argonaute2 are unable to mount an experimental response to
siRNAs. Mutations within a cryptic ribonuclease H domain within
Argonaute2, as identified by comparison with the structure of an archeal
Argonaute protein, inactivate RISC. Thus, our evidence supports a model
in which Argonaute contributes “Slicer” activity to RISC, providing the catalytic engine for RNAi.
1
Cold Spring Harbor Laboratory, Watson School of Biological Sciences, 1 Bungtown
Road, Cold Spring Harbor, NY 11724, USA. 2Program in Genetics, Stony Brook
University, Stony Brook, NY 11794, USA. 3Department of Cell and Developmental
Biology, University of North Carolina, Chapel Hill, NC 27599, USA.
*These authors contributed equally to this work.
†To whom correspondence should be addressed. E-mail: [email protected]
Page 8
The presence of double-stranded RNA (dsRNA) in most eukaryotic cells provokes
a sequence-specific silencing response known as RNA interference (RNAi) (1, 2).
The dsRNA trigger of this process can be derived from exogenous sources or transcribed
from endogenous noncoding RNA genes that produce microRNAs (miRNAs) (1, 3).
RNAi begins with the conversion of dsRNA silencing triggers into small RNAs of
~21 to 26 nucleotides (nts) in length (4). This is accomplished by the processing of triggers by specialized ribonuclease III (RNase III)–family nucleases, Dicer and Drosha
(5, 6). Resulting small RNAs join an effector complex, known as RISC (RNA-induced
silencing complex) (7). Silencing by RISC can occur through several mechanisms. In
flies, plants, and fungi, dsRNAs can trigger chromatin remodeling and transcriptional
gene silencing (8–11). RISC can also interfere with protein synthesis, and this is the
predominant mechanism used by miRNAs in mammals (12, 13). However, the best studied mode of RISC action is mRNA cleavage (14, 15). When programmed with a small
RNA that is fully complementary to the substrate RNA, RISC cleaves that RNA at a
discrete position, an activity that has been attributed to an unknown RISC component,
“Slicer” (16, 17). Whether or not RISC cleaves a substrate can be determined by the
degree of complementarity between the siRNA and mRNA, as mismatched duplexes
are often not processed (16). However, even for mammalian miRNAs, which normally
repress at the level of protein synthesis, cleavage activity can be detected with a substrate
that perfectly matches the miRNA sequence (18). This result prompted the hypothesis
that all RISCs are equal, with the outcome of the RISC-substrate interaction being
determined largely by the character of the interaction between the small RNA and its
substrate.
RISC contains two signature components. The first is the small RNA, which cofractionated with RISC activity in Drosophila S2 cell extracts (7), and whose presence
correlated with dsRNA-programmed mRNA cleavage in Drosophila embryo lysates
(14, 15). The second is an Argonaute (Ago) protein, which was identified as a component of purified RISC in Drosophila (19). Subsequent studies have suggested that
Argonautes are also key compnents of RISC in mammals, fungi, worms, protozoans,
and plants (17, 20).
Argonautes are often present as multiprotein families and are identified by two characteristic domains, PAZ and PIWI (21). These proteins mainly segregate into two subfamilies, comprising those that are more similar to either Arabidopsis Argonaute1 or
Drosophila Piwi. The Argonaute family was first linked to RNAi through genetic studies
in Caenorhabditis elegans, which identified Rde-1 as a gene essential for silencing (22).
Our subsequent placement of a Drosophila Argonaute protein in RISC (19) prompted
us to explore the roles of this protein family. Toward this end, we have undertaken both
biochemical and genetic studies of the Ago1 subfamily proteins in mammals.
Mammals contain four Argonaute1 subfamily members, Ago1 to Ago4 [nomenclature as in (23); see fig. S1]. We have previously shown that different Argonaute family
members in Drosophila preferentially associate with different small RNAs, with Ago1
preferring miRNAs and Ago2 siRNAs (24). Recent studies of Drosophila melanogaster
(dm) Ago1 and dmAgo2 mutants have strengthened these conclusions (25). To assess
whether mammalian Ago proteins specialized in their interactions with small RNAs, we
examined Ago-associated miRNA populations by microarray analysis. Ago1-, Ago2and Ago3- associated RNAs were hybridized to microarrays that report the expression
status of 152 human microRNAs. Patterns of associated RNAs were identical within experimental error in each case (Fig. 1A). Additionally, each of the tagged Ago proteins associated similarly with a cotransfected siRNA (Fig. 1C).
Previous studies have used tagged siRNAs to affinity purify Argonaute-containing RISC (17). These preparations, containing mixtures of at least two mammalian
Argonautes, were capable of cleaving synthetic mRNAs that were complementary to
the tagged siRNA. We examined the ability of purified complexes containing individual Argonaute proteins to catalyze similar cleavages. Unexpectedly, irrespective of the
siRNA sequence, only Ago2-containing RISC was able to catalyze cleavage (Fig. 1B
Page 9
Fig. 1. Only mammalian Ago2 can
form cleavage-competent RISC. (A)
The miRNA populations associated with
Ago1, Ago2, and Ago3 were measured by
microarray analysis as described in (44).
The heat map shows normalized log-ratio
values for each data set, with yellow representing increased relative amounts
and blue indicating decreased amounts
relative to the median. The top 25 log
ratios are shown in the expanded region.
In each panel, “control” indicates parallel
analysis of cells transfected with a vector
control. (B) The 293T cells were transfected with a control vector or with vectors
encoding myc-tagged Ago1, Ago2, or
Ago3, along with an siRNA that targets
firefly luciferase. Immunoprecipitates were
tested for siRNA-directed mRNA cleavage as described in (44). Positions of 5´
and 3´ cleavage products are shown. (C)
Immunoprecipitates as in (B) were tested
for in vivo siRNA binding by Northern blotting of Ago immunoprecipitates (44). (D)
Western blots of transfected cell lysates
show similar levels of expression for each
recombinant Argonaute protein.
and fig. S2). All three Ago proteins were similarly expressed and bound similar amounts
of transfected siRNA (Fig. 1, C and D).
These results demonstrated that mammalian Argonaute complexes are biochemically distinct, with only a single family member being competent for mRNA cleavage. To
examine the possibility that Ago proteins might also be biologically specialized, we disrupted the mouse Ago2 gene by targeted insertional mutagenesis (fig. S3 and Fig. 2A)
(26). Intercrosses of Ago2 heterozygotes produced only wild-type and heterozygous
offspring, strongly suggesting that disruption of Ago2 produced an embryonic-lethal
phenotype.
Ago2-deficient mice display several developmental abnormalities beginning approximately halfway through gestation. Both gene-trap and in situ hybridization data of day
9.5 embryos show broad expression of Ago2 in the embryo, with some hot spots of expression in the forebrain, heart, limb buds, and branchial arches (Fig. 2, F and G). The
most prominent phenotype is a defect in neural tube closure (Fig. 2, D and E), often
accompanied by apparent mispatterning of anterior structures, including the forebrain
(Fig. 2, C and D). Roughly half of the embryos display complete failure of neural tube
closure in the head region (Fig. 2E), while all embryos display a wavy neural tube in
more caudal regions. Mutant embryos also suffer from apparent cardiac failure. The
hearts are enlarged and often accompanied by pronounced swelling of the pericardial
cavity (Fig. 2C). By day 10.5, mutant embryos are severely developmentally delayed
compared with wild-type and heterozygous littermates (Fig. 2B). This large difference
in size, like the apparent cardiac failure, may be accounted for by a general nutritional
deficiency caused by yolk sac and placental defects (27), as histological analysis reveals
abnormalities in these tissues.
Not all Argonaute proteins are required for successful mammalian development (28,
29). Thus, it is unclear why Ago2 should be required for development, while other Ago
proteins are dispensable. Ago subfamily members are expressed in overlapping patterns
in humans (30). In situ hybridization demonstrates overlapping expression patterns for
Page 10
Fig. 2. Argonaute2 is essential
for mouse development. (A)
Total RNA from wild-type or mutant embryos was tested for expression of Ago1, Ago2, or Ago3
by RT-PCR. Actin was also examined as a control. (B) At day
E10.5, Ago2-null embryos show
severe developmental delay as
compared with heterozygous
and wild-type littermates. These
embryos also show a variety of
developmental defects, including swelling inside the pericardial membrane (indicated by arrow) (h, heart) (C) and failure to close the neural tube
(D and E). Arrows in (D) indicate the edges of the neural tube that has failed to close.
In caudal regions, where the neural tube does close, it has an abnormal appearance,
being wavy as compared with wild-type embryos (E) (compare wild-type and Ago2
–/–). Ago2 is expressed in most tissues of the developing embryo as measured by in
situ hybridization (F) or by analysis of an Ago2 gene-trap animal (G). In (F), f is forebrain, b is branchial arches, h is heart, and lb is limb bud, all of which are relative hot
spots for Ago2 mRNA. In (G), the left embryo shows similar patterns when staining for
the gene-trap marker, β-galactosidase, proceeds for only a short period. Longer incubation (G, right) gives uniform staining throughout the embryo.
Ago2 and Ago3 in mouse embryos (Fig. 2F and fig. S4). Considered together with
the essentially identical patterns of miRNA binding, our results suggest the possibility that the ability of Ago2 to assemble into catalytically active complexes might be
critical for mouse development. Although most miRNAs regulate gene expression at
the level of protein synthesis, recently miR196 has been demonstrated to cleave the
mRNA encoding HoxB8, a developmental regulator (31). Evolutionary conservation of
an essential cleavage-competent RISC in organisms in which miRNAs predominantly
act by translational regulation raises the possibility that target cleavage by mammalian
miRNAs might be more important and widespread than previously appreciated.
Numerous studies have indicated that experimentally triggered RNAi in mammalian cells proceeds through siRNA-directed mRNA cleavage because in many, but not all,
cases, reiterated binding sites are necessary for repression at the level of protein synthesis [see, for example (13, 32, 33)]. If Ago2 were uniquely capable of assembling into
cleavage-competent complexes in mice, then embryos or cells lacking Ago2 might be
resistant to experimental RNAi. To address this question, we prepared mouse embryo
fibroblasts (MEFs) from E10.5 embryos from Ago2 heterozygous intercrosses. Reverse
transcription polymerase chain reaction (RT-PCR) analysis and genotyping revealed
that we were able to obtain wild-type, mutant, and heterozygous MEF populations.
Importantly, MEFs also express other Ago proteins, including Ago1 and Ago3 (Fig. 3A).
Ago2-null MEFs were unable to repress gene expression in response to an siRNA (Fig.
3B and fig. S5). This defect could be rescued by the addition of a third plasmid that encoded human Ago2 but not by a plasmid encoding human Ago1 (Fig. 3B). In contrast,
responses were intact for a reporter of repression at the level of protein synthesis, mediated by an siRNA binding to multiple mismatched sites (32) (Fig. 3C).
Page 11
Fig. 3. Argonaute2 is essential
for RNAi in MEFs. (A) RT-PCR
of mRNA prepared from wildtype or Ago2⫺/⫺ MEFs reveals
consistent expression of Ago1
and Ago3 but a specific lack of
Ago2 expression in the null
Fig. 3. Argonaute2 is essential for RNAi in MEFs. (A) RT-PCR of mRNA prepared from
wild-type or Ago2–/– MEFs reveals consistent expression of Ago1 and Ago3 but a specific lack of Ago2 expression in the null MEF. Actin mRNA serves as a control. (B) Wildtype and mutant MEFs were cotransfected with plasmids encoding Renilla and firefly
luciferases, either with or without firefly siRNA. Ratios of firefly to Renilla activity, normalized to 1 for the no-siRNA control, were plotted. For each genotype, the ability of Ago1
and Ago2 to rescue suppression was tested by cotransfection with expression vectors
encoding each protein as indicated. (C) NIH-3T3 cells, wild-type MEFs, or Ago2 mutant MEFs were tested as described in (B) (except that Renilla/firefly ratios are plotted)
for their ability to suppress a reporter of repression at the level of protein synthesis. In
this case, the Renilla luciferase mRNA contains multiple imperfect binding sites for
a CXCR4 siRNA. Cells were transfected with a mixture of firefly and Renilla luciferase
plasmids with or without the siRNA.
Because Ago2 is exceptional in its ability to form cleavage-competent complexes, we
set out to map the determinants of this capacity. Deletion analysis indicated that an intact Ago2 was required for RISC activity (fig. S6). We therefore used the sequence of
highly conserved but cleavage-incompetent Ago proteins as a guide to the construction
of Ago2 mutants. A series of point mutations included H634P, H634A, Q633R, Q633A,
H682Y, L140W, F704Y, and T744Y. Whereas all of these mutations retain siRNA-binding activity and most retain cleavage activity, changes at Q633 and H634 have a profound effect on target cleavage (Fig. 4). Both the Q633R and H634P mutations, in which
residues were changed to corresponding residues in Ago1 and Ago3, abolished catalysis.
Changing H634 to A also inactivated Ago2, whereas a similar change, Q633A, was permissive for cleavage. Thus, even relatively conservative changes can negate the ability
of Ago2 to form cleavage-competent RISC.
Several possibilities could explain a lack of cleavage activity for Ago2 mutants. Such
mutations could interfere with the proper folding of Ago2. However, this seems unlikely because those same residues presumably permit proper folding in closely related Argonaute proteins, and mutant Ago2 proteins retained the ability to interact with
siRNAs. Alternatively, cleavage-incompetent Ago2 mutants could lose the ability to interact with the putative Slicer. Finally, Ago2 itself might be Slicer, with our conservative
substitutions altering the active center of the enzyme in a way that prevents cleavage.
The last possibility predicted that we might reconstitute an active enzyme with relatively pure Ago2 protein. We immunoaffinity purified Ago2 from 293T cells and attempted to reconstitute RISC in vitro. Incubation with the double-stranded siRNA produced no appreciable activity, whereas Ago2 could be successfully programmed with
single-stranded siRNAs to cleave a complementary substrate (Fig. 5A). Formation of
the active enzyme was unaffected by first washing the immunoprecipitates with up to
2.5 M NaCl or 1 M urea. A 21-nt single-stranded DNA was unable to direct cleavage
(Fig. 5A). Programming could be accomplished with different siRNAs that direct activity against different substrates (fig. S7). RISC is formed though a concerted assembly
process in which the RISC-loading complex (RLC) acts in an adenosine triphosphate
(ATP)–dependent manner to place one strand of the small RNA into RISC (34–36). In
vitro reconstitution occurs in the absence of ATP, which suggests that Ago2 could be
Page 12
programmed with siRNAs without a need for the normal assembly process (Fig. 5A).
However, in vitro reconstitution of RISC still requires the essential characteristics of
an siRNA. For example, single-stranded siRNAs that lack a 5´ phosphate group cannot
reconstitute an active enzyme.
Although consistent with the possibility that the catalytic activity of RISC is carried within Ago2, these results do not rule out the possibility that a putative Slicer copurifies with Ago2. To demonstrate more conclusively that Ago2 is Slicer, we turned to the
crystal structure of an Argonaute protein from an archebacterium, Pyrococcus furiosus
(37). This structure revealed that the PIWI domain folds into a structure analogous to
the catalytic domain of RNase H and avian sarcoma virus (ASV) integrase. The notion
that such a domain would lie at the center of RISC cleavage is consistent with previous
observations. RNase H and integrases cleave their substrates, leaving 5´ phosphate and
3´ hydroxyl groups through a metal-catalyzed cleavage reaction (38, 39). Notably, previous studies have strongly indicated that the scissile phosphate in the targeted mRNA is
Fig. 4. Mapping the requirements for assembly of cleavagecompetent RISC.Ago1, Ago2, or
mutants of Ago2 were expressed
as myc-tagged fusion proteins in
293T cells. In all cases, expression constructs were cotransfected with a luciferase siRNA.
Western blotting indicated similar expression for each mutant.
Immunoprecipitates containing
individual proteins
were tested
for cleavage activity against a luciferase mRNA (44). Positions of
5´ and 3´ cleavage products are
indicated. SiRNA
binding was
examined for each mutant by
Northern blotting of immunoprecipitates or by staining of immunoprecipitates with Sybr Gold
(Molecular Probes, Eugene, Oregon). Representatives for these assays are shown. In
no case did we detect a defect in interaction of mutants with siRNAs.
cleaved via a metal ion in RISC to give the same phosphate polarity (40). Our in vitro
data are consistent with the reconstituted RISC also requiring a divalent metal (fig. S8).
The active center of RNase H and its relatives consists of a catalytic triad of three
carboxylate groups contributed by aspartic or glutamic acid (38, 39). These amino acid
residues coordinate the essential metal and activate water molecules for nucleolytic attack. Reference to the known structure of RNase H reveals two aspartate residues in
the archeal Ago protein present at the precise spatial locations predicted for formation
of an RNase H–like active site (37). These align with identical residues in the human
Ago2 protein (fig. S9). Therefore, to test whether the PIWI domain of Ago2 provides
catalytic activity to RISC, we changed the two conserved aspartates, D597 and D669,
to alanine, with the prediction that either mutation would inactivate RISC cleavage.
Consistent with our hypothesis, the mutant Ago2 proteins were incapable of assembling
into a cleavage-competent RISC in vitro or in vivo, despite retaining the ability to bind
siRNAs (Fig. 5, B to D).
Considered together, our data provide strong support for the notion that
Argonaute proteins are the catalytic components of RISC. First, the ability to form an
active enzyme is restricted to a single mammalian family member, Ago2. This conclusion is supported both by biochemical analysis and by genetic studies in mutant MEFs.
Page 13
Fig. 5. Argonaute2 is a candidate for Slicer. (A) Ago2 protein was immunoaffinity purified from transiently transfected 293T cells. The preparation contained two major proteins (protein gel), in addition to heavy and light chains. These were identified by mass
spectrometry as Ago2 and HSP90. Immunoprecipitates were mixed (44) in vitro with
single- or double-stranded siRNAs or with a 21-nt DNA having the same sequence
as the siRNA. Reconstituted RISC was tested for cleavage activity with a uniformly labeled synthetic mRNA. Positions of 5´ and 3´ cleavage products are noted. Where
indicated, the siRNA was not 5´ phosphorylated and, in one case, ATP was not added
to the reconstitution reaction. (B) Ago2 or Ago2 mutants were assembled into RISC
in vivo by cotransfection with siRNAs, followed by immunoaffinity purification or by in
vitro reconstitution, mixing affinity-purified proteins with single-stranded siRNAs. These
mutants were tested for activity against a complementary mRNA substrate. 5´ and
3´ cleavage products are as in (A). (C and D) Both mutant proteins were expressed at
levels similar to wild-type Ago2 and bound siRNAs as readily. Ago2 (H634P) and Ago2
(Q633R) behave similarly in this assay.
Second, single amino acid substitutions within Ago2 that convert residues to those
present in closely related proteins negate RISC cleavage. Third, the structure of the P.
furiosis Argonaute protein reveals provocative structural similarities between the PIWI
domain and the RNase H domains, providing a hypothesis for the method by which
Argonaute cleaves its substrates. We tested this hypothesis by introducing mutations in
the predicted Ago2 active site. It is extremely unlikely that such mutations could affect
interactions with other proteins, because they are buried within a cleft of Ago.
Our studies indicate that the Argonaute proteins that are unable to form cleavagecompetent RISC differ from Ago2 at key positions that do not include the putative metalcoordinating residues themselves. However, we cannot yet, based either on biochemical
or structural studies, provide a precise explanation for the catalytic defects in these proteins. It is conceivable that Ago1 and Ago3 fail to coordinate the catalytic metal or that
the structure of the active site is distorted sufficiently that a bound metal is unable to access the scissile phosphate. Alternatively, catalytic mechanisms with two metal ions have
been proposed for RNase H (38, 39), which leaves open the possibility that catalytically
inert Ago family members might lack structures essential to bind the second metal ion.
The relationship between the nuclease domain in PIWI and conserved nuclease domains in viral reverse transcriptases, transposases, and viral integrases has potential evolutionary implications. In Drosophila, plants, and C. elegans, the RNAi pathway has a
major role in controlling parasitic nucleic acids such as viruses and transposons (41–43).
The fact that the RNAi machinery shares a core structural domain with viruses and transposons suggests that this nucleic acid immune system may have arisen in part by pirating components from the replication and movement machineries of the very elements
Page 14
that RNAi protects against. This hypothesis is made even more poignant by considering
the role of RNA-dependent RNA polymerases in RNAi, their functional relationship to
viral replicases, and the possibility that the siRNAs themselves might first have served as
primers that enable such replicases to duplicate primordial genomes.
1.
2.
3.
4.
5.
6.
7.
8.
9.
10.
11.
12.
13.
14.
15.
16.
17.
18.
19.
20.
21.
22.
23.
24.
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27.
28.
29.
30.
31.
32.
33.
34.
35.
36.
37.
38.
39.
40.
41.
42.
43.
44.
45.
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Materials and methods are available as supporting material on Science Online.
The authors thank members of the Hannon lab for helpful discussions, Alea Mills for
advice on ES cell work and for providing the library of targeting constructs, Sang
Page 15
Yong Kim for generating chimeras, Kathryn Anderson for insightful discussions and
advice, and Phil Sharp for providing the CXCR4 constructs. M.C. is supported by
the U.S. Army Breast Cancer Research Program, F.V.R. by the Jane Coffin Childs
Memorial Fund, and J.S. by a Bristol Myers Squibb predoctoral fellowship. S.M.H. is
a General Motors Cancer Research Foundation Scholar. This work was supported in
part by grants from NIH (L.J. and G.J.H.).
Supporting Online Material
www.sciencemag.org/cgi/content/full/1102513/DC1
Materials and Methods
Figs. S1 to S9
8 July 2004; accepted 19 July 2004
Published online 29 July 2004;
10.1126/science.1102513
Include this information when citing this paper.
Structural Basis for Double-Stranded
RNA Processing by Dicer
Ian J. MacRae,1,3 Kaihong Zhou,1,3 Fei Li,1 Adrian Repic,1 Angela
N. Brooks,1 W. Zacheus Cande,1 Paul D. Adams,4 Jennifer A.
Doudna1,2,3,4*
The specialized ribonuclease Dicer initiates RNA interference by cleaving double-stranded RNA (dsRNA) substrates into small fragments about 25 nucleotides in length. In the crystal structure of an intact Dicer enzyme, the PAZ domain, a module that binds the end of dsRNA, is separated from the two catalytic
ribonuclease III (RNase III) domains by a flat, positively charged surface. The 65
angstrom distance between the PAZ and RNase III domains matches the length
spanned by 25 base pairs of RNA. Thus, Dicer itself is a molecular ruler that
recognizes dsRNA and cleaves a specified distance from the helical end.
RNA interference (RNAi) is an ancient gene-silencing process that plays a fundamental
role in diverse eukaryotic functions including viral defense (1), chromatin remodeling
(2), genome rearrangement (3), developmental timing (4), brain morphogenesis (5),
and stem cell maintenance (6). All RNAi pathways require the multidomain ribonuclease Dicer (7). Dicer first processes input dsRNA into small fragments called short interfering RNAs (siRNAs) (8), or microRNAs (miRNA) (9), which are the hallmark of
RNAi. Dicer then helps load its small RNA products into large multiprotein complexes
termed RNAinduced silencing complexes (RISC) (10). RISC and RISC-like complexes use the small RNAs as guides for the sequence-specific silencing of cognate genes
through mRNA degradation (11), translational inhibition (12), and heterochromatin formation (13).
Dicer products are typically 21 to 25 nucleotides long, which is the ideal size for a gene
silencing guide, because it is long enough to provide the sequence complexity required
1
Department of Molecular and Cell Biology, 2Department of Chemistry, 3Howard
Hughes Medical Institute, University of California, Berkeley, CA 94720, USA. 4Physical
Biosciences Division, Lawrence Berkeley National Laboratory, Berkeley, CA 94720,
USA.
*To whom correspondence should be addressed: E-mail: [email protected]
Page 16
to uniquely specify a single gene in a eukaryotic genome. Several models have been proposed for how
Dicer generates RNA fragments of this specific size
(14–16), but structural information is lacking. In an
effort to deepen our understanding of the initiation
step of RNAi, we determined the crystal structure
of an intact and fully active Dicer enzyme.
Conservation of a highly active Dicer
in Giardia intestinalis. We identified an open read-
ing frame in Giardia intestinalis that encodes the
PAZ and tandem RNase III domains characteristic
of Dicer (7), but lacks the N-terminal DExD/H
helicase, C-terminal double-stranded RNA binding domain (dsRBD), and extended interdomain
regions associated with Dicer in higher eukaryotes
(Fig. 1A). A recombinant form of this protein possesses robust dicing activity in vitro (Fig. 1B). The
RNA fragments produced by Giardia Dicer are 25
to 27 nucleotides long, which is similar to a class of
small RNAs associated with RNAi-mediated DNA
elimination in Tetrahymena (17) and RNA-direct- Fig. 1. Giardia encodes an aced DNA methylation in plants (18). dsRNA cleavage tive Dicer enzyme. (A) Schematic
by Giardia Dicer is magnesium-dependent, although representation of the primary
several other divalent cations including Mn2+, Ni2+, sequence of human and Giardia
and Co2+ also support catalytic activity (19). The Dicers. (B) Time course of in vipresence of discrete dicing intermediates sepa- tro Giardia Dicer dsRNA cleavrated by intervals of ~25 nucleotides indicates that age assay. RNA product sizes
Giardia Dicer processes dsRNA from the helical end were determined by comparison
with RNase T1 and alkaline hyin a fashion similar to human Dicer (20). However, in
drolysis (OH) sequencing ladcontrast to human Dicer, Giardia Dicer has a low af- ders (lanes 1 and 2). Dicing refinity for its small RNA product (~1 μM) (19) and quires the protein (Dcr) and Mg2+
displays multiple turnover kinetics (20).
(lanes 3 and 4).
Structural overview. We determined the crystal
structure of the full-length Giardia Dicer at 3.3 Å resolution (table S1). The structure reveals an elongated molecule that, when viewed from the front, takes on a shape resembling a hatchet; the RNase III domains form the blade and the PAZ domain makes up
the base of the handle (Fig. 2A). The PAZ domain is directly connected to the RNase
IIIa domain by a long α helix that runs through the handle of the molecule. This ‘‘connector’’ helix is encircled by the N-terminal residues of the protein, which form a platform domain composed of an antiparallel β sheet and three α helices. A large helical
domain bridges the two RNase III domains and forms the back end of the blade. Viewing
Dicer from the side reveals a contiguous flat surface that extends along one face of the
molecule.
Two–metal-ion mechanism of dsRNA cleavage. The two RNase III domains of
Dicer sit adjacent to each other in the blade region and form an internal heterodimer that
is similar to the homodimeric structure of bacterial Rnase III (fig. S1). Although previous
bacterial RNase III crystal structures revealed a single catalytic metal ion in each RNase
III domain (21), subsequent studies implicated two metal ions in the hydrolysis of each
strand of the dsRNA (22).
During our biochemical characterization of Giardia Dicer, we noticed that the enzyme is potently inhibited by trivalent lanthanide cations such as Er3+ (19). Lanthanides
often bind more tightly to cation binding sites than divalent cations do, a property previously used to identify transient Mn2+ binding sites in proteins (23). Inspection of the
anomalous difference electron density map from a crystal derivatized with ErCl3 revealed a pair of Er3+ cations in the active site of each RNase III domain of Giardia Dicer
Page 17
Fig. 2. Crystal structure of Giardia Dicer. (A) Front and side view ribbon representations of Dicer showing the N-terminal platform domain (blue), the PAZ domain (orange), the connector helix (red), the RNase IIIa domain (yellow), the RNase IIIb domain
(green) and the RNase-bridging domain (gray). Disordered loops are drawn as dotted lines. (B) Close-up view of the Dicer catalytic sites; conserved acidic residues
(sticks); erbium metal ions (purple); and erbium anomalous difference electron density
map, contoured at 20σ (blue wire mesh). Dashed lines indicate distances described
in the text.
(Fig. 2B). The prominent Er3+ metal (M1) in each domain resides between four strictly
conserved acidic residues, which make up the previously identified Mn2+ binding site of
bacterial RNase III (21). The second Er3+ binding site (M2) lies adjacent to the first, outside of the acidic residue cluster. The distances between the two Er3+ metals in the RNase
IIIa and IIIb domains are ~4.2 Å and ~5.5 Å , respectively. These distances are similar to
those previously observed in the active site of RNase H (4.1 Å ) (24), avian sarcoma virus
(ASV) integrase (3.6 Å ) (25), the restriction enzyme EcoRV (4.2 Å ) (26), and the group
I intron (3.9 Å ) (27), all of which are thought to use a two–metal-ion mechanism of
catalysis. The 17.5 Å distance between the metal-ion pairs closely matches the width of
the dsRNA major groove. We also observed Mn2+ in all M1 and some M2 sites in crystals
grown in high concentrations of MnCl2. Therefore, we propose that the Er3+ metals seen
in Giardia Dicer denote true catalytic metal-ion binding sites and that Giardia Dicer uses
a two–metal-ion mechanism of catalysis. Given the high level of sequence conservation
throughout the RNase III family, it is likely that all RNase III enzymes, including bacterial RNase III and Drosha, contain similar catalytic metal-ion binding sites.
Structural features of the Dicer PAZ domain. The PAZ domain is an RNA binding module found in Dicers and in the Argonaute family of proteins that are core components of RISC and other siRNA- and miRNA-containing complexes. Previous studies
of PAZ domains from several Argonaute proteins revealed a degenerate oligonucleotide/
oligosaccharide-binding (OB) fold that specifically recognizes dsRNA ends containing a
3´ two-base overhang (28–31). Superposition of the PAZ domains of Giardia Dicer and
human Argonaute1 reveals that the two domains share the same overall fold and 3´ twonucleotide RNA binding pocket (Fig. 3A).
The Dicer PAZ domain contains a large extended loop that is conserved among
Dicer sequences and absent in Argonaute (fig. S1). The Dicer-specific loop dramatically
changes the electrostatic potential and molecular surface surrounding the 3´ overhangbinding pocket relative to the Argonaute PAZ domain (Fig. 3B). The presence of many
basic amino acid residues in the extra loop could substantially affect the way the RNA is
recognized and perhaps handed off to other complexes by each family of proteins.
A model for siRNA formation. The structure of Giardia Dicer immediately
suggests how Dicer enzymes specify siRNA length. Measuring from the active site of
Page 18
Fig. 3. Structural features of the Dicer PAZ domain. (A) Superposition of the Cα atoms of
PAZ domains from Giardia Dicer (orange) and human Argonaute1 (white). Amino acids
forming the 3´ overhang-binding pocket are shown as sticks. (B) Electrostatic surface
representation of the PAZ domains of Giardia Dicer and Argonaute1 (hAGO1). Asterisks
denote 3´ overhang-binding pockets. The RNA in Argonaute1 PAZ structure is drawn
as green sticks.
the RNase IIIa domain to the 3´ overhang-binding pocket in the PAZ domain gives a
distance of ~65 Å (Fig. 4), which matches the length of 25 dsRNA base pairs. To produce a likely model of a Dicer-dsRNA complex, the positions of the metal-ion pairs
bound in each RNase III domain were used to anchor the two scissile phosphates of
an ideal A-form dsRNA helix into the RNase III active sites. This placement positions
the twofold symmetry element of the dsRNA coincident with the pseudo twofold symmetry axis relating the two RNase III domains, which is analogous to how restriction
enzymes typically bind dsDNA substrates (32). Bacterial RNase III has been proposed to
bind dsRNA in a similar fashion (16, 33). Outside of the RNase III region, the modeled
dsRNA extends along a flat surface formed by the platform domain. This surface contains
a large positively charged region that could interact directly with the negatively charged
phosphodiester backbone of the modeled dsRNA helix. The 3´ end of the RNA duplex
falls directly into the 3´ overhang-binding pocket of the PAZ domain, and the 5´ end lies
adjacent to the Dicer-specific PAZ domain loop. There are exactly 25 nucleotides beFig. 4. A model for dsRNA
processing by Dicer.
Front and side views of
a surface representation
of Giardia Dicer with modeled dsRNA. Red and blue
represent acidic and basic
protein surface charge,
respectively. Electrostatic
surface potentials do not
include contributions from
bound metal ions. Putative
catalytic metal ions are
shown as green spheres.
White arrows point to scissile phosphates. Asterisk
denotes PAZ domain 3´
overhang-binding pocket.
Page 19
Fig. 5. Giardia Dicer supports RNAi in vivo.
(A) Overexpression (OE) of Giardia Dicer
rescues the TBZ sensitivity of the S. pombe
Dicer delete (dcrΔ). Growth was assayed
by spotting 10-fold serial dilutions of cultures indicated. (B) Overexpression of
GiardiaDicerrestores transcriptional silencing at centromeres (cen). Transcript levels
were determined by semiquantitative
reverse-transcriptase polymerase chain
reaction. Actin (act) served as an internal
control. bp, base pair.
tween the 3' end of the helix bound to the PAZ domain and the scissile phosphate in the
RNase IIIa domain.
Thus, Dicer is a molecular ruler that measures and cleaves ~25 nucleotides from
the end of a dsRNA. The length of the small RNAs produced by Dicer is set by the
distance between the PAZ and RNase III domains, which is largely a function of the
length of the connector helix. This model of dsRNA processing is consistent with the
proposed architecture of human Dicer based on biochemical studies in which the RNase
IIIa and IIIb domains were shown to produce the siRNA 5´ and 3´ ends, respectively
(16). Furthermore, closing the ends of a dsRNA substrate by hybridization or ligation
greatly diminished dicing activity (20, 34), which may explain why circular viral dsRNA
is resistant to RNAi (35).
Giardia Dicer can support RNAi in fission yeast. Given that Giardia Dicer lacks
some of the domains commonly associated with Dicer enzymes, most notably the Nterminal helicase, we wondered if the structure represents an intact Dicer or merely the
catalytic subunit of a larger complex required for complete Dicer function in vivo. To address this question, we introduced the Giardia Dicer gene into a strain of the fission yeast
Schizosaccharomyces pombe that contains a deletion of its endogenous Dicer (dcrΔ).
Like most Dicer proteins, the S. pombe Dicer contains an N-terminal helicase domain and a C-terminal dsRBD. The S. pombe dcrΔ strain is defective in RNAi and is
hypersensitive to the microtubule-destabilizing drug thiobendazole (TBZ) because of
chromosome missegregation (36). Plasmid expression of S. pombe dcr1+ fully rescued
TBZ sensitivity of the dcrΔ cells. A partial functional rescue of TBZ sensitivity was also
achieved by episomal expression of Giardia Dicer, indicating that Giardia Dicer can
suppress the chromosome segregation defect (Fig. 5A). Furthermore, Giardia Dicer restores silencing of centromeric regions that are aberrantly transcribed in the dcrΔ mutant
(Fig. 5B). These results demonstrate that Giardia Dicer is sufficient to function as an
intact Dicer in vivo.
A conserved architecture in Dicer enzymes. Considering the structural role
played by the connector helix that links the PAZ and RNase III domains (Fig. 2), we
wondered whether larger Dicer proteins found in higher eukaryotes contain an analogous
helix. Sequence alignment of the region directly following the PAZ domain of several
evolutionarily diverse Dicer enzymes reveals a conserved pattern of hydrophobic and
hydrophilic amino acids that is predicted to form a long α helix by secondary structural analysis (fig. S2). All Dicers contain a conserved proline about 11 amino acid residues from the predicted N terminus of the helix. In the crystal structure of Giardia
Dicer, this proline induces a distinct kink that aids in directing the helix toward the
RNase IIIa domain.
Page 20
Most Dicer proteins contain a conserved region of ~100 amino acids termed ‘‘domain of unknown function 283’’ (DUF283), which lies between the helicase and PAZ
domains in the primary sequence. Low but consistent sequence homology between the
N-terminal domain of Giardia Dicer and DUF283 (fig. S3) suggests that in the Dicers of
higher eukaryotes, DUF283 forms a platform structure similar to that of Giardia Dicer.
The conserved Dicer architecture, together with the demonstration that Giardia Dicer
can substitute for S. pombe Dicer in vivo, argues that the mechanism of Dicer-catalyzed
dsRNA processing is conserved. Moreover, these results indicate that all Dicers evolved
from a common ancestral enzyme. Because Giardia is one of the most anciently diverged
members of the eukaryotic kingdom, we may consider that the earliest eukaryotic organisms had a similar Dicer enzyme and therefore were capable of RNAi-like processes.
It will be of evolutionary interest to determine the cellular function of Dicer and RNAi
in Giardia.
The structure of Giardia Dicer also provides new insight into eukaryotic RNase
III enzymes in general. This family of enzymes performs a range of specific cellular
functions involving the cleavage of dsRNA [reviewed in (37)]. The structure of Dicer illustrates how the presence of RNA binding modules, like the PAZ and platform domains,
can impart a specific function to the otherwise nonspecific double-stranded RNase activity of the RNase III dimer (38). This is likely to be the structural paradigm for all
eukaryotic RNase III enzymes that have specific activities and cellular functions.
1.
2.
3.
4.
5.
6.
7.
8.
9.
10.
11.
12.
13.
14.
15.
16.
17.
18.
19.
20.
21.
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24.
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31.
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34.
35.
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We thank members of the Doudna and Berger labs for helpful discussions, A. Fischer
for work with tissue culture, and D. King for mass spectrometry analysis. We are
grateful to C. Ralston and J. Dickert for technical support on beam lines 8.2.1 and
8.2.2 at the Advanced Light Source at the Lawrence Berkeley National Lab. I.J.M. is
a Howard Hughes Medical Institute fellow of the Life Sciences Research Foundation.
This work was supported in part by a grant from NIH (to J.A.D.). Dicer coordinates
and structure factors have been deposited in the Protein Data Bank with accession
code 2FFL.
Supporting Online Material
www.sciencemag.org/cgi/content/full/311/5758/195/DC1
Materials and Methods
Figs. S1 to S5
Table S1
21 October 2005; accepted 7 December 2005
10.1126/science.1121638
Science 311, 195-198 (2006)
Include this information when citing this paper.
RNAi-Mediated Targeting of
Heterochromatin by the RITS Complex
André Verdel,1 Songtao Jia,2 Scott Gerber,1,3
Tomoyasu Sugiyama,2 Steven Gygi,1,3 Shiv I. S. Grewal,2*
Danesh Moazed1*
RNA interference (RNAi) is a widespread silencing mechanism that acts at both
the posttranscriptional and transcriptional levels. Here, we describe the purification of an RNAi effector complex termed RITS (RNA-induced initiation of
transcriptional gene silencing) that is required for heterochromatin assembly in
fission yeast.The RITS complex contains Ago1 (the fission yeast Argonaute homolog), Chp1 (a heterochromatin-associated chromodomain protein), and Tas3
(a novel protein). In addition, the complex contains small RNAs that require
the Dicer ribonuclease for their production. These small RNAs are homologous
to centromeric repeats and are required for the localization of RITS to heterochromatic domains. The results suggest a mechanism for the role of the RNAi
machinery and small RNAs in targeting of heterochromatin complexes and epigenetic gene silencing at specific chromosomal loci.
The fission yeast Schizosaccharomyces pombe contains large stretches of heterochromatin that are associated with telomeres, repetitive DNA elements surrounding centromeres, and with the silent mating-type loci (1). Assembly of heterochromatin at these
loci involves an orchestrated array of chromatin modifications that lead to the recruit1
Department of Cell Biology, 3Taplin Biological Mass Spectrometry Facility, Harvard
Medical School, Boston, MA 02115, USA. 2Laboratory of Molecular Cell Biology,
National Cancer Institute, National Institutes of Health, Bethesda, MD 20892, USA.
*To whom correspondence should be addressed. Email: [email protected],
[email protected].
Page 22
ment of two chromodomain histone-binding proteins Swi6, a homolog of the Drosophila
and mammalian HP1 proteins, and Chp1 (2, 3). The RNAi pathway has also been implicated in regulation at the DNA and chromatin level in Arabidopsis (4–6), Drosophila
(7), and Tetrahymena (8), and in heterochromatin assembly in S. pombe (9, 10).
RNAi silencing is triggered by doublestranded RNA (dsRNA), which is cleaved
by the ribonuclease III (RNase III)–like enzyme Dicer to generate small RNA molecules
of ~22 nucleotides (nt) (11–13). These small interfering RNAs (siRNAs), load onto an
effector complex called RISC (RNA-induced silencing complex) that contains an Argonaute/PIWI family protein and targets cognate mRNAs for inactivation (12–15).
Factors involved in the RNAi pathway in other organisms are required for heterochromatin formation in S. pombe. Deletion of any of these factors, such as Dicer (dcr1+),
Argonaute (ago1+), and RNA-dependent RNA polymerase (rdp1+), disrupts heterochromatin assembly (9, 10). In support of a role for RNAi in heterochromatin assembly,
both DNA strands of the S. pombe centromeric repeats are transcribed (9), and siRNAs
have been identified that match the S. pombe centromeric repeats (16). Moreover, recent experiments suggest that artificial generation of dsRNA from a hairpin construct
can silence homologous sequences by heterochromatin formation in an RNAi-dependent
manner (17). Here, we address the key question of how small RNAs generated by the
RNAi machinery initiate heterochromatin assembly in fission yeast.
To identify factors important for RNAi-mediated targeting of heterochromatin
complexes, we reasoned that such factor(s) would act in early steps in heterochromatin assembly and would be required for the establishment of heterochromatin-specific
histone modification patterns. The Chp1 protein binds to centromeric repeats and is required for methylation of histone H3-K9 and for localization of Swi6 (3, 18). Moreover,
the phenotypes displayed by chp1Δ strains are identical to RNAi mutants. To test
whether Chp1 provides a physical and functional link between RNAi and heterochromatin assembly, we used a tandem affinity purification procedure (TAP) and a TAP tag to
identify factors that interact with Chp1 (Fig. 1). Several protein species of about 65, 90,
100, and 120 kD were specifically purified from the Chp1-TAP strain (Fig. 1A). Mass
Fig. 1. Purification of
Chp1-TAP and identification of associated
proteins. Extracts from
a Chp1-TAP strain and
an untagged control
strain were purified
by the TAP procedure and applied to
a 4 to 12% polyacrylamide gel, which
was stained with colloidal Coomassie blue
(A). The bands in the
Chp1-TAP purification
were excised from the
gel and sequenced
by
tandem
mass
spectrometry (22). The identity of each band is based on multiple sequenced peptides and is indicated on the right. *Residual GST-TEV, the protease used for elution
from the first affinity column. (B) The Chp1-TAP protein was fully functional for silencing
of a centromeric imr::ura4+ reporter gene as indicated by wild-type levels of growth
on 5-FOA medium, which only allows growth when ura4+ is silenced. N/S, nonselective medium. (C) Schematic diagram showing the subunits of the RITS complex and
their conserved motifs. The chromodomain (ChD) in Chp1, the PAZ and PIWI domains
in Ago1, and a region of sequence similarity between Tas3 and the mouse OTT (ovary
testis transcribed) protein are indicated.
Page 23
spectrometry of excised gel bands, as well as protein mixtures, identified the 120- and
100-kD bands as Chp1, the 90-kD band as Ago1, and the 65-kD band as SPBC83.03c, a
previously uncharacterized protein (Fig. 1, A and C; table S1; figs. S1 and S2), which we
named Tas3 (targeting complex subunit 3). The ratio of the 120- and 100-kD bands varies
from experiment to experiment, which suggests that the 100-kD protein is a degradation
product of Chp1.
To verify that Chp1, Ago1, and Tas3 are associated together in a complex, we constructed an S. pombe strain that produced a fully functional Tas3-TAP protein (Fig. 2, A
and B). Affinity purification followed by mass spectrometry sequencing identified Ago1
and Chp1 as Tas3-associated proteins (Fig. 2C, table S1). N- or C-terminally tagged
Ago1 proteins were not functional in centromeric silencing and were not used for purification experiments. However, identical purification profiles of Chp1-TAP and Tas3-TAP
suggests that Chp1, Ago1, and Tas3 are associated together in a complex, which we have
named RITS.
Chp1, as well as Ago1 and other components of the RNAi pathway, have previously been shown to be required for the assembly of heterochromatin and silencing of
reporter genes inserted within heterochromatic domains (9, 10, 19, 20). A tas3 deletion
strain carrying the ura4+ reporter gene inserted at innermost (imr) or outermost (otr)
Fig. 2. Purification of the RITS complex by using a Tas3-TAP strain and the requirement of
tas3+ in silencing and methylation of H3-K9 and Swi6 localization. Western blot showing that (A) the Tas3-TAP and Chp1-TAP proteins are expressed to similar levels and
(B) growth assays showing that Tas3-TAP displays wild-type levels of silencing for a centromeric imrIR::ura4+ reporter gene. (C) Tas3-TAP was purified, and silver-stained protein bands were sequenced by tandem mass spectrometry. *GST-TEV. (D) In tas3Δ cells,
silencing of a ura4+ reporter gene inserted at the centromeric repeats (imr1R::ura4+ and
otr1R::ura4+) is lost, but silencing of the same reporter gene at the silent mating-type interval (Kint2::ura4+) is unaffected. Loss of silencing in sir2Δ, chp1Δ, and ago1Δ is shown
for comparison. Loss of silencing results in loss of growth on counterselective 5-FOA
medium. (E) ChIP experiments showing that in tas3Δ cells methylation of histone H3-K9
and localization of Swi6 to a ura4+ reporter gene inserted at otr1R and imr1R centromeric repeats is abolished. In contrast, deletion of tas3+ has little or no effect on H3-K9
methylation and Swi6 localization (Kin2::ura4+). ChIP analysis and quantification were
performed as described previously (26). The ratios of ura4+ or cen signals to ura4DS/Eminigene signal present in the immunoprecipitated DNA(ChIP) and whole-cell extracts
(WCE) were used to calculate fold enrichment shown underneath each lane.
Page 24
Fig. 3. Dicer-dependent association of RITS with siRNAs. (A)
Small RNAs of ~22 to 25 nt copurify with Chp1-TAP. RNAs isolated from untagged control (–) and Chp1-TAP (+) strains were
3´ end-labeled with [5´-32P]pCp and separated on 15% denaturing urea polyacrylamide gel. Lane 1, [γ-32P]ATP–labeled RNA markers (Ambion);
lanes 2 and 3, labeling of RNA from whole-cell extract (WCE) (~1/2500 of input); lanes
3 and 4, labeling of RNAs after purification. Bracket on the right side indicates the position of small RNAs specifically associated with Chp1-TAP. (B) Copurification of small
RNAs with Tas3-TAP. (C) No small RNAs are associated with RITS purified from dcr1Δ
cells. Parallel purifications were performed from an untagged (control, lane 1) strain as
well as chp1-TAP, dcr1+ (lane 2) and chp1-TAP, dcr1Δ (lane 3) cells, and the associated RNAs were [5´-32P]pCp labeled (compare lanes 2 and 3, bracket). (D) Northern
blot showing that siRNAs associated with RITS hybridize to 32P-labeled probes corresponding to centromeric repeat sequences. RNA from untagged control (lane1) and
Chp1-TAP cells (lane 2), purified as described in (B), was separated on a denaturing
gel and electrotransferred to a nylon membrane (22). DNA oligonucleotides with sequence complementary to the 12 heterochromatic siRNAs identified by Reinhart and
Bartel (16) were 5´ labeled with [γ-32P]ATP and used as probes for the Northern blot.
(E) Southern blot showing that RITS contains siRNAs complementary to the outer centromeric repeats (otr). dg (lanes 2 and 4) and dh (lane 3) repeats, actin (lane 5), and
LTRs (lane 6) were amplified by polymerase chain reaction (PCR) from genomic DNA,
separated on 1.1% agarose gel, and transferred to nylon membrane. 32P-labeled RITS
siRNAs, obtained by labeling RNAs as described in (A), were separated on a denaturing urea gel, eluted, and used as probes for the blot.
centromeric repeats of chromosome 1 (imr1R::ura4+ and otr1R::ura4+, respectively)
displayed a loss of silencing of both reporter genes (Fig. 2D) to an extent similar to
that of the deletion of sir2, chp1, or ago1 (Fig. 2D) (9, 10, 19, 21). Further, chromatin
immunoprecipitation (ChIP) showed that Tas3 was required for H3-K9 methylation and
Swi6 localization of a ura4+ reporter gene inserted at each of the above loci (Fig. 2E).
As is the case for RNAi mutants (10), deletion of tas3+ had little or no effect on silencing or localization of H3-K9 methylation and Swi6 to the ura4+ reporter gene inserted
at the mat locus (Kint2::ura4+) (Fig. 2, D and E). The similarity in phenotypes displayed
by tas3Δ, chp1Δ, and RNAi mutants underscores the importance of Tas3 interaction with
Chp1 and the role of the RITS complex in RNAi-mediated heterochromatin assembly.
Members of the Argonaute family of proteins constitute the core subunit of
RISC, which is associated with small RNA molecules that target it to specific mRNAs
(12, 13). To determine whether the RITS complex is associated with small RNA molecules, we subjected Chp1-TAP or control purifications to phenol-chloroform extraction and precipitated the aqueous phase of the extraction containing any nucleic acid.
The precipitated material was then labeled with [5´-32P]pCp and T4 RNA ligase (22). As
Page 25
Fig. 4. The RNAi pathway is required for localization of RITS to
heterochromatin. (A) ChIP experiments showing that Tas3-TAP is localized to centromeric heterochromatin in an RNAi-dependent manner. Tas3-TAP is associated with
ura4+ inserted at the otr centromeric repeats (otr1::ura4+, left panels)
and with native centromeric repeat
sequences (cen, right panels) in
wild-type (wt) but not ago1Δ, dcr1Δ, or rdp1Δ cells. The ura4DS/E-minigene at the
endogenous euchromatic location is used as a control. (B) The RNAi pathway is required for the localization of Chp1-(Flag)3 to centromeric heterochromatin. (C) Tas3 is
required for the localization of Chp1-(Flag)3 to heterochromatin. Immunoprecipitations
were carried out using a Flag-specific antibody from tas3+ and tas3Δ cells. (D) Tas3 is
associated with ura4+ inserted at the imr centromeric region (imr1::ura4+). WCE, wholecell extract. Fold enrichment values are shown underneath each lane.
shown in Fig. 3A, Chp1-TAP is specifically associated with small RNA molecules ranging in size from ~22 to 25 nt. In contrast, the predominant RNA species prepared from a
whole-cell extract (total RNA) are 70 to 100 nt in size, most likely representing transfer
RNA (tRNA) and 5S RNA (Fig. 3A). RNA species, mainly in the size range of abundant
tRNAs, as well as a small amount of an RNA species of ~25 nt, were present in both
the untagged control and Chp1-TAP purification and represent nonspecific background
binding (Fig. 3A, lanes 2 to 4). Similar results were obtained when the RITS complex
was purified from a strain producing Tas3-TAP (Fig. 3B).
siRNAs are produced by the ribonuclease Dicer (12, 13).We purified the RITS complex
from a strain that carried a deletion of dcr1+, the only S. pombe gene that codes for Dicer.
Deletion of dcr1+ resulted in a loss of small RNA species that specifically copurify with
Chp1-TAP but had no effect on the presence of nonspecific RNA species, which were
also present in the untagged control purification (Fig. 3C). These results indicated that
the small RNA species specifically associated with RITS are siRNAs that are produced
in a Dcr1-dependent manner.
Sequencing of small RNAs from S. pombe has identified a series of small RNA
species that are complementary to the centromeric repeat sequences (16). These small
RNAs have been termed heterochromatic siRNAs and are clustered at two regions within the centromeric repeats, the dh repeats and a region immediately downstream of the
dg repeats. Centromeric siRNAs have been proposed to function in sequence-specific
targeting of homologous DNA regions (i.e., centromeric repeats) for heterochromatin
assembly. To determine whether siRNAs associated with RITS originate from centromeric repeats, we first analyzed RITS-associated RNAs on a Northern blot probed
with a mixture of oligonucleotides derived from the centromeric repeats. These oligonucleotides were specifically designed to hybridize to siRNAs previously identified by
Reinhart and Bartel (16). The 32P-labeled oligonucleotide probes specifically hybridized
Page 26
to RNA species of ~22 to 25 nt in size present in the Chp1-TAP purification but not with
nonspecific RNAs present in the untagged control purification (Fig. 3D).
As a second test for the identities of the siRNAs associated with RITS, we labeled RITS-associated siRNAs with [5´-32P]pCp, then gel purified and used them to
probe a Southern blot containing equal amounts of DNA fragments (ranging in size from
300 to 700 base pairs) corresponding to the dg and dh centromeric repeats, the region
downstream of dg repeats to which siRNAs map (designated dg-D), retrotransposon long
terminal repeats (LTRs) that have been shown to mediate RNAi-dependent gene silencing (17), and DNA fragments corresponding to actin and molecular size markers. The
labeled siRNAs specifically hybridized to dg, dh, and dg-D centromeric sequences (Fig.
3E). No hybridization was detected to LTR, actin, or DNA size markers (Fig. 3E). Our
inability to detect hybridization of RITS-associated siRNAs with LTR sequences may be
due to a relatively lower abundance of LTR siRNAs compared with siRNAs that originate from the centromeric repeats. Together, these experiments show that RITS is associated with siRNAs that originate from processing of centromeric dsRNA transcripts.
We next used S. pombe strains that produced either Tas3-TAP or Chp1-Flag to determine the in vivo chromatin localization of the RITS complex and the requirement for the
RNAi pathway in its localization. It has previously been shown that Chp1 localizes to the
centromeric repeat regions and together with the Clr4 methyltransferase is required for
H3-K9 methylation and Swi6 localization (3). ChIP experiments showed that Tas3-TAP
is similarly localized to a ura4+ reporter gene inserted within in the otr centromeric repeat
region (otr1::ura4+) and centromeric repeat sequences but not to the control mini-ura4
(ura4DS/E) gene at the endogenous euchromatic location (Fig. 4). Tas3-TAP, like Chp1
(18), is also localized to the imr centromeric repeats (Fig. 4D). Furthermore, deletion of
ago1+, dcr1+, or rdp1+ abolished the association of Chp1-Flag and Tas3-TAP with
otr1::ura4+, as well as with centromeric repeat sequences (Fig. 4, A and B). These results
indicated that the RNAi pathway is required for association of the Chp1 and Tas3 subunits of RITS with heterochromatic DNA regions. Our purification of the RITS complex
from dcr1Δ cells showed that the protein subunits of the complex remained associated
together in the absence of siRNAs (fig. S4). The purification results, together with the
ChIP analysis, indicate that the “empty” RITS complex is inactive and can only associate
with its chromosomal target after it is programmed by siRNAs.
We further tested whether Tas3 was required for the localization of Chp1-Flag
to each of the above regions. Deletion of tas3+ abolished the association of Chp1-Flag
with otr1::ura4+, as well as with native cen sequences (Fig. 4C). These results support
the biochemical identification of Tas3 as an integral subunit of RITS and indicate that
it plays an essential role in localizing the complex to heterochromatin.
Our analysis suggests a remarkably direct role for the RNAi machinery in heterochromatin assembly. By analogy to RISC complexes, which use small RNAs as guides
to target specific mRNAs for degradation or translational repression, we propose
that RITS uses siRNAs to recognize and to bind to specific chromosome regions so as
to initiate heterochromatic gene silencing (Fig. 5). Four lines of evidence support this
view. First, RITS contains Ago1, the S. pombe homolog of the Argonaute family of
proteins, which form the common subunit of RISC complexes purified from different
organisms and are thought to be directly responsible for target recognition (12). Second,
RITS is associated with siRNAs that require Dcr1 for their formation and originate
from heterochromatin repeat regions. Thus, this complex contains the expected specificity determinants, i.e., siRNAs, which in other systems have been shown to direct target
recognition (14, 15, 23, 24). Third, at least two subunits of the RITS complex, Chp1
and Tas3, are specifically associated with the expected heterochromatic DNA regions,
which suggests that the complex localizes directly to its target DNA. Fourth, in addition
to Ago1, RITS contains a chromodomain protein, Chp1, which is localized throughout
heterochromatic DNA regions (18) (Fig. 4) and requires the methyltransferase Clr4 and
histone H3-K9 methylation for localization to chromatin (3, 18). Thus, RITS contains
Page 27
Fig. 5. A model for siRNA-dependent
initiation of heterochromatin assembly by
RITS. The RITS complex is programmed
by Dcr1-produced siRNAs to target specific chromosome regions by sequencespecific interactions involving either siRNA-DNA or siRNA-nascent transcript
(blue arrows) base pairing. Nuc, nucleosome; red triangle, K9-methylation on the
amino terminus of histone H3. See text
for further discussion and references.
both a subunit (Ago1) that binds to siRNAs and can function in RNA or DNA targeting by sequence-specific pairing interaction and a subunit (Chp1) that associates with
specifically modified histones and may be involved in further stabilizing its association
with chromatin (Fig. 5).
Mechanisms analogous to the RITS-mediated targeting of heterochromatin complexes are likely to be conserved in other systems. For example, in Tetrahymena, genomewide DNA elimination during macronucleus development requires an Argonaute family
protein, Twi1, and a chromodomain protein, Pdd1, both of which are also required for
H3-K9 methylation and accumulation of small RNAs corresponding to target sequences
(8, 25). Similarly, in Drosophila repeat-induced transcriptional gene silencing requires
an Argonaute family protein, Piwi, and a chromodomain protein, Polycomb (7). Our
results support the hypothesis that Argonaute proteins form the core subunit of a number of different effector complexes that use sequence-specific recognition to target either
RNA or DNA.
1.
2.
3.
4.
5.
6.
7.
8.
9.
10.
11.
12.
13.
14.
15.
16.
17.
18.
19.
20.
References and Notes
S. I. S. Grewal, J. Cell. Physiol. 184, 311 (2000).
S. I. S. Grewal, D. Moazed, Science 301, 798 (2003).
J. F. Partridge, K. S. Scott, A. J. Bannister, T. Kouzarides, R. C. Allshire, Curr. Biol. 12,
1652 (2002).
D. Zilberman, X. Cao, S. E. Jacobsen, Science 299, 716 (2003).
M. Matzke, A. J. M. Matzke, J. M. Kooter, Science 293, 1080 (2001).
F. E. Vaistij, L. Jones, D. C. Baulcombe, Plant Cell 14, 857 (2002).
M. Pal-Bhadra, U. Bhadra, J. A. Birchler, Mol. Cell 9, 315 (2002).
K. Mochizuki, N. A. Fine, T. Fujisawa, M. A. Gorovsky, Cell 110, 689 (2002).
T. A. Volpe et al., Science 297, 1833 (2002).
I. M. Hall et al., Science 297, 2232 (2002).
A. Fire et al., Nature 391, 806 (1998).
G. J. Hannon, Nature 418, 244 (2002).
P. D. Zamore, Science 296, 1265 (2002).
S. M. Hammond, S. Boettcher, A. A. Caudy, R. Kobayashi, G. J. Hannon, Science
293, 1146 (2001).
G. Hutvágner, P. D. Zamore, Science 297, 2056 (2002).
B. J. Reinhart, D. P. Bartel, Science 297, 1831 (2002).
V. Schramke, R. Allshire, Science 301, 1069 (2003).
J. F. Partridge, B. Borgstrom, R. C. Allshire, Genes Dev. 14, 783 (2000).
G. Thon, J. Verhein-Hansen, Genetics 155, 551 (2000).
C. L. Doe et al., Nucleic Acids Res. 26, 4222 (1998).
Page 28
21. G. D. Shankaranarayana, M. R. Motamedi, D. Moazed, S. I. S. Grewal, Curr. Biol. 13,
1240 (2003).
22. Materials and methods are available as supporting material on Science Online.
23. D. S. Schwarz, G. Hutvágner, B. Haley, P. D. Zamore, Mol. Cell 10, 537 (2002).
24. J. Martinez, A. Patkaniowska, H. Urlaub, R. Luhrmann, T. Tuschl, Cell 110, 563
(2002).
25. S. D. Taverna, R. S. Coyne, C. D. Allis, Cell 110, 701 (2002).
26. J. Nakayama, J. C. Rice, B. D. Strahl, C. D. Allis, S. I. S. Grewal, Science 292, 110
(2001).
27. We thank M. Ohi, K. Gould, C. Hoffman, and D. Wolf for gifts of strains and plasmids; members of the Moazed, Grewal, and Reed laboratories for support and
encouragement; R Ohi and El C. Ibrahim for advice; El C. Ibrahim and M. Wahi
for comments on the manuscript; and C. Centrella for technical help. A.V. was
supported by a postdoctoral fellowship from INSERM and is now a fellow of the
Human Frontier Science Programme. This work was supported by grants from the
NIH (S.I.S.G. and D.M.) and a Carolyn and Peter S. Lynch Award in Cell Biology and
Pathology (D.M.). D.M. is a scholar of the Leukemia and Lymphoma Society.
Supporting Online Material
www.sciencemag.org/cgi/content/full/1093686/DC1
Materials and Methods
Figs. S1 to S4
Tables S1 and S2
14 November 2003; accepted 5 December 2003
Published online 2 January 2004;
10.1126/science.1093686
Include this information when citing this paper.
Page 29
MISSION™ TRC shRNA Library: Next Generation RNA Interference
By Stephanie Uder, Henry George, and Betsy Boedeker
Sigma-Aldrich Corporation, St. Louis, MO, USA
Introduction
The technology of RNA interference emerged
in its earliest form following a 1998 study in
Caenorhabditis elegans1 and has since rapidly
evolved to its current form as a revolutionary tool
for studying gene function, biological pathways,
and the physiology of disease. Work and refinement
of the RNAi technology has exploded in recent
years. We now know the basic mechanism of the
endogenous RNAi pathway,2,3 that the pathway is
present in most eukaryotes,4 and how cellular
machinery can be harnessed to silence gene
expression without triggering the cell’s antiviral
response mechanism.5 Further advances have
shown that siRNAs can be expressed from DNA
vectors within the host cell, providing methods for
longer term silencing, inducible silencing, and a
plasmid DNA format that can be replicated for
unlimited supply (compared to synthetic siRNA). In
addition, these vector-based RNAi platforms may be
integrated with viral delivery systems6 allowing the
researcher to perform gene knockdown in a myriad
of cell lines. Studies of endogenous microRNAs
(miRNAs) suggested that synthetic or expressed
miRNA mimics could be used to induce the RNAi
pathway rather than directly using the standard 21
bp siRNA sequence. Short hairpin RNAs (shRNAs)7
are structurally related to miRNA and can be
expressed from pol II or pol III promoters.
solutions. shRNA expression vectors may be
propagated in Escherichia coli and thus provide an
unlimited supply of DNA for transfection. In addition,
such vectors provide selectable markers for stable
shRNA expression and gene silencing. One of the
most attractive features of plasmid-based systems is
the coupling of the technology to viral delivery systems.
Vectors containing appropriate viral packaging
signals and regulatory elements may be used to
package the shRNA sequence into infectious
virions. When appropriately pseudotyped, these
viral particles can transduce a broader spectrum of
cell lines and overcome issues faced in standard
transfection methods. Viral delivery systems have
been extensively studied for gene therapy research
and have thus undergone numerous modifications
for safety and use.
The lentiviral system, pseudotyped with the VSV-G
envelope protein, presents one of the most
attractive systems for viral packaging and delivery
of shRNA constructs. This is due to its broad
tropism and receptor independent delivery, its
ability to integrate into the genome for stable gene
Sense Strand
U6
sigma.com/rnai
cppt
RRE
puroR
SIN/3’ LTR
pLKO.1-puro
(Ψ) Psi
7,091 bp
RSV/5' LTR
f1 ori
pUC ori
ampR
5' -
Sense Strand
3' - UU
Loop
For those facing the above hurdles, DNA vectorbased shRNA methods provide the necessary
Antisense Strand
hPGK
Discussion
As the portfolio of RNAi methods continues to expand,
options become available for even the most complex
systems being studied. Until recently, synthetic siRNA
was the RNAi vehicle most broadly applicable to a
wide variety of systems and applications. However,
obstacles for using synthetic siRNA include being a
non-renewable resource, the transient nature of
silencing, and the difficulty faced in transfecting
primary cells and non-dividing cell lines such as
neurons, lymphocytes and macrophages. In addition,
in vivo knockdown studies are particularly cumbersome.
Loop
CCGGNNNNNNNNNNNNNNNNNNNNNCTCGAGNNNNNNNNNNNNNNNNNNNNNT T T T T
GGCCNNNNNNNNNNNNNNNNNNNNNGAGCTCNNNNNNNNNNNNNNNNNNNNNAAAAA
Antisense Strand
Figure 1. The pLKO.1-puro Vector for transient or stable expression of shRNA.
INNOVATION @ WORK
silencing, and the fact that it does not require a
mitotic event for integration into the genome, which
extends its use to both dividing and non-dividing
cell lines. The lentiviral system is also not known to
elicit immune responses minimizing concerns of offtarget effects and use in in vivo applications.
In an effort to help further the development and
distribution of tools for RNAi research, SigmaAldrich announced its membership and sponsorship
of The RNAi Consortium (TRC) in March 2005. TRC
is a research collaboration between The Broad
Institute and renowned scientists and their
Silencing of MAPK1 Using MISSION shRNA Constructs
Quantitative RT-PCR Results Normalized to GAPDH
72 hours
144 hours
110
Percentage Expression Level
100
90
80
70
60
50
40
30
20
10
0
1
2
3
4
5
Control
Figure 2. The Human Mitogen-Activated Protein Kinase 1 (MAPK 1, NM_138957)
MISSION shRNA set (5 individual hairpins) was used to achieve gene silencing
of MAPK 1.
laboratories from Harvard Medical School,
Massachusetts Institute of Technology, The
Whitehead Institute, Dana Farber Cancer Institute,
Massachusetts General Hospital, Washington
University, and Columbia University. Four other
leading life science research organizations are
sponsoring members in addition to Sigma-Aldrich.
The mission of TRC is to create comprehensive tools
for genomics medicine, make them broadly
available to scientists worldwide, and to pioneer
applications of these tools to the study of disease.
The consortium is creating a comprehensive library
of RNAi reagents with an anticipated completion
date of March 2007. As a scientific collaborator
and commercial partner, Sigma-Aldrich is assisting
the development, manufacturing, and global
distribution of TRC’s human and mouse lentiviral
vector-based shRNA libraries (MISSION™ TRC). The
collection is designed by the Broad Institute of MIT
and Harvard and is in the process of being
expanded to 150,000 clones targeting 15,000
annotated human genes (MISSION TRC-Hs1.0) and
15,000 annotated mouse genes (MISSION TRC-Mm1.0).
Approximately 72,600 clones targeting 10,500
human and 5,300 mouse genes are currently
available. The libraries include a broad range of
gene families, functional classes, and druggable
targets. MISSION shRNA constructs are designed
using a rules-based algorithm for efficient
knockdown and to minimize off-target effects.
Up to five shRNA sequences are individually cloned
into pLKO.1-puro (Figure 1) for broad coverage of
each target gene and varying degrees of knockdown
(Figure 2). The hairpin structure includes an
intramolecular 21 bp stem and 6 base loop that is
recognized and cleaved by the enzyme Dicer upon
expression via the U6 (pol III) promoter in the host
cell. The resulting siRNA duplex then continues in
the RNAi pathway by association with the RNAiinduced Silencing Complex (RISC). The puromycin
resistance marker is present for stable selection in
mammalian cells while the ampicillin resistance
marker provides for plasmid propagation in E. coli.
The constructs may be used for transient or stable
transfection of mammalian cells. In addition,
pLKO.1-puro allows the generation of lentiviral
particles to infect a wide variety of cells, enabling
stable long-term expression of the shRNA.
Acknowledgements
The authors would like to thank David Root and
colleagues at the Broad Institute, Sheila Stewart of
Washington University, and all of TRC for their
scientific consultation and collaboration.
References
1.Fire, A., et al., Potent and specific genetic interference by double-stranded
RNA in Caenorhabditis elegans. Nature, 391, 806-811 (1998).
2.Bernstein, E., et al., Role for a bidentate ribonuclease in the initiation step
of RNA interference. Nature, 409, 363-366 (2001).
3.Hammond, S.M., et al., An RNA-directed nuclease mediates posttranscriptional gene silencing in Drosophila cells. Nature, 404, 293-296 (2000).
4. Hannon, G.J., RNA Interference. Nature, 418, 244-251 (2002).
5. Williams, B.R., PKR; a sentinel kinase for cellular stress. Oncogene, 18,
6112-6120 (1999).
6. Stewart, S.A., et al., Lentivirus-delivered stable gene silencing by RNAi in
primary cells. RNA, 9, 493-501 (2003).
7. Brummelkamp, T.R., et al., A system for stable expression of short
interfering RNAs in mammalian cells. Science, 296, 550-553 (2002).
MISSION is a trademark belonging to Sigma-Aldrich Co. and its affiliate
Sigma-Aldrich Biotechnology LP.
The RNAi Consortium shRNA library is produced and distributed under
license from the Massachusetts Institute of Technology.
Accelerating Customers' Success through Leadership in Life Science, High Technology and Service
S I G M A - A L D R I C H C O R P O R AT I O N • B O X 1 4 5 0 8 • S T. L O U I S • M I S S O U R I 6 3 1 7 8 • U S A
The future of
is here.
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Innovate!
INNOVATION @ WORK
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Sigma is committed to help you innovate.
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Whether you are determining gene function, analyzing signal
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S I G M A - A L D R I C H C O R P O R AT I O N • B O X 1 4 5 0 8 • S T. L O U I S • M I S S O U R I 6 3 1 7 8 • U S A
Member of the RNAi Consortium
MISSION is a trademark belonging to
Sigma-Aldrich Co. and its affiliate
Sigma-Aldrich Biotechnology LP.
The RNAi Consortium shRNA library
is produced and distributed under
license from the Massachusetts Institute
of Technology.
Create!
INNOVATION @ WORK
with Sigma, the new leader in RNAi
create your advantage
Faster siRNA manufacturing? 100% transduction efficiency of shRNA
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• Taking siRNA manufacturing to
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• MISSION™ TRC shRNA libraries,
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Accelerating Customers' Success through Leadership in Life Science, High Technology and Service
S I G M A - A L D R I C H C O R P O R AT I O N • B O X 1 4 5 0 8 • S T. L O U I S • M I S S O U R I 6 3 1 7 8 • U S A
Member of the RNAi Consortium
MISSION is a trademark belonging to
Sigma-Aldrich Co. and its affiliate
Sigma-Aldrich Biotechnology LP.
The RNAi Consortium shRNA library
is produced and distributed under
license from the Massachusetts Institute
of Technology.