Cardiovasc Drugs Ther (2011) 25:151–159 DOI 10.1007/s10557-011-6290-z
An Introduction to Small Non-coding RNAs: miRNA and snoRNA Christopher L. Holley & Veli K. Topkara
Published online: 6 April 2011 # Springer Science+Business Media, LLC 2011
Abstract Research into small non-coding RNAs (ncRNA) has fundamentally transformed our understanding of gene regulatory networks, especially at the post-transcriptional level. Although much is now known about the basic biology of small ncRNAs, our ability to recognize the impact of small ncRNA in disease states is preliminary, and the ability to effectively target them in vivo is very limited. However, given the larger and growing focus on targeting RNAs for disease therapeutics, what we do know about the intrinsic biology of these small RNAs makes them potentially attractive targets for pharmacologic manipulation. With that in mind, this review provides an introduction to the biology of small ncRNA, using microRNA (miRNA) and small nucleolar RNA (snoRNA) as examples. Key words Small Noncoding RNA . MicroRNAs . Small Nucleolar RNA . Post-Transcriptional RNA Processing
Introduction With due respect to the importance of RNA for encoding proteins, the last 20 years have produced an C. L. Holley (*) Diabetic Cardiovascular Disease Center, Division of Cardiology, Department of Internal Medicine, Washington University School of Medicine, 660 South Euclid Ave. Campus, PO Box 8086, St Louis, MO 63110-1093, USA e-mail:
[email protected] V. K. Topkara Center for Cardiovascular Research, Division of Cardiology, Department of Internal Medicine, Washington University School of Medicine, 660 South Euclid Ave. Campus, PO Box 8086, St Louis, MO 63110-1093, USA
incredible bounty of discoveries highlighting the importance RNAs that do not encode proteins. Moreover, where the world of non-coding RNA (ncRNA) was once dominated by transfer RNAs (tRNA), the new world of ncRNAs is one with a seemingly limitless diversity of long and short ncRNAs, influencing almost all areas of cellular biology with a wide variety of molecular mechanisms. Perhaps the most exciting families of small ncRNA are small interfering RNA (siRNA) and microRNA (miRNA), with their celebrated roles in the process of RNA interference. However, siRNA and miRNA are not alone—there are many other families of small noncoding RNA that are necessary for regulatory roles in the cell, such as snRNAs, snoRNAs, scaRNAs, and gRNAs, to name a few (Table 1). Because these small ncRNA are so important to fundamental biologic processes and contribute significantly to certain pathophysiologic states, they may therefore be attractive new targets for medical therapeutics. In order to facilitate an understanding of how small ncRNAs might be manipulated therapeutically, we provide here a brief overview of two very distinct small ncRNA families: miRNA and snoRNA. On one hand, much has been learned about RNA interference from siRNA and miRNA, and exploiting miRNA as therapeutics or drug targets is foreseeable. On the other hand, the biology of snoRNAs is somewhat less well understood, but early evidence suggests that this class of ncRNA may also contribute significantly to post-transcriptional regulatory networks in health and disease. For both miRNA and snoRNA, we intend that this review will provide an adequate introduction to the basic biology involved, which will in turn facilitate a better appreciation of ncRNA in general and its potential for therapeutic manipulation.
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Table 1 Selected families of small non-coding RNAs Family
Role
tRNA (transfer RNA) siRNA (small interfering RNA) miRNA (microRNA) snRNA (small nuclear RNA) snoRNA (small nucleolar RNA) scaRNA (small Cajal body RNA) gRNA (guide RNA) piRNA (Piwi-interacting RNA)
Translation RNA silencing RNA silencing RNA splicing RNA modification RNA modification RNA editing Gene silencing
miRNA MicroRNAs (miRNAs) are endogenous, single-stranded, short RNA sequences (~22 nucleotides) that regulate gene expression at the post-transcriptional level by base-pairing with target mRNA sequences. Since the first discovery of miRNAs in C. elegans in 1993 [1], hundreds of miRNAs in eukaryotes have been identified that are involved in orchestrating a wide range of physiological and pathological processes such as development, growth, differentiation, immune reaction, and adaptation to stress [2–4]. Indeed, a variety of disease states such as cancer and heart failure are associated with distinct miRNA signatures suggesting that specific miRNA programs are activated in various pathophysiological processes [5, 6]. Given that a single miRNA can target hundreds of mRNAs and a single mRNA can be targeted by multiple miRNAs, miRNA-mRNA pairs operate highly complex signaling networks.
miRNA biogenesis Genomic location analysis of miRNA genes has suggested that the majority of mammalian miRNAs are transcribed from an intronic location, although some are transcribed from an exonic location [7]. Expression of these miRNAs largely coincides with the expression of the host transcription units, indicating that intronic miRNAs and their host genes are derived from a common transcript [8]. Alternatively, miRNAs can reside in intergenic locations and act as independent transcription units. Animal miRNA genes are often clustered in polycistronic units that generate long primary transcripts containing multiple miRNAs. In addition, miRNAs may also have multiple copies in the genome and such miRNAs are transcribed from multiple loci at different genomic location. The first step of miRNA biogenesis involves transcription of miRNA genes, which is regulated by conventional transcriptional factors and largely mediated by the RNA Polymerase II enzyme (Fig. 1) [9]. Transcription of a
Fig. 1 miRNA biogenesis and function. The primary transcripts of miRNAs, called pri-miRNAs, are transcribed as individual miRNA genes, from introns of protein-coding genes, or from polycistronic transcripts. The RNase Drosha further processes the pri-miRNA into 70–100 nucleotide, hairpin-shaped precursors, called pre-miRNA, which are exported from the nucleus by exportin 5. In the cytoplasm, the pre-miRNA is cleaved by Dicer into an miRNA:miRNA* duplex. Assembled into the RISC, the mature miRNA negatively regulates gene expression by either translational repression or mRNA degradation, which is dependent on sequence complementarity between the miRNA and the target mRNA. ORF, open reading frame. Reproduced from Van Rooij E. et al. [59] with permission
miRNA gene generates a primary transcript miRNA (primiRNA) which is approximately 2 kb in length, has characteristic hairpin structure, and contains the mature miRNA sequence in the stem portion near the loop (Fig. 1). There pri-miRNAs are further processed in the nucleus by Drosha (a member of highly conserved RNase III enzyme family, bound to co-factor DiGeorge Syndrome Critical Region 8 (DGCR8)/Pasha) leading to generation of a 70– 100 nucleotide long hairpin-shaped precursor miRNA (premiRNA). Like other non-coding RNAs, pre-miRNAs are exported to cytoplasm by a karyopherin family nuclear transport receptor, Exportin-5 [10]. Once in the cytoplasm, pre-miRNA is further processed by a RNAse Type III enzyme Dicer which removes the terminal loop portion of the molecule to form a 22 nucleotide long miRNA-
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miRNA* duplex [11]. Subsequently, one strand of the duplex - named as ‘guide strand’—is incorporated into a multiprotein RNA-silencing complex (RISC), while the other strand—named as ‘passenger strand’—is stripped away and targeted for degradation [12]. Factors influencing strand selection include thermodynamic stability and the position of the stem-loop; however, in some cases both miRNA strands can become functional and target different mRNA populations [13–15]. Silencing of target RNAs is achieved by the multiprotein RISC complex that is composed of the mature miRNA attached to Argonaute proteins and accessory factors such as GW182 that are needed for miRNA-mRNA interaction [16, 17].
miRNA mechanism of action In animals, miRNA mediated gene silencing is generally accomplished by imperfect base pairing of 5′ region of miRNAs with the target mRNA sequence, leading to translational repression and/or mRNA degradation [18]. miRNAs base-pair primarily with the 3′ untranslated region (3′UTR) of the target gene; however, recent studies have demonstrated that miRNAs may also target 5′UTRs, open reading frames (ORFs), and even DNA sequences to regulate transcription [19]. Translation repression was suggested as the predominant mechanism of miRNA induced silencing in animals with little or no change in target mRNA levels [20, 21], but recent evidence utilizing ribosome profiling has demonstrated that mRNA degradation may be the predominant reason for reduced protein output [22]. Moreover, miRNAs have been paradoxically shown to upregulate gene expression by enhancing translation under specific conditions [23, 24]. Approximately 1,000 miRNA genes are predicted to exist in the human genome, each of which could potentially target hundreds or thousands of mRNAs and regulate up to 30% of transcriptome [25]. In addition, the majority of mRNAs contain multiple binding sites for miRNAs, generating a highly complex regulatory network system. Distinct miRNA signatures at various developmental and disease states suggests that in addition to “fine-tuning” mRNA expression, miRNAs can act as on and off switches to either generate or eliminate certain target genes at various stages. The 5′ region of a miRNA from nucleotide positions 2–7 (termed as ‘seed region’) is highly conserved and thought to confer much of the target recognition specificity [26]. This is supported by studies demonstrating that miRNAlike regulation was most sensitive to substitutions that disrupt base pairing within the seed region [27, 28]. Indeed, miRNAs with identical seed regions are commonly grouped into miRNA families and are likely to target similar sets of
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mRNAs. However, since miRNAs are evolutionarily conserved through the whole sequence, it is likely that the remainder of the miRNA may also contribute to target specificity. In this regard, it is important to note that pairing to the 3′ region of the miRNA can supplement seed pairing to enhance target recognition or even compensate for a seed mismatch (termed as 3′-supplementary sites and 3′-compensatory sites respectively), suggesting that members of miRNA families may have distinct functions [26, 29].
miRNA targets Identification of miRNA targets remains a major challenge in the field of miRNA function. A simple approach to contrast miRNA and mRNA expression signatures is to detect reciprocally regulated miRNA-mRNA pairs. However, this method is of limited use given that miRNAs can indirectly regulate mRNAs that are not direct targets. In addition, a number of bioinformatic tools with varying methodologies have recently become available including TargetScan, miRanda, PicTar, Diana-T, and RNA22 [30– 34]. The search algorithms generally incorporate information on seed complementarity, evolutional conservation, predicted binding energy of miRNA-mRNA duplexes, and total number of binding site in 3′UTR for target prediction. However, bioinformatic approach remains problematic, particularly since animal miRNAs bind to mRNA with imperfect complementarity. It is important to emphasize that each of these algorithms is expected to have a sizable degree of false positive and false negative predictions, and experimental validation is essential for confirming target genes. More recently, biochemical target prediction has been suggested as alternative method for identification of miRNA targets [35, 36]. This technique includes immunoprecipitation of RISC followed by mRNA profiling through either microarray technology or RNA sequencing, and therefore provides in vivo targets bound to RISC complex under various experimental conditions. However, targets that are identified by overexpression or knockdown of a particular miRNA may not be relevant in a physiological context.
snoRNA snoRNAs, or small nucleolar RNAs, are best known as guide molecules for site-specific methylation and pseudouridylation of other RNAs. As their name implies, they are typically localized to the nucleolus, where they play a canonical role in the modification and processing of rRNA (Fig. 2). Fully processed human rRNA contains around 200 such snoRNA-guided nucleotide modifications, with 91
154 Fig. 2 snoRNAs localize to the nucleolus where they guide sequence-specific modifications of rRNA. Most snoRNAs are contained within introns (of pre-mRNA or rRNA), and are released during splicing. scaRNAs localize to the Cajal body and guide the same modifications of snRNA. * denotes RNA base modification with either 2′-O-methylation or pseudouridylation
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ribosome (with rRNA*)
rRN snoRNA
A*
rRNA
nucleolus DNA
pre-mRNA
cytoplasm snoRNA
scaRNA Cajal body
snRNA spliceosome (with snRNA*) mRNA nucleus
pseudouridines and 106 2′-O-methyl groups [37]. Importantly, these chemical alterations are required for proper rRNA processing and ribosome function [38]. Similarly, methylation and pseudouridylation of snRNAs are also required for proper function of the spliceosome, and these modifications are guided by a subset of snoRNAs referred to as scaRNA (Fig. 2). In addition to these core functions, certain snoRNAs have now been shown to influence mRNA splicing or even to serve as precursors for miRNA. In fact, the hundreds of predicted snoRNAs in mammals is several times the number predicted in yeast, and these newly characterized roles outside of the nucleolus hint that this large family of ncRNA may be even more important than previously recognized.
snoRNA: General features and biogenesis The snoRNAs are grouped together based on some very general features. As noted above, they are small (60– 220 nt) non-coding RNAs with primarily nucleolar localization. Conserved sequence motifs bind a core complement of proteins to form a snoRNP (ribonucleoprotein). With this, the protein component provides an enzymatic activity for target RNA modification (either methylation or pseudouridylation; see Table 2). In addition to serving as a scaffold for the enzymatic complex, each snoRNA also contains at least one antisense element to recruit a target RNA. In general, the presence of a snoRNA and its target RNA, along with the relevant RNA-modifying enzyme complex is sufficient to catalyze target RNA methylation or pseudouridylation in vitro.
Another common feature in higher organisms is that almost all snoRNAs are encoded by introns (in contrast to yeast, where most snoRNAs are encoded by independent transcripts). In fact, a single transcript may encode a protein and also host multiple intronic snoRNAs. While there is no firm association between the host genes and the function of the snoRNAs that they contain, many host genes are loci that encode ribosomal proteins. Other transcripts have only noncoding exons surrounding introns that contain the snoRNAs. In these cases, the exons are usually not evolutionarily conserved, and they function simply as structural placeholders to ensure correct splicing of the snoRNAs. Following transcription, snoRNA-containing introns are removed from the pre-mRNA by the typical spliceosome, which itself contains snRNAs that are scaRNA targets (Fig. 2). The intron lariats with snoRNAs then undergo specific processing that includes debranching of the lariat, followed by exonuclease trimming of the intron ends to form the mature snoRNA. This entire process appears to be spatially restricted to the nucleus. It is important to note that the interactions between snoRNAs and their associated proteins are highly conserved from archaea to human. This has greatly facilitated molecular analysis, including in vitro assays and x-ray crystallography to resolve the protein-protein and proteinRNA interactions [see review by [39]]. However, because the study of snoRNAs and snoRNPs has been largely carried out in archaea and yeast (with a few functional studies in Xenopus and mice), there are currently a variety of names for each component. In this review, we refer to the orthologous human nomenclature for the snoRNAs and their associated proteins, in order to focus more directly on implications for human disease and therapeutics.
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Table 2 C/D Box and H/ACA Box snoRNAs Associated RNA modification Associated enzyme (human) Other core binding proteins
C/D Box
H/ACA Box
2′-O-methylation Fibrillarin 15.5 K/NHPX, NOP56, NOP58
pseudouridylation Dyskerin NHP2, GAR1, NOP10
snoRNA classes: C/D and H/ACA boxes
C/D box snoRNAs
The snoRNAs themselves are broadly divided into two classes, known as the C/D Box and H/ACA Box snoRNAs. These designations correspond to conserved sequence motifs and the enzyme complexes that they recruit. The C/D box and H/ACA box motifs themselves are necessary and sufficient for the nucleolar localization of these RNAs [40, 41]. C/D Box snoRNAs serve as a scaffold to bring together the 2′-O-methyltransferase (fibrillarin; a.k. a. NOP1 in yeast) and a target RNA (Fig. 3). In a similar manner, H/ACA Box snoRNAs bring together the pseudouridine synthase (dyskerin; Cbf5 in yeast) with a target RNA (Fig. 4). The assembly of the ribonucleoprotein (RNP) complex occurs co-transcriptionally, and loss of snoRNA-associated proteins leads to reduced levels of the snoRNA itself [42], probably due to the fact that the snoRNA-associated proteins serve to protect the snoRNA from exonuclease cleavage (this “RNase protection” effect also seems to determine the size of a mature snoRNA).
As shown in Fig. 2, C/D box snoRNAs form a single stemloop secondary structure and are identified at the sequence level by their characteristic C-boxes (RUGAUGA, where R is a purine) and D-boxes (CUGA). The stem is formed by short inverted repeats at the 5′ and 3′ termini of the snoRNA, which bring the 5′ C-box in approximation with the 3′ D-box (Fig. 3). Typically, the snoRNA will also have a second internal set of less-well conserved C/D boxes (C’ and D’) such that the stem loop exhibits symmetry. The apposed C and D boxes then serve to recruit the core proteins of the methyl transferase complex. The 15.5 K protein is the first to bind, interacting directly with the primary C/D box motif at the stem. It then nucleates the assembly of the proteins NOP58 and NOP56, which interact with the C- and C’-boxes, respectively [43]. The NOP proteins then each recruit a fibrillarin methyl transferase enzyme to interact with the D boxes (Fig. 3) [39]. An antisense guide sequence to recruit and position a specific RNA target for methylation is immediately 5′ to
Target RNA
Target RNA
’-b D
C ’-b
ox
’-b D
ox
C ’-b
ox
ox
-2’-O-Me
-2’-O-Me
D-box guide
D’-box guide
N
op
56
fibrillarin
2’-O-Me-
N
C
ox
D
ox -b
C 3’
5’
-b
-b ox
5’
3’
ox -b
5’
fibrillarin
5’
3’
C-box: RUGAUGA (R=purine) D-box: CUGA
Fig. 3 Schematic representations of a typical C/D box snoRNA, without and with associated ribonucleoproteins
D
op
58
2’-O-Me-
15.5K protein
3’
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H/ACA box snoRNAs
guide sequences
5’ H-box 5’
3’ ACA
3’
H-box: ANANNA, N is any nucleotide ACA-box: ACA; is psuedouridylation site
Fig. 4 Schematic representation of a typical H/ACA box snoRNA, with a target RNA in one of the two pseudouridylation pockets
each D-box. Specifically, the 2′-O-methylation occurs on the target RNA base that binds exactly five nucleotides proximal to the D box. As noted above, fully processed human rRNA contains 101 of these 2′-O-methylations. Although mutations in 2′-O-methylation can lead to ribosome dysfunction [38], the exact molecular mechanism is unclear, as ribosomes assemble normally and are capable of translation. It seems most likely that rRNA requires 2′-O-methylations to make appropriate intramolecular bonds with ribosomal protein components, such that the efficiency of the complex is maximized. Alternatively, 2′-O-methylation makes RNA more resistant to nucleases, and these modifications may substantially increase the stability of the rRNA itself. The most prominent human disorder associated with C/ D-box snoRNAs is probably not a defect of 2′-O-methylation, but appears to highlight a non-canonical snoRNA function in which snoRNAs affect alternative splicing. Prader-Willi syndrome is an inherited disorder that manifests in childhood with hyperphagia, obesity, hypogonadism, and mental retardation. The affected genetic locus encodes many snoRNAs, including clusters of two C/D box snoRNAs that are highly expressed in the brain: HBII-52 and HBII-85. Of these, HBII52 has a guide sequence that targets RNA encoding a serotonin receptor (5-HT2cR), such that the snoRNA regulates splicing of the receptor message [44]. Many affected individuals fail to express the HBII-52 snoRNA, resulting in altered ratios of 5-HT2cR isoforms and apparently abnormal serotonergic signaling responses. Because splicing occurs in the nucleoplasm, this demonstrates one role for snoRNAs outside of the nucleolus. The molecular role of the HBII-85 C/D box snoRNA is less clear, but its deficiency has been implicated as the primary defect in some cases of PWS [45].
The H/ACA snoRNAs are somewhat longer than the C/D box snoRNAs and form two stem loops (Fig. 4). They are identified at the sequence level by their H-boxes (ANANNA, N is any nucleotide) and ACA-boxes (ACA), where the ACA box is exactly three nucleotides proximal to the 3′ terminus of the RNA. As with the C/D box motifs, the H/ACA boxes direct the nucleolar localization of the snoRNA. Each stem-loop forms a pseudouridylation pocket, which contains guide sequences that recruit the target RNA by antisense binding (Fig. 4). The specific nucleotide targeted for pseudouridylation is located 14–16 nt upstream from the H or ACA box. The pseudouridylation reaction itself is an isomerization of uridine that is catalyzed by the enzyme dyskerin. Like the C/D-box snoRNAs, the RNP complex if composed of four evolutionarily conserved proteins: dyskerin, Nhp2, Nop10, and Gar1 (Table 2). For this RNP, the dyskerin and Nhp2 components bind directly with the H/ACA RNA, with the ACA motif being required for interaction with dyskerin. Nop10 and Gar1 complete the snoRNP complex via protein-protein interactions with dyskerin [46]. Targets for pseudouridylation primarily include the rRNAs and some snRNAs. As noted above, fully processed human rRNA contains 91 pseudouridines. The importance of pseudouridylation is evident from genetic studies in yeast [38], as well as from the human genetic disorder dyskeratosis congenita. This disorder, which is characterized by dysfunctional telomerase, manifests clinically with bone marrow failure, premature aging, increased incidence of malignancy, and premature death. While it is not a typical snoRNA, the telomerase RNA component (TERC) in vertebrates is a chimeric molecule, with a 5′ reverse transcriptase domain and a 3′ H/ACA domain. Both domains are required for telomerase activity in vivo. As expected, the H/ACA domain recruits fibrillarin, and pseudouridylation activity is required for normal telomere maintenance. Accordingly, dyskeratosis congenita can be due to mutations in the pseudouridylation machinery (dyskerin, Nhp2, or Nop10), or components of human telomerase itself (TERT, TERC, or Tin2). The most severe form of the disease is X-linked and is due to mutations of dyskerin [46].
scaRNAs The scaRNAs are a related subsetset of small ncRNA that are retained in nucleoplasmic domains called Cajal bodies and have sequence, structural, and functional features of either or both classes of snoRNAs. These ncRNAs are responsible for 2-O-methylation and pseudouridylation of the snRNAs, which associate with
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snRNPs to form the spliceosome. In addition to their C/D box or H/ACA box motifs, the scaRNAs contain an additional localization motif called the CAB box (Cajal body box, consensus: UGAG) that mediates accumulation in the Cajal body.
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by antisense binding or snoRNAs that serve as precursors for RISC-mediated gene silencing are exciting new avenues for research into the biology of small ncRNA.
Future perspectives Non-canonical snoRNA functions In addition to guiding methylation and pseudouridylation, some snoRNAs have recently been shown to have very different roles outside of the nucleolus. These intriguing new roles include regulation of alternative splicing and serving as precursors for miRNA. The HBII-52 C/D box snoRNA associated with Prader-Willi syndrome is one prominent example. As described above, this snoRNA regulates alternative splicing of a serotonin receptor. More recent studies have shown that the mouse ortholog of this snoRNA (MBII-52) can actually influence the splicing of at least five other transcripts as well [47]. The mechanism appears to involve further processing of the snoRNA to remove the 5′ and 3′ inverted repeats that usually stabilize the C/D box snoRNA stem. MBII-52 snoRNA that has been processed in this way loses its ability to associate with canonical C/D box proteins (including fibrillarin), and instead associates with hnRNPs that are involved with splice-site selection [47]. This sort of processing of snoRNAs to smaller fragments has now been reported several times, with convincing evidence that some snoRNAs act as precursors for functional miRNA [48–51]. The first such report identified fragments of the H/ACA box snoRNA ACA45 in association with Ago proteins from the miRNA RISC complex. Like other miRNAs, the processing of ACA45 is dependent on Dicer, and furthermore, the resulting 20–22 nt fragments were validated to function as a repressive miRNA on the expression of an identified target (CDC2L6).[48] While only a small fraction of the total cellular ACA45 is processed by Dicer to act as miRNA, this clearly represents a role for snoRNA derivatives in the cytoplasm. A more recent publication has identified another 11 C/D box snoRNAs that are processed to miRNA-sized species and have verified gene-silencing activity [51]. Clearly, snoRNAs represent a large group of small ncRNA with many diverse functions, many of which are only just beginning to be understood. We do not yet precisely understand the utility of snoRNA-guided rRNA modifications, though they are evolutionarily conserved and clearly affect ribosome function. snoRNA-guided modifications of mRNAs seem possible and could affect splicing [52], translation, or mRNA stability, but this area is essentially unstudied. Finally, newly recognized noncanonical roles such as regulation of alternative splicing
One of the most exciting aspects of these ncRNA is the apparent ease with which they can be depleted or “knocked down” both in cell culture and in vivo using antisense oligonucleotide technologies. Since some miRNA and snoRNA are redundantly encoded by multiple genetic loci, they can be difficult to knockout with traditional genetic approaches in mice. In vivo knockdown should therefore significantly improve our ability to characterize the physiologic roles of these ncRNA. For miRNA, the development of “antagomirs” and similar chemically-modified antisense oligonucleotides for targeted knockdown of specific miRNA in vivo has been quite successful [53– 56]. As an example, antagomirs are antisense oligonucleotides synthesized with 2′-O-Me ribose nucleotides and phosphorothioate linkages to resist nuclease digestion, plus a 3′-cholesterol moiety to improve cellular uptake. The initial report using this chemistry in 2005 showed excellent knockdown of miRNA-122 in mice across all tissues tested except brain. Furthermore, since miRNA-122 regulates HMG-CoA reductase levels, the knockdown was shown to be physiologically relevant, with antagomir-mediated knockdown leading to decreased cholesterol levels [53]. Subsequent publications have shown similar success in non-human primates [55], as well as in models of cardiac hypertrophy [54] and breast cancer [56], Several companies have already been formed with the goal of developing novel therapeutics targeting miRNA, including Regulus Therapeutics, miRagen, and Mirna Therapeutics. Antisense oligonucleotide approaches to modulating snoRNA levels have been successful only recently, with an in vitro technique amenable to cell culture being introduced in 2009 [57], and the first in vivo knockdown in mice reported in late 2010 [58]. In summary, this brief review highlights the fact that much has been learned about the fundamental biology of ncRNA, but that there are many more challenges remaining. For example, identification of these small ncRNA has been considerably easier than identifying their targets, despite the fact that target interactions are largely directed by sequence complementarity. The problem is complicated by apparently non-canonical mechanisms of action, at least for some snoRNAs. Thus, while bioinformatic approaches have been useful, there is much work to be done at the bench in order to truly identify and verify targets. Presumably, the use of primary biochemical approaches to identify relevant
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interactors will speed this process. Beyond that, there will be much more to be learned about just how these small ncRNA actually influence so many aspects of cellular and organismal biology. The study of ncRNA is still really only in its infancy, but it promises to be a rich field of study for years to come. Acknowledgements This study was supported by funds from National Institutes of Health (T32-HL007081).
References 1. Lee RC, Feinbaum RL, Ambros V. The C. elegans heterochronic gene lin-4 encodes small RNAs with antisense complementarity to lin-14. Cell. 1993;75:843–54. 2. Zhao Y, Ransom JF, Li A, et al. Dysregulation of cardiogenesis, cardiac conduction, and cell cycle in mice lacking miRNA-1-2. Cell. 2007;129:303–17. 3. Xiao C, Calado DP, Galler G, et al. MiR-150 controls B cell differentiation by targeting the transcription factor c-Myb. Cell. 2007;131:146–59. 4. van Rooij E, Sutherland LB, Qi X, Richardson JA, Hill J, Olson EN. Control of stress-dependent cardiac growth and gene expression by a microRNA. Science. 2007;316:575–9. 5. Calin GA, Ferracin M, Cimmino A, et al. A MicroRNA signature associated with prognosis and progression in chronic lymphocytic leukemia. N Engl J Med. 2005;353:1793–801. 6. Matkovich SJ, Van Booven DJ, Youker KA, et al. Reciprocal regulation of myocardial microRNAs and messenger RNA in human cardiomyopathy and reversal of the microRNA signature by biomechanical support. Circulation. 2009;119:1263–71. 7. Rodriguez A, Griffiths-Jones S, Ashurst JL, Bradley A. Identification of mammalian microRNA host genes and transcription units. Genome Res. 2004;14:1902–10. 8. Baskerville S, Bartel DP. Microarray profiling of microRNAs reveals frequent coexpression with neighboring miRNAs and host genes. RNA. 2005;11:241–7. 9. Cai X, Hagedorn CH, Cullen BR. Human microRNAs are processed from capped, polyadenylated transcripts that can also function as mRNAs. RNA. 2004;10:1957–66. 10. Yi R, Qin Y, Macara IG, Cullen BR. Exportin-5 mediates the nuclear export of pre-microRNAs and short hairpin RNAs. Genes Dev. 2003;17:3011–16. 11. Hutvagner G, McLachlan J, Pasquinelli AE, Balint E, Tuschl T, Zamore PD. A cellular function for the RNA-interference enzyme Dicer in the maturation of the let-7 small temporal RNA. Science. 2001;293:834–8. 12. Khvorova A, Reynolds A, Jayasena SD. Functional siRNAs and miRNAs exhibit strand bias. Cell. 2003;115:209–16. 13. Krol J, Krzyzosiak WJ. Structural aspects of microRNA biogenesis. IUBMB Life. 2004;56:95–100. 14. Lin SL, Chang D, Ying SY. Asymmetry of intronic pre-miRNA structures in functional RISC assembly. Gene. 2005;356:32–8. 15. Okamura K, Phillips MD, Tyler DM, Duan H, Chou YT, Lai EC. The regulatory activity of microRNA* species has substantial influence on microRNA and 3' UTR evolution. Nat Struct Mol Biol. 2008;15:354–63. 16. Liu J, Carmell MA, Rivas FV, et al. Argonaute2 is the catalytic engine of mammalian RNAi. Science. 2004;305:1437–41. 17. Liu J, Rivas FV, Wohlschlegel J, Yates III JR, Parker R, Hannon GJ. A role for the P-body component GW182 in microRNA function. Nat Cell Biol. 2005;7:1261–6.
Cardiovasc Drugs Ther (2011) 25:151–159 18. Ambros V. The functions of animal microRNAs. Nature. 2004;431:350–5. 19. Kim DH, Saetrom P, Snove Jr O, Rossi JJ. MicroRNA-directed transcriptional gene silencing in mammalian cells. Proc Natl Acad Sci U S A. 2008;105:16230–5. 20. Wightman B, Ha I, Ruvkun G. Posttranscriptional regulation of the heterochronic gene lin-14 by lin-4 mediates temporal pattern formation in C. elegans. Cell. 1993;75:855–62. 21. Olsen PH, Ambros V. The lin-4 regulatory RNA controls developmental timing in Caenorhabditis elegans by blocking LIN-14 protein synthesis after the initiation of translation. Dev Biol. 1999;216:671–80. 22. Guo H, Ingolia NT, Weissman JS, Bartel DP. Mammalian microRNAs predominantly act to decrease target mRNA levels. Nature. 2010;466:835–40. 23. Vasudevan S, Tong Y, Steitz JA. Switching from repression to activation: microRNAs can up-regulate translation. Science. 2007;318:1931–4. 24. Orom UA, Nielsen FC, Lund AH. MicroRNA-10a binds the 5'UTR of ribosomal protein mRNAs and enhances their translation. Mol Cell. 2008;30:460–71. 25. Berezikov E, Guryev V, van de Belt J, Wienholds E, Plasterk RH, Cuppen E. Phylogenetic shadowing and computational identification of human microRNA genes. Cell. 2005;120:21–4. 26. Bartel DP. MicroRNAs: target recognition and regulatory functions. Cell. 2009;136:215–33. 27. Doench JG, Sharp PA. Specificity of microRNA target selection in translational repression. Genes Dev. 2004;18:504–11. 28. Lai EC, Tam B, Rubin GM. Pervasive regulation of Drosophila Notch target genes by GY-box-, Brd-box-, and K-box-class microRNAs. Genes Dev. 2005;19:1067–80. 29. Brennecke J, Stark A, Russell RB, Cohen SM. Principles of microRNA-target recognition. PLoS Biol. 2005;3:e85. 30. Lewis BP, Shih IH, Jones-Rhoades MW, Bartel DP, Burge CB. Prediction of mammalian microRNA targets. Cell. 2003;115:787– 98. 31. Griffiths-Jones S, Grocock RJ, van DS, Bateman A, Enright AJ. miRBase: microRNA sequences, targets and gene nomenclature. Nucleic Acids Res. 2006;34:D140–44 32. Krek A, Grun D, Poy MN, et al. Combinatorial microRNA target predictions. Nat Genet. 2005;37:495–500. 33. Kiriakidou M, Nelson PT, Kouranov A, et al. A combined computational-experimental approach predicts human microRNA targets. Genes Dev. 2004;18:1165–78. 34. Miranda KC, Huynh T, Tay Y, et al. A pattern-based method for the identification of MicroRNA binding sites and their corresponding heteroduplexes. Cell. 2006;126:1203–17. 35. Karginov FV, Conaco C, Xuan Z, et al. A biochemical approach to identifying microRNA targets. Proc Natl Acad Sci U S A. 2007;104:19291–6. 36. Matkovich SJ, Van Booven DJ, Eschenbacher WH, Dorn GW. RISC RNA Sequencing for Context-Specific Identification of In Vivo MicroRNA Targets. Circ Res. 2010. 37. Filipowicz W, Pelczar P, Pogacic V, Dragon F. Structure and biogenesis of small nucleolar RNAs acting as guides for ribosomal RNA modification. Acta Biochim Pol. 1999;46:377– 89. 38. Liang XH, Liu Q, Fournier MJ. Loss of rRNA modifications in the decoding center of the ribosome impairs translation and strongly delays pre-rRNA processing. RNA. 2009;15:1716–28. 39. Matera AG, Terns RM, Terns MP. Non-coding RNAs: lessons from the small nuclear and small nucleolar RNAs. Nat Rev Mol Cell Biol. 2007;8:209–20. 40. Narayanan A, Lukowiak A, Jady BE, et al. Nucleolar localization signals of box H/ACA small nucleolar RNAs. EMBO J. 1999;18:5120–30.
Cardiovasc Drugs Ther (2011) 25:151–159 41. Samarsky DA, Fournier MJ, Singer RH, Bertrand E. The snoRNA box C/D motif directs nucleolar targeting and also couples snoRNA synthesis and localization. EMBO J. 1998;17:3747–57. 42. Aftab MN, He H, Skogerbo G, Chen R. Microarray analysis of ncRNA expression patterns in Caenorhabditis elegans after RNAi against snoRNA associated proteins. BMC Genomics. 2008;9:278. 43. Reichow SL, Hamma T, Ferre-D'Amare AR, Varani G. The structure and function of small nucleolar ribonucleoproteins. Nucleic Acids Res. 2007;35:1452–64. 44. Kishore S, Stamm S. The snoRNA HBII-52 regulates alternative splicing of the serotonin receptor 2 C. Science. 2006;311:230–2. 45. Sahoo T, del GD, German JR, et al. Prader-Willi phenotype caused by paternal deficiency for the HBII-85 C/D box small nucleolar RNA cluster. Nat Genet. 2008;40:719–21. 46. Kiss T, Fayet-Lebaron E, Jady BE. Box H/ACA small ribonucleoproteins. Mol Cell. 2010;37:597–606. 47. Kishore S, Khanna A, Zhang Z, et al. The snoRNA MBII-52 (SNORD 115) is processed into smaller RNAs and regulates alternative splicing. Hum Mol Genet. 2010;19:1153–64. 48. Ender C, Krek A, Friedlander MR, et al. A human snoRNA with microRNA-like functions. Mol Cell. 2008;32:519–28. 49. Taft RJ, Glazov EA, Lassmann T, Hayashizaki Y, Carninci P, Mattick JS. Small RNAs derived from snoRNAs. RNA. 2009;15:1233–40.
159 50. Scott MS, Avolio F, Ono M, Lamond AI, Barton GJ. Human miRNA precursors with box H/ACA snoRNA features. PLoS Comput Biol. 2009;5:e1000507. 51. Brameier M, Herwig A, Reinhardt R, Walter L, Gruber J. Human box C/D snoRNAs with miRNA like functions: expanding the range of regulatory RNAs. Nucleic Acids Res. 2010 52. Ge J, Liu H, Yu YT. Regulation of pre-mRNA splicing in Xenopus oocytes by targeted 2′-O-methylation. RNA. 2010;16:1078–85. 53. Krutzfeldt J, Rajewsky N, Braich R, et al. Silencing of microRNAs in vivo with ‘antagomirs’. Nature. 2005;438:685–9. 54. Care A, Catalucci D, Felicetti F, et al. MicroRNA-133 controls cardiac hypertrophy. Nat Med. 2007;13:613–18. 55. Elmen J, Lindow M, Schutz S, et al. LNA-mediated microRNA silencing in non-human primates. Nature. 2008;452:896–9. 56. Ma L, Reinhardt F, Pan E, et al. Therapeutic silencing of miR-10b inhibits metastasis in a mouse mammary tumor model. Nat Biotechnol. 2010;28:341–7. 57. Ideue T, Hino K, Kitao S, Yokoi T, Hirose T. Efficient oligonucleotide-mediated degradation of nuclear noncoding RNAs in mammalian cultured cells. RNA. 2009;15:1578–87. 58. Liang XH, Vickers TA, Guo S, Crooke ST. Efficient and specific knockdown of small non-coding RNAs in mammalian cells and in mice. Nucleic Acids Res 2010. 59. van Rooij E, Olson EN. MicroRNAs: powerful new regulators of heart disease and provocative therapeutic targets. J Clin Invest. 2007;117:2369–76.
