Analysis of small RNA in fission yeast; centromeric siRNAs are potentially generated through a structured RNA
2009; Springer Nature; Volume: 28; Issue: 24 Linguagem: Inglês
10.1038/emboj.2009.351
ISSN1460-2075
AutoresIngela Djupedal, Isabelle C. Kos‐Braun, Rebecca A. Mosher, Niklas Söderholm, Femke Simmer, Thomas J. Hardcastle, Aurélie Fender, Nadja Heidrich, Alexander Kagansky, Elizabeth H. Bayne, E. Gerhart H. Wagner, David C. Baulcombe, Robin C. Allshire, Karl Ekwall,
Tópico(s)Plant Disease Resistance and Genetics
ResumoArticle26 November 2009free access Analysis of small RNA in fission yeast; centromeric siRNAs are potentially generated through a structured RNA Ingela Djupedal Ingela Djupedal Department of Biosciences and Nutrition, Center for Biosciences, Karolinska Institutet, Sweden/School of Life Sciences, University College Sodertorn, NOVUM, Huddinge, Sweden Search for more papers by this author Isabelle C Kos-Braun Isabelle C Kos-Braun Wellcome Trust Centre for Cell Biology, Institute of Cell Biology, School of Biological Sciences, The University of Edinburgh, Edinburgh, Scotland, UK Biochemie Zentrum BZH, Universität Heidelberg, Im Neuenheimer Feld, Heidelberg, Germany Search for more papers by this author Rebecca A Mosher Rebecca A Mosher Department of Plant Sciences, University of Cambridge, Cambridge, UK Search for more papers by this author Niklas Söderholm Niklas Söderholm Department of Biosciences and Nutrition, Center for Biosciences, Karolinska Institutet, Sweden/School of Life Sciences, University College Sodertorn, NOVUM, Huddinge, Sweden Search for more papers by this author Femke Simmer Femke Simmer Wellcome Trust Centre for Cell Biology, Institute of Cell Biology, School of Biological Sciences, The University of Edinburgh, Edinburgh, Scotland, UK Search for more papers by this author Thomas J Hardcastle Thomas J Hardcastle Department of Plant Sciences, University of Cambridge, Cambridge, UK Search for more papers by this author Aurélie Fender Aurélie Fender Department of Cell and Molecular Biology, Microbiology Program, Biomedical Center, Uppsala University, Uppsala, Sweden Search for more papers by this author Nadja Heidrich Nadja Heidrich Department of Cell and Molecular Biology, Microbiology Program, Biomedical Center, Uppsala University, Uppsala, Sweden Search for more papers by this author Alexander Kagansky Alexander Kagansky Wellcome Trust Centre for Cell Biology, Institute of Cell Biology, School of Biological Sciences, The University of Edinburgh, Edinburgh, Scotland, UK Search for more papers by this author Elizabeth Bayne Elizabeth Bayne Wellcome Trust Centre for Cell Biology, Institute of Cell Biology, School of Biological Sciences, The University of Edinburgh, Edinburgh, Scotland, UK Search for more papers by this author E Gerhart H Wagner E Gerhart H Wagner Department of Cell and Molecular Biology, Microbiology Program, Biomedical Center, Uppsala University, Uppsala, Sweden Search for more papers by this author David C Baulcombe David C Baulcombe Department of Plant Sciences, University of Cambridge, Cambridge, UK Search for more papers by this author Robin C Allshire Corresponding Author Robin C Allshire Wellcome Trust Centre for Cell Biology, Institute of Cell Biology, School of Biological Sciences, The University of Edinburgh, Edinburgh, Scotland, UK Search for more papers by this author Karl Ekwall Corresponding Author Karl Ekwall Department of Biosciences and Nutrition, Center for Biosciences, Karolinska Institutet, Sweden/School of Life Sciences, University College Sodertorn, NOVUM, Huddinge, Sweden Search for more papers by this author Ingela Djupedal Ingela Djupedal Department of Biosciences and Nutrition, Center for Biosciences, Karolinska Institutet, Sweden/School of Life Sciences, University College Sodertorn, NOVUM, Huddinge, Sweden Search for more papers by this author Isabelle C Kos-Braun Isabelle C Kos-Braun Wellcome Trust Centre for Cell Biology, Institute of Cell Biology, School of Biological Sciences, The University of Edinburgh, Edinburgh, Scotland, UK Biochemie Zentrum BZH, Universität Heidelberg, Im Neuenheimer Feld, Heidelberg, Germany Search for more papers by this author Rebecca A Mosher Rebecca A Mosher Department of Plant Sciences, University of Cambridge, Cambridge, UK Search for more papers by this author Niklas Söderholm Niklas Söderholm Department of Biosciences and Nutrition, Center for Biosciences, Karolinska Institutet, Sweden/School of Life Sciences, University College Sodertorn, NOVUM, Huddinge, Sweden Search for more papers by this author Femke Simmer Femke Simmer Wellcome Trust Centre for Cell Biology, Institute of Cell Biology, School of Biological Sciences, The University of Edinburgh, Edinburgh, Scotland, UK Search for more papers by this author Thomas J Hardcastle Thomas J Hardcastle Department of Plant Sciences, University of Cambridge, Cambridge, UK Search for more papers by this author Aurélie Fender Aurélie Fender Department of Cell and Molecular Biology, Microbiology Program, Biomedical Center, Uppsala University, Uppsala, Sweden Search for more papers by this author Nadja Heidrich Nadja Heidrich Department of Cell and Molecular Biology, Microbiology Program, Biomedical Center, Uppsala University, Uppsala, Sweden Search for more papers by this author Alexander Kagansky Alexander Kagansky Wellcome Trust Centre for Cell Biology, Institute of Cell Biology, School of Biological Sciences, The University of Edinburgh, Edinburgh, Scotland, UK Search for more papers by this author Elizabeth Bayne Elizabeth Bayne Wellcome Trust Centre for Cell Biology, Institute of Cell Biology, School of Biological Sciences, The University of Edinburgh, Edinburgh, Scotland, UK Search for more papers by this author E Gerhart H Wagner E Gerhart H Wagner Department of Cell and Molecular Biology, Microbiology Program, Biomedical Center, Uppsala University, Uppsala, Sweden Search for more papers by this author David C Baulcombe David C Baulcombe Department of Plant Sciences, University of Cambridge, Cambridge, UK Search for more papers by this author Robin C Allshire Corresponding Author Robin C Allshire Wellcome Trust Centre for Cell Biology, Institute of Cell Biology, School of Biological Sciences, The University of Edinburgh, Edinburgh, Scotland, UK Search for more papers by this author Karl Ekwall Corresponding Author Karl Ekwall Department of Biosciences and Nutrition, Center for Biosciences, Karolinska Institutet, Sweden/School of Life Sciences, University College Sodertorn, NOVUM, Huddinge, Sweden Search for more papers by this author Author Information Ingela Djupedal1,‡, Isabelle C Kos-Braun2,5,‡, Rebecca A Mosher3,‡, Niklas Söderholm1, Femke Simmer2, Thomas J Hardcastle3, Aurélie Fender4, Nadja Heidrich4, Alexander Kagansky2, Elizabeth Bayne2, E Gerhart H Wagner4, David C Baulcombe3, Robin C Allshire 2 and Karl Ekwall 1 1Department of Biosciences and Nutrition, Center for Biosciences, Karolinska Institutet, Sweden/School of Life Sciences, University College Sodertorn, NOVUM, Huddinge, Sweden 2Wellcome Trust Centre for Cell Biology, Institute of Cell Biology, School of Biological Sciences, The University of Edinburgh, Edinburgh, Scotland, UK 3Department of Plant Sciences, University of Cambridge, Cambridge, UK 4Department of Cell and Molecular Biology, Microbiology Program, Biomedical Center, Uppsala University, Uppsala, Sweden 5Biochemie Zentrum BZH, Universität Heidelberg, Im Neuenheimer Feld, Heidelberg, Germany ‡These authors contributed equally to this work *Corresponding authors: Department of Biosciences and Medical Nutrition, Karolinska Institutet, University College Sodertorn, Novum, Stockholm, Huddinge S-141 57, Sweden. Tel.: +46 8608 9133; Fax: +46 8774 5538; E-mail: [email protected] Trust Centre for Cell Biology, Institute of Cell Biology, School of Biological Sciences, The University of Edinburgh, Edinburgh, Scotland, UK. Tel.: +44 131 650 7117; Fax: +44 845 280 2340; E-mail: [email protected] The EMBO Journal (2009)28:3832-3844https://doi.org/10.1038/emboj.2009.351 PDFDownload PDF of article text and main figures. ToolsAdd to favoritesDownload CitationsTrack CitationsPermissions ShareFacebookTwitterLinked InMendeleyWechatReddit Figures & Info The formation of heterochromatin at the centromeres in fission yeast depends on transcription of the outer repeats. These transcripts are processed into siRNAs that target homologous loci for heterochromatin formation. Here, high throughput sequencing of small RNA provides a comprehensive analysis of centromere-derived small RNAs. We found that the centromeric small RNAs are Dcr1 dependent, carry 5′-monophosphates and are associated with Ago1. The majority of centromeric small RNAs originate from two remarkably well-conserved sequences that are present in all centromeres. The high degree of similarity suggests that this non-coding sequence in itself may be of importance. Consistent with this, secondary structure-probing experiments indicate that this centromeric RNA is partially double-stranded and is processed by Dicer in vitro. We further demonstrate the existence of small centromeric RNA in rdp1Δ cells. Our data suggest a pathway for siRNA generation that is distinct from the well-documented model involving RITS/RDRC. We propose that primary transcripts fold into hairpin-like structures that may be processed by Dcr1 into siRNAs, and that these siRNAs may initiate heterochromatin formation independent of RDRC activity. Introduction Small RNA molecules are the effectors of RNA interference (RNAi), in which gene expression is regulated either on the transcriptional or the post-transcriptional level (Fire et al, 1998; Hamilton and Baulcombe, 1999; Volpe et al, 2002). RNAi typically involves the enzymes Dicer and Argonaute, and in some systems RNA-directed RNA polymerases. Animals and plants encode several isoforms of the Dicer and Argonaute enzymes, and small RNAs can be classified depending on their origin and the pathway in which they function (for review, see Djupedal and Ekwall (2009)). The three major classes are microRNA (miRNA), short interfering RNA (siRNA) and Piwi-interacting RNA (piRNA). In general, post-transcriptional silencing of genes is accomplished by miRNA, whereas siRNA can induce silencing of genes by recruitment of factors required for heterochromatin formation. The mechanism of transcriptional silencing has been extensively studied in the yeast Schizosaccharomyces pombe. This organism is a well-established model organism for the study of heterochromatin and has single genes coding for Dicer (dcr1+) and Argonaute (ago1+) (Volpe et al, 2002), which function in both transcriptional and post-transcriptional gene silencing (Sigova et al, 2004). The current model for the RNAi-dependent formation of heterochromatin at the centromeres of S. pombe has been described as a self-reinforced feedback loop (Noma et al, 2004; Sugiyama et al, 2005) with siRNA, Ago1, Dcr1 and the RNA-directed RNA polymerase, Rdp1, as integral components (Volpe et al, 2002; Verdel et al, 2004). The RNA-induced initiation of transcriptional silencing (RITS) complex, which includes siRNA, Ago1, Tas3 and Chp1 (Verdel et al, 2004), is targeted to centromeres by dual mechanisms: through the recognition of nascent RNA transcripts by complementary siRNA and through Chp1 binding to the canonical heterochromatin mark, H3K9me. The RITS complex permits recruitment of the Clr4 complex (ClrC) (Motamedi et al, 2004; Zhang et al, 2008) that contains the histone methyl transferase (HMTase) Clr4KMT1, which is specific for lysine 9 of histone H3 (H3K9me) (Rea et al, 2000). ClrC creates new H3K9me marks that are specifically bound by chromodomain proteins, such as Swi6 and Chp2, homologues of metazoan heterochromatin protein 1 (Lorentz et al, 1994; Bannister et al, 2001), in addition to Chp1 (RITS) and Clr4 itself. The RITS complex also allows association of the RNA-directed RNA polymerase complex (RDRC) containing Rdp1, which can produce double-stranded (ds) RNA from nascent transcripts (Motamedi et al, 2004; Sugiyama et al, 2005). Dcr1 cleaves dsRNA into additional siRNAs, causing amplification of siRNAs and further increasing H3K9me levels. In this manner, both siRNAs and H3K9me are required for assembly of heterochromatin. Heterochromatin, characterized by binding of chromodomain proteins to H3K9me and transcriptional silencing of embedded genes, is necessary at the centromeres of S. pombe for proper segregation of chromosomes during cell division (Ekwall et al, 1995; Bernard et al, 2001). The DNA sequences of the three centromeres consist of a central core domain (cnt) flanked by arrays of non-coding, inverted repeats of low GC content, which are interspersed with tRNA genes (Clarke et al, 1986; Nakaseko et al, 1986, 1987; Fishel et al, 1988; Chikashige et al, 1989; Takahashi et al, 1991, 1992; Wood et al, 2002). The repeats are divided into innermost repeats (imr) and outer repeats (otr), which are further subdivided into dg and dh elements. The smallest endogenous S. pombe centromere, cen I, has single copies of dg and dh elements on each chromosome arm, whereas cen III is estimated to have 12 copies of dg and dh elements in total (Wood et al, 2002). The copy number of dg and dh elements at the centromeres can vary in distinct S. pombe isolates as well as in different laboratory strains (Steiner et al, 1993). The use of plasmid-based constructs demonstrated that cnt together with a 2.1-kb fragment from the dg element is required to allow the formation of a mitotically functional centromere (Baum et al, 1994; Wood et al, 2002). Analyses of RITS- and Ago1-associated siRNAs from S. pombe have revealed that a sizeable fraction of the siRNA originates from the dg and dh elements at the centromeres (Cam et al, 2005; Buhler et al, 2008). According to the current model for RNAi-dependent heterochromatin formation, RITS is guided by the encapsulated siRNAs to nascent RNA polymerase II transcripts from the centromeres. Transcription of the dg and dh elements has been shown to occur predominantly in the S-phase of the cell cycle when the non-permissive, heterochromatic structure is dispersed after DNA replication (Chen et al, 2008; Kloc et al, 2008). Hence, RNAi is involved in the maintenance of heterochromatin after DNA replication in each cell cycle. In unsynchronized, wild-type (WT) cells, transcripts derived from the 'reverse' strand of the DNA are more easily detected (Volpe et al, 2002) and a strong promoter upstream of this transcript has been mapped (Djupedal et al, 2005). As a result of heterochromatin formation the expression of reporter genes inserted into the dg and dh elements is silenced in WT cells (Allshire et al, 1995). In strains defective for RNAi, such as ago1Δ and dcr1Δ, transcripts derived from both strands accumulate and reporter genes in dg or dh elements are not silenced (Provost et al, 2002; Volpe et al, 2002), probably due to the absence of heterochromatin at the centromeric region. Here, we have performed high-throughput pyrosequencing of S. pombe size-selected small RNA to gain insight into the role of RNAi in this organism. We found that most centromeric small RNA are produced from two types of small RNA clusters that share a high degree of sequence identity, and we present a detailed description of these loci. Most small RNAs are found within regions of perfect sequence identity between repeats from all three centromeres, indicating functional conservation of these sequences. The predominant sequence class of small RNAs was validated by northern blots and has the characteristics of bona fide siRNAs. In addition, we have determined the in vitro secondary structure of a portion of the transcript from one of the small RNA clusters and have shown that it forms a partially double-stranded secondary structure that is processed by recombinant human Dicer. Furthermore, we demonstrate that a small fraction of centromeric small RNAs is synthesized independently of Rdp1, including small RNA corresponding to the experimentally determined hairpin. Therefore, we propose that the ability of nascent centromeric transcripts to fold into double-stranded 'hairpin' structures may permit their Dcr1-dependent processing into siRNAs, which in turn contributes to the establishment of heterochromatin at the centromeres. Results Small RNA in S. pombe With high-throughput 454 sequencing, 21 776 sequenced small RNAs were determined from WT S. pombe cells (Table I). In addition, sequences of small RNA were determined from rpb7G150D cells that carry a point mutation in the RNA polymerase II subunit Rpb7, which causes lower levels of transcription of centromeric siRNA precursors (Djupedal et al, 2005). A large percentage of sequences mapped to the transcribed region of tRNA or rRNA genes and were, together with reads without perfect match to the S. pombe genome, excluded from further analysis. Nearly half of the remaining 3552 WT sequences begin with a uracil (U), with lesser representation of A and C as a first base and strong selection against G (Figure 1A). Although this is a significant deviance from the genome average, Ago1-associated siRNAs (Buhler et al, 2008; subjected to the same matching and filtering as WT) show a strong bias for uracil in the first position (>85%; Figure 1A). Similarly, Ago1-associated siRNAs are primarily 22–23 nucleotides (nt) in length (Figure 1B; Buhler et al, 2008), whereas our WT small RNAs range in size from 20 to 30nt, with most being between 22 and 25nt (Figure 1B). These data indicate that Ago1 has a preference for 22–23-nt siRNAs that begin with U, and that it selects these siRNAs from a more diverse population generated in the cell. Interestingly, in the rpb7G150D mutant, there is a shift in the size of small RNAs with a larger proportion of short small RNAs than in the WT sample, combined with a lower preference for small RNAs that begin with U (Figure 1A and B). Figure 1.Characteristics of the small RNA populations of wild-type and rpb7G150D mutant cells in comparison to Ago1-associated siRNAs (Buhler et al, 2008). Small RNAs without a perfect match in the S. pombe genome or those matching tRNA and rRNA were removed. (A) Start nucleotide distribution of small RNA. The 5′ nucleotide on the x-axis and the relative abundance of small RNAs on the y-axis in wild-type and rpb7G150D mutant cells compared with Ago1-associated siRNAs. (B) Size distribution of small RNA, length in nucleotides on the x-axis and relative abundance on the y-axis in wild-type, rpb7G150D mutant cells and Ago1-associated siRNAs. (C) Percentages of genomic distributions of small RNA reads in wild-type cells, rpb7G150D mutant cells and Ago1-associated siRNAs. Download figure Download PowerPoint Table 1. Compilation of 454 deep sequencing analysis of small RNA from wild-type (WT) and the rpb7G150D mutant strain Total number of reads Genome- matching readsa Filtered matching readsb Genome matches/ read 972 (WT) 21776 16341 3531 4.9 rpb7G150D 51894 36990 11081 1.5 Ago1-associated siRNAsc 349251 257629 197170 6.3 a Reads with a perfect match in the Schizosaccharomyces pombe genome. b Genome-matching reads after removal of structural RNAs (tRNA, rRNA, snoRNA, and snRNA). c Data are extracted from a study by Buhler et al (2008); GEO accession number: GSE12416. Excluding sequences from transcribed structural RNAs, which are likely degradation products, the genomic regions that generate most small RNAs are centromeric repeats and protein-coding genes (Figure 1C). At the centromeres, hundreds of small RNAs cluster within the dg and the dh elements. In WT, 100 genes had three matching small RNA reads or more. Nearly all of these genes (98 genes) were also retrieved in the small RNA sample from the rpb7G150D mutant. The rpb7G150D sample, which generated nearly three times as many sequences as the WT sample in combination with a depletion of centromeric small RNAs, resulted in retrieval of over 600 genes with matching small RNA (cut-off: ⩾3 small RNAs per gene). All small RNAs towards genes were of the sense orientation, with the exception of the tlh1 gene (SPAC212.11) that has homology to the dh repeat and in which small RNAs of both orientations were abundant. The most likely explanation is that these sense-oriented small RNAs represent mRNA degradation products. However, Dcr1, Ago1, and Rdp1 were shown to mediate post-transcriptional silencing of an exogenous hairpin (Sigova et al, 2004). Furthermore, investigations of the proteome of dcr1Δ cells demonstrated increased protein levels of Hsp16, Pgk1, Tpx1, and Hsp104, whereas protein levels of Hxk2, Eno1, and Thi3 were decreased, with or without concomitant alterations of mRNA levels (Gobeil et al, 2008). We detected small RNAs homologous to six of these genes (Supplementary Figure S1). Thus, it is possible that these sense small RNAs somehow contribute to gene regulation. Although we did not carry out a systematic analysis of small RNAs in intergenic regions, we did notice 14 copies of a single small RNA in the intergenic downstream region of the convergent gene pair mei4+–act1+ (Supplementary Figure S1). This locus has been reported to be coated with heterochromatin factors (Cam et al, 2005) and these small RNAs are probably involved in controlling transcription termination in this intergenic region, as suggested by Gullerova and Proudfoot (2008). Characterization of centromeric small RNA clusters All clusters abundant for small RNA overlap the dg and dh elements that constitute the centromeric outer repeats (Figure 2A). These clusters are between 2000 and 3000 bp long with up to 1000 matching sequence reads. In general, there are two recurring sequences that produce small RNAs in the centromeres. The dg cluster overlaps the 3′-end of the dg repeat, including the fragment necessary for centromere function (Baum et al, 1994). The dh cluster is situated within the second half of the dh repeat. Alignment of these sequences from all centromeres reveals a high degree of conservation across both the dg and the dh clusters (Supplementary Figure S2A and B). The dg element has previously been reported to have 97% homology between different centromeres, whereas the homology of the dh element from centromere I, II, and III was reported to have 48% identity (Wood et al, 2002). If deletions of the dh elements are taken into account, the remaining dh sequences have up to 99% sequence identity between dhI and dhII (Nakaseko et al, 1987). The clusters are found at all centromeres. A 300-bp translocation from dh to dg (Chikashige et al, 1989) is common to both the dg and dh clusters. Figure 2.Distribution of small RNA of wild-type cells at the centromeres. (A) Schematic on-scale representation of the centromeres of chromosome I, II and III based on the S. pombe GeneDB. Vertical black arrows represent tRNAs with standard one letter abbreviations representing amino-acid specificity. The green bars represent the KpnI restriction fragment that was shown to be necessary for centromere function (Baum et al, 1994). Red horizontal arrows represent RevCen; orange horizontal arrows represent the region within the dh element with a translocation similar to RevCen. A 700-bp region is deleted from the dh cluster at the left arm of centromere I and the sequence in between dg and dh siRNA clusters is deleted in centromere III. Each arrow represents a sequenced, small RNA match; thicker arrows indicate multiple sequenced small RNAs. Owing to the high degree of sequence similarity, most sequenced centromeric small RNA have perfect match to several clusters within the centromeres. These sequences have been plotted at each perfect match, that is, more than once per sequence. (B) Histogram of small RNA strand distribution at the otr2R-dh and otrR2-dg siRNA clusters. The x-axis depicts ratio of forward to reverse strand siRNAs (log10 scale). All bars are significantly different from 1; ***P<0.001; **P 25 bases. Hotspots of siRNAs have been named in roman numerals, the most abundant siRNA, 'IV', was sequenced 400 times and has been cropped. The red horizontal arrow depicts the location of the RevCen sequence and roman numerals in red indicate the siRNA hotspots that match to both dg and dh siRNA clusters. Download figure Download PowerPoint Although small RNAs match both strands of the centromeric clusters, there is a preference for small RNAs from the reverse strand of the repeat, with stronger bias at the dh cluster than at the dg cluster (Figure 2B). No strand bias is expected according to the current models for RNAi-dependent heterochromatin formation at the centromeres in S. pombe because cleavage of Rdp1-derived dsRNA will produce equal amounts of plus- and minus-stranded siRNAs. Interestingly, a direct comparison with Ago1-associated siRNAs (Buhler et al, 2008) from these clusters showed no minus strand bias. Instead there was a small but significant enrichment for the positive strand. Consistent with this, 25nt small RNAs from WT, which are unlikely to associate with Ago1 due to the size, show the strongest minus strand bias (>10-fold). The accumulation of small RNAs is not equal across the clusters, resulting in several hotspots of small RNAs. The 26 most abundant small RNA hotspots were named with roman numerals (Figure 2C). The most abundant centromeric small RNA hotspot, small RNA IV, was sequenced 400 times, with the length of the small RNA varying by a few nucleotides. The GC content of the small RNAs is significantly higher than in the surrounding sequence (Supplementary Figure S3). As the small RNAs are homologous to regions in which the sequence is identical in most or all copies of the dg and dh elements, it is not possible to determine from which centromeric repeat copy these small RNAs originate. Hotspot small RNAs are Dcr1 dependent, carry a 5′-monophosphate, and are associated with Ago1 To verify the occurrence of small RNA hotspots, single oligonucleotide probes corresponding to four of the most abundant sequenced small RNAs were synthesized and used as probes on northern blots of small RNA preparations. In addition, control sense oligonucleotides or oligonucleotides homologous to neighbouring sequence with few matching small RNAs were used. In accordance with the small RNA distribution determined by sequencing, the four hotspot small RNA probes (anti-VII, anti-XII, anti-XXII, and anti-VI) readily detected small RNAs by northern analyses, whereas little or no signal was detected with sense probes (sense XII and sense XXII) or nearby control probes (+35-anti-XII and 50-anti-XXII). No signals were detected in small RNA preparations from dcr1Δ cells (Figure 3A). Figure 3.Validation and analyses of siRNAs by northern blots. (A) Single oligonucleotide probes, antisense, sense, or nearby to siRNAs IV, XII, XXII and VI were used for detection of small RNAs from wild-type or dcr1Δ cells. SnoRNA 58 was used as loading control. (B) and (C) Analyses of 5′-termini of small RNAs by enzymatic reactions followed by northern blots. (B) Terminator exonuclease digests monophosphorylated 5′-termini and (C) guanylyltransferase (GTase) caps di- or triphosphorylated 5′-termini. The control oligonucleotides are 5′-triphosphorylated RNA oligonucleotide (PPP—) and 5′-monophosphorylated RNA oligonucleotide with a blocked 3′-end (P—X) (Ule et al, 2005). The blots were probed sequentially with specific oligonucleotide probes and with a random-primed probe spanning the whole siRNA cluster (dh +dg siRNAs). (D) Analysis of Ago1-purified siRNA 5′-ends by enzymatic reactions followed by northern blot as described above. Download figure Download PowerPoint Sequencing of Ago1-associated siRNAs in S. pombe indicates that 5′-monophosphate siRNAs are present (Buhler et al, 2008). Our analysis corroborated this as we prepared small RNA libraries in a 5′-monophosphate-dependent manner. However, in Caenorhabditis elegans, 5′-monophosphate siRNAs are a minority. The majority of siRNAs have been reported to have 5′-triphosphates in accordance with being products of an RNA-directed RNA polymerase (Pak and Fire, 2007; Sijen et al, 2007). To determine if 5′-triphosphate siRNAs were also present in S. pombe, we treated small RNAs with Terminator exonuclease, a 5′-exonuclease that digests RNA with 5′-monophosphates. Small RNAs completely disappear from the centromere after treatment with Terminator exonuclease, whereas a 5′-triphosphate control oligonucleotide was unaffected (Figure 3B). Furthermore, small RNAs were not capped by guanylyltransferase (GTase) that caps 5′-di- or triphosphorylated RNA and produces approximately two-nucleotide slower gel migration. In addition, GTase-treated small RNAs were digested with Terminator exonuclease, which cannot digest if a 5′-cap is present (Figure 3C). Finally, in RNA samples purified from Ago1–FLAG (Buhler et al, 2007), single oligonucleotide probes, as well as random-primed probes that detect all small RNAs, detected small RNA with 5′-monophosphates (Figure 3D). These data indicate that sequenced S. pombe centromeric small RNAs reported here are Dcr1 dependent, possess 5′-monophosphates, and associate with Ago1 in vivo and thus seem to be true siRNAs. The 5′-end of the transcript from the dg cluster forms a partially double-stranded RNA structure and is processed by human recombinant Dicer in vitro The high degree of sequence identity at the centromeric dg and dh elements could be caused by frequent events of homologous recombination. Alternatively, functional conservation could maintain important features of the sequence, such as the ability to form secondary structures. Within the transcripts from the dg and dh clusters, siRNAs are derived from several
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