Small RNAs serve as the specificity determinant for a collection of regulatory mechanisms known as RNA silencing. Plants use these mechanisms to control the expression of endogenous genes and to suppress unwanted foreign nucleic acids. Several gene families implicated in silencing have undergone expansion and evidence exists for multiple RNA silencing pathways. Recent progress in defining the components of a number of these pathways is examined here.

Introduction

The ability to process double-stranded RNA into small 21–28 nt fragments by an RNaseIII-based enzyme called Dicer [or Dicer-like (DCL)] is widespread among eukaryotic cells. The products of cleavage, termed siRNA (short interfering RNA) or miRNA (micro-RNA) depending on the nature of the double-stranded substrate, carry 5′ phosphates and 2 nt 3′ overhangs, which license them for incorporation into protein effector complexes that regulate gene expression in a sequence-specific manner [1,2]. The known evolutionary diversity speaks of the remarkable number of ways in which this regulation can occur. Small RNAs have been implicated in directing endonucleolytic cleavage of mRNA [3], suppressing protein expression [4], directing double-stranded RNA synthesis by RDRs (RNA-dependent RNA polymerases) [5], mediating DNA methylation/heterochromatin formation [68] and even guiding programmed deletion of DNA [9].

Plants are an ideal system to study small-RNA-mediated regulatory mechanisms because many of the genes implicated in silencing have undergone amplification and specialization. Uncovering the various silencing pathways represented by these gene families is critical for understanding how this proliferation of genes contributes to plant biology. The existence of similar non-redundant parallel pathways leads to the additional question of how much cross-talk occurs between the different avenues of regulation. Are small RNAs destined for a particular application from their inception? If so how is fate enforced? Is there a sharing of small RNA-based information between different pathways? If so, is this sharing regulated? Continued characterization of small RNAs in plants and the genetic requirements for their production/accumulation are proving useful in defining different silencing pathways and their potential points of intersection.

From miRNAs to RISC

miRNAs are encoded by dispersed loci within the genome and arise from precursors (pri-miRNA, primary miRNA), which fold back on themselves to create a partial dsRNA substrate for a DCL enzyme. Plants have genes for four DCL proteins (Table 1), each with an N-terminal RNA-helicase domain followed by tandem RNase III domains (reviewed in [10]). They also have a PAZ (Piwi/Argonaute/Zwille) domain in the central part of the protein, which is a standard feature of animal Dicers (and the signature of the PAZ family of proteins discussed below) and/or two dsRNA-binding domains at the C-terminus. DCL1 is required for most, if not all, miRNA processing as shown by decreased miRNA levels in DCL1 mutants. Null mutations in DCL1 are embryonic lethal [11] and partial loss of function mutants show severe pleiotropic defects consistent with misregulation of miRNA-controlled developmental pathways [12]. A second gene required for miRNA accumulation is HEN1 (Hua enhancer 1), which displays overlapping phenotypes with DCL1 mutants [13].

Table 1
Proven and possible RNA silencing genes in Arabidopsis
 Gene name Locus Function 
Argonautes (ago) AGO1 At1g48410  
 AGO2 At1g31280  
 AGO3 At1g31290  
 AGO4 At2g27040 Chromatin silencing 
 AGO5 At2g27880  
 AGO6 At2g32940  
 AGO7/ZIPPY At1g69440 Developmental timing 
 AGO8 At5g21030  
 AGO9 At5g21150  
 PINHEAD/ZWILLE At5g43810 Meristem identity 
DCL DCL1/CAF/SIN1 At1g01040 miRNA processing 
 DCL2 At3g03300 Viral resistance 
 DCL3 At3g43920 Chromatin silencing 
 DCL4 At5g20320  
RDR RDR1 At1g14700 Viral defence 
 RDR2 At4g11140 Chromatin silencing 
 RDR3 At2g19910  
 RDR4 At2g19920  
 RDR5 At2g19930  
 RDR6/SDE1/SGS2 At3g49500 PTGS viral defence 
Two tandem dsRNA-binding domains HYL1 At1g07900 miRNA biogenesis 
 At2g28380   
 At5g41070   
 At3g62800   
Chromatin level silencing MET1 At5g49160 CpG methylation 
 DDM1 At5g66750 Chromatin remodelling 
 DRM1 At1g28330 CpNpN methylation 
 DRM2 At1g15380 CpNpN methylation 
 CMT3 At1g69770 CpNpG methylation 
Others SGS3/SDE2 At5g23570 PTGS viral defence 
 SDE3 At1g05460 PTGS viral defence 
 SDE4  PTGS/chromatin silencing 
 HEN1 At4g20910 miRNA processing/stability, PTGS, viral defence, chromatin silencing 
 HASTY At3g05040 Exportin 5, miRNA transport? 
 Gene name Locus Function 
Argonautes (ago) AGO1 At1g48410  
 AGO2 At1g31280  
 AGO3 At1g31290  
 AGO4 At2g27040 Chromatin silencing 
 AGO5 At2g27880  
 AGO6 At2g32940  
 AGO7/ZIPPY At1g69440 Developmental timing 
 AGO8 At5g21030  
 AGO9 At5g21150  
 PINHEAD/ZWILLE At5g43810 Meristem identity 
DCL DCL1/CAF/SIN1 At1g01040 miRNA processing 
 DCL2 At3g03300 Viral resistance 
 DCL3 At3g43920 Chromatin silencing 
 DCL4 At5g20320  
RDR RDR1 At1g14700 Viral defence 
 RDR2 At4g11140 Chromatin silencing 
 RDR3 At2g19910  
 RDR4 At2g19920  
 RDR5 At2g19930  
 RDR6/SDE1/SGS2 At3g49500 PTGS viral defence 
Two tandem dsRNA-binding domains HYL1 At1g07900 miRNA biogenesis 
 At2g28380   
 At5g41070   
 At3g62800   
Chromatin level silencing MET1 At5g49160 CpG methylation 
 DDM1 At5g66750 Chromatin remodelling 
 DRM1 At1g28330 CpNpN methylation 
 DRM2 At1g15380 CpNpN methylation 
 CMT3 At1g69770 CpNpG methylation 
Others SGS3/SDE2 At5g23570 PTGS viral defence 
 SDE3 At1g05460 PTGS viral defence 
 SDE4  PTGS/chromatin silencing 
 HEN1 At4g20910 miRNA processing/stability, PTGS, viral defence, chromatin silencing 
 HASTY At3g05040 Exportin 5, miRNA transport? 

The DCL1-mediated mechanism of miRNA biogenesis in plants may differ somewhat from that in flies and mammals. In these organisms, miRNA formation begins in the nucleus with Drosha, which contains tandem RNase III domains but lacks an RNA helicase domain [14,15]. The stem loop containing the miRNA is excised from the rest of the RNA and exported to the cytoplasm as a pre-miRNA, where a second cleavage by Dicer liberates the dsRNA–miRNA duplex [16]. Subsequent unwinding of the duplex then allows incorporation of the mature miRNA into a downstream effector complex as well as degradation of the other strand. Plants do not encode an equivalent enzyme to Drosha and DCL1 localizes to the nucleus [17] raising the possibility that miRNA biogenesis is a one-step process that is completed before miRNA export from the nucleus.

Export of pre-miRNAs from the nucleus in animal cells requires exportin-5 [1820]. Although exportin-5 binds RNA directly, associated proteins can be transported across the nuclear membrane as well [2123]. Consistent with inefficient transport of small RNA regulators from the nucleus, mutation of the exportin-5 homologue in plants (HASTY) causes developmental timing defects [24]. However, HASTY is unlikely to be the only transporter of miRNAs, because the mutants do not display a DCL1-like phenotype.

When liberated from the precursor, plant miRNAs load into a complex capable of directing endonucleolytic cleavage [5,25], which is analogous to the RISC (RNA-induced silencing complex) found in animal cells [3,26,27]. Unlike animal miRNAs, which pair imperfectly with their target RNAs and suppress protein expression, most plant miRNAs match their target well and direct mRNA cleavage. miRNA-containing fractions from animal cells, which suppress protein expression, are capable of directing cleavage if presented with a homologous target, which suggests that in animal cells siRNAs and miRNAs feed into at least the same kind of complex [28,29].

Loading of siRNAs into RISC in Drosophila requires R2D2, a protein with tandem dsRNA-binding domains, shown to directly bind siRNAs and to associate with the Dicer (DCR-2) that generates siRNAs. Arabidopsis HYL1 encodes a nuclear-localized protein with tandem dsRNA-binding domains, produces overlapping dcl1-like phenotypes when defective and is required for the accumulation of a number of miRNAs, suggesting that the protein may play a role analogous to R2D2 in miRNA trafficking [30,31]. If miRNA processing is nuclear, perhaps HYL1 is shuttled out of the nucleus along with miRNA. There are other uncharacterized small proteins with tandem dsRNA-binding domains in Arabidopsis that may also play a role (Table 1). Others may function more like RDE4, an R2D2-like essential component of RNAi in worms, which associates with Dicer but binds longer dsRNA rather than siRNA duplexes [32].

Animal RISC complexes from flies and human cells require argonaute [or PPD (PAZ and PIWI domain)] proteins [33,34]. A series of recent structural studies indicates that the PAZ domain binds the ends of RNase III cleavage products [3538]. The 2 nt 3′ overhang is nestled in a binding pocket, whereas extensive contacts are made with the adjacent seven nucleotides of that strand [38]. The extent of contacts to this strand, as well as binding studies to ssRNA, suggests that the PAZ domain would continue to associate with the RNA after unwinding of the duplex. In the crystal structure, minor contacts are also made with a non-phosphorylated 5′ nucleotide of the opposite strand [38]. Although the contribution of a phosphorylated 5′ nucleotide was not assessed, it is known from other work to be required for incorporation into RISC [2].

AGO1, the founding member of the PPD family, was originally identified in plants from a mutant with pleiotropic effects on development [39]. Subsequent genetic studies demonstrated that both severe and hypomorphic ago1 alleles compromised PTGS (post-transcriptional gene silencing), in which a single-stranded transcript was converted into dsRNA [40,41]. A hypomorphic ago1 allele did not affect silencing triggered by transcription of an inverted repeat, which suggested that AGO1 acted upstream of DCL [42]. Because Arabidopsis carries ten PPD genes (Table 1), it seemed possible that AGO1 helped to produce dsRNA while a different PPD anchored RISC. The only nagging problem was how to explain the rather severe developmental defects of strong ago1 alleles. New data now show that these alleles significantly elevate a number of different miRNA targets [43]. Functionally, this places AGO1 in plant RISC, but another interpretation is that AGO1 has a role in directing miRNA traffic. One line of evidence for this idea is that the stability of some, but not all, miRNAs is compromised in strong ago1 alleles. A second line comes from the observation that in the developing embryo, AGO1 restricts miR165 to the abaxial surface, where it down-regulates PHABULOSA and PHAVOLUTA to control abaxial/adaxial patterning [44]. A strong ago1 allele leads to ectopic adaxial expression of miR165. In spite of increased overlap with its targets, miRNA is unable to guide mRNA degradation. It is as if miR165 has wandered away from the silencing pathway. The two ideas are not mutually exclusive and a role for stabilizing miRNA association with RISC may best explain the influence of AGO1 in miRNA localization.

If AGO1 acts in RISC, how do inverted repeats silence genes in ago1 [42]? The ago1 allele used for this test is hypomorphic and contains residual AGO1-mediated RISC activity as shown by looking at miRNA target levels [43]. This activity, coupled with the high levels of siRNA derived from the inverted repeat, is probably sufficient for silencing. In the PTGS line where silencing was compromised by the same ago1 mutant, the steady-state siRNA levels may be naturally lower or the conversion of sense transcripts into dsRNA and siRNAs may be dependent on AGO1 function. Thus, a working model of the miRNA pathway in plants begins with transcription of the pri-miRNA, processing by DCL1, transport by HYL1, export out of the nucleus and incorporation into AGO1-containing RISC.

A network of siRNA-generating pathways in nucleic acid immunity

siRNAs serve as the specificity determinant of nucleic acid immunity in plants. They were first discovered in virus-infected plant tissue and from plants undergoing PTGS [45]. Sense and anti-sense RNAs accumulated along the length of the silenced RNA, implying that long dsRNA from the virus or transgene was being degraded by a DCL. Northern blots and direct sequencing of siRNAs revealed two size classes, centred on 21 and 24 nt [6,46], which appeared to have different functions in resistance. Intriguingly, viral proteins that block a long-distance sequence-specific silencing signal that moves through the vasculature also block production of 24 nt RNA [6].

Analysis of small RNAs from plants demonstrated the presence of endogenous siRNAs of the same two size classes [5,46]. Some of the silenced loci produced exclusively 24-nt-long siRNAs [6,47], indicating that the two size classes represent distinct pathways. Further evidence for two different processing pathways came from wheat-germ extracts, which produce both size classes of siRNA when seeded with long dsRNA. Production of a 24 nt RNA, but not a 21 nt RNA, was inhibited in the presence of excess 24 nt ds siRNA, which is consistent with product inhibition and two different DCL enzymes [5]. Recent analysis of dcl mutants in Arabidopsis indicates that DCL3 produces the 24 nt siRNA, whereas DCL2 produces a 21 nt RNA from virus infections [47].

In addition to multiple processing pathways there are multiple routes to dsRNA in plants. One pathway of dsRNA production was revealed by screens for Arabidopsis mutants impaired in PTGS [48,49]. In addition to AGO1, mutations were obtained in the RNA-dependent RNA polymerase gene, RDR6 (SDE1/SGS2) [48,50], a coiled-coil domain protein (SGS3/SDE2) [48,50], a DEAD-box RNA helicase (SDE3) [51], and HEN1 [52]. As with AGO1, these genes were proposed to be involved in the production of dsRNA on the basis of two criteria. First, PTGS-specific 21 and 24 nt siRNAs were lost when these genes were mutant and second, virus-induced gene silencing or inverted repeat silencing was still effective [42,48,52].

Plants encode five other RDR genes besides RDR6 (Table 1). The ability to isolate mutations in one implies that the others may produce dsRNA from different templates. In keeping with this, RDR1, which is up-regulated in the plant innate immune response, is required for resistance to TMV and TRV [53], whereas RDR6 is not [48,50]. No evidence currently exists regarding the role of RDR3-5 subfamily, but recent work suggests RDR2 silences endogenous transposons and other repetitive DNA [47].

The evidence for this chromatin level pathway began with the finding that SDE4, a gene initially described as having a partial role in PTGS (Figure 1) [48], was required for methylation and 24 nt siRNA production from the retroelement, AtSN1 [6]. Subsequently, AGO4 [54] RDR2, DCL3 and HEN1 have all been implicated in AtSN1 silencing [47]. Additional silenced loci affected by this pathway have also been identified [47,55], although only a subset requires HEN1 and A604. Viroids, viruses and certain transgenes have long been known to direct methylation and silencing of DNA in plants (termed RNA-directed DNA methylation) [5658]. The hallmark of RNA-directed DNA methylation is methylation of both symmetrical (CpG and CpNpG) and non-symmetrical cytosines (CpNpN, where N is any nucleotide). Methylation at CpG and CpNpG sites can be inherited into the next generation through the action of the maintenance methyl-transferase (MET1) [5961] and the chromodomain containing protein (CMT3) [62] respectively. Methylation at non-symmetrical sites, however, requires a continual source of dsRNA and the de novo methyl-transferase DRM [63,64].

Silencing of GFP (green fluorescent protein) in sde4-1 is partially compromised

Figure 1
Silencing of GFP (green fluorescent protein) in sde4-1 is partially compromised

The 4-week-old plant was photographed under UV light. Tissue that is silenced for GFP appears red due to autofluorescence of chlorophyll.

Figure 1
Silencing of GFP (green fluorescent protein) in sde4-1 is partially compromised

The 4-week-old plant was photographed under UV light. Tissue that is silenced for GFP appears red due to autofluorescence of chlorophyll.

Consistent with a connection between chromatin and silencing, ago4, rdr2 and dcl3 mutants have all been shown to shift modification of histone H3 at AtSN1 from a repressed state to one associated with active chromatin [47,54]. A direct link between the RNAi machinery and modification of the histone code has also been shown from work on Schizosaccharomyces pombe. S. pombe Ago, Dicer and RdRp genes are required for silencing of centromeric repeats [7,8] and silencing of sequences homologous to inverted repeats [65,66]. Recently, a complex (termed RITS for RNA-induced transcriptional gene silencing) has been purified containing siRNA corresponding to silenced loci, Ago, the chromodomain protein, Chp1 [67], and a novel protein termed Tas3. This suggests that PPD-based effector complexes may act like DNA-binding proteins to recruit chromatin-remodelling complexes to specific loci. Intriguingly, these complexes, guided by small RNA, may in turn lead to the production of more small RNA. MET1 and the chromatin-remodelling factor, DDM1, are required for maintenance of PTGS [68] and for AtSN1 siRNA production [69].

Cross-talk between pathways

An immune system that shares information and amplifies a defence response has the best chance of eliminating a pathogen. Like SDE4, RDR2 is partially required for PTGS, which suggests that there is cross-talk between the RDR6 and RDR2 pathways (A.J. Herr and D.C. Baulcombe, unpublished work). The RDR2/SDE4 contribution to silencing must be dependent on a signal from the RDR6 pathway because silencing is completely lost in the rdr6 mutant. Movement of a signal from the cytoplasm to the nucleus has precedence in the observation that cytoplasmically replicating viruses guide methylation of non-transcribed promoters [58]. The traffic may not necessarily go both ways. Abundant 24 nt siRNA directed against a Sine element in Nicotiana benthamiana are unable to confer resistance to a virus carrying a portion of the same Sine element [6].

If an immune system can be saturated, improperly maintaining a strong defence against one pathogen may make an individual plant more susceptible to a different infection. One way to avoid this problem is to condition amplification of the defence response to the presence of the pathogen. One way to avoid this problem is to condition amplification of the defence response to the presence of the pathogen. In keeping with this idea, RDR6 amplifies the silencing response by converting target RNA into a long dsRNA substrate for DCL; not, it seems, by amplifying dsRNA or siRNAs directly [70]. The evidence for this comes from experiments that demonstrate that a dsRNA trigger of silencing corresponding to only a portion of a target gene, leads to RDR6-dependent production of secondary siRNAs from the entire length of the gene – even if the trigger is targeted at the 5′ end of the gene. This does not favour a simple primer-dependent model of RDR6 activity, because antisense siRNAs from the trigger can only prime dsRNA synthesis from target RNA sequences that are 5′ of the trigger [70]. Studies in wheat-germ extracts indicate that ssRNA is readily converted into siRNA even in the absence of a primer [5]. One possibility is that an initial siRNA recognition event might license the entire mRNA for conversion into dsRNA.

An RDR–DCL cycle not only provides a continual supply of siRNAs for RISC, but also destroys some proportion of the target RNA. Whether this cycle is robust enough to eliminate the need for endonucleolytic cleavage by RISC is unclear. One potential danger with a robust RDR–DCL cycle is recruitment of miRNA targets by RDR pathways. To have miRNA-directed cleavage co-exist harmoniously alongside the defence response, there must be mechanisms to ensure that transitivity does not run wild on miRNA targets, where amplification of silencing signal might disrupt the timing or spatial constraints of regulation. Examination of the siRNA database and ability to detect miRNA cleavage products suggest that RDR-dependent transitivity does not take place on miRNA targets [25,47]. Furthermore, transitivity on endogene mRNA does not proceed with the same efficiency as on transgene mRNA [70], suggesting that there are controls in place that distinguish self from non-self.

Perspective

The evidence for at least one miRNA pathway and three different RDR-dependent pathways indicates the diversity of silencing mechanisms in plants (Figure 2). The observations that PINHEAD (AGO10) maintains the proper number of meristem cells [71,72] and ZIPPY (AGO7) ensures proper timing of adult vegetative structures [73] are reminders that there may be two additional small RNA-guided effector complexes that we know little about. In fact, there are probably others, the activities of six more PPD proteins remain unaccounted for. The cloning of small RNAs and their analysis in silencing mutants has been critical for giving shape to the current silencing landscape in plants. One way forward may be defining the small-RNA pathways more rigorously by systematic cloning of small RNAs associated with specific effector complexes.

A model for RNA silencing in Arabidopsis

Figure 2
A model for RNA silencing in Arabidopsis

Evidence exists for at least three branches of RNA silencing: one that acts at the chromatin level, one that processes and effects miRNA regulation and one involved in viral defence and PTGS.

Figure 2
A model for RNA silencing in Arabidopsis

Evidence exists for at least three branches of RNA silencing: one that acts at the chromatin level, one that processes and effects miRNA regulation and one involved in viral defence and PTGS.

Note added in proof (received 3 September 2004)

Two recent papers provide convincing biochemical evidence that argonaute proteins are the sequence-specific endonuclease of RISC [74,75]. A third, reporting the crystal structure of an archaeal argonaute shows that, while the PAZ domain binds the siRNA, the PIWI domain directs cleavage by an RNase-H-like mechanism [76].

Genes: Regulation, Processing and Interference: A Focus Topic at BioScience2004, held at SECC Glasgow, U.K., 18–22 July 2004. Edited by I. McEwan (Aberdeen, U.K.), B. White (Glasgow, U.K.), S. Graham (Glasgow, U.K.), S. Roberts (Manchester, U.K.), A. Sharrocks (Manchester, U.K.), D. Black (Organon, U.K.), S. Newbury (Oxford, U.K.), J. Sayers (Sheffield, U.K.) and A. Lloyd (University College London, U.K.).

Abbreviations

     
  • DCL

    Dicer-like

  •  
  • PAZ

    Piwi/Argonaute/Zwille

  •  
  • PPD

    PAZ and PIWI domain

  •  
  • PTGS

    post-transcriptional gene silencing

  •  
  • RDR

    RNA-dependent RNA polymerases

  •  
  • RISC

    RNA-induced silencing complex

I thank David Baulcombe and past and present members of the Baulcombe Laboratory for stimulating discussions. I am also grateful for the generous financial support of the Gatsby Charitable Trust and a post-doctoral fellowship from Burrough's Wellcome Fund.

References

References
1
Elbashir
S.M.
Harborth
J.
Lendeckel
W.
Yalcin
A.
Weber
K.
Tuschl
T.
Nature (London)
2001
, vol. 
411
 (pg. 
494
-
498
)
2
Schwarz
D.S.
Hutvagner
G.
Haley
B.
Zamore
P.D.
Mol. Cell
2002
, vol. 
10
 (pg. 
537
-
548
)
3
Zamore
P.D.
Tuschl
T.
Sharp
P.A.
Bartel
D.P.
Cell (Cambridge, Mass.)
2000
, vol. 
101
 (pg. 
25
-
33
)
4
Olsen
P.H.
Ambros
V.
Dev. Biol.
1999
, vol. 
216
 (pg. 
671
-
680
)
5
Tang
G.
Reinhart
B.J.
Bartel
D.P.
Zamore
P.D.
Genes Dev.
2003
, vol. 
17
 (pg. 
49
-
63
)
6
Hamilton
A.
Voinnet
O.
Chappell
L.
Baulcombe
D.
EMBO J.
2002
, vol. 
21
 (pg. 
4671
-
4679
)
7
Volpe
T.A.
Kidner
C.
Hall
I.M.
Teng
G.
Grewal
S.I.
Martienssen
R.A.
Science
2002
, vol. 
297
 (pg. 
1833
-
1837
)
8
Hall
I.M.
Shankaranarayana
G.D.
Noma
K.
Ayoub
N.
Cohen
A.
Grewal
S.I.
Science
2002
, vol. 
297
 (pg. 
2232
-
2237
)
9
Mochizuki
K.
Fine
N.A.
Fujisawa
T.
Gorovsky
M.A.
Cell (Cambridge, Mass.)
2002
, vol. 
110
 (pg. 
689
-
699
)
10
Schauer
S.E.
Jacobsen
S.E.
Meinke
D.W.
Ray
A.
Trends Plant Sci.
2002
, vol. 
7
 (pg. 
487
-
491
)
11
Golden
T.A.
Schauer
S.E.
Lang
J.D.
Pien
S.
Mushegian
A.R.
Grossniklaus
U.
Meinke
D.W.
Ray
A.
Plant Physiol.
2002
, vol. 
130
 (pg. 
808
-
822
)
12
Jacobsen
S.E.
Running
M.P.
Meyerowitz
E.M.
Development
1999
, vol. 
126
 (pg. 
5231
-
5243
)
13
Park
W.
Li
J.
Song
R.
Messing
J.
Chen
X.
Curr. Biol.
2002
, vol. 
12
 (pg. 
1484
-
1495
)
14
Filippov
V.
Solovyev
V.
Filippova
M.
Gill
S.S.
Gene
2000
, vol. 
245
 (pg. 
213
-
221
)
15
Lee
Y.
Ahn
C.
Han
J.
Choi
H.
Kim
J.
Yim
J.
Lee
J.
Provost
P.
Radmark
O.
Kim
S.
, et al. 
Nature (London)
2003
, vol. 
425
 (pg. 
415
-
419
)
16
Lee
Y.
Jeon
K.
Lee
J.T.
Kim
S.
Kim
V.N.
EMBO J.
2002
, vol. 
21
 (pg. 
4663
-
4670
)
17
Papp
I.
Mette
F.
Aufsatz
W.
Daxinger
L.
Schauer
S.E.
Ray
A.
van der Winden
J.
Matzke
M.
Matzke
A.J.M.
Plant Physiol.
2003
, vol. 
132
 (pg. 
1382
-
1390
)
18
Lund
E.
Guttinger
S.
Calado
A.
Dahlberg
J.E.
Kutay
U.
Science
2004
, vol. 
303
 (pg. 
95
-
98
)
19
Yi
R.
Qin
Y.
Macara
I.G.
Cullen
B.R.
Genes Dev.
2003
, vol. 
17
 (pg. 
3011
-
3016
)
20
Bohnsack
M.T.
Czaplinski
K.
Gorlich
D.
RNA
2004
, vol. 
10
 (pg. 
185
-
191
)
21
Brownawell
A.M.
Macara
I.G.
J. Cell Biol.
2002
, vol. 
156
 (pg. 
53
-
64
)
22
Calado
A.
Treichel
N.
Muller
E.C.
Otto
A.
Kutay
U.
EMBO J.
2002
, vol. 
21
 (pg. 
6216
-
6224
)
23
Bohnsack
M.T.
Regener
K.
Schwappach
B.
Saffrich
R.
Paraskeva
E.
Hartmann
E.
Gorlich
D.
EMBO J.
2002
, vol. 
21
 (pg. 
6205
-
6215
)
24
Bollman
K.M.
Aukerman
M.J.
Park
M.Y.
Hunter
C.
Berardini
T.Z.
Poethig
R.S.
Development
2003
, vol. 
130
 (pg. 
1493
-
1504
)
25
Llave
C.
Xie
Z.
Kasschau
K.D.
Carrington
J.C.
Science
2002
, vol. 
297
 (pg. 
2053
-
2056
)
26
Hammond
S.M.
Bernstein
E.
Beach
D.
Hannon
G.
Nature (London)
2000
, vol. 
404
 (pg. 
293
-
296
)
27
Tuschl
T.
Zamore
P.D.
Lehmann
R.
Bartel
D.P.
Sharp
P.A.
Genes Dev.
1999
, vol. 
13
 (pg. 
3191
-
3197
)
28
Hutvagner
G.
Zamore
P.D.
Science
2002
, vol. 
297
 (pg. 
2056
-
2060
)
29
Yekta
S.
Shih
I.H.
Bartel
D.P.
Science
2004
, vol. 
304
 (pg. 
594
-
596
)
30
Han
M.-H.
Goud
S.
Song
L.
Fedoroff
N.
Proc. Natl. Acad. Sci. U.S.A.
2004
, vol. 
101
 (pg. 
1093
-
1098
)
31
Vazquez
F.
Gasciolli
V.
Crete
P.
Vaucheret
H.
Curr. Biol.
2004
, vol. 
14
 (pg. 
346
-
351
)
32
Tabara
H.
Yigit
E.
Siomi
H.
Mello
C.C.
Cell (Cambridge, Mass.)
2002
, vol. 
109
 (pg. 
861
-
871
)
33
Hammond
S.M.
Boettcher
S.
Caudy
A.A.
Kobayashi
R.
Hannon
G.J.
Science
2001
, vol. 
293
 (pg. 
1146
-
1150
)
34
Martinez
J.
Patkaniowska
A.
Urlaub
H.
Luhrmann
R.
Tuschl
T.
Cell (Cambridge, Mass.)
2002
, vol. 
110
 (pg. 
563
-
574
)
35
Yan
K.S.
Yan
S.
Farooq
A.
Han
A.
Zeng
L.
Zhou
M.M.
Nature (London)
2003
, vol. 
426
 (pg. 
468
-
474
)
36
Lingel
A.
Simon
B.
Izaurralde
E.
Sattler
M.
Nature (London)
2003
, vol. 
426
 (pg. 
465
-
469
)
37
Song
J.J.
Liu
J.
Tolia
N.H.
Schneiderman
J.
Smith
S.K.
Martienssen
R.A.
Hannon
G.J.
Joshua-Tor
L.
Nat. Struct. Biol.
2003
, vol. 
10
 (pg. 
1026
-
1032
)
38
Ma
J.B.
Ye
K.
Patel
D.J.
Nature (London)
2004
, vol. 
429
 (pg. 
318
-
322
)
39
Bohmert
K.
Camus
I.
Bellini
C.
Bouchez
D.
Caboche
M.
Benning
C.
EMBO J.
1998
, vol. 
17
 (pg. 
170
-
180
)
40
Fagard
M.
Boutet
S.
Morel
J.B.
Bellini
C.
Vaucheret
H.
Proc. Natl. Acad. Sci. U.S.A.
2000
, vol. 
97
 (pg. 
11650
-
11654
)
41
Morel
J.B.
Godon
C.
Mourrain
P.
Beclin
C.
Boutet
S.
Feuerbach
F.
Proux
F.
Vaucheret
H.
Plant Cell
2002
, vol. 
14
 (pg. 
629
-
639
)
42
Beclin
C.
Boutet
S.
Waterhouse
P.
Vaucheret
H.
Curr. Biol.
2002
, vol. 
12
 (pg. 
684
-
688
)
43
Vaucheret
H.
Vazquez
F.
Crete
P.
Bartel
D.P.
Genes Dev.
2004
, vol. 
18
 (pg. 
1187
-
1197
)
44
Kidner
C.A.
Martienssen
R.A.
Nature (London)
2004
, vol. 
428
 (pg. 
81
-
84
)
45
Hamilton
A.J.
Baulcombe
D.C.
Science
1999
, vol. 
286
 (pg. 
950
-
952
)
46
Llave
C.
Kasschau
K.D.
Rector
M.A.
Carrington
J.C.
Plant Cell
2002
, vol. 
14
 (pg. 
1605
-
1619
)
47
Xie
Z.
Johansen
L.K.
Gustafson
A.M.
Kasschau
K.D.
Lellis
A.D.
Zilberman
D.
Jacobsen
S.E.
Carrington
J.C.
PLoS Biol.
2004
, vol. 
2
 pg. 
E104
 
48
Dalmay
T.
Hamilton
A.
Rudd
S.
Angell
S.
Baulcombe
D.C.
Cell (Cambridge, Mass.)
2000
, vol. 
101
 (pg. 
543
-
553
)
49
Elmayan
T.
Balzergue
S.
Beon
F.
Bourdon
V.
Daubremet
J.
Guenet
Y.
Mourrain
P.
Palauqui
J.C.
Vernhettes
S.
Vialle
T.
, et al. 
Plant Cell
1998
, vol. 
10
 (pg. 
1747
-
1758
)
50
Mourrain
P.
Beclin
C.
Elmayan
T.
Feuerbach
F.
Godon
C.
Morel
J.B.
Jouette
D.
Lacombe
A.M.
Nikic
S.
Picault
N.
, et al. 
Cell (Cambridge, Mass.)
2000
, vol. 
101
 (pg. 
533
-
542
)
51
Dalmay
T.
Horsefield
R.
Braunstein
T.H.
Baulcombe
D.C.
EMBO J.
2001
, vol. 
20
 (pg. 
2069
-
2078
)
52
Boutet
S.
Vazquez
F.
Liu
J.
Beclin
C.
Fagard
M.
Gratias
A.
Morel
J.B.
Crete
P.
Chen
X.
Vaucheret
H.
Curr. Biol.
2003
, vol. 
13
 (pg. 
843
-
848
)
53
Yu
D.
Fan
B.
MacFarlane
S.A.
Chen
Z.
Mol. Plant Microbe Interact.
2003
, vol. 
16
 (pg. 
206
-
216
)
54
Zilberman
D.
Cao
X.
Jacobsen
S.E.
Science
2003
, vol. 
299
 (pg. 
716
-
719
)
55
Chan
S.W.
Zilberman
D.
Xie
Z.
Johansen
L.K.
Carrington
J.C.
Jacobsen
S.E.
Science
2004
, vol. 
303
 pg. 
1336
 
56
Wassenegger
M.
Heimes
S.
Riedel
L.
Sanger
H.L.
Cell (Cambridge, Mass.)
1994
, vol. 
76
 (pg. 
567
-
576
)
57
Matzke
A.J.
Neuhuber
F.
Park
Y.D.
Ambros
P.F.
Matzke
M.A.
Mol. Gen. Genet.
1994
, vol. 
244
 (pg. 
219
-
229
)
58
Jones
L.
Hamilton
A.J.
Voinnet
O.
Thomas
C.L.
Maule
A.J.
Baulcombe
D.C.
Plant Cell
1999
, vol. 
11
 (pg. 
2291
-
2301
)
59
Kishimoto
N.
Sakai
H.
Jackson
J.
Jacobsen
S.E.
Meyerowitz
E.M.
Dennis
E.S.
Finnegan
E.J.
Plant Mol. Biol.
2001
, vol. 
46
 (pg. 
171
-
183
)
60
Jones
L.
Ratcliff
F.
Baulcombe
D.C.
Curr. Biol.
2001
, vol. 
11
 (pg. 
747
-
757
)
61
Aufsatz
W.
Mette
M.F.
van der Winden
J.
Matzke
A.J.
Matzke
M.
Proc. Natl. Acad. Sci. U.S.A.
2002
, vol. 
99
 
Suppl. 4
(pg. 
16499
-
16506
)
62
Lindroth
A.M.
Cao
X.
Jackson
J.P.
Zilberman
D.
McCallum
C.M.
Henikoff
S.
Jacobsen
S.E.
Science
2001
, vol. 
292
 (pg. 
2077
-
2080
)
63
Cao
X.
Jacobsen
S.E.
Curr. Biol.
2002
, vol. 
12
 (pg. 
1138
-
1144
)
64
Cao
X.
Aufsatz
W.
Zilberman
D.
Mette
M.F.
Huang
M.S.
Matzke
M.
Jacobsen
S.E.
Curr. Biol.
2003
, vol. 
13
 (pg. 
2212
-
2217
)
65
Schramke
V.
Allshire
R.
Science
2003
, vol. 
301
 (pg. 
1069
-
1074
)
66
Raponi
M.
Arndt
G.M.
Nucleic Acids Res.
2003
, vol. 
31
 (pg. 
4481
-
4489
)
67
Verdel
A.
Jia
S.
Gerber
S.
Sugiyama
T.
Gygi
S.
Grewal
S.I.
Moazed
D.
Science
2004
, vol. 
303
 (pg. 
672
-
676
)
68
Morel
J.B.
Mourrain
P.
Beclin
C.
Vaucheret
H.
Curr. Biol.
2000
, vol. 
10
 (pg. 
1591
-
1594
)
69
Lippman
Z.
May
B.
Yordan
C.
Singer
T.
Martienssen
R.
PLoS Biol.
2004
, vol. 
1
 (pg. 
420
-
428
)
70
Vaistij
F.E.
Jones
L.
Baulcombe
D.C.
Plant Cell
2002
, vol. 
14
 (pg. 
857
-
867
)
71
Moussian
B.
Schoof
H.
Haecker
A.
Jurgens
G.
Laux
T.
EMBO J.
1998
, vol. 
17
 (pg. 
1799
-
1809
)
72
Moussian
B.
Haecker
A.
Laux
T.
Dev. Genes Evol.
2003
, vol. 
213
 (pg. 
534
-
540
)
73
Hunter
C.
Sun
H.
Poethig
R.S.
Curr. Biol.
2003
, vol. 
13
 (pg. 
1734
-
1739
)
74
Meister
G.
Landthaler
M.
Patkaniowska
A.
Dorsett
Y.
Teng
G.
Tuschl
T.
Mol. Cell
2004
, vol. 
15
 (pg. 
185
-
197
)
75
Liu
J.
Carmell
M.A.
Rivas
F.V.
Marsden
C.G.
Thomson
J.M.
Song
J.J.
Hammond
S.M.
Joshua-Tor
L.
Hannon
G.J.
Science
2004
, vol. 
305
 (pg. 
1437
-
1441
)
76
Song
J.J.
Smith
S.K.
Hannon
G.J.
Joshua-Tor
L.
Science
2004
, vol. 
305
 (pg. 
1434
-
1437
)