INSIGHT REGULATORY RNA
INSIGHT 16 February 2012 / Vol 482 / Issue No 7385
R E G U L AT O RY R N A
REVIEWS 322 Functional complexity and regulation through RNA dynamics The conformational changes of coding and non-coding RNA provide an important source of complexity that drives many of the fundamental processes of life. The unique properties of RNA allow changes in its structure to occur in a robust and biologically specific way. A wide range of cellular inputs are used to interface with RNA dynamics, and various mechanisms are used to guide these dynamics. The conventional view that one sequence codes for one structure and function is now being replaced with a more nuanced view of how RNA conformational states can lead to a broad range of functional outcomes. Studying RNA dynamics within an in vivo environment will be an important goal for the future. Elizabeth A Dethoff, Jeetender Chugh, Anthony M Mustoe & Hashim M Al-Hashimi
331 RNA-guided genetic silencing systems in bacteria and archaea In response to viral challenge and plasmid transfer, bacteria and archaea integrate short fragments of foreign nucleic acid into the host chromosome at a locus known as CRISPR (clustered regularly interspaced short palindromic repeat). Such elements contain a record of all encounters with foreign invaders. After transcription, the long primary CRISPR transcript is processed into short RNAs that contain sequences complementary to previously encountered invading nucleic acids. Recognition of complementary sequences in invading DNA leads to their elimination, providing immunity to further infection. Blake Wiedenheft, Samuel H Sternberg &Â Jennifer A Doudna
have been discovered still needs to be explained. Studies have begun to explore the biological significance and mechanism of large non-coding RNAs, and classic examples of these can be used to form a framework for understanding the principles of RNA interactions. In the model suggested in this review, long noncoding RNAs are proposed to contain
339 Modular regulatory principles of large non-coding RNAs Although it is clear that RNAs have diverse roles, the functional significance of some of the classes of large and small transcripts that
Ratcheting motion of a ribosome.
discrete domains that interact with specific proteins to form a flexible modular scaffold. Targeting of the non-coding RNAâ€“protein complex to achieve its function is probably facilitated by interactions with DNA or other RNA. Mitchell Guttman & John L Rinn
347 The microcosmos of cancer MicroRNAs are small, non-coding RNAs that modulate the expression of a large fraction of genes. Some microRNAs, on the basis of the genes they regulate, can have tumour-suppressor or oncogenic properties. Alterations in microRNA sequence or expression can drive cancer initiation and progression. Inhibiting the function of these oncogenic microRNAs, or engineering synthetic microRNAs to suppress oncogenes, are new strategies for therapeutic intervention. In addition, profiling of microRNAs may be a way to classify cancers, and they may eventually find a place alongside conventional treatments for cancer. Amaia Lujambio & Scott W Lowe
1 6 F E B R UA RY 2 0 1 2 | VO L 4 8 2 | N AT U R E | 2 7 3
INSIGHT REGULATORY RNA 16 February 2012 / Vol 482 / Issue No 7385
A Cover illustration by Nik Spencer
Editor, Nature Philip Campbell Publishing Nick Campbell Insights Editor Ursula Weiss Production Editor Nicola Bailey Art Editor Nik Spencer Sponsorship Gerard Preston Production Emilia Orviss Marketing Elena Woodstock Hannah Phipps Editorial Assistant Hazel Mayhew
The Macmillan Building 4 Crinan Street London N1 9XW, UK Tel: +44 (0) 20 7833 4000 e: email@example.com
lthough proponents of RNA might beg to differ, in the hierarchy of popular interest, DNA has historically held more sway. Being able to decipher genomes was seen as a milestone on the way to understanding life itself. What genome-wide RNA sequencing studies have revealed, however, is the unexpected complexity of RNA species encoded by DNA, most of which do not code for a protein. We now appreciate that such non-coding RNAs exert important regulatory controls on many biological processes. The reviews in this Insight illustrate some of these principles. RNA is synthesized as a single-stranded molecule, but it is able to base-pair with itself, other RNA molecules or DNA. Hashim Al-Hashimi and colleagues discuss how secondary and tertiary structures of RNA are influenced by external cues to elicit a specific functional output. The cell exploits this dynamism to regulate processes such as transcription, post-transcriptional processing and translation. Jennifer Doudna and colleagues review a microbial adaptive immune system, CRISPR (clustered regularly interspaced short palindromic repeat). This system incorporates small pieces of invading viral or plasmid sequences into the bacterial genome as CRISPR loci; when future invasions occur, the expressed CRISPR RNAs recognize the foreign nucleic acids and mediate their degradation. The physiological function of many long non-coding RNAs remains undetermined, but Mitchell Guttman and John Rinn propose a model in which these molecules act in a modular fashion to bind different proteins or hybridize to various DNAs or RNAs; this modularity expands the scope of a single RNA’s function. Finally, Amaia Lujambio and Scott Lowe highlight the role of another class of much shorter, non-coding RNA — microRNAs — in cancer development and suppression, and as a target for therapeutic intervention. We hope these reviews provide a flavour of how the inherent properties of RNA make it a robust species to regulate cellular processes. Angela K. Eggleston, Alex Eccleston, Barbara Marte & Claudia Lupp Senior Editors
REVIEWS 322 Functional complexity and regulation through RNA dynamics Elizabeth A. Dethoff, Jeetender Chugh, Anthony M. Mustoe & Hashim M. Al-Hashimi
331 RNA-guided genetic silencing systems in bacteria and archaea Blake Wiedenheft, Samuel H. Sternberg & Jennifer A. Doudna
339 Modular regulatory principles of large non-coding RNAs Mitchell Guttman & John L. Rinn
347 The microcosmos of cancer Amaia Lujambio & Scott W. Lowe TSP
1 6 F E B R UA RY 2 0 1 2 | VO L 4 8 2 | N AT U R E | 3 2 1
Functional complexity and regulation through RNA dynamics Elizabeth A. Dethoff1, Jeetender Chugh1, Anthony M. Mustoe1 & Hashim M. Al-Hashimi1
Changes to the conformation of coding and non-coding RNAs form the basis of elements of genetic regulation and provide an important source of complexity, which drives many of the fundamental processes of life. Although the structure of RNA is highly flexible, the underlying dynamics of RNA are robust and are limited to transitions between the few conformations that preserve favourable base-pairing and stacking interactions. The mechanisms by which cellular processes harness the intrinsic dynamic behaviour of RNA and use it within functionally productive pathways are complex. The versatile functions and ease by which it is integrated into a wide variety of genetic circuits and biochemical pathways suggests there is a general and fundamental role for RNA dynamics in cellular processes.
nalysis of the first X-ray structure of the protein myoglobin1 prompted researchers to ask the question: how do ligands reach the deeply buried haem group centre? This simple, but powerful, observation has inspired decades of investigation into the dynamic behaviour of proteins, so that we now know protein structures are in constant motion, and that these fluctuations in structure are crucial to, and sometimes drive, function. Early X-ray structures of RNA contained indications of the importance of conformational dynamics: large changes in the helical arms of transfer RNA were observed on the binding of tRNA synthetase2, and changes in the conformation of ribozymes needed to be invoked to envision catalytically active states3–5. However, no one could have anticipated the existence of new genetic circuits that are based on RNA conformational switches, or that the ‘acrobatic’ nature of a biopolymer that consists of only four chemically similar nucleotides would be at the centre of a complex macromolecular structure such as the ribosome. The dynamic changes that occur in the structure of RNA serve an ever-increasing range of functionality that generally follows a common two-step process (see Supplementary Information for more reviews on RNA dynamics). The process involves a cellular signal that triggers RNA dynamics, which are then transduced into a specific biological output. This Review provides a critical account of RNA dynamics as a regulatory mechanism and source of functional complexity. We review the known dynamic properties of RNA structure and emphasize the unique properties that allow large changes in structure to take place in a biologically specific and robust manner. We then examine the wide range of cellular inputs used to interface with RNA dynamics and the various mechanisms that are used to guide the dynamics to achieve a broad spectrum of functional outputs.
RNA free-energy landscape It is important to distinguish between the two types of RNA dynamics: ‘equilibrium fluctuations’ and ‘conformational transitions’. Equilibrium fluctuations are related to the thermal activated motions that occur in RNA. Conformational transitions arise when cellular cues, such as an increase in the concentration of a metabolite, create a nonequilibrium state that then relaxes back to equilibrium. This Review is focused principally on conformational transitions because of their dominant role in regulatory mechanisms; however, the two motions are intricately related, as highlighted by studies of RNA and protein 1
dynamics6,7. This, and other aspects of RNA dynamic behaviour that are relevant to function, is best understood by looking at the free-energy landscape of RNA8,9. The free-energy landscape specifies the free energy of every possible RNA conformation (Fig. 1a). Equilibrium fluctuations correspond to the spontaneous jumps that occur between various conformers along the free-energy landscape. The population of a given conformer depends on its free energy, whereas the transition rate between conformers depends on the free-energy barrier of separation (Fig. 1a). Conformational transitions arise when cellular cues perturb the freeenergy landscape, which leads to a redistribution of conformational states (Fig. 1a). The RNA free-energy landscape is punctuated by deep local minima, or conformational wells, in which conformations within a well are highly similar and conformations from different wells are structurally distinct. These are the conformations that are significantly sampled by equilibrium motions and are stabilized by cellular cues to effect conformational transitions10–12 (Fig. 1a). For example, the degeneracy of base-pairing and stacking interactions, together with the high stability of RNA duplexes, results in deep local minima that correspond to different but energetically equal secondary structures that are separated by large kinetic barriers13 (Fig. 1b). As few as two secondary structures can dominate the RNA dynamic landscape because the loss of energy that accompanies the disruption of just one base-pair can markedly destabilize alternative conformations. In addition, RNA of a given secondary structure can undergo more facile dynamic excursions in tertiary structure, which involve smaller energetic barriers. These dynamics are commonly dominated by large changes in the relative orientation of helical domains, which carry motifs involved in tertiary contacts, and occur around flexible pivot points consisting of bulges, internal loops and higher-order junctions (Fig. 1c). Although these excursions can lead to very large changes in tertiary structure, they are limited to a narrow set of conformations. For example, calculating the set of conformations that are accessible to two helices that are connected by a three-residue bulge reveals that the interhelical bend angle, when combined with interhelical twisting, can range from 0° to 180°. Despite this large accessible range, the connectivity constraints that are imposed by the bulge junction and the steric forces that act on the two helices direct changes in the interhelical orientations along a highly directional pathway and therefore restrict the conformational space to less than 20% of what
Department of Chemistry and Biophysics, The University of Michigan, 930 North University Avenue, Ann Arbor, Michigan 48109-1055, USA.
3 2 2 | N AT U R E | VO L 4 8 2 | 1 6 F E B R UA RY 2 0 1 2
REVIEW INSIGHT a
b Free energy
U C C A A G C G U U G U G U C C G C
Te tra rtiar ns y itio n
ary nd co ition e S ns tra
Bistable RNA G
G U C C
C G G G U U C G 3′
5′ A C A G G U U
G G G U
A A G C G U U G U
U U C G C G G A C A 5′
U C G 3′
C U G
U C C G C U U G G A C A 5′
U G U G U C G A A C C U G C G G G U U C G 3′
Figure 1 | Shape and form of RNA dynamics. a, The secondary and tertiary RNA conformations of different low-lying energy states are shown above the RNA free-energy landscape (green line). The relative populations of each conformation are shown within the landscape (red balls). Cellular effectors (bolts) can modify the energy landscape to favour an alternative secondary structure (top), or preferentially stabilize an alternate tertiary conformation (bottom). b, Exchange between alternative, isoenergetic secondary structures (A and B) that are separated by large energetic barriers owing to disruption of base pairs in the transition state13. c, The accessible range of interhelical
conformations for an RNA two-way junction consisting of a trinucleotide bulge, with the possible paths of the bulge, which were excluded during the modelling, illustrated in red14,15. The allowed range of conformations is restricted towards a specific and directed conformational pathway by steric and stereochemical forces. The structure is rotated 90° to illustrate the bending (left) and twisting (right) motion. d, Flipping out of a residue (red) participating in a noncanonical base pair within an RNA internal loop is illustrated, progressing from an intrahelical stacked to an extrahelical unstacked conformation. The motion occurs without perturbing flanking Watson–Crick pairs (green).
is theoretically possible10,14–16 (Fig. 1c). In addition, owing to the high stability of duplexes, residues participating in non-canonical base pairs can loop-out from intra- to extrahelical conformations without significantly disturbing the structure of the flanking helices17,18 (Fig. 1d). Precise control over these dynamics is encoded within the sequence, and small sequence variations can greatly alter the relative populations of different RNA structures and their rates of interconversion11,19. For example, distinct interhelical orientations can be sampled by changing the length and asymmetry of junctions10,14,15, and the tendency of residues to loop-out can be modulated on the basis of sequence-specific stacking interactions20,21 (see Supplementary Information for links to movies and animations of experimentally determined RNA dynamics). These features can help to explain the three remarkable aspects of RNA conformational transitions that are of fundamental importance for regulatory functions. First, the landscape is hierarchical due to the height of the energy barriers that separate alternative secondary structures. Changes in tertiary contacts rarely involve changes in the secondary structure and the two types of conformational changes can be used to serve different functions. Throughout this review, we will use ‘secondary’ and ‘tertiary’ conformational changes to distinguish between these two types of dynamics. Second, the limited landscape of energetically favourable conformations allows RNA to undergo very large changes in structure, but to be directed towards a very specific set of conformations from a vast number of possibilities. Third,
there is increasing evidence that RNA dynamics are determined by the underlying RNA free-energy landscape, and to lesser extent by cellular cues7,22,23. Thus, conformational transitions can be considered perturbations that guide pre-existing equilibrium fluctuations towards specific functionally productive pathways. In this way, even an imperfect force or cellular signal will drive changes in the RNA structure along a predetermined pathway, which makes the transitions highly robust.
Triggers of RNA conformational transitions RNA dynamics can be triggered by a remarkably diverse set of molecular effectors and environmental cues through several different mechanisms. This provides many different points of entry for integrating RNA conformational transitions into biological circuits and biochemical pathways. Specific protein binders The most common effectors are proteins that bind their target RNA specifically through well-defined structural features, thereby stabilizing one or a subset of conformations from the pre-existing energy landscape. For example, the mitochondrial tyrosyl-tRNA synthetase CYT18 from Neurospora crassa binds specifically to group-I introns (a class of large self-splicing ribozymes that catalyse their own excision from messenger RNA, tRNA and ribosomal RNA precursors) and stabilizes the conformation required for catalytic activity24. Protein binding often leads to large changes in the overall orientation of RNA helices around 1 6 F E B R UA RY 2 0 1 2 | VO L 4 8 2 | N AT U R E | 3 2 3
INSIGHT REVIEW junctions such as bulges25, three-way junctions26 and other motifs such as the K-turn27. For example, the spliceosomal U4 small nuclear RNA (snRNA) undergoes a sharp transition in the interhelical bend angle, from approximately 69° to 25°, around a K-turn motif, when it binds to its cognate protein target28 (Fig. 2a). These changes in interhelical conformation are driven in part by nonspecific electrostatic interactions between basic amino acids and the high negative-charge density that builds up at interhelical junctions, and are often observed as equilibrium dynamics in the absence of an effector29–31. For example, unbound HIV-1 transactivating response RNA (TAR) dynamically samples the many different interhelical orientations that are observed when it is bound to seven distinct ligands, including peptide mimics of its cognate protein, Tat31 (Fig. 2b). In an increasing number of cases, protein binding does not involve the stabilization of a specific minimum of the RNA free-energy landscape. Instead, binding selectively lowers the surrounding energy barriers to accentuate or alter the equilibrium dynamics of the RNA. For example, binding of the U1A protein to its cognate RNA target does not cause the pre-existing equilibrium interhelical motions to stop, but rather induces mobility in regions of the RNA that are in direct contact with the protein32. The CBP2 protein from yeast mitochondria binds specifically to the bI5 group-I intron and activates large-scale RNA equilibrium motions33. Even simple small-molecule ligands lead a
U4 - U6 stem II
U2 - U6 U6 stem stem I
5´ 5´ exon
RNA chaperones and helicases As is often the case in RNAs that possess alternative secondary structures, the large energy barriers associated with base-pair melting can limit the dynamics between RNA conformational wells. In this way, the RNA can become kinetically trapped in a metastable, non-equilibrium conformation. In response to this, a variety of proteins have evolved that possess the RNA ‘chaperone’ activity needed to efficiently drive RNA secondary structural-transitions over the large energy barriers35,36. For example, the HIV nucleocapsid protein uses non-specific interactions between the RNA and protein to destabilize the RNA helices37. This lowers the energetic barrier to conformational exchange, accelerating relaxation to equilibrium and allowing metastable RNAs to convert to conformations that are more thermodynamically favourable. Other RNA chaperones, such as RNA helicases, help RNA traverse high energy barriers by unwinding helices and disrupting RNA structure, as well as promoting the formation of new RNA duplexes to accelerate conformational transitions in RNAs and ribonucleoprotein (RNP)
U4 - U6 stem I 2
to the reorganization of the TAR RNA equilibrium dynamics34. These observations highlight the importance of embracing a broader view of trigger factors as elements that perturb the entire energy landscape and guide RNA dynamics rather than simply stabilize a single conformation from a dynamic range.
DExD or H-box U2 - U6 helicases stem II ATP U6
P3 P1 5´
Transcription on RNAP
e Transcription off
5´ 5´ exon
gly gly (Un)
!" ! U20 C75
Catalytic core conformation
Figure 2 | Triggering RNA conformational transitions. a, Conformational changes in the spliceosomal U4 snRNA K-turn motif (Protein Data Bank (PDB) ID 2KR8) triggered when it binds to a complex of the human protein PRP31 and the 15.5K protein (PDB ID 2OZB)28. b, Similarity between the TAR RNA interhelical conformations that are triggered by binding to small molecules, Tat peptide derivatives and divalent ions (grey helices) and those that are sampled by equilibrium dynamics (green helices labelled as 1,2 and 3) in the unbound state shown as a horizontal and vertical view. The path of helix II (HII) as it moves from one unbound, equilibrium conformer to the next is shown by the orange arrows. HI, helix 1. Figure modified, with permission, from ref. 31. c, RNA conformation transitions during spliceosome assembly on pre-mRNA (dashed line) in the presence of DExD or H-box helicases and ATP. Sections are colour-coded to indicate base-pairing after helicase action. d, The RNA structure is modulated by steering of the co-transcriptional folding pathway. The adenine transcriptionterminating riboswitch is a typical example. The progression of co-transcriptional 3 2 4 | N AT U R E | VO L 4 8 2 | 1 6 F E B R UA RY 2 0 1 2
folding with and without the ligand (adenine) is shown. Adenine binds to the apatamer domain and stabilizes the structure, allowing transcription to be turned on. RNAP, RNA polymerase. e, Two examples of tandem riboswitch architectures. Left, cooperative binding of glycine by the glycine riboswitch using tandem aptamer domains and one expression platform. Right, tandem SAM and AdoCbl riboswitches in which either of the two ligands triggers the conformational switch and yields an output of gene repression. Sequences that can form transcription terminator stems are shown in red. f, Conformations of HDV ribozyme precleavage (PDB ID 1VC7)53 and post-cleavage (PDB ID 1DRZ)52 states. Enlarged details of the catalytic core (dashed box) of the two structures are shown, with the bound substrate (green) and the magnesium ion (yellow) present only in the precleavage state. g, Melting of the secondary structure around the ribosome-binding site of virulence genes in the pathogen is triggered by an increase in temperature that makes the Shine-Dalgarno sequence (SD, blue) available for ribosome binding and translation initiation.
REVIEW INSIGHT complexes38. These chaperone proteins are important for remodelling the structure of RNA and RNP complexes because they can anneal or unwind RNA strands depending on the environmental cues39. For example, helicases are essential in the assembly of the spliceosome, which is a complex RNP that consists of five RNAs and multiple proteins that catalyses excision of introns from a nuclear pre-mRNA40,41. Assembly proceeds through a series of transitions that involve the melting and annealing of RNA duplexes that are catalysed by DExD/H-box ATPase helicases (Fig. 2c). For example, the U4 RNA escorts the U4–U6–U5 triple small nuclear RNP complex (tri-snRNP) to the pre-mRNA, but is subsequently released by the DExD/H-box helicase Brr2, which catalyses the melting of the two stems within U4 and U6. This frees the U6 stem to base-pair with U2 snRNA and leads to a new RNA structure that is required for the first transesterification reaction42 (Fig. 2c). In addition, DExD/H-box proteins are involved in the release of mRNA produced in pre-mRNA splicing reactions. For example, the DEAH-box splicing factor Prp22 is deposited on spliced mRNA downstream of the exon–exon junction and catalyses the disruption of contacts between mRNA and U5 snRNP, thereby releasing the spliced mRNA from the U5–U6–U2 spliceosomal assembly43. In another example of the variety of functions of RNA chaperones, the DExD/H-box protein CYT19 unfolds native and misfolded conformations of a group-I catalytic RNA in an ATP-dependent process. A large free-energy gap between the native and misfolded conformers directs CYT-19 to unfold misfolded conformers more frequently than native conformers. In the process, CYT-19 redistributes the two conformation populations, which allows native RNA to populate a wider range of conformations than would otherwise be possible44. Metabolites and physiochemical conditions Another ingenious strategy is used to modulate RNA structure in response to a wide range of metabolite-based effectors, including small molecules (such as amino-acids, coenzymes and nucleotides23,45) and changes in physiochemical conditions (such as magnesium ion concentration46 and pH47). It would be difficult, if not impossible, for these smaller effectors and cellular cues to possess the chaperone activity needed to efficiently drive secondary structural transitions over the associated large energy barriers. Instead, this strategy operates on the initial RNA-folding process itself, intervening while the energy barriers are still low. Specifically, these effectors and cues act by directing the RNA to different folding pathways during RNA co-transcriptional folding. This process is made possible by the unidirectional and comparatively slow rate with which RNA is transcribed from the 5ʹ to the 3ʹ direction relative to RNA folding and effector binding. Each pathway favours one of two distinct secondary structures, where each secondary structure is associated with an alternative biological outcome (Fig. 2d). This trigger mechanism is implicated in a growing list of other RNA switches, although it has been best described for metabolite-sensing riboswitches23,45. Riboswitches are RNA-based genetic elements typically embedded in the 5ʹ untranslated region of bacterial genes that regulate expression of metabolic genes in response to changes in cellular metabolite concentration23,45. In a prototypical metabolite riboswitch, a metabolite, such as adenine, binds to the aptamer domain with exceptional affinity and selectivity. This stabilizes an otherwise shallow energy well, which induces a redistribution of the aptamer conformational states towards one state that, in most riboswitches, sequesters an RNA element into a helix of the aptamer domain48 (Fig. 2d). In turn, the unavailability of the RNA element changes the folding pathway of a downstream decisionmaking expression platform, directing it towards structures that turn off (and in some cases, turn on) gene expression, either by forming a transcription-terminating helix (Fig. 2d) or by sequestering the ShineDalgarno sequence (a ribosome binding site located eight base pairs upstream of the start codon in mRNA), thereby inhibiting translation. This system also keeps the number of spontaneous conformational transitions, or premature switching in the absence of ligands, to a minimum because very large energy barriers separate the two alternative
secondary structural forms of the expression platform. More complex functionality can be achieved by coupling multiple riboswitches together. For example, the glycine riboswitch uses two aptamer domains in tandem to cooperatively bind glycine, thereby increasing responsiveness to changes in ligand concentrations 49 (Fig. 2e). The tandem arrangement of two entire riboswitches that respond to two distinct ligands allows the construction of more sophisticated genetic circuits such as two-input Boolean NOR logic gates, in which either of the two ligands can trigger the conformational switch and yield an output of gene repression50 (Fig. 2e). In another example, the c-di-GMP-sensing riboswitch and a GTP-dependent self-splicing group-I ribozyme in the 5ʹ untranslated region of a putative Clostridium difficile virulence gene work in tandem to regulate translation51. In the presence of c-di-GMP and GTP, the riboswitch and ribozyme form a structure that stabilizes a 5ʹ splice site, and the ribozyme self-splices to yield an RNA transcript with a perfect ribosome-binding site located upstream of the start codon. Conversely, in the presence of GTP alone, alternative base pairing between the riboswitch and ribozyme occurs to form a structure that promotes splicing at an alternative site, which results in a splicing product without a ribosome-binding site, and thus downregulates translation. This RNA arrangement represents the first natural example of an allosteric ribozyme. Chemical reactions Chemical reactions, such as cleavage of the RNA phosphodiester backbone, can also reshape the underlying RNA energy landscape so that a state that was previously in equilibrium becomes a non-equilibrium state, which triggers changes in RNA secondary and tertiary structure. For example, X-ray analysis of the structures of precursor and product states of the hepatitis delta virus (HDV) ribozyme, which catalyses site-specific self-cleavage of the viral RNA phosphodiester backbone, reveal changes in the local arrangement of catalytic groups, as well as the ejection of a catalytically important magnesium ion52. These conformational changes may help to accelerate product release53,54 (Fig. 2f). Another example is seen in the secondary structural switch triggered by cleavage of the 3ʹ end of the pre-18S rRNA during eukaryotic ribosome maturation, which is used to enforce a sequential order to the maturation process55. Thermal and mechanical triggers Other energy-dependent processes can induce the complete ‘melting’ of RNA helices. RNA thermosensors alter expression of genes during heatshock response and pathogenic invasion in response to increases in temperature56 (Fig. 2g). For example, when Listeria monocytogenes invades an animal host, the pathogen enters a warmer environment, which activates a thermosensor located at the 5ʹ untranslated region of the prfA mRNA57. The higher host temperature causes a shift in the energy landscape from one that favours the formation of the thermosensor hairpin to one that favours the melted, single-stranded conformation. This melting transition exposes ribosome-binding sites, which are required for translation. Mechanical triggers can also induce the unfolding of RNA hairpins. One example is translation-induced unfolding of mRNA hairpins, which is thought to slow the rate of ribosome elongation to allow the folding of autonomous-folding proteins and protein domains58.
Functions of secondary structural transitions Secondary structural transitions are widely used in gene regulation as binary switches that are activated by cellular cues. The switch can be transduced into a range of outputs by sequestering or exposing key RNA regulatory elements. Transcription Many RNA switches regulate gene expression at the transcriptional level by producing transcription-terminating helices. In addition to metabolite-sensing riboswitches, other RNA switches use the same strategy to regulate gene expression in response to more complex molecules 23,45. For example, in the T-box mechanism, 1 6 F E B R UA RY 2 0 1 2 | VO L 4 8 2 | N AT U R E | 3 2 5
INSIGHT REVIEW non-aminoacylated or uncharged tRNA can activate transcription of the cognate gene that encodes its aminoacyl-tRNA synthetase. The interaction between the acceptor end of an uncharged tRNA and residues in the antiterminator bulge in the 5ʹ untranslated region of the mRNA (Fig. 3a) promotes formation of an antiterminator helix during co-transcriptional folding that allows transcription to continue. However, the acceptor end of aminoacylated or charged tRNA cannot interact with the antiterminator helix residues, which results in formation of the more stable terminator stem that aborts synthetase gene transcription59 (Fig. 3a). Only a few proteins have been identified that modulate transcription by influencing folding of transcription-terminating helices. One example is the tryptophan-activated RNA-binding attenuation protein (TRAP), which binds trp mRNA to regulate gene expression at both the transcriptional and translational level by several processes (for example, promoting the formation of a terminator hairpin that terminates transcription60). Translation There is an increasing list of protein- and RNA-triggered61 RNA switches that regulate translation by sequestering or exposing ribosome-binding sites or by affecting the structure of ribosomal RNA, and therefore blocking translation. For example, a protein-dependent RNA switch has recently been identified in the 3ʹ untranslated region
of VEGFA mRNA in myeloid cells that regulates translation of VEGFA in response to proteins associated with two disparate stress stimuli (Fig. 3b). The interferon-γ (IFN-γ)-activated inhibitor of translation (GAIT)-complex binds a structural GAIT element within a family of inflammatory mRNAs and silences their translation by promoting the formation of a translational-silencing (TS) conformer62. During oxidative stress, the heterogeneous nuclear ribonucleoprotein L (hnRNP L) overrides GAIT silencing by triggering a secondary structural RNA switch to a translation-permissive (TP) conformer, in which the GAIT element is occluded. The RNA alternates between two mutuallyexclusive conformers in response to the binding of the GAIT complex or hnRNP L, thereby functioning as an AND NOT Boolean logic-gate switch in which the presence of one protein, but not the other, yields an output of gene repression (Fig. 3b). Post-transcriptional processing An increasing number of RNA switches are involved in regulating posttranscriptional processing; for example, splicing, gene silencing by microRNA (miRNA) and RNA editing. Although the detailed mechanics of many of these systems are still unknown, in all cases the RNA switch exposes, occludes or modulates the structure of the processing sites to regulate post-transcriptional processes. For example, one of the thiamine pyrophosphate riboswitches discovered in eukaryotes
b Stem I
Translation of VEGFA GAIT element
aa GAIT hnRNPL
hnRNPL TP conformer
Splicing 1st 5′ SS
2nd 5′ SS
Repression of p27 translation
e Diploid packaging
TPP uORF uORF
Splicing 1st 5′ SS uORF uORF
2nd 5′ SS
3′ SS NMT1 !"#$%&' ORF
Figure 3 | Functional outputs of secondary structural changes. a, Transcriptional activation of the aminoacyl-tRNA synthetase gene by uncharged tRNA (no aminoacylation (aa)). Binding of uncharged tRNA induces formation of an antiterminator helix during co-transcriptional folding99. b, Translation control of VEGFA expression through a dual proteindependent RNA secondary structural switch that responds to interferon-γ (IFN-γ) by binding the IFN-γ-activated inhibitor of translation (GAIT) complex (green) to form a translational-silencing (TS) conformer (on the left) and to hypoxic stress that results in hnRNP L binding and causes a switch to a translation permissive (TP) conformer (on the right). c, Thiamine pyrophosphate (TPP) riboswitch-regulated alternative splicing and gene expression of NMT1. In the absence of TPP, the aptamer domain base-pairs (red dotted line) to the sequence surrounding a proximal 5ʹ splice site (SS, 3 2 6 | N AT U R E | VO L 4 8 2 | 1 6 F E B R UA RY 2 0 1 2
shown as coloured diamonds: green, activation or red, repression) to block it from the SS machinery. Instead, a distal SS is selected. On binding TPP, the aptamer domain undergoes a conformational change to expose the second proximal 5ʹ SS. The resultant spliced mRNA contains decoy upstream open reading frames (uORFs), thus reducing expression of the NMT1 ORF. d, Pumilio protein-mediated mRNA secondary structural switch controls accessibility of microRNA-binding sites and regulates expression of p27 protein. Binding of PUM1 induces a conformational change to expose the miR-211 and miR-222 binding site to allow p27 silencing. RISC, RNA-induced silencing complex. e, Secondary structural switch couples dimerization and diploid genome packaging of the Moloney murine leukaemia virus. Dimerization leads to a coupled frame-shift that exposes nucleocapsid protein binding sites (green) required for genome packaging. NC, nucleocapsid.
REVIEW INSIGHT regulates alternative splicing63 (Fig. 3c). Here, changes in the secondary structure sequester or expose splice sites (Fig. 3c). An RNA switch has recently been identified in the 3ʹ untranslated region of p27 mRNA that simultaneously sequesters both an miRNA target site from cleavage by the RNA-induced silencing complex (RISC) and a Pumilio-recognition element (PRE), which binds a Pumilio RNA-binding protein (PUM1)64. Binding of PUM1 to the PRE region triggers a secondary structural switch that exposes the miRNA target site, leading to miRNA silencing (Fig. 3d). In another example, HDV genotype III editing levels are determined by a pre-existing equilibrium between two secondary structures of the antigenome RNA, involving a kinetically trapped conformation and a thermodynamically more favourable state65. These initial discoveries suggest RNA switches have a range of functions in post-transcriptional processing.
RNA switches can also couple distinct processes within a given step. For example, an RNA switch is used to couple dimerization and selective encapsidation of two copies of the Moloney murine leukaemia virus RNA genome. Dimerization of two RNA genomes induces a shift in the base-pairing pattern within the ψ-RNA packaging signal, which exposes conserved UCUG elements that bind the nucleocapsid protein with high affinity, thereby promoting genome packaging67 (Fig. 3e). These elements are base-paired and bind nucleocapsid protein weakly in the monomeric RNA (Fig. 3e).
Functions of tertiary conformational changes RNA tertiary conformational changes can range from large global changes in the orientation of helices to more subtle local changes in the structure of motifs that are involved in tertiary interactions. These conformational transitions allow RNA molecules to bind adaptively to a wide range of molecular partners and can help to direct the assembly of RNPs.
Viral replication RNA genomes of retroviruses take advantage of RNA secondary structural switches to transition between the different functions required for the various steps of the viral replication cycle. For example, there is evidence that the 5ʹ untranslated region of the HIV-1 genome can form two mutually exclusive secondary structures: a metastable branched multiple-hairpin conformation, which is involved in dimerization and packaging; and a more energetically favourable long-distance interaction conformation, which is involved in transcription and translation. The transition from the long-distance interaction to the branched multiple hairpin conformation is catalysed by the RNA chaperone nucleocapsid protein66. a
Polyvalent binding Some of the first solved structures of RNA–protein complexes revealed a remarkable ability of RNA to undergo adaptive changes in conformation2,25 that had the potential to allow the optimization of intermolecular interactions with disparate targets. In a classic example, these conformational changes allow tRNAs to interact with many diverse partners, including ribonuclease P (RNase P), various nucleotide modifying enzymes, tRNA synthetase, EF-Tu, the ribosome and other RNA elements. High-resolution structures of tRNA, tRNA–protein and
S6 S18 S15
S6 S18 S15
0 –1 –5
30S body rotation (°)
Figure 4 | Functional outputs of tertiary conformational changes. a, Different X-ray structures of tRNAPhe in the unbound state (black, PDB ID 1EHZ), in complex with RNaseP (blue, engineered anticodon stem removed, PDB ID 3Q1Q), the ribosome in the P/E state (green, PDB ID 3R8N), isopentenyl-tRNA transferase (red, PDB ID 3FOZ), and phenyalanyltRNA synthetase (yellow, PDB ID 1EIY). The structures are superimposed by the acceptor stem. b, Hierarchical assembly of the central domain of the 30S ribosomal subunit by successive protein-induced changes in the conformation of 16S rRNA. S15 changes the orientation of the helical domains to favour the binding of S6 and S18. c, Enzymatic cycle of the hairpin ribozyme. d, Ratcheting motions of the ribosome seen through X-ray crystallography. The degree of 30S subunit atomic displacement between the unratcheted and R2 ratcheted states with the 50S subunit as a reference (not shown) are colour-coded by Å. Atomic displacement vectors and
1.0 0.8 0.6 0.4 0.2 0.0 40
Time (s) 1,200 4,000
e 20 Å
30S head movement (a.u.)
0 0.0 0.2 0.4 0.6 0.8 1.0
0 0.0 0.2 0.4 0.6 0.8 1.0
arrows (on the right) indicate the direction of the change. Figure reprinted, with permission, from ref. 82. e, The free-energy landscape of ribosomal ratcheting, as calculated from subclassification of cryoelectron microscopy particles. Movements of the 30S subunit body and head domains in relation to the 50S subunit are shown in units of degrees and arbitrary units (a.u.), respectively, with corresponding tRNA translocation intermediates (Pre1 and so on) outlined in black. Figure reprinted, with permission, from ref. 84. f, Dynamics of the 50S ribosomal L1 stalk monitored by single-molecule fluorescence resonance energy transfer (smFRET). Representative smFRET trace (top) and histogram (bottom left) of the L1 stalk dynamically sampling open and closed conformations in A- and P-site tRNA-bound ribosome complexes. Translocation by EF-G and tRNA occupation of the E- and P-sites causes the L1 stalk conformation to shift dramatically (bottom right). Figure modified, with permission, from ref. 100. 1 6 F E B R UA RY 2 0 1 2 | VO L 4 8 2 | N AT U R E | 3 2 7
INSIGHT REVIEW tRNAâ€“RNP complexes show that binding is often accompanied by significant conformational changes, which range from the reorientation of helical domains to finer changes in local structure, all of which optimize intermolecular interactions68 (Fig.Â 4a). Ordering RNP assembly RNA tertiary conformational changes that are induced by successive protein-binding events are thought to help direct the order of assembly of complex RNP machines, including the 30S ribosome69,70, the signal recognition particle71 and telomerase72. For example, the binding of ribosomal protein S15 to 16S rRNA initiates the ordered assembly of the central domain in the 30S ribosomal subunit73, and leads to a change in the orientation of helical domains that favours the binding of ribosomal proteins S6 and S18 (ref. 74) (Fig.Â 4b). Premature binding of S6 and S18 to the unbound 16S rRNA may be disfavoured, in part, because of the entropic penalty that is associated with the partial freezing-out of interhelical motions. Even in the simpler telomerase RNP (consisting of one RNA and two protein components), the binding of the first protein p65 induces a conformational change in the RNA that facilitates the binding of telomerase reverse transcriptase, thereby ordering assembly72. Assembly can also involve coupled protein binding that induces changes in both RNA secondary and tertiary structure. For example, coupled binding of the maturase and Mrs1 protein cofactors to the RNA of the bI3 group-I intron RNP stabilizes both the native tertiary contacts and induces a reorganization of a non-native intermediate secondary structure75. Although both Mrs1 dimers and maturase can independently bind and stabilize portions of the bI3 tertiary structure, binding by both proteins is required to induce the secondary structure rearrangement and assembly to the native, active state. Ribozyme catalysis Tertiary conformational transitions involving large changes in the orientation of helical arms are often observed in small ribozymes, such as the hairpin and HDV, and are thought to be important for the transition between the different steps of the catalytic cycles. Typically, an undocked (inactive) conformation binds the substrate, promoting the transition into a docked (active) conformation, which is required for catalysis. After catalysis, another undocking transition allows the release of the product (Fig.Â 4c). The importance of these motions is demonstrated by the fact that the junction motions can accelerate the rate of folding of the active conformation76. Similarly, large hinge-like motions of the J2a/b bulge in human telomerase have been proposed to help with dynamic telomere repeat synthesis77. A more exceptional example is the Tetrahymena group-I ribozyme that has been shown to interconvert between alternative tertiary conformations, which have a range of substrate binding affinities but similar enzymatic activities78. The rates of interconversion between these states are slower than the rate of catalysis, implying the existence of multiple native states. Such long-lived heterogeneities have been observed in the tertiary folds of many other RNAs, although some of these may be the result of RNA purification side-products 79. The atomic level structural differences between these species and the source of the severe heterogeneity are still unknown but they may constitute yet another mechanism used by RNA to define a narrow set of differentiated conformations and this should be an exciting topic for future research. Protein synthesis Perhaps the best example of the cell manipulating the intrinsic dynamic landscape of RNA to achieve a desired biological outcome is ribosome catalysis. Large-scale ratcheting motions are required for translation. The small and large subunits reorient with respect to one another through numerous structural intermediates that are driven by changes in the conformation of both the ribosomal RNAs and proteins80â€“84 (Fig.Â 4d). Data strongly indicate that all of these 3 2 8 | N AT U R E | VO L 4 8 2 | 1 6 F E B R UA RY 2 0 1 2
intermediates are relatively low-lying energy states that readily interconvert, which has been highlighted by the ability of the ribosome to spontaneously undergo full tRNA retrotranslocation84,85 (Fig.Â 4e). This has led to a â€˜Brownian machineâ€™ model of the ribosome, where the ribosomeâ€™s functionality is derived in part by its ability to harness thermally driven equilibrium fluctuations and bias them to promote the translation process22 (Fig.Â 4f). The cell combines these intrinsic ribosome dynamics with numerous effectors to achieve tight control over the complex transactions that are required by translation. One such transaction is the selection and proofreading of incoming tRNAs that are responsible for the ribosomeâ€™s remarkable ability to consistently discriminate between cognate and near- or non-cognate tRNAs, in which small differences between the minihelices of incorrect and correct anticodonâ€“ codon pairs will lead to tRNA accommodation or rejection. Here, the formation of a cognate minihelix results in a kinked tRNA structure and triggers a 30S â€˜domain closureâ€™ motion86â€“88. This stabilizes tRNAâ€“ribosome interactions and in turn promotes conformational SFBSSBOHFNFOUTJOUIF&'5VQSPUFJOPGUIF&'5Vr(51U3/"UFSnary complex that delivered tRNA to the ribosome; this results in &'5Vr(51IZESPMZTJT SFMFBTFPGU3/"GSPN&'5VBOEJOJUJBMU3/" selection89,90. The second proofreading step that follows EF-Tu dissociation is thought to be driven by relaxation of the kinked tRNA. In cognate tRNAs, the strong interactions between the codon and the anticodon cause a bias of tRNA relaxation towards a conformation that is fully accommodated within the A-site. However, for nearcognate tRNAs, which have weak codonâ€“anticodon interactions, the relaxation of the kinked tRNA can occur through other pathways that lead to rejection91,92. Following tRNA accommodation, other factors, including EF-G93, other initiation factors94, recycling factors95, release factors96, and even the identity and acylation state of the tRNA occupying the neighbouring ribosomal P-site97, act on the translation process, manipulating the ribosomeâ€™s dynamic landscape to drive efficient synthesis of the mRNA-encoded protein. Owing to the overwhelming complexity of the ribosome, the mechanisms and atomic level details of the many conformational transitions involved in the translation process remain unclear. Among these unresolved questions are how the ribosomeâ€™s RNA and protein components cooperate to confer dynamic specificity and robustness on ribosome dynamics98. Research into this process is another exciting area of future study, and we can confidently predict that this will be yet another biological system shown to rely heavily on the virtuosity of RNA dynamics.
Outlook The conventional view that one sequence codes for one structure and one function is being replaced by a dynamic view of RNA as a pre-existing superposition of conformational states that can be resolved into a directed and synchronized motion by dedicated cellular machinery, leading to a broad range of functional outcomes. This makes it all the more important to study RNA dynamics within the complex in vivo environment of living cells, an important goal for the future. We also need to increase our basic understanding of RNA dynamic behaviour, even within the simpler in vitro environment. It is remarkable that, even for well-studied molecules such as tRNA, there is very little experimental data available regarding the equilibrium fluctuations in tRNA at the atomic level; the same is also true for catalytically important motions in ribozymes. Similarly, little is known about the structure and dynamics of large RNAs, such as eukaryotic mRNAs. This will require the combined development of computational and experimental tools to move towards developing atomic-level movies of RNA in dynamic action within living cells as well as a better predictive understanding of RNA dynamic behaviour. In the meantime, great advances can be made by simply embracing this new dynamic view of RNA and always being on the lookout for another myoglobin. â–
REVIEW INSIGHT 1. 2. 3. 4. 5.
6. 7. 8. 9. 10. 11. 12. 13. 14.
16. 17. 18. 19. 20. 21. 22. 23. 24. 25. 26. 27. 28. 29. 30. 31.
Kendrew, J. C. et al. A three-dimensional model of the myoglobin molecule obtained by X-ray analysis. Nature 181, 662–666 (1958). Rould, M. A., Perona, J. J., Söll, D. & Steitz, T. A. Structure of E. coli glutaminyltRNA synthetase complexed with tRNA(Gln) and ATP at 2.8 Å resolution. Science 246, 1135–1142 (1989). Pley, H. W., Flaherty, K. M. & McKay, D. B. Three-dimensional structure of a hammerhead ribozyme. Nature 372, 68–74 (1994). Scott, W. G., Finch, J. T. & Klug, A. The crystal structure of an all-RNA hammerhead ribozyme: a proposed mechanism for RNA catalytic cleavage. Cell 81, 991–1002 (1995). Wang, S., Karbstein, K., Peracchi, A., Beigelman, L. & Herschlag, D. Identification of the hammerhead ribozyme metal ion binding site responsible for rescue of the deleterious effect of a cleavage site phosphorothioate. Biochemistry 38, 14363–14378 (1999). Boehr, D. D., Nussinov, R. & Wright, P. E. The role of dynamic conformational ensembles in biomolecular recognition. Nature Chem. Biol. 5, 789–796 (2009). Al-Hashimi, H. M. & Walter, N. G. RNA dynamics: it is about time. Curr. Opin. Struct. Biol. 18, 321–329 (2008). Frauenfelder, H., Sligar, S. G. & Wolynes, P. G. The energy landscapes and motions of proteins. Science 254, 1598–1603 (1991). Cruz, J. A. & Westhof, E. The dynamic landscapes of RNA architecture. Cell 136, 604–609 (2009). Bailor, M. H., Mustoe, A. M., Brooks, C. L. 3rd & Al-Hashimi, H. M. Topological constraints: using RNA secondary structure to model 3D conformation, folding pathways, and dynamic adaptation. Curr. Opin. Struct. Biol. 21, 296–305 (2011). Schultes, E. A., Spasic, A., Mohanty, U. & Bartel, D. P. Compact and ordered collapse of randomly generated RNA sequences. Nature Struct. Mol. Biol. 12, 1130–1136 (2005). Schultes, E. A., Hraber, P. T. & LaBean, T. H. Estimating the contributions of selection and self-organization in RNA secondary structure. J. Mol. Evol. 49, 76–83 (1999). Fürtig, B., Wenter, P., Pitsch, S. & Schwalbe, H. Probing mechanism and transition state of RNA refolding. ACS Chem. Biol. 5, 753–765 (2010). Bailor, M. H., Sun, X. & Al-Hashimi, H. M. Topology links RNA secondary structure with global conformation, dynamics, and adaptation. Science 327, 202–206 (2010). This article reports the simple topological constraints that are governed by steric and stereochemical forces severely restrict the allowed orientation of helices across two-way junctions. Mustoe, A. M., Bailor, M. H., Teixeira, R. M., Brooks, C. L. 3rd & Al-Hashimi, H. M. New insights into the fundamental role of topological constraints as a determinant of two-way junction conformation. Nucleic Acids Res. 40, 892–904 (2012). Chu, V. B. et al. Do conformational biases of simple helical junctions influence RNA folding stability and specificity? RNA 15, 2195–2205 (2009). Venditti, V., Clos, L. 2nd, Niccolai, N. & Butcher, S. E. Minimum-energy path for a U6 RNA conformational change involving protonation, base-pair rearrangement and base flipping. J. Mol. Biol. 391, 894–905 (2009). Fourmy, D., Yoshizawa, S. & Puglisi, J. D. Paromomycin binding induces a local conformational change in the A-site of 16S rRNA. J. Mol. Biol. 277, 333–345 (1998). Le, S. Y., Zhang, K. & Maizel, J. V. Jr. RNA molecules with structure dependent functions are uniquely folded. Nucleic Acids Res. 30, 3574–3582 (2002). Stelzer, A. C., Kratz, J. D., Zhang, Q. & Al-Hashimi, H. M. RNA dynamics by design: biasing ensembles towards the ligand-bound state. Angew. Chem. Int. Ed. Engl. 49, 5731–5733 (2010). Shankar, N. et al. NMR reveals the absence of hydrogen bonding in adjacent UU and AG mismatches in an isolated internal loop from ribosomal RNA. Biochemistry 46, 12665–12678 (2007). Frank, J. & Gonzalez, R. L., Jr. Structure and dynamics of a processive Brownian motor: the translating ribosome. Annu. Rev. Biochem. 79, 381–412 (2010). Haller, A., Souliere, M. F. & Micura, R. The dynamic nature of RNA as key to understanding riboswitch mechanisms. Acc. Chem. Res. 44, 1339–1348 (2011). Paukstelis, P. J., Chen, J. H., Chase, E., Lambowitz, A. M. & Golden, B. L. Structure of a tyrosyl-tRNA synthetase splicing factor bound to a group I intron RNA. Nature 451, 94–97 (2008). Puglisi, J. D., Tan, R., Calnan, B. J., Frankel, A. D. & Williamson, J. R. Conformation of the TAR RNA-arginine complex by NMR spectroscopy. Science 257, 76–80 (1992). Orr, J. W., Hagerman, P. J. & Williamson, J. R. Protein and Mg2+-induced conformational changes in the S15 binding site of 16S ribosomal RNA. J. Mol. Biol. 275, 453–464 (1998). Turner, B., Melcher, S. E., Wilson, T. J., Norman, D. G. & Lilley, D. M. Induced fit of RNA on binding the L7Ae protein to the kink-turn motif. RNA 11, 1192–1200 (2005). Falb, M., Amata, I., Gabel, F., Simon, B. & Carlomagno, T. Structure of the K-turn U4 RNA: a combined NMR and SANS study. Nucleic Acids Res. 38, 6274–6285 (2010). Kim, H. D. et al. Mg2+-dependent conformational change of RNA studied by fluorescence correlation and FRET on immobilized single molecules. Proc. Natl Acad. Sci. USA 99, 4284–4289 (2002). Zacharias, M. & Hagerman, P. J. The influence of symmetric internal loops on the flexibility of RNA. J. Mol. Biol. 257, 276–289 (1996). Zhang, Q., Stelzer, A. C., Fisher, C. K. & Al-Hashimi, H. M. Visualizing spatially correlated dynamics that directs RNA conformational transitions. Nature 450, 1263–1267 (2007).
32. Shajani, Z., Drobny, G. & Varani, G. Binding of U1A protein changes RNA dynamics as observed by 13C NMR relaxation studies. Biochemistry 46, 5875–5883 (2007). 33. Bokinsky, G. et al. Two distinct binding modes of a protein cofactor with its target RNA. J. Mol. Biol. 361, 771–784 (2006). 34. Bardaro, M. F. Jr., Shajani, Z., Patora-Komisarska, K., Robinson, J. A. & Varani, G. How binding of small molecule and peptide ligands to HIV-1 TAR alters the RNA motional landscape. Nucleic Acids Res. 37, 1529–1540 (2009). 35. Herschlag, D., Khosla, M., Tsuchihashi, Z. & Karpel, R. L. An RNA chaperone activity of non-specific RNA binding proteins in hammerhead ribozyme catalysis. EMBO J. 13, 2913–2924 (1994). 36. Pyle, A. M. & Green, J. B. RNA folding. Curr. Opin. Struct. Biol. 5, 303–310 (1995). 37. Treiber, D.K. & Williamson, J.R. Beyond kinetic traps in RNA folding. Curr. Opin. Struct. Biol. 11, 309–314 (2001). 38. Hirling, H., Scheffner, M., Restle, T. & Stahl, H. RNA helicase activity associated with the human p68 protein. Nature 339, 562–564 (1989). 39. Yang, Q. & Jankowsky, E. ATP- and ADP-dependent modulation of RNA unwinding and strand annealing activities by the DEAD-box protein DED1. Biochemistry 44, 13591–13601 (2005). 40. Will, C. L. & Lührmann, R. Spliceosome structure and function. Cold Spring Harb. Perspect. Biol. 3, a003707 (2011). 41. Kosowski, T. R., Keys, H. R., Quan, T. K. & Ruby, S. W. DExD/H-box Prp5 protein is in the spliceosome during most of the splicing cycle. RNA 15, 1345–1362 (2009). 42. Maeder, C., Kutach, A. K. & Guthrie, C. ATP-dependent unwinding of U4/U6 snRNAs by the Brr2 helicase requires the C terminus of Prp8. Nature Struct. Mol. Biol. 16, 42–48 (2009). 43. Schwer, B. A conformational rearrangement in the spliceosome sets the stage for Prp22-dependent mRNA release. Mol. Cell 30, 743–754 (2008). 44. Bhaskaran, H. & Russell, R. Kinetic redistribution of native and misfolded RNAs by a DEAD-box chaperone. Nature 449, 1014–1018 (2007). 45. Winkler, W., Nahvi, A. & Breaker, R. R. Thiamine derivatives bind messenger RNAs directly to regulate bacterial gene expression. Nature 419, 952–956 (2002). This article reports the discovery of an RNA switch in the 5ʹ untranslated region of bacterial mRNA that regulates gene expression in response to ligands without assistance from proteins. 46. Cromie, M. J., Shi, Y., Latifi, T. & Groisman, E. A. An RNA sensor for intracellular Mg2+. Cell 125, 71–84 (2006). 47. Nechooshtan, G., Elgrably-Weiss, M., Sheaffer, A., Westhof, E. & Altuvia, S. A pH-responsive riboregulator. Genes Dev. 23, 2650–2662 (2009). 48. Greenleaf, W. J., Frieda, K. L., Foster, D. A. Woodside, M. T. & Block, S. M. Direct observation of hierarchical folding in single riboswitch aptamers. Science 319, 630–633 (2008). 49. Mandal, M. et al. A glycine-dependent riboswitch that uses cooperative binding to control gene expression. Science 306, 275–279 (2004). 50. Sudarsan, N. et al. Tandem riboswitch architectures exhibit complex gene control functions. Science 314, 300–304 (2006). 51. Lee, E. R., Baker, J. L., Weinberg, Z., Sudarsan, N. & Breaker, R. R. An allosteric self-splicing ribozyme triggered by a bacterial second messenger. Science 329, 845–848 (2010). 52. Ferre-D’Amare, A. R., Zhou, K. & Doudna, J. A. Crystal structure of a hepatitis delta virus ribozyme. Nature 395, 567–574 (1998). 53. Ke, A., Zhou, K., Ding, F., Cate, J. H. D. & Doudna, J. A. A conformational switch controls hepatitis delta virus ribozyme catalysis. Nature 429, 201–205 (2004). This article reports a significant local conformational change in the active site of the HDV ribozyme is observed post-cleavage and is associated with ejection of the substrate and a catalytically critical divalent metal ion. 54. Harris, D. A., Rueda, D. & Walter, N. G. Local conformational changes in the catalytic core of the trans-acting hepatitis delta virus ribozyme accompany catalysis. Biochemistry 41, 12051–12061 (2002). 55. Lamanna, A. C. & Karbstein, K. An RNA conformational switch regulates pre18S rRNA cleavage. J. Mol. Biol. 405, 3–17 (2011). 56. Nocker, A. et al. A mRNA-based thermosensor controls expression of rhizobial heat shock genes. Nucleic Acids Res. 29, 4800–4807 (2001). 57. Johansson, J. et al. An RNA thermosensor controls expression of virulence genes in Listeria monocytogenes. Cell 110, 551–561 (2002). 58. Watts, J. M. et al. Architecture and secondary structure of an entire HIV-1 RNA genome. Nature 460, 711–716 (2009). 59. Grundy, F. J., Winkler, W. C. & Henkin, T. M. tRNA-mediated transcription antitermination in vitro: codon-anticodon pairing independent of the ribosome. Proc. Natl Acad. Sci. USA 99, 11121–11126 (2002). 60. Babitzke, P. & Yanofsky, C. Reconstitution of Bacillus subtilis trp attenuation in vitro with TRAP, the trp RNA-binding attenuation protein. Proc. Natl Acad. Sci. USA 90, 133–137 (1993). 61. Diaz-Toledano, R., Ariza-Mateos, A., Birk, A., Martinez-Garcia, B. & Gomez, J. In vitro characterization of a miR-122-sensitive double-helical switch element in the 5´ region of hepatitis C virus RNA. Nucleic Acids Res. 37, 5498–5510 (2009). 62. Ray, P. S. et al. A stress-responsive RNA switch regulates VEGFA expression. Nature 457, 915–919 (2009). This article reports that the 3ʹ untranslated region of human VEGFA mRNA undergoes a binary conformational switch in response to inflammatory and hypoxic protein stress signals to regulate VEGFA expression. 63. Cheah, M. T., Wachter, A., Sudarsan, N. & Breaker, R. R. Control of alternative RNA splicing and gene expression by eukaryotic riboswitches. Nature 447, 497–500 (2007). 1 6 F E B R UA RY 2 0 1 2 | VO L 4 8 2 | N AT U R E | 3 2 9
64. 65. 66. 67.
68. 69. 70. 71. 72. 73. 74. 75. 76. 77. 78.
79. 80. 81. 82. 83. 84.
This article reports a secondary structural change in a eukaryotic thiamine pyrophosphate riboswitch regulates gene expression through the control of alternative splicing. Kedde, M. et al. A Pumilio-induced RNA structure switch in p27-3ʹ untranslated region controls miR-221 and miR-222 accessibility. Nature Cell Biol. 12, 1014–1020 (2010). Casey, J. L. Control of ADAR1 editing of hepatitis delta virus RNAs. Curr. Top. Microbiol. Immunol. 353, 123–143 (2012). Abbink, T. E., Ooms, M., Haasnoot, P. C. & Berkhout, B. The HIV-1 leader RNA conformational switch regulates RNA dimerization but does not regulate mRNA translation. Biochemistry 44, 9058–9066 (2005). Miyazaki, Y. et al. An RNA structural switch regulates diploid genome packaging by Moloney murine leukemia virus. J. Mol. Biol. 396, 141–152 (2010). This article reports that dimerization of the 5ʹ untranslated region of the Moloney murine leukaemia virus results in a secondary structural change that promotes genome packaging. Giege, R. Toward a more complete view of tRNA biology. Nature Struct. Mol. Biol. 15, 1007–1014 (2008). Mulder, A. M. et al. Visualizing ribosome biogenesis: parallel assembly pathways for the 30S subunit. Science 330, 673–677 (2010). Adilakshmi, T., Bellur, D. L. & Woodson, S. A. Concurrent nucleation of 16S folding and induced fit in 30S ribosome assembly. Nature 455, 1268–1272 (2008). Menichelli, E., Isel, C., Oubridge, C. & Nagai, K. Protein-induced conformational changes of RNA during the assembly of human signal recognition particle. J. Mol. Biol. 367, 187–203 (2007). Stone, M. D. et al. Stepwise protein-mediated RNA folding directs assembly of telomerase ribonucleoprotein. Nature 446, 458–461 (2007). Held, W. A., Ballou, B., Mizushima, S. & Nomura, M. Assembly mapping of 30S ribosomal proteins from Escherichia coli. Further studies. J. Biol. Chem. 249, 3103–3111 (1974). Agalarov, S. C., Prasad, G. S., Funke, P. M., Stout, C. D. & Williamson, J. R. Structure of the S15,S6,S18-rRNA complex: assembly of the 30S ribosome central domain. Science 288, 107–112 (2000). Duncan, C. D. & Weeks, K. M. Nonhierarchical ribonucleoprotein assembly suggests a strain-propagation model for protein-facilitated RNA folding. Biochemistry 49, 5418–5425 (2010). Wilson, T. J., Nahas, M., Ha, T. & Lilley, D. M. Folding and catalysis of the hairpin ribozyme. Biochem. Soc. Trans. 33, 461–465 (2005). Zhang, Q., Kim, N. K., Peterson, R. D., Wang, Z. & Feigon, J. Structurally conserved five nucleotide bulge determines the overall topology of the core domain of human telomerase RNA. Proc. Natl Acad. Sci. USA 107, 18761–18768 (2010). Solomatin, S. V., Greenfeld, M., Chu, S. & Herschlag, D. Multiple native states reveal persistent ruggedness of an RNA folding landscape. Nature 463, 681–684 (2010). This article reports the observation of slowly interconverting catalytically active states in a ribozyme, thereby establishing the coexistence of multiple native states. Greenfeld, M., Solomatin, S. V. & Herschlag, D. Removal of covalent heterogeneity reveals simple folding behavior for P4–P6 RNA. J. Biol. Chem. 286, 19872–19879 (2011). Frank, J. & Agrawal, R. K. A ratchet-like inter-subunit reorganization of the ribosome during translocation. Nature 406, 318–322 (2000). Valle, M. et al. Locking and unlocking of ribosomal motions. Cell 114, 123–134 (2003). Zhang, W., Dunkle, J. A. & Cate, J. H. Structures of the ribosome in intermediate states of ratcheting. Science 325, 1014–1017 (2009). Ratje, A. H. et al. Head swivel on the ribosome facilitates translocation by means of intra-subunit tRNA hybrid sites. Nature 468, 713–716 (2010). Fischer, N., Konevega, A. L. Wintermeyer, W., Rodnina, M. V. & Stark, H. Ribosome dynamics and tRNA movement by time-resolved electron cryomicroscopy. Nature 466, 329–333 (2010).
3 3 0 | N AT U R E | VO L 4 8 2 | 1 6 F E B R UA RY 2 0 1 2
This article demonstrates the cryo-electron microscopy observation of thermally driven tRNA retrotranslocation on the ribosome. 85. Shoji, S., Walker, S. E. & Fredrick, K. Reverse translocation of tRNA in the ribosome. Mol. Cell 24, 931–942 (2006). 86. Ogle, J. M., Murphy, F. V., Tarry, M. J. & Ramakrishnan, V. Selection of tRNA by the ribosome requires a transition from an open to a closed form. Cell 111, 721–732 (2002). 87. Valle, M. et al. Incorporation of aminoacyl-tRNA into the ribosome as seen by cryo-electron microscopy. Nature Struct. Biol. 10, 899–906 (2003). 88. Lee, T. H., Blanchard, S. C., Kim, H. D., Puglisi, J. D. & Chu, S. The role of fluctuations in tRNA selection by the ribosome. Proc. Natl Acad. Sci. USA 104, 13661–13665 (2007). 89. Schmeing, T. M. et al. The crystal structure of the ribosome bound to EF-Tu and aminoacyl-tRNA. Science 326, 688–694 (2009). 90. Voorhees, R. M., Schmeing, T. M., Kelley, A. C. & Ramakrishnan, V. The mechanism for activation of GTP hydrolysis on the ribosome. Science 330, 835–838 (2010). 91. Pape, T., Wintermeyer, W. & Rodnina, M. V. Conformational switch in the decoding region of 16S rRNA during aminoacyl-tRNA selection on the ribosome. Nature Struct. Mol. Biol. 7, 104–107 (2000). 92. Blanchard, S. C., Gonzalez, R. L., Kim, H. D., Chu, S. & Puglisi, J. D. tRNA selection and kinetic proofreading in translation. Nature Struct. Mol. Biol. 11, 1008–1014 (2004). This important single-molecule FRET study directly observes the dynamics of tRNA initial selection and proofreading by the ribosome. 93. Fei, J. et al., Allosteric collaboration between elongation factor G and the ribosomal L1 stalk directs tRNA movements during translation. Proc. Natl Acad. Sci. USA 106, 15702–15707 (2009). 94. Blaha, G., Stanley, R. E. & Steitz, T. A. Formation of the first peptide bond: the structure of EF-P bound to the 70S ribosome. Science 325, 966–970 (2009). 95. Dunkle, J. A. et al. Structures of the bacterial ribosome in classical and hybrid states of tRNA binding. Science 332, 981–984 (2011). 96. Laurberg, M. et al. Structural basis for translation termination on the 70S ribosome. Nature 454, 852–857 (2008). 97. Cornish, P. V., Ermolenko, D. N., Noller, H. F. & Ha, T. Spontaneous intersubunit rotation in single ribosomes. Mol. Cell 30, 578–588 (2008). 98. Tama, F., Valle, M., Frank, J. & Brooks, C. L 3rd. Dynamic reorganization of the functionally active ribosome explored by normal mode analysis and cryoelectron microscopy. Proc. Natl Acad. Sci. USA 100, 9319–9323 (2003). 99. Green, N. J., Grundy, F. J. & Henkin, T. M. The T box mechanism: tRNA as a regulatory molecule. FEBS Lett. 584, 318–324 (2010). 100. Cornish, P. V. et al. Following movement of the L1 stalk between three functional states in single ribosomes. Proc. Natl Acad. Sci. USA 106, 2571–2576 (2009). Supplementary Information is linked to the online version of the paper at www.nature.com/nature. Acknowledgements E.A.D. and J.C. contributed equally to this Review. We thank C. Eichhorn and Q. Zhang for their input and assistance in the preparation of figures, and S. Butcher and S. Serganov for their comments on this Review. A.M.M. is supported by an NSF graduate research fellowship. The authors gratefully acknowledge the Michigan Economic Development Cooperation and the Michigan Technology Tri-Corridor for their support in the purchase of a 600 MHz spectrometer. This work was supported by the US National Institutes of Health (R01 AI066975 and R01 GM089846) and the US National Science Foundation (NSF Career Award CHE-0918817). Author Information Reprints and permissions information is available at www.nature.com/reprints. The authors declare no competing financial interests. Readers are welcomed to comment on the online version of this article at www.nature.com/nature. Correspondence should be addressed to author H.M.A.H. (firstname.lastname@example.org).
RNA-guided genetic silencing systems in bacteria and archaea Blake Wiedenheft1,2†, Samuel H. Sternberg3 & Jennifer A. Doudna1–4
Clustered regularly interspaced short palindromic repeat (CRISPR) are essential components of nucleic-acid-based adaptive immune systems that are widespread in bacteria and archaea. Similar to RNA interference (RNAi) pathways in eukaryotes, CRISPR-mediated immune systems rely on small RNAs for sequence-specific detection and silencing of foreign nucleic acids, including viruses and plasmids. However, the mechanism of RNA-based bacterial immunity is distinct from RNAi. Understanding how small RNAs are used to find and destroy foreign nucleic acids will provide new insights into the diverse mechanisms of RNA-controlled genetic silencing systems.
acteria and archaea are the most diverse and abundant organisms on the planet, thriving in habitats that range from hot springs to humans. However, viruses outnumber their microbial hosts in every ecological setting, and the selective pressures imposed by these rapidly evolving parasites has driven the diversification of microbial defence systems1–3. Historically, our understanding of antiviral immunity in bacteria has focused on restriction-modification systems, abortive-phage phenotypes, toxin–antitoxins and other innate defence systems4,5. More recently, bioinformatic, genetic and biochemical studies have revealed that many prokaryotes use an RNA-based adaptive immune system to target and destroy genetic parasites (reviewed in refs 6–12). Such adaptive immunity, previously thought to occur only in eukaryotes, provides an example of RNA-guided destruction of foreign genetic material by a process that is distinct from RNA interference (RNAi) (Fig. 1). In response to viral and plasmid challenges, bacteria and archaea integrate short fragments of foreign nucleic acid into the host chromosome at one end of a repetitive element known as CRISPR (clustered regularly interspaced short palindromic repeat)13–15. These repetitive loci serve as molecular ‘vaccination cards’ by maintaining a genetic record of prior encounters with foreign transgressors. CRISPR loci are transcribed, and the long primary transcript is processed into a library of short CRISPR-derived RNAs (crRNAs)16–21 that each contain a sequence complementary to a previously encountered invading nucleic acid. Each crRNAs is packaged into a large surveillance complex that patrols the intracellular environment and mediates the detection and destruction of foreign nucleic acid targets15,22–27. CRISPRs were originally identified in the Escherichia coli genome in 1987, when they were described as an unusual sequence element consisting of a series of 29-nucleotide repeats separated by unique 32-nucleotide ‘spacer’ sequences28. Repetitive sequences with a similar repeat–spacer–repeat pattern were later identified in phylogenetically diverse bacterial and archaeal genomes, but the function of these repeats remained obscure until many spacer sequences were recoginized as being identical to viral and plasmid sequences29–31. This observation led to the hypothesis that CRISPRs provide a genetic memory of infection29, and the detection of short CRISPR-derived RNA transcripts suggested that there may be functional similarities between CRISPRbased immunity and RNAi30,32. In this Insight, we review three stages of CRISPR-based adaptive immunity and compare mechanistic aspects of these immune systems to other RNA-guided genetic silencing pathways.
Architecture and composition of CRISPR loci The defining feature of CRISPR loci is a series of direct repeats (approximately 20–50 base pairs) separated by unique spacer sequences of a similar length 11,33,34 (Fig. 2). The repeat sequences within a CRISPR locus are conserved, but repeats in different CRISPR loci can vary in both sequence and length. In addition, the number of repeat–spacer units in a CRISPR locus varies widely within and among organisms35. The sequence diversity of these repetitive loci initially limited their detection and obscured their relationship, but computational methods have been developed for detecting repeat patterns rather than related sequences33,34,36–38. One of the first-generation pattern-recognition algorithms identified the repeat–spacer–repeat architecture in phylogenetically diverse bacterial and archaeal genomes, but related structures were not identified in eukaryotic chromosomes39. Comparative analyses of the sequences adjacent to the CRISPR loci have revealed an (A+T)-rich ‘leader’ sequence that has been shown to serve as a promoter element for CRISPR transcription39–42. In addition to the leader sequence, Jansen et al.39 identified a set of four CRISPR-associated (cas) genes known as cas1–4 that are found exclusively in genomes containing CRISPRs. Based on sequence similarity to proteins of known function, Cas3 was predicted to be a helicase and Cas4 a RecB-like exonuclease39. Subsequent bioinformatic analyses have shown that CRISPR loci are flanked by a large number of extremely diverse cas genes32,43. The cas1 gene is a common component of all CRISPR systems, and phylogenetic analyses of Cas1 sequences indicate there are several versions of the CRISPR system. Providing additional evidence for the classification of distinct CRISPR types, neighbourhood analysis has identified conserved arrangements of between four and ten cas genes that are found in association with CRISPR loci harbouring specific repeat sequences35. These distinct immune systems have been divided into three major CRISPR types on the basis of gene conservation and locus organization10. More than one CRISPR type is often found in a single organism, indicating that these systems are probably mutually compatible and could share functional components10. Despite the variation in number and diversity of cas genes, the distinguishing feature of all type I systems is that they encode a cas3 gene. The Cas3 protein contains an N-terminal HD phosphohydrolase domain and a C-terminal helicase domain32,39,43,44. In some type I systems, the Cas3 nuclease and helicase domains are encoded by separate genes (cas3ʹʹ and cas3ʹ, respectively), but in each case they are thought to participate in degrading foreign nucleic acids22,44–46 (Fig. 2).
Howard Hughes Medical Institute, 4000 Jones Bridge Road, Chevy Chase, Maryland 20815-6789, USA. 2Department of Molecular and Cell Biology, University of California, Berkeley, California 94720, USA. 3Department of Chemistry, University of California, Berkeley, California 94720, USA. 4Physical Biosciences Division, Lawrence Berkeley National Laboratory, Berkeley, California 94720, USA. †Present address: Department of Immunology and Infectious Diseases, Montana State University, Bozeman, Montana, USA 1 6 F E B R UA RY 2 0 1 2 | VO L 4 8 2 | N AT U R E | 3 3 1
INSIGHT REVIEW Type II CRISPR systems consist of just four cas genes, one of which is always cas9 (formerly referred to as csn1). Cas9 is a large protein that includes both a RuvC-like nuclease domain and an HNH nuclease domain. Studies in Streptococcus pyogenes and Streptococcus thermophilus have indicated that Cas9 may participate in both CRISPR RNA processing and target destruction14,15,17. Two variations of the type III system have been identified (known as III-A and III-B). This division is supported by the functional differences reported in Staphylococcus epidermidis and Pyrococcus furiosus47,48. The immune system in S. epidermidis (type III-A) targets plasmid DNA in vivo, whereas the purified components of the type III-B system in P. furiosus have been found to cleave only single-stranded RNA substrates in vitro. The functional distinction between these two closely related systems suggests there could be other mechanistic differences between the distinct CRISPR subtypes.
Integration of new information into CRISPR loci Acquisition of foreign DNA is the first step of CRISPR-mediated immunity (Fig. 2 and 3). During this stage, a short segment of DNA from an invading virus or plasmid (known as the protospacer) is integrated preferentially at the leader end of the CRISPR locus14,15. Although metagenomic studies performed on environmental samples indicate that CRISPRs evolve rapidly in dynamic equilibrium with resident phage populations13,49,50, the type II system in S. thermophilus is currently the only CRISPR system that has been shown to robustly acquire new phage or plasmid sequences in a pure culture. Phage-challenge experiments in S. thermophilus have indicated that a small proportion of the cells in a population will typically incorporate a single virus-derived sequence at the leader end of a CRISPR locus14,15,51,52. The CRISPR-repeat sequence is duplicated for each new spacer seqenced added, thus maintaining the repeat–spacer–repeat architecture. Although the mechanism of spacer integration and replication of the repeat sequence is still unknown, studies in S. thermophilus and E. coli have indicated that several Cas proteins are involved in the process14,15,22,53. Mutational analysis of the
cas genes in S. thermophilus demonstrated that csn2 (previously known as cas7) is required for new spacer sequence acquisition14. This gene is not conserved in other CRISPR types, which suggests that either the mechanism of adaptation in S. thermophilus is distinct from the other types or that there are functional orthologues of Csn2 in other systems. Furthermore, gene deletion experiments in both S. thermophilus and E. coli have shown that neither cas1 nor cas2 genes are required for CRISPR RNA processing or targeted interference22,53,54. These genetic studies suggest a role for Cas1 and Cas2 in the integration of foreign DNA into the CRISPR. The role of Cas1 in CRISPR-mediated immunity is still uncertain; however, biochemical and structural data indicate a function for Cas1 in new–spacer–sequence acquisition54–56. Cas1 proteins from Pseudomonas aeruginosa56, E. coli54 and Sulfolobus solfataricus55 have been purified and studied biochemically. The Cas1 protein from S. solfataricus has been shown to bind nucleic acids with high affinity (Kd ranging from 20 to 50 nM), but without sequence preference55. The Cas1 protein from E. coli also binds to DNA with a preference for mismatched or abasic substrates57. This observation is consistent with a recent study showing a physical and genetic interaction between E. coli Cas1 and several proteins associated with DNA replication and repair54. Activity assays with Cas1 from P. aeruginosa and E. coli indicate that Cas1 is a metal-dependent nuclease. The Cas1 protein from P. aeruginosa is a DNA-specific nuclease, whereas the Cas1 protein from E. coli had a nuclease activity on a wider range of nucleic acid substrates54,56. These in vitro assays suggest that Cas1 proteins interact with nucleic acids in a non-sequence-specific manner. Crystal structures for five different Cas1 proteins are currently available (Protein Data Bank (PDB) identifiers: 3GOD, 3NKD, 3LFX, 3PV9 and 2YZS)54,56. Although the amino acid sequences for these proteins are extremely diverse (less than 15% sequence identity), their tertiary and quaternary structures are similar. All Cas1 proteins seem to share a twodomain architecture consisting of an N-terminal β-strand domain and a
Foreign RNA Nucleus
CRISPR locus Source Repeat
Repeat CRISPR transcription
Cas or RNase III
crRNA 5′ 3′ crRNA-guided surveillance complex
siRNA 3′ Dicer
Figure 1 | Parallels and distinctions between CRISPR RNA-guided silencing systems and RNAi. CRISPR systems and RNAi recognize long RNA precursors that are processed into small RNAs, which act as sequence-specific guides for targeting complementary nucleic acids. In CRISPR systems, foreign DNA is integrated into the CRISPR locus, and long transcripts from these loci are processed by a CRISPR-associated (Cas) or RNase III family nuclease16–21,64. The short CRISPR-derived RNAs (crRNAs) assemble with Cas proteins into large surveillance complexes that target destruction of invading genetic material15,22,24–27,48. In some eukaryotes, long double-stranded RNAs are recognized as foreign, and a specialized RNase III family endoribonuclease (Dicer) cleaves these RNAs into short-interfering RNAs (siRNAs) that guide the immune system to invading RNA viruses76. PIWI-interacting RNAs (piRNAs) are transcribed from repetitive clusters in the genome that often contain many copies of retrotransposons and primarily 3 3 2 | N AT U R E | VO L 4 8 2 | 1 6 F E B R UA RY 2 0 1 2
RNA-induced silencing complex
Cas protein(s) 5′
Seed 5′ Target interference
act by restricting transposon mobility76–78. The biogenesis of piRNAs is not yet fully understood. MicroRNAs (miRNAs) are also encoded on the chromosome, and primary miRNA transcripts form stable hairpin structures that are sequentially processed (shown by red triangles) by two RNase III family endoribonucleases (Drosha and Dicer)79. miRNAs do not participate in genome defence but are major regulators of endogenous gene expression80. Like crRNAs, eukaryotic piRNAs, siRNAs and miRNAs associate with proteins that facilitate complementary interactions with invading nucleic acid targets27,60,69,79. In eukaryotes, the Argonaute proteins pre-order the 5ʹ region of the guide RNA into a helical configuration, reducing the entropy penalty of interactions with target RNAs69. This high-affinity binding site, called the ‘seed’ sequence, is essential for target sequence interactions. Recent studies indicate that the CRISPR system may use a similar seed-binding mechanism for enhancing target sequence interactions26,27,53,60.
REVIEW INSIGHT C-terminal α-helical domain (Fig. 3). The C-terminal domain contains a conserved divalent metal-ion binding site, and alanine substitutions of the metal-coordinating residues inhibit Cas1-catalysed DNA degradation54,56. The metal ion is surrounded by a cluster of basic residues that form a strip of positive charge across the surface of the C-terminal domain. This positively charged surface may serve as an electrostatic snare to position nucleic-acid substrates near the catalytic metal ions56 (Fig. 3). The Cas1 protein forms a stable homodimer that is formed through interactions between the two β-strand domains, which are related by a pseudo-twofold axis of symmetry54,56. This organization creates a saddle-like structure that can be modelled onto double-stranded DNA without steric clashing. β-hairpins, one from each of the two symmetrically related molecules, hang on opposite faces of the double-stranded DNA (like stirrups on a
saddle). Although this feature of the Cas1 structure did not initially stand out as a potential DNA-binding site, comparative analysis of the available Cas1 structures reveals a conserved set of positively charged residues along each of the β-hairpins that could contact the phosphate backbone. The two β-hairpins, which are symmetrically related, might participate in sequence-specific interactions with the CRISPR repeat, whereas the large positively charged surface on the C-terminal α-helical domain could account for the high-affinity, non-sequence-specific interactions that have been observed in vitro. In spite of these structural studies and biochemical results, it is still only possible to speculate on the role of Cas1 in the integration of new spacer sequences, and many steps associated with the integration process still need to be explained. For example, new spacer sequences are inserted
cas gene casette
ot M Pr PA * *
New spacer sequence
Stage 1: Acquisition CRISPR trancription
Repeat duplication Stage 2: CRISPR RNA biogenesis Type I
Type III CRISPR transcription
CRISPR transcription RNase III
Leader Leader CRISPRspecific endoribonuclease
5′ 3′3′ ∼30-nt spacer
5′ crRNA trimming
No crRNA trimming
3′ 3′ crRNA trimming
3′ PAM 5′
Target DNA Target either RNA or DNA
Target DNA Stage 3: Interference
Figure 2 | Diversity of CRISPR-mediated adaptive immune systems in bacteria and archaea. A diverse set of CRISPR-associated (cas) genes (grey arrows) encode proteins required for new spacer sequence acquisition (Stage 1), CRISPR RNA biogenesis (Stage 2) and target interference (Stage 3). Each CRISPR locus consists of a series of direct repeats separated by unique spacer sequences acquired from invading genetic elements (protospacers). Protospacers are flanked by a short motif called the protospacer adjacent motif (PAM, **) that is located on the 5ʹ (type I) or 3ʹ (type II) side in foreign DNA10,51,52,59,67. Long CRISPR transcripts are processed into short crRNAs by distinct mechanisms. In type I and III systems, a CRISPR-specific endoribonuclease (yellow ovals and green circles, respectively) cleaves 8 nucleotides upstream of each spacer sequence16,18–21,64. In type III systems, the repeat sequence on the 3ʹ
end of the crRNA is trimmed by an unknown mechanism (green pacman, right). In type II systems, a trans-acting antisense RNA (tracrRNA) with complementarity to the CRISPR RNA repeat sequence forms an RNA duplex that is recognized and cleaved by cellular RNase III (brown ovals)17. This cleavage intermediate is further processed at the 5ʹ end resulting in a mature, approximately 40-nucleotide crRNA with an approximately 20-nucleotide 3ʹ-handle. In each system, the mature crRNA associates with one or more Cas proteins to form a surveillance complex (green rectangles). Type I systems encode a Cas3 nuclease (blue pacman), which may be recruited to the surveillance complex following target binding24,27,44. A short high-affinity binding site called a seed-sequence has been identified in some type I systems27,60, and genetic experiments suggest that type II systems have a seed sequence located at the 3ʹ end of the crRNA spacer sequence53. 1 6 F E B R UA RY 2 0 1 2 | VO L 4 8 2 | N AT U R E | 3 3 3
INSIGHT REVIEW preferentially at the leader end of the CRISPR, but the mechanism of leader end recognition is unknown. One simple model suggests that the leader sequence contains a recognition element that recruits the integration machinery. It is equally possible that integration relies on singlestranded regions of the CRISPR DNA that are made available during transcription. Transcription-associated recombination is involved in genome stability58, and a mechanism that couples integration together with transcription would link the process of adaptation to CRISPR RNA expression, ensuring that spacers from the most recent virus or plasmid are transcribed first. The integration machinery must be able to distinguish foreign DNA from that of the host genome. The molecular cues that are involved in the distinction of ‘self ’ from ‘non-self ’ are still unknown, but sequencing of CRISPR loci following phage challenge suggests that spacer sequences are not selected at random15,29,51,52,59,60. Mapping spacer sequences onto viral genomes reveals a short sequence motif proximal to the protospacer, which is referred to as the protospacer adjacent motif (PAM). PAM sequences are only a few nucleotides long, and the precise sequence varies depending on the CRISPR system type59. This variation suggests that one or more of the Cas proteins associated with each immune system is involved in PAM recognition, but the mechanism governing this specificity is unknown.
CRISPR RNA biogenesis Spacer acquisition is the first step of immunization, but successful protection from bacteriophage or plasmid challenge requires the CRISPR to be transcribed and processed into short CRISPR-derived RNAs (crRNAs). crRNAs were first detected by small RNA profiling in Archaeoglobus fulgidus61 and S. solfataricus62. Northern-blot analysis using probes against the repeat sequence of the CRISPR revealed a ‘ladder-like’ pattern of RNA consistent with a long precursor CRISPR RNA transcript (pre-crRNA) that was processed at approximately 60-nucleotide intervals. In fact, the 3ʹ ends of cloned crRNAs were mapped to the middle of the CRISPR repeat61, which suggested that the repeat sequence was recognized and cleaved. The need for crRNAs in CRISPR-mediated defence was demonstrated initially by investigation of a CRISPR-specific endoribonuclease in E. coli called Cas6e (formerly known as Cse3 or CasE)22. Cas6e specifically binds and cleaves within each repeat sequence of the long pre-crRNA, resulting in a library of crRNAs that each contain a unique spacer sequence flanked by fragments of the adjacent repeats. Mutation of a conserved histidine blocks crRNA biogenesis and leaves the cell susceptible to phage infection22. The Cas6e protein consists of a double ferredoxin-like fold that selectively associates with specific RNA repeats and does not associate with DNA or CRISPR RNAs containing a non-cognate repeat sequence 18,20,22,63 (Fig. 4). Crystal structures of Cas6e bound to a CRISPR RNA repeat reveal a combination of sequence- and structure-specific interactions that explain the molecular mechanism of substrate recognition18,20. The repeat sequence of the E. coli CRISPR is partially palindromic, and the RNA forms a stable (approximately 20-nucleotide) stem loop22,35. A positively charged β-hairpin in Cas6e interacts with the major groove of the RNA duplex, which positions the 3ʹ strand of the crRNA stem along a conserved, positively charged cleft on one face of the protein18,20 (Fig. 4). RNA binding induces a conformational change that disrupts the bottom base pair of the stem and positions the scissile phosphate within the enzyme active site for site-specific cleavage20. CRISPR RNA cleavage occurs 8 nucleotides upstream of the spacer sequence, which results in 61-nucleotide mature crRNAs consisting of a 32-nucleotide spacer flanked by 8 nucleotides of the repeat sequence on the 5ʹ end (known as the 5ʹ-handle) and 21 nucleotides of the remaining repeat sequence on the 3ʹ end (Fig. 4). Cas6e remains tightly bound to the 3ʹ stem-loop20 and may serve as a nucleation point for assembly of a large effector complex, Cascade (CRISPR-associated complex for antiviral defence), that is required for phage silencing in the next stage of the immune system22,24,26 (discussed later). 3 3 4 | N AT U R E | VO L 4 8 2 | 1 6 F E B R UA RY 2 0 1 2
Crystal structures of CRISPR-specific endoribonucleases from two other immune systems offer additional insights into the co-evolutionary relationship between these specialized enzymes and their cognate RNAs16,19,21 (Fig. 4). In P. aeruginosa, Cas6f (previously known as Csy4) specifically binds and cleaves the CRISPR-RNA-repeat 8 nucleotides upstream of the spacer sequence, which leaves a similar 8-nucleotide 5ʹ-handle on mature crRNAs19. The co-crystal structure of Cas6f bound to its cognate RNA reveals interesting parallels between the method of RNA binding used by Cas6f and Cas6e18–20. Like Cas6e, the P. aeruginosa Cas6f protein recognizes the sequence and shape of a stable stem-loop in the crRNA repeat sequence by interacting extensively with the major groove of the double-stranded RNA. However, the structural elements responsible for this interaction are distinct between the two proteins18–20 (Fig. 4). The Cas6f protein has a two-domain architecture, which consists of an N-terminal ferredoxin-like fold similar to that in Cas6e, but its C-terminal domain is structurally distinct. An arginine-rich helix in the C-terminal domain of Cas6f inserts into the major groove of the crRNA duplex, and the bottom of the crRNA is positioned for sequencespecific hydrogen-bonding contacts in the RNA major groove. These contacts position the scissile phosphate of the crRNA in the enzyme active site so that cleavage occurs 8 nucleotides upstream of the spacer sequence18–20 (Fig. 4). Although Cas6f and Cas6e recognize the sequence and shape of the crRNA hairpin in their respective systems, CRISPR RNA repeats in other CRISPR systems are thought to be unstructured35. For example, the Cas6 protein from P. furiosus associates with CRISPR transcripts that are expected to contain unstructured repeats64. The specific recognition of an unstructured RNA repeat requires a distinct mechanistic solution for RNA substrate discrimination. Remarkably, crystallographic studies of the Cas6 protein from P. furiosus have revealed the same duplicated ferredoxin-like fold observed in the Cas6e protein, but with a different mode of RNA recognition involving the opposite face of the protein (Fig. 4). In Cas6, the two ferredoxin-like folds clamp the 5ʹ end of the single-stranded RNA repeat sequence in place21. Although the RNA in this structure is disordered in the enzyme active site, biochemical studies have shown that cleavage occurs 8 nucleotides upstream of the spacer sequence16,64. While the nucleotide sequences at the cleavage site vary for each of the different Cas6 proteins, all Cas6 family endoribonucleases cleave their cognate RNA 8 nucleotides upstream of the spacer sequence using a metal-ion-independent mechanism. Despite advances in our understanding of crRNA biogenesis, the diversity of cas genes has obscured identification of the protein factors responsible for CRISPR RNA processing in some systems. Type II immune systems consist of four cas genes, none of which have a detectable sequence similarity to known CRISPR-specific endoribonucleases. Recently, a different CRISPR RNA processing mechanism has been reported that involves RNase-III-mediated cleavage of double-stranded regions of the CRISPR RNA repeats17. The first indication of this mechanism came from deep sequencing of RNA from S. pyogenes. An abundant transcript containing a 25-nucleotide sequence that was complementary to the CRISPR repeat was identified. This RNA, termed tracrRNA (trans-activating CRISPR RNA), is coded on the opposite strand and just upstream of the CRISPR locus. Genetic and biochemical experiments demonstrated that tracrRNA and pre-crRNA are co-processed by RNase III, which produces cleavage products with a 2 nucleotide 3ʹ overhang17. In vivo processing of CRISPR RNAs required Cas9 (previously known as Csn1), although a precise role for this enzyme in RNA processing has not yet been defined. The essential role of cellular proteins that are not solely involved in CRISPR-mediated defence, such as RNase III, indicates that different host factors may be involved as ancillary components of these immune systems.
crRNA-guided interference The third stage of CRISPR-mediated immunity is target interference (Fig. 2). Here crRNAs associate with Cas proteins to form large CRISPRassociated ribonucleoprotein complexes that can recognize invading
REVIEW INSIGHT a
b Leader recognition Leader CRISPR DNA
Viral DNA DNA binding?
10 20 Base pairs
Recombination/repair New ‘spacer’ acquisition Leader CRISPR DNA
Figure 3 | Steps leading to new spacer integration. a, The Cas1 protein forms a stable homodimer where the two molecules (green and grey) are related by a pseudo-two-fold axis of symmetry (PBD ID: 3GOD)54,56. This organization creates a saddle-like structure in the N-terminal domain, in which β-hairpins (blue) from each symmetrically related molecule hang (like stirrups) that are separated by approximately 20 Å, and may interact with the phosphodiester backbone of double-stranded DNA. An electrostatic surface representation (bottom) reveals a cluster of basic residues (blue) that form a positively charged strip across the metalbinding surface of the C-terminal domain. This strip may serve as an electrostatic trap that positions DNA substrates proximally to catalytic
metal ions (green sphere). b, CRISPR adaptation occurs by integrating fragments of foreign nucleic acid preferentially at the leader end of the CRISPR, forming new repeat-spacer units in the process. Protospacers are chosen non-randomly and may be selected from regions flanking the protospacer adjacent motif (PAM). Coordinated cleavage of the foreign DNA and integration of the protospacer into the leader-end of the CRISPR occurs through a mechanism that duplicates the repeat sequence and thus preserves the repeat-spacer-repeat architecture of the CRISPR locus. Although the protein components required for this process have not been conclusively identified, Cas1 and other general recombination or repair factors have been implicated (blue ovals)32,54,56.
nucleic acids. Foreign nucleic acids are identified by base-pairing interactions between the crRNA spacer sequence and a complementary sequence from the intruder. Phage- and plasmid-challenge experiments performed in several model systems have demonstrated that crRNAs complementary to either the coding or the non-coding strand of the invading DNA can provide immunity14,22,47,60,65,66. This is indicative of an RNA-guided DNAtargeting system, and indeed a pathway for DNA silencing has recently been demonstrated in S. thermophilus15. DNA sequencing and Southern blots indicated that both strands of the target DNA are cleaved within the region that is complementary to the crRNA spacer sequence15. This mechanism efficiently eliminates foreign DNA sequences, which have been specified by the spacer region of the crRNA, but avoids targeting the complementary DNA sequences in the CRISPR region of the host chromosome. The mechanism for distinguishing self from non-self is built into the crRNA. The spacer sequence of each crRNA is flanked by a portion of the adjacent CRISPR repeat sequence, and any complementarity beyond the spacer into the adjacent repeat region signals self and prevents the destruction of the host chromosome67. However, not all CRISPR systems target DNA. In vitro experiments using enzymes from the type III-B CRISPR system of P. furiosus have shown that this system cleaves target RNA rather than DNA48. All DNA targeting systems encode a complementary DNA sequence for each crRNA in the CRISPR locus and therefore require a mechanism for distinguishing self (CRISPR locus) from non-self (invading DNA). In contrast, systems that target RNA may not be required to make this distinction because most CRISPR loci are transcribed only in one direction and thus do not generate complementary RNA targets. CRISPR systems that target RNA may be uniquely capable of defending against viruses that have RNA-based genomes. However, adaptation of the CRISPR in response to a challenge by an RNA-based virus will probably require the invading RNA to be reverse-transcribed into DNA before it can be integrated into the CRISPR locus.
Cas proteins directly participate in target binding. Recent biochemical studies have shown that CRISPR-associated complexes facilitate target recognition by enhancing sequence-specific hybridization between the CRISPR RNA and complementary target sequences27. A short high-affinity binding site located at one end of the crRNA spacer sequence governs the efficiency of target binding, and viruses that acquired a single mismatch in this region were able to escape detection by the immune system60. This high-affinity binding site is functionally analogous to the ‘seed’ sequence (Fig. 1) that has been identified in eukaryotic microRNAs (miRNAs)68. Structural and biochemical studies have shown that Argonaute proteins facilitate target recognition by pre-ordering the nucleotides at the 5ʹ end of the miRNA in a helical configuration69. This pre-ordering reduces the entropic penalty that is associated with helix formation and provides a thermodynamic advantage for target binding within this region. A similar mechanism may occur during crRNA target binding, providing an interesting example of convergent evolution between CRISPR-based immunity in prokaryotes and RNAi in eukaryotes (Fig. 1). Structural and biochemical studies have been performed on CRISPR-associated complexes isolated from three different type I CRISPR systems24–27,48. These complexes seem to share some general morphological features, but the precise special arrangement of the Cas proteins and their interactions with the crRNA have been unclear. Sub-nanometre-resolution structures of the CRISPR-associated complex from E. coli (Cascade) have recently been determined using cryo-electron microscopy26. This complex is comprised of an unequal stoichiometry of 5 functionally essential Cas proteins and a 61-nucleotide crRNA22,24,26. The structure reveals a sea-horseshaped architecture in which the crRNA is displayed along a helical arrangement of protein subunits that protect the crRNA from degradation26. The 5ʹ and 3ʹ ends of the crRNA form unique structures 1 6 F E B R UA RY 2 0 1 2 | VO L 4 8 2 | N AT U R E | 3 3 5
INSIGHT REVIEW Type I
crRNAs: 8-nt 5′-handle
Figure 4 | Diverse mechanisms of CRISPR RNA biogenesis. CRISPR RNA repeats are specifically recognized and cleaved by diverse mechanisms. In type I CRISPR systems, Cas6e (PDB ID: 2Y8W) and Cas6f (PDB ID: 2XLK) recognize the major groove of the crRNA stem-loop primarily through electrostatic interactions using a β-hairpin and α-helix, respectively18,19,20. Cleavage occurs at the double-stranded–single-stranded junction (black arrows), leaving an 8-nt 5ʹ-handle on mature crRNAs. In type II CRISPR systems, tracrRNA hybridizes to the pre-crRNA repeat to form duplex RNAs that are substrates for endonucleolytic cleavage by host RNase III (PDB ID: 2EZ6), an activity that may also require Cas9 (ref. 17). Subsequent trimming (red arrows) by an unidentified nuclease
removes leftover repeat sequences from the 5ʹ end. Cas6 (PDB ID: 3PKM) in type III-B CRISPR systems specifically recognizes single-stranded RNA, upstream of the scissile phosphate, on a face of the protein opposite that of the previously identified active site residues16,21,64. The remainder of the repeat substrate probably wraps around the protein (red dashed line) to allow cleavage 8 nucleotides upstream of the repeat-spacer junction. Subsequent 3ʹ trimming (red arrows) generates mature crRNAs of two discrete lengths. The N-terminal domain of all Cas 6 family proteins adopts a ferredoxin-like fold (light blue).The C-terminal domain of Cas6 and Cas6e also adopts a ferredoxin-like fold but the C-terminal domain of Cas6f is structurally distinct (dark blue).
that are anchored at opposite ends of the Cascade complex, displaying the 32-nucleotide spacer sequence for base-pairing with complementary targets. The structure of Cascade bound to a 32-nucleotide target sequence26 reveals a concerted conformational change that could be a signal for recruiting Cas3. Cas3 — the trans-acting nuclease of type I CRISPR systems — may function as a target ‘slicer’ in a similar way to Argonaute in RNAi pathways22,44,46,70. Although Cas3 was implicated previously in the process of self versus non-self discrimination, recent studies have demonstrated that Cascade recognizes the PAM directly and that mutations in the PAM decrease Cascade’s affinity for the target60. The importance of the PAM is highlighted by the recovery of phage and plasmid escape mutants, which frequently contain a single mutation in the PAM15,51–53,60. The structure of Cascade indicates that the PAM is positioned near the ‘tail’ of the sea-horse-shaped complex. High-resolution structures and mutational analysis of the nucleic acid and protein components in this and related systems are needed to determine the mechanisms of target authentication and degradation.
Laboratory strains of bacteria are grown in high-density bioreactors for many different applications in the food industry, and they are becoming increasingly important in the production of biofuels. CRISPR systems offer a natural mechanism for adapting economically important bacteria for resistance against multiple phages. The biochemical activities of various Cas proteins may have useful applications in molecular biology in much the same way that DNA restriction enzymes have revolutionized cloning and DNA manipulation. A wide range of CRISPR-specific endoribonucleases that recognize small RNA motifs with high affinity expand the number of tools available for manipulating nucleic acids. In addition, a crRNA-guided ribonucleoprotein complex in P. furiosus was shown to cleave target RNAs48. Site-specific cleavage of target RNA molecules could have a range of uses, from generating homogeneous termini after in vitro transcription to targeting a specific intracellular messenger RNA for inactivation in a similar way to RNAi. CRISPRs also provide a new mechanism for limiting the spread of antibiotic resistance or the transfer of virulence factors by blocking horizontal gene transfer15,47. In addition, CRISPRs participate in a regulatory mechanism that alters biofilm formation in P. aeruginosa74,75. Although the clinical relevance of CRISPRs remains to be demonstrated, the opportunities for creative implementation of this new gene-regulation system are perceivably vast.
Applications of CRISPR structure and function The sequence diversity of CRISPR loci, even within closely related strains, has been used for high-resolution genotyping and forensic medicine. This technique, known as spoligotyping (spacer oligotyping), has been applied successfully to the analysis of human pathogens, including Mycobacterium tuberculosis71, Corynebacterium diphtheriae72 and Salmonella enterica73. Spoligotyping was developed long before the function of CRISPRs was understood, but now that studies have begun to reveal the biological function and mechanism of CRISPR-mediated genetic silencing, new opportunities for creative applications have emerged. 3 3 6 | N AT U R E | VO L 4 8 2 | 1 6 F E B R UA RY 2 0 1 2
Future directions of CRISPR biology The discovery of some of the fundamental mechanisms of CRISPRbased adaptive immunity has raised new questions and highlighted the areas with the greatest potential for future research. Although CRISPR RNA processing and targeting steps are now understood in some detail, how and when target sequences are identified during a phage infection
REVIEW INSIGHT or plasmid transformation are still unclear. Furthermore, why DNA or RNA target sequences are chosen, and their fate once they are bound to a crRNA-targeting complex is not well understood. In addition, the mechanisms by which foreign sequences are selected and integrated into CRISPR loci are almost entirely unknown. Some CRISPR loci seem to be considerably more active than others, at least under laboratory conditions, so selection of the model organisms will be important. The diversity and prevalence of CRISPR systems throughout microbial communities ensures that new findings and applications in this field will be forthcoming in the years ahead. ■ 1. 2. 3. 4. 5. 6. 7. 8. 9. 10. 11. 12. 13. 14.
18. 19. 20. 21. 22.
23. 24. 25.
Hoskisson, P. A. & Smith, M. C. Hypervariation and phase variation in the bacteriophage ‘resistome’. Curr. Opin. Microbiol. 10, 396–400 (2007). Rodriguez-Valera, F. et al. Explaining microbial population genomics through phage predation. Nature Rev. Microbiol. 7, 828–836 (2009). Weinbauer, M. G. Ecology of prokaryotic viruses. FEMS Microbiol. Rev. 28, 127–181 (2004). Labrie, S. J., Samson, J. E. & Moineau, S. Bacteriophage resistance mechanisms. Nature Rev. Microbiol. 8, 317–327 (2010). Stern, A. & Sorek, R. The phage-host arms race: shaping the evolution of microbes. Bioessays 33, 43–51 (2010). Al-Attar, S., Westra, E. R., van der Oost, J. & Brouns, S. J. Clustered regularly interspaced short palindromic repeats (CRISPRs): the hallmark of an ingenious antiviral defense mechanism in prokaryotes. Biol. Chem. 392, 277–289 (2011). Deveau, H., Garneau, J. E. & Moineau, S. CRISPR/Cas system and its role in phage-bacteria interactions. Annu. Rev. Microbiol. 64, 475–493 (2010). Horvath, P. & Barrangou, R. CRISPR/Cas, the immune system of bacteria and archaea. Science 327, 167–170 (2010). Karginov, F. V. & Hannon, G. J. The CRISPR system: small RNA-guided defense in bacteria and archaea. Mol. Cell 37, 7–19 (2010). Makarova, K. S. et al. Evolution and classification of the CRISPR-Cas systems. Nature Rev. Microbiol. 9, 467–477 (2011). Marraffini, L. A. & Sontheimer, E. J. CRISPR interference: RNA-directed adaptive immunity in bacteria and archaea. Nature Rev. Genet. 11, 181–190 (2010). Sorek, R., Kunin, V. & Hugenholtz, P. CRISPR-a widespread system that provides acquired resistance against phages in bacteria and archaea. Nature Rev. Microbiol. 6, 181–186 (2008). Andersson, A. F. & Banfield, J. F. Virus population dynamics and acquired virus resistance in natural microbial communities. Science 320, 1047–1050 (2008). Barrangou, R. et al. CRISPR provides acquired resistance against viruses in prokaryotes. Science 315, 1709–1712 (2007). This study provided the first direct evidence for CRISPR immune system function, demonstrating that new spacer acquisition provides resistance against future phage infection in a cas gene-dependent manner. Garneau, J. E. et al. The CRISPR/Cas bacterial immune system cleaves bacteriophage and plasmid DNA. Nature 468, 67–71 (2010). This study showed that CRISPRs can acquire new spacers upon plasmid challenge and provided the first example of crRNA-guided cleavage of double-stranded DNA in vivo. Carte, J., Wang, R., Li, H., Terns, R. M. & Terns, M. P. Cas6 is an endoribonuclease that generates guide RNAs for invader defense in prokaryotes. Genes Dev. 22, 3489–3496 (2008). Deltcheva, E. et al. CRISPR RNA maturation by trans-encoded small RNA and host factor RNase III. Nature 471, 602–607 (2011). This study identified a new CRISPR RNA processing pathway that involves a trans-acting RNA complementary to the CRISPR repeat sequence and demonstrated that processing of this duplex is mediated by cellular RNase III. Gesner, E. M., Schellenberg, M. J., Garside, E. L., George, M. M. & MacMillan, A. M. Recognition and maturation of effector RNAs in a CRISPR interference pathway Nature Struct. Mol. Biol. 18, 688–692 (2011). Haurwitz, R. E., Jinek, M., Wiedenheft, B., Zhou, K. & Doudna, J. A. Sequenceand structure-specific RNA processing by a CRISPR endonuclease. Science 329, 1355–1358 (2010). Sashital, D. G., Jinek, M. & Doudna, J. A. An RNA induced conformational change required for CRISPR RNA cleavage by the endonuclease Cse3. Nature Struct. Mol. Biol. 18, 680–687 (2011). Wang, R., Preamplume, G., Terns, M. P., Terns, R. M. & Li, H. Interaction of the Cas6 riboendonuclease with CRISPR RNAs: recognition and cleavage. Structure 19, 257–264 (2011). Brouns, S. J. et al. Small CRISPR RNAs guide antiviral defense in prokaryotes. Science 321, 960–964 (2008). This paper revealed that long CRISPR RNA transcripts are processed into small RNAs (crRNAs) by a dedicated endoribonuclease and that the processed RNAs are bound by a Cas protein complex (Cascade), which together with Cas3 are required for phage protection. Hale, C., Kleppe, K., Terns, R. M. & Terns, M. P. Prokaryotic silencing (psi)RNAs in Pyrococcus furiosus. RNA 14, 2572–2579 (2008). Jore, M .M. et al. Structural basis for CRISPR RNA-guided DNA recognition by Cascade. Nature Struct. Mol. Biol. 18, 529–536 (2011). Lintner, N. G. et al. Structural and functional characterization of an archaeal clustered regularly interspaced short palindromic repeat (crispr)-associated complex for antiviral defense (CASCADE). J. Biol. Chem. 286, 21643–21656 (2011).
26. Wiedenheft, B. et al. Structures of the RNA-guided surveillance complex from a bacterial immune system. Nature 477, 486–489 (2011). This paper presented the first sub-nanometer resolution structures of Cascade, showing how the crRNA is accommodated within a large ribonucleoprotein complex that is involved in foreign nucleic acid surveillance. 27. Wiedenheft, B. et al. RNA-guided complex from a bacterial immune system enhances target recognition through seed sequence interactions. Proc. Natl Acad. Sci. USA 108, 10092–10097 (2011). 28. Ishino, Y., Shinagawa, H., Makino, K., Amemura, M. & Nakata, A. Nucleotide sequence of the iap gene, responsible for alkaline phosphatase isozyme conversion in Escherichia coli, and identification of the gene product. J. Bacteriol. 169, 5429–5433 (1987). 29. Bolotin, A., Ouinquis, B., Sorokin, A. & Ehrlich, S. D. Clustered regularly interspaced short palindrome repeats (CRISPRs) have spacers of extrachromosomal origin. Microbiology 151, 2551–2561 (2005). 30. Mojica, F. J., Diez-Villasenor, C., Garcia-Martinez, J. & Soria, E. Intervening sequences of regularly spaced prokaryotic repeats derive from foreign genetic elements. J. Mol. Evol. 60, 174–182 (2005). 31. Pourcel, C., Salvignol, G. & Vergnaud, G. CRISPR elements in Yersinia pestis acquire new repeats by preferential uptake of bacteriophage DNA, and provide additional tools for evolutionary studies. Microbiology 151, 653–663 (2005). 32. Makarova, K. S., Grishin, N. V., Shabalina, S. A., Wolf, Y. I. & Koonin, E. V. A putative RNA-interference-based immune system in prokaryotes: computational analysis of the predicted enzymatic machinery, functional analogies with eukaryotic RNAi, and hypothetical mechanisms of action. Biol. Direct 1, 7 (2006). 33. Grissa, I., Vergnaud, G. & Pourcel, C. CRISPRFinder: a web tool to identify clustered regularly interspaced short palindromic repeats. Nucleic Acids Res. 35, W52–W57 (2007). 34. Rousseau, C., Gonnet, M., Le Romancer, M. & Nicolas, J. CRISPI: a CRISPR interactive database. Bioinformatics 25, 3317–3318 (2009). 35. Kunin, V., Sorek, R. & Hugenholtz, P. Evolutionary conservation of sequence and secondary structures in CRISPR repeats. Genome Biol. 8, R61 (2007). 36. Bland, C. et al. CRISPR recognition tool (CRT): a tool for automatic detection of clustered regularly interspaced palindromic repeats. BMC Bioinformatics 8, 209 (2007). 37. Dsouza, M., Larsen, N. & Overbeek, R. Searching for patterns in genomic data. Trends Genet. 13, 497–498 (1997). 38. Edgar, R. C. PILER-CR: fast and accurate identification of CRISPR repeats. BMC Bioinformatics 8, 18 (2007). 39. Jansen, R., Embden, J. D., Gaastra, W. & Schouls, L. M. Identification of genes that are associated with DNA repeats in prokaryotes. Mol. Microbiol. 43, 1565–1575 (2002). 40. Pougach, K. et al. Transcription, processing and function of CRISPR cassettes in Escherichia coli. Mol. Microbiol. 77, 1367–1379 (2010). 41. Pul, U. et al. Identification and characterization of E. coli CRISPR-cas promoters and their silencing by H-NS. Mol. Microbiol. 75, 1495–1512 (2010). 42. Westra, E. R. et al. H-NS-mediated repression of CRISPR-based immunity in Escherichia coli K12 can be relieved by the transcription activator LeuO. Mol. Microbiol. 77, 1380–1393 (2010). 43. Haft, D. H., Selengut, J., Mongodin, E. F. & Nelson, K. E. A guild of 45 CRISPRassociated (Cas) protein families and multiple CRISPR/Cas subtypes exist in prokaryotic genomes. PLoS Comput. Biol. 1, e60 (2005). 44. Sinkunas, T. et al. Cas3 is a single-stranded DNA nuclease and ATP-dependent helicase in the CRISPR/Cas immune system. EMBO J. 30, 1335–1342 (2011). 45. Han, D. & Krauss, G. Characterization of the endonuclease SSO2001 from Sulfolobus solfataricus P2. FEBS Lett. 583, 771–776 (2009). 46. Mulepati, S. & Bailey, S. Structural and biochemical analysis of the nuclease domain of the clustered regularly interspaced short palindromic repeat (CRISPR) associated protein 3(CAS3). J. Biol. Chem. 286, 31896–31903 (2011). 47. Marraffini, L. A. & Sontheimer, E. J. CRISPR interference limits horizontal gene transfer in staphylococci by targeting DNA. Science 322, 1843–1845 (2008). 48. Hale, C. R. et al. RNA-guided RNA cleavage by a CRISPR RNA-Cas protein complex. Cell 139, 945–956 (2009). Most CRISPR systems appear to target DNA, but this article reported the isolation of a multisubunit ribonucleoprotein complex (Cmr-complex) from P. furriosus that cleaves target RNAs (not DNA) at a fixed distance from the 3΄ end of the crRNA-guide. 49. Tyson, G. W. & Banfield, J. F. Rapidly evolving CRISPRs implicated in acquired resistance of microorganisms to viruses. Environ. Microbiol. 10, 200–207 (2008). 50. Snyder, J. C., Bateson, M. M., Lavin, M. & Young, M. J. Use of cellular CRISPR (clusters of regularly interspaced short palindromic repeats) spacer-based microarrays for detection of viruses in environmental samples. Appl Environ Microbiol 76, 7251–7258 (2010). 51. Deveau, H. et al. Phage response to CRISPR-encoded resistance in Streptococcus thermophilus. J. Bacteriol. 190, 1390–1400 (2008). 52. Horvath, P. et al. Diversity, activity, and evolution of CRISPR loci in Streptococcus thermophilus. J. Bacteriol. 190, 1401–1412 (2008). 53. Sapranauskas, R. et al. The Streptococcus thermophilus CRISPR/Cas system provides immunity in Escherichia coli. Nucleic Acids Res. 39, 9275–9282 (2011). 54. Babu, M. et al. A dual function of the CRISPR-Cas system in bacterial antivirus immunity and DNA repair. Mol. Microbiol. 79, 484–502 (2011). 55. Han, D., Lehmann, K. & Krauss, G. SSO1450--a CAS1 protein from Sulfolobus 1 6 F E B R UA RY 2 0 1 2 | VO L 4 8 2 | N AT U R E | 3 3 7
56. 57. 58. 59. 60. 61. 62. 63. 64. 65. 66. 67.
68. 69. 70.
solfataricus P2 with high affinity for RNA and DNA. FEBS Lett. 583, 1928– 1932 (2009). Wiedenheft, B. et al. Structural basis for DNase activity of a conserved protein implicated in CRISPR-mediated antiviral defense. Structure 17, 904–912 (2009). Chen, C. S. et al. A proteome chip approach reveals new DNA damage recognition activities in Escherichia coli. Nature Methods 5, 69–74 (2008). Aguilera, A. The connection between transcription and genomic instability. EMBO J. 21, 195–201 (2002). Mojica, F. J., Diez-Villasenor, C., Garcia-Martinez, J. & Almendros, C. Short motif sequences determine the targets of the prokaryotic CRISPR defence system. Microbiology 155, 733–740 (2009). Semenova, E. et al. Interference by clustered regularly interspaced short palindromic repeat (CRISPR) RNA is governed by a seed sequence. Proc. Natl Acad. Sci. USA 108, 10098–10103 (2011). Tang, T. H. et al. Identification of 86 candidates for small non-messenger RNAs from the archaeon Archaeoglobus fulgidus. Proc. Natl Acad. Sci. USA 99, 7536–7541 (2002). Tang, T. H. et al. Identification of novel non-coding RNAs as potential antisense regulators in the archaeon Sulfolobus solfataricus. Mol. Microbiol. 55, 469–481 (2005). Ebihara, A. et al. Crystal structure of hypothetical protein TTHB192 from Thermus thermophilus HB8 reveals a new protein family with an RNA recognition motif-like domain. Protein Sci. 15, 1494–1499 (2006). Carte, J., Pfister, N. T., Compton, M. M., Terns, R. M. & Terns, M. P. Binding and cleavage of CRISPR RNA by Cas6. RNA 16, 2181–2188 (2010). Gudbergsdottir, S. et al. Dynamic properties of the Sulfolobus CRISPR/Cas and CRISPR/Cmr systems when challenged with vector-borne viral and plasmid genes and protospacers. Mol. Microbiol. 79, 35–49 (2011) Manica, A., Zebec, Z., Teichmann, D. & Schleper, C. In vivo activity of CRISPRmediated virus defence in a hyperthermophilic archaeon. Mol. Microbiol. 80, 481–491 (2011). Marraffini, L. A. & Sontheimer, E. J. Self versus non-self discrimination during CRISPR RNA-directed immunity. Nature 463, 568–571 (2010). This study identified a mechanism for distingishing self from non-self, which relies on base-pairing potential in regions outside the crRNA spacer sequence. Bartel, D. P. MicroRNAs: genomics, biogenesis, mechanism, and function. Cell 116, 281–297 (2004). Parker, J. S., Parizotto, E. A., Wang, M., Roe, S. M. & Barford, D. Enhancement of the seed-target recognition step in RNA silencing by a PIWI/MID domain protein. Mol. Cell 33, 204–214 (2009). Beloglazova, N. et al. Structure and activity of the Cas3 HD nuclease MJ0384, an
3 3 8 | N AT U R E | VO L 4 8 2 | 1 6 F E B R UA RY 2 0 1 2
effector enzyme of the CRISPR interference. EMBO J. 30, 4616–4627 (2011). 71. Groenen, P. M., Bunschoten, A. E., van Soolingen, D. & van Embden, J. D. Nature of DNA polymorphism in the direct repeat cluster of Mycobacterium tuberculosis; application for strain differentiation by a novel typing method. Mol. Microbiol. 10, 1057–1065 (1993). 72. Mokrousov, I., Limeschenko, E., Vyazovaya, A. & Narvskaya, O. Corynebacterium diphtheriae spoligotyping based on combined use of two CRISPR loci. Biotechnol. J. 2, 901–906 (2007). 73. Liu, F. et al. Novel virulence gene and clustered regularly interspaced short palindromic repeat (CRISPR) multilocus sequence typing scheme for subtyping of the major serovars of Salmonella enterica subsp. enterica. Appl. Environ. Microbiol. 77, 1946–1956 (2011). 74. Cady, K. C. & O’Toole, G. A. Non-identity targeting of yersinia-subtype CRISPRprophage interaction requires the Csy and Cas3 proteins. J. Bacteriol. 193, 3433–3445 (2011). 75. Zegans, M. E. et al. Interaction between bacteriophage DMS3 and host CRISPR region inhibits group behaviors of Pseudomonas aeruginosa. J. Bacteriol. 191, 210–219 (2009). 76. Obbard, D. J., Gordon, K. H. J., Buck, A. H. & Jiggins, F. M. The evolution of RNAi as a defence against viruses and transposable elements. Philos. Trans. R. Soc. B Biol. Sci. 364, 99–115 (2009). 77. Aravin, A. A., Hannon, G. J. & Brennecke, J. The Piwi-piRNA pathway provides an adaptive defense in the transposon arms race. Science 318, 761–764 (2007). 78. Aravin, A. A. et al. Double-stranded RNA-mediated silencing of genomic tandem repeats and transposable elements in the D-melanogaster germline. Curr Biol 11, 1017–1027 (2001). 79. Bartel, D. P. MicroRNAs: target recognition and regulatory functions. Cell 136, 215–233 (2009). 80. Guo, H., Ingolia, N. T., Weissman, J. S. & Bartel, D. P. Mammalian microRNAs predominantly act to decrease target mRNA levels. Nature 466, 835–840 (2010). Acknowledgements B.W. is a Howard Hughes Medical Institute Fellow of the Life Sciences Research Foundation. S.H.S. acknowledges support from the National Science Foundation and National Defense Science & Engineering Graduate Research Fellowship programs. J.A.D. is an Investigator of the Howard Hughes Medical Institute. Author Information Reprints and permissions information is available at www.nature.com/reprints. The authors declare no competing financial interests. Readers are welcome to comment on the online version of this article at www.nature.com/nature. Correspondence should be addressed to J.A.D. (email@example.com).
Modular regulatory principles of large non-coding RNAs Mitchell Guttman1, 2 & John L. Rinn1,3
It is clear that RNA has a diverse set of functions and is more than just a messenger between gene and protein. The mammalian genome is extensively transcribed, giving rise to thousands of non-coding transcripts. Whether all of these transcripts are functional is debated, but it is evident that there are many functional large non-coding RNAs (ncRNAs). Recent studies have begun to explore the functional diversity and mechanistic role of these large ncRNAs. Here we synthesize these studies to provide an emerging model whereby large ncRNAs might achieve regulatory specificity through modularity, assembling diverse combinations of proteins and possibly RNA and DNA interactions.
ore than half a century after being placed as the central component in the flow of genetic information from gene to protein, it is now accepted that RNA can perform diverse roles. Shortly after the discovery of messenger RNA, a large class of heteronuclear RNAs (hnRNAs)1 was described, which did not include mRNA or associate with polyribosomes2. Following years of sifting through these hnRNAs, the first RNA subfamilies were identified. These included small nuclear RNAs involved in splicing regulation3 and small nucleolar RNAs involved in ribosome biogenesis4, as well as the ribosomal RNAs and transfer RNAs involved in translation5,6. The world of RNA genes became even more complex with the discovery of RNAs that resembled mRNA in length and splicing structure but did not code for proteins. The first example was H19, which was identified as an RNA that was induced during liver development in the mouse7. The mouse H19 transcript contained no large open reading frames (ORFs), but instead only small sporadic ORFs that were not evolutionarily conserved, did not template translation in vivo and did not produce an identifiable protein product8. Shortly afterwards, another non-coding RNA (ncRNA), termed XIST, was found to be expressed exclusively from the inactive X chromosome9 and later demonstrated to be required for X inactivation in mammals10. Over the next two decades, more large ncRNA genes were discovered including Airn11, Tug1 (ref. 12), NRON13 and HOTAIR14. With the availability of a draft sequence of the human genome, it became clear that much of the mammalian genome is transcribed15–18. These transcripts were mapped to discrete loci throughout the genome. Over the next 10 years, both large and small RNA transcripts were discovered at an unprecedented rate15,17–20; however, the functional significance of most of these transcripts was unclear. Although some of these could be considered noise21,22, there are still many large ncRNAs that are known to have diverse functions23–29. This Review focuses on the classic examples of large ncRNAs that have helped to form the basis of more recent global studies of coding potential, function and mechanism. We discuss the concepts that have emerged from these examples that provide a framework for understanding the principles of RNA interactions. We propose that by assembling distinct regulatory components, large ncRNAs could produce intricate functional specificity, which is suggestive of a possible modular RNA code.
RNA maps After the sequencing of the human genome, the next major hurdle was to define the genes it encoded. To do this, several research groups developed tiling microarrays17,19,20 and complementary DNA sequencing
methods15 to investigate transcriptional activity across the human genome, which led to the observation of widespread transcription of the genome. These studies, although limited to specific tissues and cell types, demonstrated that the mammalian genome encodes many thousands of non-coding transcripts including both short (<200 nucleotides in length) and long (>200 nucleotides in length) transcripts. In this Review, we focus on large ncRNAs produced from long transcripts, including those that originate from intergenic loci or overlapping protein-coding genes. Dramatic innovations in sequencing technologies have allowed the deep sequencing of cDNAs, known as RNA-Seq30; this deep sequencing, coupled with new computational methods for assembling the transcriptome31, has identified non-coding transcripts across many different cell types and tissues31,32. It is now clear that there are thousands of wellexpressed large ncRNAs with exquisite cell-type and tissue specificity31–33. As the numbers of identified non-coding transcripts increased, so did the uncertainty regarding their function; this led some authors to express concern that many of these transcripts may be just transcriptional noise21,22 with no function or incidental by-products of transcription from enhancer regions34,35. These concerns are supported by the observations that many of these transcripts are expressed at extremely low levels32,36 and they have lower levels of evolutionary conservation than protein-coding genes25,31,37. Although some of these transcripts may indeed be transcriptional noise21, the remaining transcripts consist of many distinct subclasses, including processed small RNAs18,29,38, promoter-associated RNAs18,39, transcripts from enhancer regions34,35 and functional large ncRNAs14,23; each class varies in its expression and conservation properties31,37. Distinguishing between these classes of RNA transcripts requires additional biological information including the coding potential of the RNA and the chromatin modifications of the corresponding genomic region (Fig. 1a).
Chromatin signatures Genomic DNA is wrapped around histone proteins and packaged into higher-order structures termed chromatin40. These histones can be modified in different ways that are indicative of the underlying DNA functional state. Advances in sequencing technologies have allowed the comprehensive characterization of the chromatin-modification landscape of mammalian genomes41–44. These studies revealed combinations of histone modifications (termed chromatin signatures) that correspond to various gene properties, including a signature for active
Broad Institute of MIT and Harvard, 7 Cambridge Center, Cambridge, Massachusetts 02142, USA. 2Department of Biology, Massachusetts Institute of Technology, Cambridge, Massachusetts 02139, USA. 3Stem Cell and Regenerative Biology, Harvard University, Cambridge, Massachusetts 02138, USA. 1 6 F E B R UA RY 2 0 1 2 | VO L 4 8 2 | N AT U R E | 3 3 9
INSIGHT REVIEW transcription41,44. This signature consists of a short stretch of trimethylation of histone protein H3 at the lysine in position 4 (H3K4me3), which corresponds to promoter regions, followed by a longer stretch of trimethylation of histone H3 at the lysine in position 36 (H3K36me3), which covers the entire transcribed region41,44 (Fig. 1a). Chromatin maps revealed that, similar to protein-coding genes, many ncRNA genes also contain a ‘K4–K36’ signature44. By searching
for K4–K36 domains that do not overlap with known genes, chromatin signatures revealed approximately 1,600 regions in the mouse genome and approximately 2,500 regions in the human genome that were actively transcribed25,45. The vast majority of these intergenic K4– K36 domains produce multi-exonic RNAs that have little capability to encode a conserved protein25,31. RNAs expressed from these K4–K36 domains were termed large intergenic ncRNAs (lincRNAs) because
a RNA map
Chromatin signature (K4–K36)
K4me1 and p300
SIRT1 (protein-coding gene)
Tarsal-less (short peptides)
1A 2A 3A AA ORF length = 576 aa
Figure 1 | Layering of genomic regions. a, Genomic regions are colour-coded by the presence of different genomic annotations. RNA transcription of a locus (grey), K4–K36 chromatin signature (red), K4me1 modification and transcriptional activator p300 (green) and protein-coding potential (blue). By overlaying this information, distinct transcripts are revealed, including ncRNAs (red), protein-coding genes (purple) and transcripts from enhancer regions (green). b, A cross-species alignment of a coding and a non-coding gene. Boxes represent codons, and each row represents a different aligned species. Blue boxes represent mutations that cause a synonymous substitution, and red boxes represent mutations that cause a non-synonymous substitution. A score capturing the coding potential of a sequence across species aligns sequences in all frames and scores mutations that maintain coding potential (blue boxes) 3 4 0 | N AT U R E | VO L 4 8 2 | 1 6 F E B R UA RY 2 0 1 2
Max ORF length = 172 aa
ORF length = 11 aa = 32 aa
relative to mutations that break coding potential (that is, non-synonymous mutations, stop codons and frameshifting insertions or deletions) (red boxes). c, The coding potential score is shown for three gene types, SIRT1 (a protein-coding gene), XIST (ncRNA gene) and tarsal-less (smallpeptide coding gene), in which positive scores represent coding regions (blue) and negative scores represent non-coding regions (red). In each example, the gene structure is shown, where blue boxes represent known protein-coding exons and red boxes represent non-coding exons. SIRT1 with an ORF length of 576 amino acids (aa) contains a positive score over each coding exon but not the non-coding regions. XIST with an ORF length of 172 amino acids contains negative scores over the entire transcribed region. tarsal-less with an ORF of 11 and 32 amino acids, contains positive scores over all known small peptides.
REVIEW INSIGHT identification by this chromatin signature required the RNAs to be contained within the intergenic regions25. Similarly, chromatin-state maps revealed that active enhancer regions contained short stretches of H3 lysine 4 monomethylation (H3K4me1) (ref. 43) and the transcriptional coactivator p300 (ref. 42), as well as additional modifications46 (Fig. 1a). By coupling RNA sequencing and chromatin maps, many of the already identified non-coding transcripts were observed to be transcribed from active enhancers34,35. However, lincRNAs and transcripts from enhancer regions are distinct classes, which are marked by different chromatin signatures25,34. Although it needs to be determined whether transcripts originating from enhancers have a function34,35, the functional importance of lincRNAs is becoming clearer14,23,24,26,28,47. Several of these lincRNAs have been shown to have enhancer-like functions as they activate the expression of neighbouring genes24,28.
Coding potential Determining whether a transcript is non-coding is challenging because a long non-coding transcript is likely to contain an ORF purely by chance48. Accordingly, the evidence for the absence of coding potential for the XIST and H19 genes came from the lack of evolutionary conservation of the identified ORFs, the lack of homology to known protein domains and the inability to template significant protein production8,49. These principles have been generalized to classify coding potential across thousands of transcripts by scoring conserved ORFs across dozens of species50,51, by searching for homology in large protein-domain databases52, and by sequencing RNA associated with polyribosomes53. Computational methods such as the ‘codon substitution frequency’ algorithm50,51 leverage evolutionary information to determine whether an ORF is conserved across species and provide a general strategy for determining coding potential (Fig. 1b, c). Owing to the large number of available genome sequences, these methods have been used to accurately determine conserved coding potential in regions as small as 5 amino acids25, which makes them extremely sensitive to the potentially small peptides, such as the 11 amino acid peptide encoded by the tarsal-less gene54,55 (Fig. 1c). Despite their sensitivity, conservation-based methods may fail to detect newly evolved proteins because they do not contain a conserved ORF50,51. However, because many ncRNAs show clear evolutionary constraint25,31,37 but no evolutionarily conserved ORF, this indicates that the observed evolutionary selection is not due to a newly evolved protein. Experimental methods, such as ribosome profiling, have provided a strategy for identifying ribosome occupancy on RNA, which have been proposed as a method for distinguishing between coding and noncoding transcripts53. However, this still needs to be tested because noncoding transcripts that show an association with the ribosome have not been shown to have a protein product53,56. Importantly, an association of RNA with a ribosome alone cannot be taken as evidence of proteincoding potential because both the ncRNAs of H19 and TUG1 can be detected in the ribosome53,57 despite having clear roles as ncRNAs8,45,58,59. An alternative explanation for these observed associations is ‘translational noise’, spurious association that may lead to non-functional translation products22. Consistent with this, virtually all of the transcripts that have been suggested to encode small peptides by ribosome profiling53 lack the evolutionary conservation of their proposed coding regions25,31, which is in striking contrast to almost all known proteincoding genes60, including the few well-characterized functional small peptides56,61,62 (Fig. 1c). Accordingly, identification of any new proteincoding gene requires the clear demonstration of the function of the protein product in vivo54,55.
Global identification of ncRNA function Identifying the functional role of an ncRNA requires direct perturbation experiments, such as loss-of-function and gain-of-function. Individual ncRNAs involved in specific processes have been functionally characterized (see ref. 63 for a review). For example, XIST is crucial for random
inactivation of the X chromosome10; Air is crucial for imprinting control at the Igf2r locus11; HOTAIR affects expression of the HOXD gene family14, as well as other genes throughout the genome45,64,65; HOTTIP affects expression of the HOXA gene family28; lincRNA-RoR affects reprogramming efficiency47; NRON affects NFAT transcription factor activity13; and Tug1 affects retina development through the regulation of the cell cycle12. Although there are now many examples of large ncRNAs that are required for the correct regulation of gene expression, this is just one of many functions in which they are involved; ranging from telomere replication66 to translation67. The global characterization of ncRNA function has proved to be challenging because, in most cases, it is unclear which phenotype to investigate13. One approach to classifying the putative function of ncRNAs uses ‘guilt-by-association’25. This approach associates ncRNAs with biological processes based on a common expression pattern across cell types and tissues (Fig. 2a) and can therefore identify groups of ncRNAs that are associated with specific cellular processes (Fig. 2b). This approach has been used to predict roles for hundreds of ncRNAs in diverse biological processes such as stem cell pluripotency, immune responses, neural processes and cell-cycle regulation25,27,36. Although these correlations cannot prove that ncRNAs have a function in these processes, they do provide a hypothesis for targeted loss-offunction experiments. For example, lincRNA-p21 was predicted to be associated with the p53-mediated DNA damage response25, and indeed lincRNA-p21 was found to be a target of p53 and on perturbation was shown to regulate apoptosis in response to DNA damage26. In the same way, the ncRNA PANDA (p21 associated ncRNA DNA damage activated) was implicated, and was demonstrated to have a function, in the regulation of apoptosis27. Another ncRNA, lincEnc1 (ref. 25), was predicted to have a role in cell-cycle regulation in embryonic stem (ES) cells and has been shown in a separate study to affect the proliferation of ES cells68. Alternatively, global approaches can be used to determine function, such as systematic RNA interference (RNAi) knockdown followed by gene-expression profiling. Unlike correlation analysis, these perturbation-based experiments provide evidence for the function of an ncRNA23. Methods to classify function using this approach are conceptually similar to guilt-by-association because the function can be inferred on the basis of the genes that are affected by loss of function of ncRNAs23. A systematic perturbation study demonstrated that knockdown of the vast majority of lincRNAs expressed in ES cells had a major effect on gene expression23. The gene-expression signatures revealed dozens of lincRNAs that block key lineage-commitment programs within ES cells and function in crucial ES cell regulatory and signalling pathways. Importantly, this study also identified 26 lincRNAs that are required to maintain the pluripotent state23. Not all non-coding transcripts are functional RNA molecules. Several examples of intergenic transcription have been identified in which the process of transcription alone changes the chromatin- and transcriptionfactor-binding landscape to allow activation and repression of neighbouring genes69,70. Methods that degrade RNA after its transcription, such as RNAi, can distinguish between a functional RNA molecule and the process of transcription, on which there should be no observable effect after RNA degradation. Collectively, the genome-wide guilt-byassociation approach and targeted and global perturbation studies have demonstrated that large ncRNAs have a crucial regulatory role in diverse biological processes23,25–27,32,47.
cis- versus trans-regulatory mechanisms The discovery that the XIST product was an ncRNA, led immediately to the suggestion of a model for how it could function in an allelespecific manner9. In theory, an ncRNA has an intrinsic cis-regulatory capacity because it can function while remaining tethered to its own locus9,71 (Fig. 2c), whereas an mRNA must be dissociated, exported and translated for it to function. Here we define a cis-regulator as one that exerts its function on a neighbouring gene on the same allele from which it is transcribed, and define a trans-regulator as one that 1 6 F E B R UA RY 2 0 1 2 | VO L 4 8 2 | N AT U R E | 3 4 1
INSIGHT REVIEW have used correlation analysis and identified a significant correlation of expression between ncRNAs and their neighbouring protein-coding genes21,72. However, several of these cases have been demonstrated to be trans-regulatory models, and the apparent correlations are due to shared upstream regulation (such as, lincRNAp21 (ref. 26) and lincRNA-Sox2 (ref. 25)), positional correlation
Cell type n
Cell type 1 Cell type 2 Cell type 3
does not meet this criterion. Owing to the unique cis-regulatory capability of ncRNAs, it has been speculated that cis-regulation could be a common mechanism for large ncRNAs24,71. However, global functional evidence strongly suggests that this is not the case (Box 1). To distinguish cis- from trans-regulatory models, initial studies b
For example, cell differentiation
For example, cancer cell formation
? Polymerase Mark
Figure 2 | Classification of ncRNA function. a, Illustration of an ncRNA with expression patterns related to the NFкB pathway. Each row represents a gene, and a positive association (red box) is assigned between the ncRNA and the pathway based on the correlation of the genes in the process. Similarly, the ncRNA is assigned negative association (blue box) with the p53 pathway based on anticorrelation with the genes in the process. b, The scores for each functional term and ncRNA can be clustered to identify classes of ncRNAs. In this example (adapted, with permission, from ref. 25) each column represents a different ncRNA, and each row represents a different functional term. c, A model of ncRNAs that have a cis-function by remaining tethered to their site of transcription. In this model, RNA 3 4 2 | N AT U R E | VO L 4 8 2 | 1 6 F E B R UA RY 2 0 1 2
polymerase (green) transcribes an RNA (red), which can associate with regulatory proteins (purple) to affect neighbouring regions, as proposed for XIST9,71. d, One model for ncRNA trans-regulation. In this model an ncRNA can associate with DNA-binding proteins (blue) and regulatory proteins to localize and affect the expression of the targets, as proposed for HOTAIR64. e, A model for ncRNAs that bind regulatory proteins and change their activity, in this case leading to a change in modification state and expression of the target gene, as proposed for the CCND1 ncRNAs, which interact with the TLS protein89. f, A model for ncRNAs that act as ‘decoys’. In this model, ncRNAs bind protein complexes and prevent them from binding to their proper regulatory targets, as proposed for GAS5 and PANDA27.
REVIEW INSIGHT BOX 1
Distinguishing cis- from trans-regulation If an ncRNA is a cis-regulator, then several observations will be true: (i) the geneexpression levels of a neighbouring gene will be correlated with the RNA expression across all conditions; (ii) loss-of-function of the RNA would affect expression of a neighbouring gene, and (iii) the ncRNA would affect expression of a neighbouring gene on the same allele that it is expressed from. The absence of any of these criteria supports trans-regulation. We illustrate this point using five common regulatory models. The figure shows what would be observed using specific computational and experimental methods for each regulatory model. The boxes with a tick indicate observed effects on neighbouring genes for each method, and boxes with a cross indicate no observed effect on neighbouring genes. Known ncRNA examples of each of these regulatory models are shown to the right of the figure.
Known ncRNA examples HOTAIR
trans lincRNA-P21 Cdkn1a/P21 trans
Allele 1 Allele 2
Allele 1 Allele 2
JPX Allele 1 Allele 2
Allele 1 Allele 2
(such as, HOTAIR14), transcriptional ‘ripple effects’21 and indirect regulation of neighbouring genes (Box 1). Consistent with these explanations, a recent study showed that an increased correlation of expression between ncRNAs and their neighbouring genes is comparable to that observed for protein-coding genes32. Recently, loss-of-function experiments have been used to investigate cis- versus trans-effects of lincRNAs. One study knocked down seven lincRNAs and identified no effects on neighbouring genes but did show an effect on other genes 45. A second study knocked down 12 lincRNAs, 7 of which had modest effects on some of the genes within a wide genomic neighbourhood24. More recently, a systematic study knocked down approximately 150 lincRNAs and identified no effect on the neighbouring genes for about 95% of the lincRNAs, which is similar to that observed for protein-coding genes23. Although perturbation experiments can demonstrate that an RNA functions as a trans-regulator, evidence for RNA acting as a cis-regulator is more difficult to obtain (Box 1). For example, perturbation experiments demonstrated that the ncRNA from JPX affects the expression of the neighbouring XIST gene, but as a transregulator73. Conclusive proof of cis-regulation requires the demonstration that an RNA regulates a neighbouring gene on the same allele (Box 1). So far, few studies have performed this test, and it is unclear what percentage of ncRNAs that are suggested to have a cis-function by loss-of-function experiments24,28 will pass this test. Together, these studies indicate that although some ncRNAs are cis-regulators9,11,74–76, the vast majority, which have been identified and characterized so far, function as trans-regulators14,23,26,45,73,77.
Formation of RNA–protein interactions The precise mechanism by which ncRNAs function remains poorly understood. However, one emerging theme is the interaction between ncRNAs and protein complexes. The functional importance of many ncRNA–protein interactions for correct transcriptional regulation has been demonstrated14,23,45,78–81, including several ncRNAs that are required for the correct localization of chromatin proteins to genomic DNA targets79–83.
XIST Neighbour affected
The XIST ncRNA is a key example demonstrating that RNA can play a direct role in silencing large genomic regions81 by physically interacting with the polycomb complex84, leading to the condensation of chromatin and transcriptional repression of an entire X chromosome85 (Fig. 2c). Similar to XIST, many ncRNAs have been identified that physically associate with chromatin-regulatory complexes and ‘guide’ the associated complexes to specific genomic DNA regions, including HOTAIR14, AIR86, KCNQ1ot1 (ref. 75) and lincRNA-p21 (ref. 26) (Fig. 2d). Biochemical evidence has demonstrated that many large ncRNAs interact with chromatin regulators23,45,87,88. The precise numbers vary depending on the experimental approach45,87, but a conservative estimate suggests that at least 30% of lincRNAs associate with at least 1 of 12 distinct chromatin-regulatory complexes, which include readers, writers and erasers of chromatin modifications23. Importantly, lincRNAs can provide regulatory specificity to these complexes because the knockdown of these lincRNAs affects a subset of the genes that are normally regulated by these complexes23,45. One hypothesis is that ncRNAs provide regulatory specificity by localizing chromatin-regulatory complexes to genomic DNA targets14,26,28,45,78,86. Several methods have been developed to generate maps of RNA–DNA proximity82,83, but it still needs to be determined what percentage of ncRNAs localize to genomic DNA regions and how these interactions occur. In addition to their role in chromatin regulation, ncRNAs can also modulate the regulatory activity of protein complexes (Fig. 2e). As an example, an ncRNA upstream of cyclin D1 can bind to the TLS (translocation in liposarcoma) RNA-binding protein, which changes it from an inactive to an active state89. Similarly, the NRON ncRNA can bind to the NFAT (nuclear factor of activated T cells)-transcription factor rendering it inactive because it prevents nuclear accumulation13. ncRNAs can also function as molecular ‘decoys’ by preventing correct regulation through competitive binding (Fig. 2f). For example, the GAS5 ncRNA binds to the glucocorticoid receptor and prevents the receptor from binding to its correct regulatory elements90, and the PANDA ncRNA can prevent NF-Y localization, which leads to apoptosis27. Similarly, several studies have shown that ncRNAs can function as decoys to other RNA species, such as miRNAs, to control miRNA levels91,92. 1 6 F E B R UA RY 2 0 1 2 | VO L 4 8 2 | N AT U R E | 3 4 3
INSIGHT REVIEW a 1. RNA–Protein
1 + 2 = DHFR
1 + 3 = Hotair and Xist
1 + 4 = Ribosome
a telomerase RNA component (TERC)94, which serves as a template for telomeric regulation and as a molecular scaffold for the polymerase enzyme around the RNA95 (Fig. 3b). Importantly, genetic studies demonstrated that TERC plays a modular functional role, as genetically swapping particular domains of TERC retained the overall function66. This indicated that TERC was made up of discrete functional modules to bring multiple proteins into the proximity of a protein66. More recently, HOTAIR was shown to contain distinct proteininteraction domains that can associate with polycomb repressive complex 2 (PRC2) (ref. 14) and the CoREST–LSD1 complex64, which together are required for correct function (Fig. 3b). XIST also has discrete functional domains. Through a series of genetic deletions XIST was shown to contain at least two discrete domains that are responsible for silencing (RepA) and localization (RepC)81 (Fig. 3b). These functional domains could be independently deleted without affecting the role of the other domain, which suggests the modular nature of the XIST ncRNA81. These functional domains of XIST also interact with discrete proteins; the silencing domain (RepA) binds to PRC2 and the localization domain (RepC) binds to YY1 (ref. 96) and hnRNPU97. These examples show that large ncRNAs can function as molecular scaffolds of protein complexes. Importantly, this phenomenon is likely to be a general one because approximately 30% of ES cell lincRNAs associate with multiple regulatory complexes23. In addition to interacting with multiple proteins, in several examples, ncRNAs have been shown to interact directly with both DNA and RNA. ncRNAs for example form triplex structures with DNA98,99 (Fig. 3a) such as a ncRNA that binds to the ribosomal DNA promoter and interacts with the DNMT3b protein to silence expression98. Furthermore, RNA can form traditional duplex base-pairing interactions with DNA, a property that has long been speculated for large ncRNAs71. Finally, RNA can form base-pair interactions with RNA (Fig. 3a), which are crucial for processes such as tRNA–mRNA anticodon recognition5, ribonuclease P recognition of pre-tRNAs5, miRNA targeting100, ribosome structure as a ribozyme67 and splicing regulation6. Despite these examples, the interactions between large ncRNAs, genomic DNA and other RNAs are not well characterized.
A potential modular RNA code 1 + 2 + 3 = Telomerase
Figure 3 | Modular principles of large ncRNA genes. a, The four principles of nucleic acid and protein interactions. (1) RNA–protein interactions, (2) DNA–RNA hybridization-based interactions, (3) DNA–protein interactions and (4) RNA–RNA hybridization based interactions. b, Each of these principles can be combined to build distinct complexes. For example, combining RNA– protein and RNA–DNA interactions can localize a protein complex to a specific DNA sequence in an RNA-dependent manner; as has been implicated for the DHFR99 promoter and localization of DNMT3b98. Combining RNA–protein and protein–DNA principles can also localize a diverse set of proteins, which have a molecular scaffold created by RNA, to a specific DNA sequence in a protein-dependent manner. The ribosome is a multifaceted combination of RNA–protein interactions that facilitate correct RNA–RNA interactions for the ribozyme activity of the ribosome. The telomere replication activity of telomerase is an example of combining RNA–protein, RNA–DNA and protein–DNA interactions.
Large ncRNAs as molecular scaffolds of proteins One emerging theme common to many large ncRNAs is the formation of multiple distinct RNA–protein interactions that are used to carry out their function (Fig. 3). The first indication of this phenomenon came from the discovery of telomerase93. Telomerase activity requires 3 4 4 | N AT U R E | VO L 4 8 2 | 1 6 F E B R UA RY 2 0 1 2
Collectively, the studies reviewed here suggest an intriguing hypothesis: large ncRNAs are flexible modular scaffolds23,64,66,81. In this model, RNA contains discrete domains that interact with specific protein complexes. These RNAs, through a combination of domains, bring specific regulatory components into proximity with each other, which results in the formation of a unique functional complex. These RNA regulatory complexes can include interactions with proteins but can also extend to RNA–DNA and RNA–RNA regulatory interactions. RNA is well-suited for this role because it is a malleable evolutionary substrate compared with a protein, allowing for the selection of discrete interaction domains5. Specifically, RNA can be easily mutated, tested and selected without breaking its core functionality5. This model of modular interactions can explain the observation that there are highly conserved ‘patches’ within large ncRNA genes25,31,37 that could have evolved for specific protein interactions26,81,84. The remaining regions may be more evolutionarily flexible, allowing the formation of new functional domains by random mutation and selection. This is consistent with the observation that non-constrained regions of telomerase are dispensable66. The model of RNA as a modular scaffold is not limited to protein interactions. RNA can also base-pair with DNA, which might be used to guide complexes to specific DNA sequences. Alternatively, RNAs might guide complexes by bridging together sets of DNA-binding proteins. Such a model could explain how the same protein complexes are guided to different DNA loci in distinct cell types. Large ncRNAs can also form RNA–RNA interactions, raising intriguing possibilities for future investigations. For example, two large RNA molecular scaffolds might be linked through RNA–RNA interactions. Another possibility is that RNA–RNA interactions could result in
REVIEW INSIGHT unique RNA structures that can interact with protein complexes that are not attainable by the individual units. This has been observed in the ribosome, where the combination of RNA–RNA and RNA–protein interactions are required for correct complex formation.
Outlook We are only beginning to understand the mechanism by which large ncRNAs carry out their regulatory function. A modular RNA regulatory code is an attractive hypothesis but remains to be tested; in particular, the way in which large ncRNAs, and proteins interact, and the underlying molecular principles are still unknown. Understanding these principles will require the identification of the sites of the RNA–protein interactions and the exact RNA-binding proteins in vivo. Furthermore, the way in which large ncRNAs localize to their target genes is unknown but could involve direct RNA–DNA interactions (Fig. 3a) or interactions with proteins that contain DNA recognition elements, which has been suggested for XIST96 and HOTAIR64. To gain insight into these processes, it will be important to catalogue the interactions that ncRNAs form with genomic DNA and RNAs. These data will help elucidate the rules that guide these interactions as well as the functional implications of these associations, which can then be tested experimentally. If large ncRNAs are truly modular, then each individual domain would have a unique function that is independent of other domains. Demonstrating modularity will require the genetic deletion of domains and spacer regions, as well as domain-swapping experiments. Learning these principles would result in a defined ‘modular RNA code’ for how RNAs can affect cell states. By truly understanding this modular RNA code, it may be possible to create synthetically engineered RNAs that could interact with both nucleic acids and protein modules to carry out engineered regulatory roles. However, at present, it is premature to dismiss the possibility of large ncRNAs having other mechanisms of action that may not fit neatly into this modular RNA code. In the meantime, it is clear that mammalian genomes encode a diverse set of large important ncRNAs. ■ 1.
3. 4. 5. 6. 7. 8.
9. 10. 11. 12. 13. 14. 15.
Warner, J. R., Soeiro, R., Birnboim, H. C., Girard, M. & Darnell, J. E. Rapidly labeled HeLa cell nuclear RNA. I. Identification by zone sedimentation of a heterogeneous fraction separate from ribosomal precursor RNA. J. Mol. Biol. 19, 349–361 (1966). Salditt-Georgieff, M., Harpold, M. M., Wilson, M. C. & Darnell, J. E., Jr. Large heterogeneous nuclear ribonucleic acid has three times as many 5΄ caps as polyadenylic acid segments, and most caps do not enter polyribosomes. Mol. Cell. Biol. 1, 179–187 (1981). This paper demonstrates an abundant class of RNA species that do not enter polyribosomes. Weinberg, R. A. & Penman, S. Small molecular weight monodisperse nuclear RNA. J. Mol. Biol. 38, 289–304 (1968). Zieve, G. & Penman, S. Small RNA species of the HeLa cell: metabolism and subcellular localization. Cell 8, 19–31 (1976). Gesteland, R. F., Cech, T. & Atkins, J. F. The RNA World : The Nature of Modern RNA Suggests a Prebiotic RNA World. 3rd edn (Cold Spring Harbor Laboratory Press, 2006). Eddy, S. R. Non-coding RNA genes and the modern RNA world. Nature Rev. Genet. 2, 919–929 (2001). Pachnis, V., Brannan, C. I. & Tilghman, S. M. The structure and expression of a novel gene activated in early mouse embryogenesis. EMBO J. 7, 673–681 (1988). Brannan, C. I., Dees, E. C., Ingram, R. S. & Tilghman, S. M. The product of the H19 gene may function as an RNA. Mol. Cell. Biol. 10, 28–36 (1990). This paper was the first report of a large ncRNA showing that the H19 transcript lacked conserved ORFs and did not make a protein product in vivo. Brown, C. J. et al. A gene from the region of the human X inactivation centre is expressed exclusively from the inactive X chromosome. Nature 349, 38–44 (1991). Penny, G. D., Kay, G. F., Sheardown, S. A., Rastan, S. & Brockdorff, N. Requirement for Xist in X chromosome inactivation. Nature 379, 131–137 (1996). Sleutels, F., Zwart, R. & Barlow, D. P. The non-coding Air RNA is required for silencing autosomal imprinted genes. Nature 415, 810–813 (2002). Young, T. L., Matsuda, T. & Cepko, C. L. The noncoding RNA taurine upregulated gene 1 is required for differentiation of the murine retina. Curr. Biol. 15, 501–512 (2005). Willingham, A. T. et al. A strategy for probing the function of noncoding RNAs finds a repressor of NFAT. Science 309, 1570–1573 (2005). Rinn, J. L. et al. Functional demarcation of active and silent chromatin domains in human HOX loci by noncoding RNAs. Cell 129, 1311–1323 (2007). Carninci, P. et al. The transcriptional landscape of the mammalian genome. Science 309, 1559–1563 (2005).
16. 17. 18. 19. 20. 21. 22. 23. 24. 25.
26. 27. 28. 29. 30. 31. 32. 33. 34. 35. 36. 37. 38. 39. 40. 41. 42. 43. 44. 45.
46. 47. 48. 49. 50.
This paper describes the large-scale cDNA sequencing efforts in the mouse genome and reveals many thousands of non-coding transcripts. Birney, E. et al. Identification and analysis of functional elements in 1% of the human genome by the ENCODE pilot project. Nature 447, 799–816 (2007). Bertone, P. et al. Global identification of human transcribed sequences with genome tiling arrays. Science 306, 2242–2246 (2004). Kapranov, P. et al. RNA maps reveal new RNA classes and a possible function for pervasive transcription. Science 316, 1484–1488 (2007). Rinn, J. L. et al. The transcriptional activity of human Chromosome 22. Genes Dev. 17, 529–540 (2003). Kapranov, P. et al. Large-scale transcriptional activity in chromosomes 21 and 22. Science 296, 916–919 (2002). Ebisuya, M., Yamamoto, T., Nakajima, M. & Nishida, E. Ripples from neighbouring transcription. Nature Cell Biol. 10, 1106–1113 (2008). Struhl, K. Transcriptional noise and the fidelity of initiation by RNA polymerase II. Nature Struct. Mol. Biol. 14, 103–105 (2007). Guttman, M. et al. lincRNAs act in the circuitry controlling pluripotency and differentiation. Nature 477, 295–300 (2011). Orom, U. A. et al. Long noncoding RNAs with enhancer-like function in human cells. Cell 143, 46–58 (2010). Guttman, M. et al. Chromatin signature reveals over a thousand highly conserved large non-coding RNAs in mammals. Nature 458, 223–227 (2009). This paper applied a chromatin signature to identify lincRNAs and used a guilt-by-association approach to classify their likely functions in diverse biological processes. Huarte, M. et al. A large intergenic noncoding RNA induced by p53 mediates global gene repression in the p53 response. Cell 142, 409–419 (2010). Hung, T. et al. Extensive and coordinated transcription of noncoding RNAs within cell-cycle promoters. Nature Genet. 43, 621–629 (2011). Wang, K. C. et al. A long noncoding RNA maintains active chromatin to coordinate homeotic gene expression. Nature 472, 120–124 (2011). Wilusz, J. E., Freier, S. M. & Spector, D. L. 3΄ end processing of a long nuclear-retained noncoding RNA yields a tRNA-like cytoplasmic RNA. Cell 135, 919–932 (2008). Mortazavi, A., Williams, B. A., McCue, K., Schaeffer, L. & Wold, B. Mapping and quantifying mammalian transcriptomes by RNA-Seq. Nature Methods 5, 621–628 (2008). Guttman, M. et al. Ab initio reconstruction of cell type-specific transcriptomes in mouse reveals the conserved multi-exonic structure of lincRNAs. Nature Biotechnol. 28, 503–510 (2010). Cabili, M. N. et al. Integrative annotation of human large intergenic noncoding RNAs reveals global properties and specific subclasses. Genes Dev. 25, 1915–1927 (2011). Mercer, T. R., Dinger, M. E., Sunkin, S. M., Mehler, M. F. & Mattick, J. S. Specific expression of long noncoding RNAs in the mouse brain. Proc. Natl Acad. Sci. USA 105, 716–721 (2008). De Santa, F. et al. A large fraction of extragenic RNA Pol II transcription sites overlap enhancers. PLoS Biol. 8, e1000384 (2010). Kim, T. K. et al. Widespread transcription at neuronal activity-regulated enhancers. Nature 465, 182–187 (2010). Ravasi, T. et al. Experimental validation of the regulated expression of large numbers of non-coding RNAs from the mouse genome. Genome Res. 16, 11–19 (2006). Ponjavic, J., Ponting, C. P. & Lunter, G. Functionality or transcriptional noise? Evidence for selection within long noncoding RNAs. Genome Res. 17, 556–565 (2007). Taft, R. J. et al. Tiny RNAs associated with transcription start sites in animals. Nature Genet. 41, 572–578 (2009). Seila, A. C. et al. Divergent transcription from active promoters. Science 322, 1849–1851 (2008). Kouzarides, T. Chromatin modifications and their function. Cell 128, 693–705 (2007). Barski, A. et al. High-resolution profiling of histone methylations in the human genome. Cell 129, 823–837 (2007). Visel, A. et al. ChIP-seq accurately predicts tissue-specific activity of enhancers. Nature 457, 854–858 (2009). Heintzman, N. D. et al. Histone modifications at human enhancers reflect global cell-type-specific gene expression. Nature 459, 108–112 (2009). Mikkelsen, T. S. et al. Genome-wide maps of chromatin state in pluripotent and lineage-committed cells. Nature 448, 553–560 (2007). Khalil, A. M. et al. Many human large intergenic noncoding RNAs associate with chromatin-modifying complexes and affect gene expression. Proc. Natl Acad. Sci. USA 106, 11667–11672 (2009). Ernst, J. et al. Mapping and analysis of chromatin state dynamics in nine human cell types. Nature 473, 43–49 (2011). Loewer, S. et al. Large intergenic non-coding RNA-RoR modulates reprogramming of human induced pluripotent stem cells. Nature Genet. 42, 1113–1117 (2010). Dinger, M. E., Pang, K. C., Mercer, T. R. & Mattick, J. S. Differentiating proteincoding and noncoding RNA: challenges and ambiguities. PLoS Comput. Biol. 4, e1000176 (2008). Brockdorff, N. et al. The product of the mouse Xist gene is a 15 kb inactive X-specific transcript containing no conserved ORF and located in the nucleus. Cell 71, 515–526 (1992). Lin, M. F., Deoras, A. N., Rasmussen, M. D. & Kellis, M. Performance and scalability of discriminative metrics for comparative gene identification in 1 6 F E B R UA RY 2 0 1 2 | VO L 4 8 2 | N AT U R E | 3 4 5
INSIGHT REVIEW 12 Drosophila genomes. PLoS Comput. Biol. 4, e1000067 (2008). 51. Lin, M. F., Jungreis, I. & Kellis, M. PhyloCSF: a comparative genomics method to distinguish protein coding and non-coding regions. Bioinformatics 27, i275–i282 (2011). 52. Finn, R. D. et al. The Pfam protein families database. Nucleic Acids Res. 38, D211–D222 (2010). 53. Ingolia, N. T., Lareau, L. F. & Weissman, J. S. Ribosome profiling of mouse embryonic stem cells reveals the complexity and dynamics of mammalian proteomes. Cell 147, 789–802(2011). 54. Galindo, M. I., Pueyo, J. I., Fouix, S., Bishop, S. A. & Couso, J. P. Peptides encoded by short ORFs control development and define a new eukaryotic gene family. PLoS Biol. 5, e106 (2007). This paper demonstrates the existence of functional small peptides within a presumed ‘non-coding’ transcript through ORF conservation, in vivo protein identification and functional analysis. 55. Kondo, T. et al. Small peptides switch the transcriptional activity of Shavenbaby during Drosophila embryogenesis. Science 329, 336–339 (2010). 56. Jiao, Y. & Meyerowitz, E. M. Cell-type specific analysis of translating RNAs in developing flowers reveals new levels of control. Mol. Syst. Biol. 6, 419 (2010). 57. Li, Y. M. et al. The H19 transcript is associated with polysomes and may regulate IGF2 expression in trans. J. Biol. Chem. 273, 28247–28252 (1998). 58. Cai, X. & Cullen, B. R. The imprinted H19 noncoding RNA is a primary microRNA precursor. RNA 13, 313–316 (2007). 59. Yang, L. et al. ncRNA- and Pc2 methylation-dependent gene relocation between nuclear structures mediates gene activation programs. Cell 147, 773–788 (2011). 60. Clamp, M. et al. Distinguishing protein-coding and noncoding genes in the human genome. Proc. Natl Acad. Sci. USA 104, 19428–19433 (2007). 61. Kastenmayer, J. P. et al. Functional genomics of genes with small open reading frames (sORFs) in S. cerevisiae. Genome Res. 16, 365–373 (2006). 62. Hanada, K., Zhang, X., Borevitz, J. O., Li, W. H. & Shiu, S. H. A large number of novel coding small open reading frames in the intergenic regions of the Arabidopsis thaliana genome are transcribed and/or under purifying selection. Genome Res 17, 632–640 (2007). 63. Mattick, J. S. The genetic signatures of noncoding RNAs. PLoS Genet. 5, e1000459 (2009). 64. Tsai, M. C. et al. Long noncoding RNA as modular scaffold of histone modification complexes. Science 329, 689–693 (2010). This paper identified multiple protein-interaction domains within HOTAIR that together allowed it to carry out its function, which demonstrated that a large ncRNA can act as a molecular scaffold. 65. Gupta, R. A. et al. Long non-coding RNA HOTAIR reprograms chromatin state to promote cancer metastasis. Nature 464, 1071–1076 (2010). 66. Zappulla, D. C. & Cech, T. R. Yeast telomerase RNA: a flexible scaffold for protein subunits. Proc. Natl Acad. Sci. USA 101, 10024–10029 (2004). This paper demonstrated that telomerase RNA can bridge proteins by showing that protein interaction domains can be swapped and spacer regions deleted with minimal impact on the function of the RNA. 67. Korostelev, A. & Noller, H. F. The ribosome in focus: new structures bring new insights. Trends Biochem. Sci. 32, 434–441 (2007). 68. Ivanova, N. et al. Dissecting self-renewal in stem cells with RNA interference. Nature 442, 533–538 (2006). 69. Martens, J. A., Laprade, L. & Winston, F. Intergenic transcription is required to repress the Saccharomyces cerevisiae SER3 gene. Nature 429, 571–574 (2004). 70. Schmitt, S., Prestel, M. & Paro, R. Intergenic transcription through a Polycomb group response element counteracts silencing. Genes Dev. 19, 697–708 (2005). 71. Lee, J. T. Lessons from X-chromosome inactivation: long ncRNA as guides and tethers to the epigenome. Genes Dev. 23, 1831–1842 (2009). 72. Ponjavic, J., Oliver, P. L., Lunter, G. & Ponting, C. P. Genomic and transcriptional co-localization of protein-coding and long non-coding RNA pairs in the developing brain. PLoS Genet. 5, e1000617 (2009). 73. Tian, D., Sun, S. & Lee, J. T. The long noncoding RNA, Jpx, is a molecular switch for X chromosome inactivation. Cell 143, 390–403 (2010). 74. Koerner, M. V., Pauler, F. M., Huang, R. & Barlow, D. P. The function of non-coding RNAs in genomic imprinting. Development 136, 1771–1783 (2009). 75. Pandey, R. R. et al. Kcnq1ot1 antisense noncoding RNA mediates lineagespecific transcriptional silencing through chromatin-level regulation. Mol. Cell 32, 232–246 (2008). 76. Bertani, S., Sauer, S., Bolotin, E. & Sauer, F. The noncoding RNA Mistral activates Hoxa6 and Hoxa7 expression and stem cell differentiation by recruiting MLL1 to chromatin. Mol. Cell 43, 1040–1046 (2011). 77. Feng, J. et al. The Evf-2 noncoding RNA is transcribed from the Dlx-5/6
3 4 6 | N AT U R E | VO L 4 8 2 | 1 6 F E B R UA RY 2 0 1 2
ultraconserved region and functions as a Dlx-2 transcriptional coactivator. Genes Dev. 20, 1470–1484 (2006). 78. Koziol, M. J. & Rinn, J. L. RNA traffic control of chromatin complexes. Curr. Opin. Genet. Dev. 20, 142–148 (2010). 79. Maison, C. et al. Higher-order structure in pericentric heterochromatin involves a distinct pattern of histone modification and an RNA component. Nature Genet. 30, 329–334 (2002). 80. Bernstein, E. et al. Mouse polycomb proteins bind differentially to methylated histone H3 and RNA and are enriched in facultative heterochromatin. Mol. Cell. Biol. 26, 2560–2569 (2006). 81. Wutz, A., Rasmussen, T. P. & Jaenisch, R. Chromosomal silencing and localization are mediated by different domains of Xist RNA. Nature Genet. 30, 167–174 (2002). This paper reported the generation of deletion mutants across the Xist locus and identified the discrete domains responsible for the silencing and localization roles of the RNA. 82. Chu, C., Qu, K., Zhong, F. L., Artandi, S. E. & Chang, H. Y. Genomic maps of long noncoding RNA occupancy reveal principles of RNA–chromatin interactions. Mol. Cell 44, 667–678 (2011). 83. Simon, M. D. et al. The genomic binding-sites of a non-coding RNA. Proc. Natl Acad. Sci. USA 108, 20497–20502 (2011). 84. Zhao, J., Sun, B. K., Erwin, J. A., Song, J. J. & Lee, J. T. Polycomb proteins targeted by a short repeat RNA to the mouse X chromosome. Science 322, 750–756 (2008). 85. Plath, K., Mlynarczyk-Evans, S., Nusinow, D. A. & Panning, B. Xist RNA and the mechanism of X chromosome inactivation. Annu. Rev. Genet. 36, 233–278 (2002). 86. Nagano, T. et al. The Air noncoding RNA epigenetically silences transcription by targeting G9a to chromatin. Science 322, 1717–1720 (2008). 87. Zhao, J. et al. Genome-wide identification of Polycomb-associated RNAs by RIP-seq. Mol. Cell 40, 939–953 (2010). 88. Kaneko, S. et al. Phosphorylation of the PRC2 component Ezh2 is cell cycleregulated and up-regulates its binding to ncRNA. Genes Dev. 24, 2615–2620 (2010). 89. Wang, X. et al. Induced ncRNAs allosterically modify RNA-binding proteins in cis to inhibit transcription. Nature 454, 126–130 (2008). 90. Kino, T., Hurt, D. E., Ichijo, T., Nader, N. & Chrousos, G. P. Noncoding RNA Gas5 is a growth arrest- and starvation-associated repressor of the glucocorticoid receptor. Sci. Signal 3, ra8 (2010). 91. Salmena, L., Poliseno, L., Tay, Y., Kats, L. & Pandolfi, P. P. A ceRNA hypothesis: the Rosetta stone of a hidden RNA language? Cell 146, 353–358 (2011). 92. Cesana, M. et al. A long noncoding RNA controls muscle differentiation by functioning as a competing endogenous RNA. Cell 147, 358–369 (2011). 93. Greider, C. W. & Blackburn, E. H. Identification of a specific telomere terminal transferase activity in Tetrahymena extracts. Cell 43, 405–413 (1985). 94. Feng, J. et al. The RNA component of human telomerase. Science 269, 1236–1241 (1995). 95. Lingner, J. et al. Reverse transcriptase motifs in the catalytic subunit of telomerase. Science 276, 561–567 (1997). 96. Jeon, Y. & Lee, J. T. YY1 tethers Xist RNA to the inactive X nucleation center. Cell 146, 119–133 (2011). 97. Hasegawa, Y., Brockdorff, N., Kawano, S., Tsutui, K. & Nakagawa, S. The matrix protein hnRNP U is required for chromosomal localization of Xist RNA. Dev. Cell 19, 469–476 (2010). 98. Schmitz, K. M., Mayer, C., Postepska, A. & Grummt, I. Interaction of noncoding RNA with the rDNA promoter mediates recruitment of DNMT3b and silencing of rRNA genes. Genes Dev. 24, 2264–2269 (2010). 99. Martianov, I., Ramadass, A., Serra Barros, A., Chow, N. & Akoulitchev, A. Repression of the human dihydrofolate reductase gene by a non-coding interfering transcript. Nature 445, 666–670 (2007). 100.Bartel, D. P. MicroRNAs: target recognition and regulatory functions. Cell 136, 215–233 (2009). Acknowledgements We thank M. Cabili, J. Engreitz, M. Garber, P. McDonel and A. Pauli for their reading and suggestions; T. Cech for comments and suggestions; E. Lander for helpful discussions and ideas; and S. Knemeyer and L. Gaffney for assistance with figures in this Review. Author Information Reprints and permissions information is available at www.nature.com/reprints. The authors declare no competing financial interests. Readers are welcome to comment on the online version of this article at www.nature.com/nature. Correspondence should be addressed to M.G. (firstname.lastname@example.org) and J.L.R. (email@example.com).
The microcosmos of cancer Amaia Lujambio1 & Scott W. Lowe1,2
The discovery of microRNAs (miRNAs) almost two decades ago established a new paradigm of gene regulation. During the past ten years these tiny non-coding RNAs have been linked to virtually all known physiological and pathological processes, including cancer. In the same way as certain key protein-coding genes, miRNAs can be deregulated in cancer, in which they can function as a group to mark differentiation states or individually as bona fide oncogenes or tumour suppressors. Importantly, miRNA biology can be harnessed experimentally to investigate cancer phenotypes or used therapeutically as a target for drugs or as the drug itself.
icroRNAs (miRNAs) are small, evolutionarily conserved, non-coding RNAs of 18–25 nucleotides in length that have an important function in gene regulation. Mature miRNA products are generated from a longer primary miRNA (pri-miRNA) transcript through sequential processing by the ribonucleases Drosha and Dicer1 (ref. 1). The first description of miRNAs was made in 1993 in Caenorhabditis elegans as regulators of developmental timing2,3. Later, miRNAs were shown to inhibit their target genes through sequences that are complementary to the target messenger RNA, leading to decreased expression of the target protein1 (Box 1). This discovery resulted in a pattern shift in our understanding of gene regulation because miRNAs are now known to repress thousands of target genes and coordinate normal processes, including cellular proliferation, differentiation and apoptosis. The aberrant expression or alteration of miRNAs also contributes to a range of human pathologies, including cancer. The control of gene expression by miRNAs is a process seen in virtually all cancer cells. These cells show alterations in their miRNA expression profiles, and emerging data indicate that these patterns could be useful in improving the classification of cancers and predicting their behaviour. In addition, miRNAs have now been shown to behave as cancer ‘drivers’ in the same way as protein-coding genes whose alterations actively and profoundly contribute to malignant transformation and cancer progression. Owing to the capacity of miRNAs to modulate tens to hundreds of target genes, they are emerging as important factors in the control of the ‘hallmarks’ of cancer4. In this Review, we summarize the findings that provide evidence for the central role of miRNAs in controlling cellular transformation and tumour progression. We also highlight the potential uses of miRNAs and miRNA-based drugs in cancer therapy and discuss the obstacles that will need to be overcome.
miRNAs are cancer genes In 2002, Croce and colleagues first demonstrated that an miRNA cluster was frequently deleted or downregulated in chronic lymphocytic leukaemia5. This discovery suggested that non-coding genes were contributing to the development of cancer, and paved the way for the closer investigation of miRNA loss or amplification in tumours. Subsequently, miRNAs were shown to be differentially expressed in cancer cells, in which they formed distinct and unique miRNA expression patterns6, and whole classes of miRNAs could be controlled directly by key oncogenic transcription factors7. In parallel, studies with mouse models established that miRNAs were actively involved in tumorigenesis8. Collectively, these findings provided the first key insights into the relevance of miRNA biology in human cancer.
Despite these results, the sheer extent of involvement of miRNAs in cancer was not anticipated. miRNA genes are usually located in small chromosomal alterations in tumours (in amplifications, deletions or linked to regions of loss of heterozygosity) or in common chromosomalbreakpoints that are associated with the development of cancer9. In addition to structural genetic alterations, miRNAs can also be silenced by promoter DNA methylation and loss of histone acetylation10. Interestingly, somatic translocations in miRNA target sites can also occur, representing a drastic means of altering miRNA function11,12. The frequent deregulation of individual or clusters of miRNAs at multiple levels mirrors the deregulation for protein-coding oncogenes or tumour suppressors (Table 1). In principle, somatic mutations that change an miRNA seed sequence could lead to the aberrant repression of tumour-suppressive mRNAs, but these seem to be infrequent13. Further sequencing could change this view, but this observation suggests that the intensity of miRNA signalling (altered by miRNA overexpression or underexpression) is more crucial than the specificity of the response. However, recent data indicate that miRNAs with an altered sequence can be produced through variable cleavage sites for Drosha and Dicer1, and that the presence of these variants can be perturbed in cancer14. Although the function of the variant ‘isomiRs’ remains unclear, in principle they could alter the quality of miRNA effects. State-of-the-art sequencing techniques will help to unmask mutations or modifications that otherwise would remain undetected. Whatever the mechanism, the widespread alteration in the expression of miRNAs is a ubiquitous feature of cancer.
miRNAs as cancer classifiers Aberrant miRNA levels reflect the physiological state of cancer cells and can be detected by miRNA expression profiling and harnessed for the purpose of diagnosis and prognosis15,16. In fact, miRNA profiling can be more accurate at classifying tumours than mRNA profiling because miRNA expression correlates closely with tumour origin and stage, and can be used to classify poorly differentiated tumours that are difficult to identify using a standard histological approach6,17. Whether or not this increased classification power relates to the biology of miRNAs or the reduced complexity of the miRNA genome still needs to be determined. The special features of miRNAs make them potentially useful for detection in clinical specimens. For example, miRNAs are relatively resistant to ribonuclease degradation, and they can be easily extracted from small biopsies, frozen samples and even formalin-fixed, paraffinembedded tissues18 . Furthermore, relatively simple and reproducible assays have been developed to detect the abundance of individual
Cancer Biology and Genetics Program, Memorial Sloan-Kettering Cancer Center (MSKCC), 1275 York Avenue, New York, New York 10065, USA. 2Howard Hughes Medical Institute, MSKCC, 1275 York Avenue, New York, New York 10065, USA. 1 6 F E B R UA RY 2 0 1 2 | VO L 4 8 2 | N AT U R E | 3 4 7
INSIGHT REVIEW BOX 1
Biogenesis and function of miRNAs miRNAs are subjected to a unique biogenesis that is closely related to their regulatory functions. As the pathway in Fig. 1 shows, in general miRNAs are transcribed by RNA polymerase II into primary transcripts called pri-miRNAs76. The primary transcripts contain a 5ʹ cap structure a poly(A)+ tail and may include introns, similar to the transcripts of protein-coding genes76. They also contain a region in which the sequences are not perfectly complementary, known as the stem–loop structure, which is recognized in the nucleus by the ribonuclease Drosha and its partner DGCR8, giving rise to the precursor miRNA (premiRNA) by cropping76. However, some intronic miRNAs (called mirtrons) bypass the Drosha processing step and, instead, use splicing machinery to generate the pre-miRNA99. The premiRNA is exported from the nucleus to the cytoplasm by XPO5 and is further cleaved by the ribonuclease Dicer1 (along with TARBP2) into a double-stranded miRNA (process known as dicing)76. Again, this cleavage can be substituted by Argonaute2-mediated processing100. After strand separation, the guide strand or mature miRNA forms, in combination with Argonaute proteins, the RNAinduced silencing complex (RISC), whereas the passenger strand is usually degraded. The mature strand is important for specific-target mRNA recognition and its consequent incorporation into the RISC1. The specificity of miRNA targeting is defined by how complementary the ‘seed’ sequence (positions 2 to 8 from the 5ʹ end of the miRNA) and the ‘seedmatch’ sequence (generally in the 3ʹ untranslated region of the target mRNA) are. The expression of the target mRNAs is silenced by miRNAs, either by mRNA cleavage (‘slicing’) or by translational repression1. In addition, miRNAs have a number of unexpected functions, including the targeting of DNA, ribonucleoproteins or increasing the expression of a target mRNA93. Overall, data indicate the complexity of miRNAmediated gene regulation and highlight the importance of a better understanding of miRNA biology.
miRNAs, and methods that combine small RNA isolation, PCR and next-generation sequencing, allow accurate and quantitative assessment of all the miRNAs that are expressed in a patient specimen, including material that has been isolated by laser capture microdissection. The detection of global miRNA expression patterns for the diagnosis of cancers has not yet been proved; however, some individual or small groups of miRNAs have shown promise. For example, in non-small cell lung cancer, the combination of high miR-155 and low let-7 expression correlates with a poor prognosis, and in chronic lymphocytic leukaemia a 13 miRNA signature is associated with disease progression15,16. Further advances in the technology of miRNA profiling could help to revolutionize molecular pathology. Perhaps the most appealing application of miRNAs as a cancer diagnostic tool comes from the discovery of circulating miRNAs in serum. For example, miR-141 expression levels in serum were significantly higher in patients with prostate cancer than in healthy control individuals19. Although the analysis of circulating miRNAs is only just beginning, the successful advancement of this technology could provide a relatively non-invasive diagnostic tool for single-point or longitudinal studies. With such diagnostic tools in place, miRNA profiling could be used to guide cancer classification, facilitate treatment decisions, monitor treatment efficacy and predict clinical outcome. 3 4 8 | N AT U R E | VO L 4 8 2 | 1 6 F E B R UA RY 2 0 1 2
When miRNA biogenesis goes awry Although the expression of some miRNAs is increased in malignant cells, the widespread underexpression of miRNAs is a more common phenomenon. Whether this tendency is a reflection of a pattern associated with specific cells of origin, is a consequence of the malignant state or actively contributes to cancer development is still unclear. Because miRNA expression generally increases as cells differentiate, the apparent underexpression of miRNAs in cancer cells may, in part, be a result of miRNAs being ‘locked’ in a less-differentiated state. Alternatively, changes in oncogenic transcription factors that repress miRNAs or variability in the expression or activity of the miRNA processing machinery could also be important. Two main mechanisms have been proposed as the underlying cause of the global downregulation of miRNAs in cancer cells. One involves transcriptional repression by oncogenic transcription factors. For example, the MYC oncoprotein, which is overexpressed in many cancers, transcriptionally represses certain miRNAs, although the extent to which this mediates its oncogenic activity or reflects a peripheral effect is still unknown20. The other mechanism proposed involves changes in miRNA biogenesis and is based on the observation that cancer cells often display reduced levels, or altered activity, of factors in the miRNA biogenesis pathway21 (Box 1, Fig. 1). In vivo studies have provided the most direct evidence of an active role for miRNA downregulation in at least some types of cancer. For example, analysis of mouse models in which the core enzymes of miRNA biogenesis have been constitutively or conditionally disrupted by different mechanisms suggests that these molecules function as haploinsufficient tumour suppressors. Thus, the repression of miRNA processing by the partial depletion of Dicer1 and Drosha accelerates cellular transformation and tumorigenesis in vivo22. Furthermore, deletion of a single Dicer1 allele in lung epithelia promotes Kras-driven lung adenocarcinomas, whereas complete ablation of Dicer1 causes lethality because of the need for miRNAs in essential processes23. Consistent with the potential relevance of these mechanisms, reduced Dicer1 and Drosha levels have been associated with poor prognosis in the clinic24. In addition to the core machinery, modulators of miRNA processing can also function as haploinsufficient tumour suppressors. Hence, point mutations that affect TARBP2 or XPO5 are correlated with sporadic and hereditary carcinomas that have microsatellite instability25,26. Other miRNA modulators that influence the processing of only a subset of miRNAs could also be important. For example, LIN28A and LIN28B can bind and repress members of the let-7 family (which are established tumour-suppressor miRNAs; Table 1), but this binding can be counteracted by KHSRP (KH-type splicing regulatory protein), also a factor involved in miRNA biogenesis; together this binding and counteracting dictate the level of mature let-7. The processing of miRNAs can be regulated by other genes including DDX5 (helicase p68) or the SMAD 1 and SMAD 5 proteins, which may contribute to cancer development through the deregulation of miRNAs27. Collectively, the global changes in miRNA expression that are seen in cancer cells probably arise through multiple mechanisms; the combined small changes in the expression of many miRNAs seem to have a large impact on the malignant state.
miRNAs as cancer drivers Functional studies show that miRNAs that are affected by somatic alterations in tumours can affect cancer phenotypes directly, therefore confirming their driver function in malignancy. As drivers of malignancy, mechanistic studies show that these miRNAs interact with known cancer networks; hence, tumour-suppressor miRNAs can negatively regulate protein-coding oncogenes, whereas oncogenic miRNAs often repress known tumour suppressors (Fig. 2a). Perhaps the best example of this is the oncogenic miR-17-92 cluster, in which individual miRNAs suppress negative regulators of phosphatidylinositol-3-OH kinase signalling or pro-apoptotic members of the BCL-2 family, which disrupts the processes that are known to influence cancer development28 (Table 1). Cancer-associated miRNAs can also alter the epigenetic landscape
REVIEW INSIGHT Table 1 | Key microRNAs involved in cancer MicroRNA
Amplification and transcriptional activation
BIM, PTEN, CDKN1A and PRKAA1
Lymphoma, lung, breast, stomach, colon and pancreatic cancer
Inhibition and Cooperates with MYC detection to produce lymphoma. Overexpression induces lymphoproliferative disease
SHIP1 and CEBPB
Chronic lymphocytic leukaemia, lymphoma, lung, breast and colon cancer
Overexpression induces pre-B-cell lymphoma and leukaemia
Inhibition and detection
Chronic lymphocytic PTEN, PDCD4 and leukaemia, acute myeloid leukaemia, glioblastoma, TPM1 pancreatic, breast, lung, prostate, colon and stomach cancer
Overexpression induces lymphoma
Inhibition and detection
Deletion, mutation and transcriptional repression
BCL2 and MCL1
Chronic lymphocytic leukaemia, prostate cancer and pituitary adenomas
Deletion causes chronic lymphocytic leukaemia
Expression with mimics and viral vectors
11 copies (multiple locations)
KRAS, MYC and HMGA2
Lung, colon, stomach, ovarian and breast cancer
Overexpression suppresses lung cancer
Expression with mimics and viral vectors
1p36 and 11q23
Epigenetic silencing, transcriptional repression and deletion
CDK4, CDK6, MYC and MET
Colon, lung, breast, kidney, bladder cancer, neuroblastoma and melanoma
No published studies
Expression with mimics and viral vectors
7q32 and 1q30
Breast cancer and indolent chronic lymphocytic leukaemia
Overexpression induces chronic lymphocytic leukaemia
No published studies
Deletion and transcriptional repression
Acute myeloid leukaemia, aggressive chronic lymphocytic leukaemia and lung cancer
No published studies
BCL2, B-cell lymphoma protein-2; BIM, BCL-2-interactiing mediator of cell death; CDKN1A, cyclin-dependent kinase inhibitor 1A; CEBPB, CCAAT/enhancer binding protein β; HMGA2, high mobility group AT-hook 2; CDK4, cyclin-dependent kinase 4; CDK6, cyclin-dependent kinase 6; DNMT, DNA methyltransferase; MCL1, myeloid cell leukaemia sequence 1; PTEN, phosphatase and tensin homologue; PRKAA1, protein kinase, AMP-activated, alpha 1 catalytic subunit; PDCD4, programmed cell death 4; SHIP1, Src homology 2 domain-containing inositol 5-phosphatase 1; TPM1, tropomyosin 1; ZFP36, zinc finger protein 36.
of cancer cells. The cancer ‘epigenome’ is characterized by global and gene-specific changes in DNA methylation, histone modification patterns and chromatin-modifying enzyme expression profiles, which impact gene expression in a heritable way29. In one way, miRNA expression can be altered by DNA methylation or histone modifications in cancer cells10,30, but miRNAs can also regulate components of the epigenetic machinery, therefore indirectly contributing to the reprogramming of cancer cells. For example, miR-29 inhibits DNMT3A and DNMT3B expression in lung cancer31, whereas miR-101 regulates the histone methyltransferase EZH2 in prostate cancer32. The presence of mature miRNAs in the nucleus33 is another indication of the potentially direct role that miRNAs have in controlling epigenetic modifications, such as DNA methylation and histone modifications — a hypothesis that has been established in plants34 but still needs to be demonstrated with certainty in mammals. In the same way as protein-coding genes, miRNAs can be oncogenes or tumour suppressors depending on the cellular context in which they are expressed, which means that defining their precise contribution to cancer can be a challenge (Fig. 2b). The fact that miRNAs show tissue-specific expression and their output, shown in the cell’s physiology, is dependent on the expression pattern of the specific mRNAs that harbour target sites could explain this apparent paradox. For example, the miR-29 family has a tumour-suppressive effect in lung tumours but appears oncogenic in breast cancer because of its ability to target the DNA methyltransferases DNMT3A and DNMT3B, and ZFP36, respectively31,35 (Table 1). To further complicate the process, some miRNAs repress several positive components of a pathway, whereas others target both positive and negative regulators, possibly to buffer against minor physiological variations that could trigger much larger changes in the cell physiology36. In cancer cells, this buffering role can mean that some miRNAs could
simultaneously target oncogenes and tumour-suppressor genes. In addition, combinations of miRNAs can cooperate to regulate one or several pathways, which increases the flexibility of regulation but confounds experimentalists37 (Fig. 2c). Consequently, the way in which miRNAs contribute to cancer development is conceptually similar to cancer-associated transcription factors such as MYC and p53, which are mediated through many targets that depend on contextual factors that are influenced by cell type and micro-environment. From a practical perspective it is crucial that miRNA targets are studied in a context that is appropriate to the environment that is being studied to determine what impact they will have on tumour cell behaviour (Fig. 2b).
Oncogenic pathways Beyond the impact of somatic genetic and epigenetic lesions, the altered expression of miRNAs in cancer can arise through the aberrant activity of transcription factors that control their expression. Interestingly, the same transcription factors are often targets of miRNA-mediated repression, which gives rise to complex regulatory circuits and feedback mechanisms. Thus, a single transcription factor can activate or repress several miRNAs and protein-coding genes; in turn, the alteration in miRNA expression can affect more protein-coding genes that then amplifies the effects of a single gene. As already mentioned, MYC directly contributes to the global transcriptional silencing of miRNAs20. This repression involves the downregulation of miRNAs with antiproliferative, antitumorigenic and pro-apoptotic activity such as, let-7, miR-15a/16-1, miR-26a or miR-34 family members38 (Fig. 2d; Table 1). Initial studies indicate that Myc uses both transcriptional and post-transcriptional mechanisms to modulate miRNA expression. This phenomenon could be due to LIN28A and LIN28B being the direct target of MYC, and that they are required for 1 6 F E B R UA RY 2 0 1 2 | VO L 4 8 2 | N AT U R E | 3 4 9
INSIGHT REVIEW a â†‘
and function of oncogenic miRNAs
TP53 is a master regulator of miRNAs
Cytoplasm - $+#regulation -ARBP2 mutation
Unwinding of miRNA duplex
Accumulation of oncogenic miRNAs
Global loss of miRNAs Loss of tumoursuppressor miRNAs
RISC mRNA cleavage
example, in pancreatic cancer with mutant KRAS, RAS-responsive element-binding protein 1 (RREB1) represses miR-143 and miR-145 promoter, and at the same time both KRAS and RREB1 are targets of miR-143 and miR-145, revealing a feedforward mechanism that increases the effect of RAS signalling41. Similarly, KRAS is a target for several miRNAs, of which the let-7 family is the most representative example42. The integration of miRNAs into key oncogenic pathways, and the generation of feedforward and feedback loops that have a balancing effect, creates intricate ways to incorporate intracellular and extracellular signals in the decisions of cell proliferation or survival, and further implicates miRNAs in the pathogenesis of cancer.
Accumulation of oncogenes
Loss of tumour-suppressor genes
Figure 1 | Mechanisms of miRNA perturbation in cancer. Cancer cells present global downregulation of miRNAs, loss of tumour-suppressor miRNAs and specific accumulation of oncogenic miRNAs. The alteration in miRNA expression patterns leads to the accumulation of oncogenes and downregulation of tumour-suppressor genes, which leads to the promotion of cancer development. a, The expression and function of oncogenic miRNAs is increased by genomic amplification, activating mutations, loss of epigenetic silencing and transcriptional activation. By contrast, tumour-suppressor miRNAs are lost by genomic deletion, inactivating mutations, epigenetic silencing or transcriptional repression. b, After transcription, global levels of miRNAs can be reduced by impaired miRNA biogenesis. Inactivating mutations and reduced expression have been described for almost all the members of the miRNA processing machinery. If there is a downreguation of DROSHA this can lead to a decrease in the cropping of primary miRNA (primiRNA) to precursor miRNA (pre-miRNA). In the case of XPO5 mutation, pre-miRNAs are prevented from being exported to the cytoplasm. Mutation of TARBP2 or downregulation of DICER1 results in a decrease in mature miRNA levels. Pol II, RNA polymerase II; RISC, RNA-induced silencing complex.
MYC-mediated repression of let-7 (ref. 38). Furthermore, MYC directly activates the transcription of miR-17-92 polycistronic cluster and, given its oncogenic role, it may contribute to MYC-induced tumorigenesis39. MYC-driven reprogramming of miRNA expression could also be a factor in hepatocellular carcinoma, because of the contribution the reprogramming has to the aggressive phenotype of tumours originating from hepatic progenitor cells40. Some miRNAs, such as let-7, also regulate MYC, closing the regulatory circuit37. miRNAs are embedded in many other oncogenic networks, including KRAS activation, which leads to the repression of several miRNAs. For 3 5 0 | N AT U R E | VO L 4 8 2 | 1 6 F E B R UA RY 2 0 1 2
The TP53 tumour suppressor is perhaps the most important and well-studied cancer gene, and it is not surprising that several studies have suggested that miRNA biology can have a role in its regulation and activity (Fig. 2e). The p53 protein acts as a sequence-specific DNAbinding factor that can activate and repress transcription. Although there is no doubt that most of the actions of p53 can be explained by its ability to control canonical protein-coding targets such as CDKN1A and PUMA, it can also transactivate several miRNAs. One of the beststudied classes is the miR-34 family (Table 1), which represses genes that can promote proliferation and apoptosis â€” plausible targets in a p53-mediated tumour-suppressor response43. In principle, the action of p53 to induce the expression of miR-34 and other miRNAs can explain some of its transcriptional repressive functions. The discovery of additional p53-regulated miRNAs, and the targeting of p53 or its pathway by other miRNAs, has provided general insights into the miRNA-mediated control of gene expression and the potential therapeutic opportunities for targeting the p53 network (Fig. 2e). Several p53-activated miRNAs, such as miR-192, miR-194, miR-215 and miR-605, can target MDM2, which is a negative regulator of p53 and a therapeutic target. These potentially relevant miRNAs can be epigenetically silenced in some types of cancer; however, their reactivation or reintroduction (see the section miRNAs as drugs and drug targets) offers an intriguing therapeutic opportunity for inhibiting MDM2 in tumours that harbour wild-type p53 (refs 44, 45). Similarly, p53 can also activate miR-107, miR-200 or miR-192, which are miRNAs that inhibit angiogenesis and epithelial-to-mesenchymal transition46â€“48. Conversely, p53 can be repressed by certain oncogenic miRNAs including miR-380-5p, which is upregulated in neuroblastomas with MYCN amplification, or miR-504, which decreases p53-mediated apoptosis and cell-cycle arrest and can promote tumorigenesis49,50. However, the extent to which these miRNAs control life and death decisions in the p53 network still needs to be shown decisively to determine whether these miRNAs are valid therapeutic targets. The studies mentioned have extended our understanding of the roles and regulation of p53 into the world of small non-coding RNAs, but the action on miRNA biology may be even more complex. For example, one study51 suggests that p53 can affect miRNA biogenesis by promoting pri-miRNA processing through association with the large Drosha complex (Fig. 2e), but the precise mechanism remains unclear51. In a more conventional way, the p53 family member p63 transcriptionally controls Dicer1 expression. Mutant TP53 can interfere with this regulation, which leads to a reduction in Dicer1 levels and reduces the levels of certain cancer-relevant miRNAs52. Thus, with the p53 network as a typical example, it is clear that miRNAs can interact with cancerrelevant pathways at multiple and unexpected levels and that a better understanding of miRNA biology will help to decipher the role and function of other important cancer genes.
Micromanagement of metastasis and beyond In addition to promoting cancer initiation, miRNAs can modulate processes that support cancer progression, including metastasis53â€“56. As indicated earlier, changes in miRNA levels can occur through effects on their transcription or by global changes in the RNA interference (RNAi)
REVIEW INSIGHT machinery, and both mechanisms seem to be important for this process. For example, in breast cancer, miR-10b and miR-9 can induce metastasis, whereas miR-126, miR-335 and miR-31 act as suppressors. The miR-200 family inhibits epithelial-to-mesenchymal transition, which influences one aspect of the metastatic process57. However, miR-200 could also promote the colonization of metastatic cells in breast cancer, which provides yet another example of the opposing activities of some miRNAs58. Conversely, in head and neck squamous-cell carcinomas, lung adenocarcinomas and breast cancers, the reduced levels of certain miRNAs that arise from Dicer1 downregulation also promote cell motility and are associated with enhanced metastasis in experimental models52,59. The pleiotropic effects of miRNA biology on cancer extend to virtually all acquired cancer traits, including cancer-associated changes in intracellular metabolism and the tissue microenvironment. For example, most cancer cells display alterations in glucose metabolism termed the Warburg effect60. miRNAs may contribute to this metabolic switch because, in glioma cells, miR-451 controls cell proliferation, migration and responsiveness to glucose deprivation, thereby allowing the cells to survive metabolic stress61. The enhanced glutaminolysis observed in cancer cells can be partially explained by MYC-mediated repression a Tumour suppressor miRNAs
of miR-23a and miR-23b (ref. 62) (Fig. 2d). In some cases, the control of these cancer-related processes by miRNAs creates an opportunity for new therapeutic approaches. Hence, miR-132, which is present in the endothelium of tumours but not in normal human endothelium, induces neovascularization by inhibition of p120RasGAP, a negative regulator of KRAS63. The delivery of a miR-132 inhibitor with nanoparticles that target the tumour vasculature suppresses angiogenesis in mice; this indicates there is a potential for the development of new antiangiogenic drugs. Further studies are likely to implicate miRNAs in the modulation of every tumour-associated pathway or trait.
Big lessons from mice Much of what we have learnt concerning the functional contribution of miRNA biology to cancer development comes from studies in genetically engineered mice. These systems provide powerful tools for the genetic and biological study of miRNAs in an in vivo context, which is particularly important given the contextual activity of most miRNAs. In addition, owing to the ability of these models to recapitulate the behaviour of some human malignancies, they are useful in preclinical studies to evaluate new therapeutics.
miR-15a/16-1 miR-34 family
Gene 2 Cancer
Gene 1 miRNA
Gene 4 Gene 5
miR-192 miR-194 miR-215 miR-605 Gene 1
Processing of tumoursuppressor miRNAs
Gene 2 Cancer miRNA
Figure 2 | Contribution of miRNAs to cancer pathways. a, Tumoursuppressor miRNAs, which repress oncogenes in healthy cells, are lost in cancer cells, leading to oncogene upregulation, whereas oncogenic miRNAs inhibit tumour-suppressor genes, giving rise to cancer. b, The presence of different target genes in different cell lines can modify the function of an miRNA, both in healthy cells and cancer cells, which can lead to the development of cancer or a different outcome. c, Two miRNAs can function together to regulate one or several pathways, which reinforces those pathways and can result in the development of cancer. d, The oncogene MYC can either
Epithelial to mesenchymal transition
repress tumour-suppressor miRNAs (in blue) or activate oncogenic miRNAs (in red) and can therefore orchestrate several different pathways. MYC can repress let-7, directly, or indirectly, through LIN28 activation. Conversely, let-7 can also repress MYC, which closes the regulatory circle. e, Tumour suppressor p53 can regulate several tumour suppressor miRNAs (blue), activating different antitumoral pathways. The regulation of MDM2 by some of these miRNAs leads to interesting feedforward loops. At the same time, p53 can be negatively regulated by oncogenic miRNAs (in red). In addition, p53 is involved in the biogenesis of several tumour suppressor miRNAs. 1 6 F E B R UA RY 2 0 1 2 | VO L 4 8 2 | N AT U R E | 3 5 1
INSIGHT REVIEW Perhaps the most widespread use of mice for characterizing miRNA biology in cancer is the validation of miRNAs that are altered in cancer cells, as bona fide oncogenes and tumour suppressor genes. As already mentioned, the first direct evidence that miRNAs have a function in cancer came from mouse models, in which it was shown that expression of the miR-17-92 cluster — which is amplified in some human B cell lymphomas — cooperates with Myc to promote B-cell lymphoma in mice8. Subsequent studies that have used genetically engineered or transplantation-based systems identified the relevant miRNA components, showing that the miR-19 family (including miR-19a and miR-19b) represents the most potent oncogenes in this cluster28,64,65. Another example is miR-155 overexpression in the lymphoid compartment, which triggers B-cell leukaemia or a myeloproliferative disorder depending on the system used to drive expression of the transgene; this was the first example of an miRNA that initiates cancer in a transgenic setting66,67 (Table 1). Gene targeting has been used extensively to delete miRNAs for the purpose of characterizing their physiological roles or action as candidate tumour suppressors. Gene targeting has suggested that miRNAs from similar families have redundant or compensatory functions, which has been shown for C. elegans68. Ablation of the miR-15a and miR-16-1 cluster, which is often deleted in human chronic lymphocytic leukaemia, predisposes mice to B-cell lymphoproliferative disease69 (Table 1). Importantly, the ability to produce mouse strains with different gene dosage through heterozygous or homozygous gene deletions has revealed that Dicer1, which if lost completely has a deleterious effect, can promote malignant phenotypes as a haploinsufficient tumour suppressor23. Such a conclusion could not be formed from studies that examined only genomic data. Conditional gene expression systems in mice have allowed researchers a Inducible in vivo miRNA expression Tet-OFF system
b Inducible in vivo miRNA inhibition Tet-OFF system TSP
Tet-ON system TSP
Figure 3 | In vivo miRNA expression or inhibition ‘á la carte’. a, Tetracycline (Tet)-mediated miRNA inactivation or activation by doxycycline administration using Tet-OFF, in which a tissue-specific promoter (TSP) is combined with a transactivator (tTA) to turn on expression of oncogenic miRNA (purple) and induce tumorigenesis (purple star) and subsequent tumour regression, revealing dependence on the oncogenic miRNA, or Tet-ON systems in which a reverse transactivator (rtTA) switches on oncogenic miRNA when the drug is applied. Drug withdrawal leads to tumour regression. b, Tet-mediated miRNA activation or inactivation by doxycycline administration using Tet-OFF or Tet-ON systems. miRNAs (green) can be inhibited by miRNA sponges (dark blue), with the same effects as miRNA expression, leading to tumorigenesis and subsequent tumour regression, which indicates a dependence on tumour-suppressor loss. 3 5 2 | N AT U R E | VO L 4 8 2 | 1 6 F E B R UA RY 2 0 1 2
to determine cancer gene dependencies, as well as whether genes that initiate cancer also participate in tumour maintenance. In many cases, withdrawal of the initiating oncogenic transgene (or restoration of the deleted or lost tumour suppressor) leads to the collapse of the tumour; this validates the transgene or pathway that is controlled by these genes, as a therapeutic target. Similar studies have also been applied to miRNAs; for example, conditional expression of miR-21, which is broadly deregulated in cancer, can promote lymphomagenesis in mice70 (Table 1). Silencing of miR-21 leads to disease regression, in part, by promoting apoptosis70 (Fig. 3a). Likewise, the use of miRNA inhibitors (for example, antagomirs) directed against miR-21 can inhibit the proliferation of human cancer cells that overexpress miR-21 (ref. 71). Together, these studies suggest that miR-21 antagonists have the potential to be effective therapies for at least some cancers. The development of new technology has meant that mouse models are increasingly used to study gene function on a large if not genome-wide scale, and miRNAs are at the forefront of this revolution. Recently, a vast collection of mouse embryonic stem-cell clones that harbour deletions that target 392 miRNA genes was generated 72. This unique and valuable toolbox, termed ‘mirKO’, will allow the creation of mice that lack specific miRNAs, express mutant miRNAs or the study of their expression. In a converse strategy, a collection of embryonic stem cells engineered to inducibly express the vast majority of known miRNAs is in production (S.W.L., Y. Park and G. Hannon, manuscript in preparation) and will allow the in vivo validation of miRNAs as oncogenes or as anticancer therapies. With a different strategy, miRNA sponges (Fig. 3b), which are oligonucleotide constructs with multiple complementary miRNA binding sites in tandem, have already been used to deplete individual miRNAs in transgenic fruitflies, in transplanted breast cancer cells in mice and in a transgenic mouse model56,73,74. Although these sponges provide a scalable strategy for miRNA loss-of-function studies, more work is needed to rule out off-target effects and assess their potency before conclusions can be made. However, the availability of such resources will help with the functional study of miRNAs in normal development and disease, and will be useful to the wider scientific community. Finally, genetically engineered mouse models of human cancers are a testing ground for preclinical studies. For example, in Myc-induced liver tumours, miR-26 delivery by adeno-associated viruses suppresses tumorigenesis by inducing apoptosis75. The increasing use of state-ofthe-art mouse models is likely to uncover new in vivo functions, such as metastasis and angiogenesis, that otherwise would have remained hidden in vitro. They will also provide key preclinical systems for testing miRNA-based therapeutics.
Constructing and deconstructing cancer The use of RNAi technology — a tool that exploits miRNA pathways — has revolutionized the study of gene function in mammalian systems and has provided a powerful means to investigate the function of any protein-coding gene. Experimental triggers of RNAi exploit different aspects of the pathway and result in the downregulation of gene expression through incorporation into the miRNA biogenesis machinery at different points76. Small-interfering RNAs (siRNAs), which function at the level of Dicer1, can transiently and potently lead to gene suppression; these RNAi triggers, or their variants, are probably the structural ‘scaffold’ for miRNA therapeutics (see the section miRNAs as drugs or drug targets). Stable RNAi can be activated by the expression of miRNA mimetics, that are either the so-called stem loop short-hairpin RNAs (shRNAs) or shRNAs that incorporate a larger miRNA fold. One example of the latter is based on miR-30 (known as miR-30-based shRNAs or ‘shRNAmirs’). These shRNAs, as occurs naturally for many miRNAs, can be embedded in non-coding sequences of protein-coding transcripts or linked in tandem, which allows, for example, the linkage of the shRNA with a fluorescent reporter or the simultaneous knockdown of two different
REVIEW INSIGHT genes77,78. Advances in the shRNAmir methodology have allowed the development of versatile vectors for the study of proliferation and survival genes, strategies for optimizing the potency of shRNAs, and rapid and effective systems for conditional shRNA expression in mice79– 81 . The last of these, together with systems based on short stem-loop shRNAs82, could eventually allow the spatial, temporal and reversible control of any gene in vivo. Regardless of the platform, RNAi technology provides an effective tool to investigate cancer phenotypes and identify therapeutic targets. For example, RNAi has been used to identify and characterize tumoursuppressor genes, which if inhibited promote cancer development. Early studies, using the same system that validated miR-17-92 as an oncogene, demonstrated that inhibition of TP53 could produce phenotypes that were consistent with TP53 loss83. Later studies showed that tumour suppressors could be identified prospectively using in vitro and in vivo shRNA screens, (for examples see refs 84 and 85). By conditionally expressing shRNAs that target tumour suppressors in mice, tumour-suppressor function in advanced tumours can be re-established by silencing the shRNA86. Tumour-suppressor reactivation leads to a marked (if not complete) tumour regression, which validates these pathways as therapeutic targets. RNAi technology can be exploited more directly to identify genotypespecific cancer drug targets. Although there may be differences in the outcome of RNAi and small-molecule-mediated protein inhibition, siRNAs and shRNAs have been widely used to determine whether a candidate target is required for the proliferation of cancer cells. Moreover, the availability of RNAi libraries that target portions of, or all, the human genome allows genetic screens to identify ‘synthetic lethal’ genes, for which, if combined, the attenuation triggers the death of the cell. In principle, the identification of an RNAi target, the inhibition of which is selectively lethal to cells harbouring a particular oncogenic alteration, should identify cancer-specific targets. Such approaches have identified potential targets for KRAS-expressing tumours87–89 and leukaemias with deregulated MYC (ref. 90). Application of these approaches could potentially be complementary to the traditional drug-target discovery approach, and possibly a systematic way to identify the combination of therapies that will ultimately be needed to combat cancer.
miRNAs as drugs and drug targets Despite advances in techniques to inhibit protein-coding genes using small molecules or biologicals, many cancers are unresponsive to the agents currently in use or become resistant to them; new and more creative approaches are therefore required for the treatment of cancer. Perhaps one of the most exciting opportunities that has arisen from our understanding of miRNA biology is the potential use of miRNA mimics or antagonists as therapeutics. Owing to the ability of miRNAs to simultaneously target multiple genes and pathways that are involved in cellular proliferation and survival38, the targeting of a single miRNA can be a form of ‘combination’ therapy that could obstruct feedback and compensatory mechanisms that would otherwise limit the effectiveness of many therapies in current use. In addition, because miRNA expression is often altered in cancer cells, agents that modulate miRNA activity could potentially produce cancer-specific effects10,91,92 . Based on this, anticancer therapies that inhibit or enhance miRNA activity are being developed (Fig. 4). Evidence for this is shown by the inhibition of oncogenic miRNAs or the expression of tumour suppressor miRNAs in mice that harbour tumours, which have a significant effect on the outcome of cancer. Oncogenic miRNAs can be blocked by using antisense oligonucleotides, antagomirs, sponges or locked nucleic acid (LNA) constructs93. The use of LNAs has achieved unexpected success in vivo, not only in mice but also for the treatment of hepatitis C in non-human primates94. The downregulation of miR-122 can lead to a significant inhibition of replication of the hepatitis C virus. This inhibition is thought to decrease the risk of chronic hepatitis and hepatocellular carcinoma in patients who are hepatitis C-positive. Early clinical studies using SPC3649, an miR-122 antagonist, in
miRNA ↑ miR-21 profiling ↓ miR-26
Tumour biopsy Patient serum
d Chemotherapy miR-21 inhibition miR-26 expression
Early detection Accurate diagnosis and prognosis Therapeutic strategy
Figure 4 | Proposed scheme for the treatment of liver cancer with combined chemotherapy and miRNA-based therapy. a, miRNA expression profiles of potential patients could be assessed by measuring circulating miRNAs in patient serum or tumoral miRNAs from a biopsy. For example, miR-21 expression and miR-26 loss could be detected in serum and tumour samples. b, This profile could be used for early detection of cancer, accurate diagnosis and prognosis, and choosing the best therapeutic strategy. The best available chemotherapeutic option could be combined with miRNA-based therapy. c, The oncomiRs detected in miRNA profiling and those present in the tumour, such as miR-21, could be inhibited by using different strategies, such as locked nucleic acid constructs. By contrast, the expression of tumour-suppressor miRNAs downregulated in the tumour could be restored and miR-26 levels could be increased with miRNA mimics. d, After treatment, the patient could be checked for relapse by periodically studying circulating miRNAs from serum in a non-invasive manner. The presence of miR-21 could indicate a potential relapse, and treatment would resume (black arrows).
healthy individuals to assess toxicity will provide valuable information about pharmacokinetics and safety of the treatment. LNAs have been optimized to target miRNAs by reducing their molecular size and this, along with developing strategies for more efficient delivery, has increased their therapeutic potential95. By contrast, another strategy involves the restoration of tumour-suppressor miRNA expression by synthetic miRNA mimics or viral delivery93. Both of these approaches have yielded positive results in mouse models of cancer75,96. Adenoassociated virus delivery of miRNAs or miRNA antagonists has the advantage of being efficient and, because the virus does not integrate into the genome, non-mutagenic. However, the delivery and safety of treatment needs to be improved before this approach can achieve widespread clinical use. In principle, the use of miRNA mimetics as therapeutics would allow ‘drugging the undruggable’ or the therapeutic inhibition of virtually any human gene. If this were possible it would undoubtedly impact many diseases including cancer by allowing the targeting of oncogenic transcription factors that are difficult to inhibit through traditional medicinal chemistry97. Furthermore, owing to the similar chemistry that is used to create drugs that target diverse molecules, the implementation of miRNA-based therapies could allow a more uniform drug development pipeline than is possible for more conventional treatments. Although experimental studies have validated the underlying biological impact of achieving miRNA modulation, there are still practical challenges that prevent the use of miRNA mimetics and antagonists clinically, including uncharacterized off-target effects, toxicities and poor agent-delivery. Concerning the last of these, most miRNA mimetics and antagonists rely on the delivery of molecules that mimic or inhibit the ‘seed’ sequence of an miRNA (typically molecules that consist of ≥6 nucleotides or related structures) across the plasma membrane — a particular challenge in the treatment of cancer, in which missing even a few cancer cells could lead to tumour relapse and progression. Extensive research is now focused on the viral and non-viral strategies required to meet this challenge, and results in the preclinical setting are promising75,94–96. Despite the considerable hurdles that have to be overcome, it 1 6 F E B R UA RY 2 0 1 2 | VO L 4 8 2 | N AT U R E | 3 5 3
INSIGHT REVIEW seems likely that miRNAs will find a place alongside more conventional approaches for the treatment of cancer. 9.
Perspectives Since the discovery of miRNAs in model organisms, miRNAs have emerged as key regulators of normal development and a diversity of normal cellular processes. Given what we know now, it is not surprising that perturbations in miRNA biogenesis or expression can contribute to disease. In cancer, the effects of miRNA alteration can be widespread and profound, and they touch on virtually all aspects of the malignant phenotype. Yet, precisely how miRNAs regulate the expression of protein-coding genes is not completely understood, and the underlying mechanism remains an important basic-science question that will have a significant impact on our understanding of gene regulation and its alteration in disease. In addition, we still lack effective approaches to understand and predict miRNA targets. New strategies to identify and characterize the targets of individual miRNAs, and to determine how they function in combination to regulate specific targets, will be required to understand their action on cell physiology. Because miRNAs can also regulate other non-coding RNAs (for example, long non-coding RNAs), which have a role in cancer development and vice versa98, these interactions will increase the complexity of gene regulation and are likely to produce regulatory processes that are currently hidden. Pioneering knowledge, gained through the study of miRNA function and regulation, will undoubtedly provide methodological and theoretical insights that will help in our understanding of the more recently identified non-coding RNA species. Understanding miRNA biology and how it contributes to cancer development is not only an academic exercise, but also provides an opportunity for the generation of new ideas for diagnosis and treatment. RNAi-based technology has allowed sophisticated loss-of-function experiments that were previously impossible and has revealed therapeutic targets that, when inhibited, can lead to cancer cell elimination. In addition, miRNAs themselves are being used directly in the diagnosis of cancer and, in the future, will probably be exploited in therapy to identify drug targets or as the drug treatment. However, cost-effective miRNA profiling strategies and larger studies are needed to determine whether miRNA profiling provides an advantage for cancer classification compared with a more traditional approach. Although drugs that function as miRNA mimetics, antagonists or synthetic siRNAs form the core of what is fundamentally a new class of drugs that are capable of targeting molecules outside the range of traditional medicinal chemistry, their clinical implementation will require improvements in drug composition and delivery; these challenges lie outside the scope of molecular biology and instead involve the fields of chemistry and nanotechnology. Nevertheless, the successful development of these technologies could ultimately translate our understanding of miRNA biology in cancer into strategies for the control of cancer. ■
10. 11. 12. 13. 14. 15. 16. 17. 18. 19. 20. 21. 22.
23. 24. 25. 26. 27. 28.
29. 30. 31. 32. 33. 34. 35.
1. 2. 3. 4. 5.
Bartel, D. P. MicroRNAs: target recognition and regulatory functions. Cell 136, 215–233 (2009). Lee, R. C., Feinbaum, R. L. & Ambros, V. The C. elegans heterochronic gene lin-4 encodes small RNAs with antisense complementarity to lin-14. Cell 75, 843–854 (1993). Wightman, B., Ha, I. & Ruvkun, G. Post-transcriptional regulation of the heterochronic gene lin-14 by lin-4 mediates temporal pattern formation in C. elegans. Cell 75, 855–862 (1993). Hanahan, D. & Weinberg, R. A. Hallmarks of cancer: the next generation. Cell 144, 646–674 (2011). Calin, G. A. et al. Frequent deletions and down-regulation of micro-RNA genes miR15 and miR16 at 13q14 in chronic lymphocytic leukemia. Proc. Natl Acad. Sci. USA 99, 15524–15529 (2002). This article reports miRNA deregulation in cancer and is the first evidence of the role of miRNAs in cancer. Lu, J. et al. MicroRNA expression profiles classify human cancers. Nature 435, 834–838 (2005). This article systematically profiles miRNAs in cancer and demonstrates their potential as classifiers. O’Donnell, K. A., Wentzel, E. A., Zeller, K. I., Dang, C. V. & Mendell, J. T. c-Mycregulated microRNAs modulate E2F1 expression. Nature 435, 839–843 (2005). He, L. et al. A microRNA polycistron as a potential human oncogene. Nature
3 5 4 | N AT U R E | VO L 4 8 2 | 1 6 F E B R UA RY 2 0 1 2
36. 37. 38. 39. 40. 41. 42.
435, 828–833 (2005). References 7 and 8 show, for the first time, that miRNAs can be actively involved in the MYC signalling pathway. Calin, G. A. et al. Human microRNA genes are frequently located at fragile sites and genomic regions involved in cancers. Proc. Natl Acad. Sci. USA 101, 2999–3004 (2004). Saito, Y. et al. Specific activation of microRNA-127 with downregulation of the proto-oncogene BCL6 by chromatin-modifying drugs in human cancer cells. Cancer Cell 9, 435–443 (2006). Mayr, C., Hemann, M. T. & Bartel, D. P. Disrupting the pairing between let-7 and Hmga2 enhances oncogenic transformation. Science 315, 1576–1579 (2007). Veronese, A. et al. Mutated β-catenin evades a microRNA-dependent regulatory loop. Proc. Natl Acad. Sci. USA 108, 4840–4845 (2011). Diederichs, S. & Haber, D. A. Sequence variations of microRNAs in human cancer: alterations in predicted secondary structure do not affect processing. Cancer Res. 66, 6097–6104 (2006). Kuchenbauer, F. et al. In-depth characterization of the microRNA transcriptome in a leukemia progression model. Genome Res. 18, 1787–1797 (2008). Yanaihara, N. et al. Unique microRNA molecular profiles in lung cancer diagnosis and prognosis. Cancer Cell 9, 189–198 (2006). Calin, G. A. et al. A microRNA signature associated with prognosis and progression in chronic lymphocytic leukemia. N. Engl. J. Med. 353, 1793–1801 (2005). Rosenfeld, N. et al. MicroRNAs accurately identify cancer tissue origin. Nature Biotechnol. 26, 462–469 (2008). Xi, Y. et al. Systematic analysis of microRNA expression of RNA extracted from fresh frozen and formalin-fixed paraffin-embedded samples. RNA 13, 1668–1674 (2007). Mitchell, P. S. et al. Circulating microRNAs as stable blood-based markers for cancer detection. Proc. Natl Acad. Sci. USA 105, 10513–10518 (2008). Chang, T. C. et al. Widespread microRNA repression by Myc contributes to tumorigenesis. Nature Genet. 40, 43–50 (2008). Thomson, J. M. et al. Extensive post-transcriptional regulation of microRNAs and its implications for cancer. Genes Dev. 20, 2202–2207 (2006). Kumar, M. S., Lu, J., Mercer, K. L., Golub, T. R. & Jacks, T. Impaired microRNA processing enhances cellular transformation and tumorigenesis. Nature Genet. 39, 673–677 (2007). Kumar, M. S. et al. Dicer1 functions as a haploinsufficient tumor suppressor. Genes Dev. 23, 2700–2704 (2009). Merritt, W. M. et al. Dicer, Drosha, and outcomes in patients with ovarian cancer. N. Engl. J. Med. 359, 2641–2650 (2008). Melo, S. A. et al. A TARBP2 mutation in human cancer impairs microRNA processing and DICER1 function. Nature Genet. 41, 365–370 (2009). Melo, S. A. et al. A genetic defect in exportin-5 traps precursor microRNAs in the nucleus of cancer cells. Cancer Cell 18, 303–315 (2010). Newman, M. A. & Hammond, S. M. Emerging paradigms of regulated microRNA processing. Genes Dev. 24, 1086–1092 (2010). Mavrakis, K. J. et al. Genome-wide RNA-mediated interference screen identifies miR-19 targets in Notch-induced T-cell acute lymphoblastic leukaemia. Nature Cell Biol. 12, 372–379 (2010). Portela, A. & Esteller, M. Epigenetic modifications and human disease. Nature Biotechnol. 28, 1057–1068 (2010). Cao, Q. et al. Coordinated regulation of Polycomb Group complexes through microRNAs in cancer. Cancer Cell 20, 187–199 (2011). Fabbri, M. et al. MicroRNA-29 family reverts aberrant methylation in lung cancer by targeting DNA methyltransferases 3A and 3B. Proc. Natl Acad. Sci. USA 104, 15805–15810 (2007). Varambally, S. et al. Genomic loss of microRNA-101 leads to overexpression of histone methyltransferase EZH2 in cancer. Science 322, 1695–1699 (2008). Hwang, H. W., Wentzel, E. A. & Mendell, J. T. A hexanucleotide element directs microRNA nuclear import. Science 315, 97–100 (2007). Khraiwesh, B. et al. Transcriptional control of gene expression by microRNAs. Cell 140, 111–122 (2010). Gebeshuber, C. A., Zatloukal, K. & Martinez, J. miR-29a suppresses tristetraprolin, which is a regulator of epithelial polarity and metastasis. EMBO Rep. 10, 400–405 (2009). Small, E. M. & Olson, E. N. Pervasive roles of microRNAs in cardiovascular biology. Nature 469, 336–342 (2011). Bueno, M. J. et al. Combinatorial effects of microRNAs to suppress the Myc oncogenic pathway. Blood 117, 6255–6266 (2011). Bui, T. V. & Mendell, J. T. Myc: maestro of microRNAs. Genes Cancer 1, 568–575 (2010). Dews, M. et al. Augmentation of tumor angiogenesis by a Myc-activated microRNA cluster. Nature Genet. 38, 1060–1065 (2006). Cairo, S. et al. Stem cell-like micro-RNA signature driven by Myc in aggressive liver cancer. Proc. Natl Acad. Sci. USA 107, 20471–20476 (2010). Kent, O. A. et al. Repression of the miR-143/145 cluster by oncogenic Ras initiates a tumor-promoting feed-forward pathway. Genes Dev. 24, 2754–2759 (2010). Johnson, S. M. et al. RAS is regulated by the let-7 microRNA family. Cell 120, 635–647 (2005). This article reports the first evidence of an oncogene, KRAS, being targeted by an miRNA. He, L., He, X., Lowe, S. W. & Hannon, G. J. microRNAs join the p53 network– another piece in the tumour-suppression puzzle. Nature Rev. Cancer 7, 819–822 (2007).
44. 45. 46. 47. 48. 49. 50. 51. 52. 53.
54. 55. 56. 57. 58. 59. 60. 61. 62. 63. 64. 65. 66.
67. 68. 69. 70. 71. 72. 73. 74.
This comprehensive review describes the regulation of the miR-34 family by the tumour suppressor p53. Pichiorri, F. et al. Downregulation of p53-inducible microRNAs 192, 194, and 215 impairs the p53/MDM2 autoregulatory loop in multiple myeloma development. Cancer Cell 18, 367–381 (2010). Xiao, J., Lin, H., Luo, X. & Wang, Z. miR-605 joins p53 network to form a p53:miR-605:Mdm2 positive feedback loop in response to stress. EMBO J. 30, 524–532 (2011). Yamakuchi, M. et al. P53-induced microRNA-107 inhibits HIF-1 and tumor angiogenesis. Proc. Natl Acad. Sci. USA 107, 6334–6339 (2010). Chang, C. J. et al. p53 regulates epithelial-mesenchymal transition and stem cell properties through modulating miRNAs. Nature Cell Biol. 13, 317–323 (2011). Kim, T. et al. p53 regulates epithelial-mesenchymal transition through microRNAs targeting ZEB1 and ZEB2. J. Exp. Med. 208, 875–883 (2011). Swarbrick, A. et al. miR-380-5p represses p53 to control cellular survival and is associated with poor outcome in MYCN-amplified neuroblastoma. Nature Med. 16, 1134–1140 (2010). Hu, W. et al. Negative regulation of tumor suppressor p53 by microRNA miR504. Mol. Cell 38, 689–699 (2010). Suzuki, H. I. et al. Modulation of microRNA processing by p53. Nature 460, 529–533 (2009). Su, X. et al. TAp63 suppresses metastasis through coordinate regulation of Dicer and miRNAs. Nature 467, 986–990 (2010). Ma, L., Teruya-Feldstein, J. & Weinberg, R. A. Tumour invasion and metastasis initiated by microRNA-10b in breast cancer. Nature 449, 682–688 (2007). This study demonstrates for the first time that miRNAs are involved in tumour invasion and metastasis. Tavazoie, S. F. et al. Endogenous human microRNAs that suppress breast cancer metastasis. Nature 451, 147–152 (2008). Ma, L. et al. miR-9, a MYC/MYCN-activated microRNA, regulates E-cadherin and cancer metastasis. Nature Cell Biol. 12, 247–256 (2010). Valastyan, S. et al. A pleiotropically acting microRNA, miR-31, inhibits breast cancer metastasis. Cell 137, 1032–1046 (2009). Cano, A. & Nieto, M. A. Non-coding RNAs take centre stage in epithelial-tomesenchymal transition. Trends Cell Biol. 18, 357–359 (2008). Korpal, M. et al. Direct targeting of Sec23a by miR-200s influences cancer cell secretome and promotes metastatic colonization. Nature Med. 17, 1101–1108 (2011). Martello, G. et al. A microRNA targeting Dicer for metastasis control. Cell 141, 1195–1207 (2010). Vander Heiden, M. G., Cantley, L. C. & Thompson, C. B. Understanding the Warburg effect: the metabolic requirements of cell proliferation. Science 324, 1029–1033 (2009). Godlewski, J. et al. MicroRNA-451 regulates LKB1/AMPK signaling and allows adaptation to metabolic stress in glioma cells. Mol. Cell 37, 620–632 (2010). Gao, P. et al. c-Myc suppression of miR-23a/b enhances mitochondrial glutaminase expression and glutamine metabolism. Nature 458, 762–765 (2009). Anand, S. et al. MicroRNA-132-mediated loss of p120RasGAP activates the endothelium to facilitate pathological angiogenesis. Nature Med. 16, 909–914 (2010). Mu, P. et al. Genetic dissection of the miR-17~92 cluster of microRNAs in Mycinduced B-cell lymphomas. Genes Dev. 23, 2806–2811 (2009). Olive, V. et al. miR-19 is a key oncogenic component of mir-17-92. Genes Dev. 23, 2839–2849 (2009). Costinean, S. et al. Pre-B cell proliferation and lymphoblastic leukemia/highgrade lymphoma in E(mu)-miR155 transgenic mice. Proc. Natl Acad. Sci. USA 103, 7024–7029 (2006). This article reports overexpression of a single miRNA can cause cancer in vivo. O’Connell, R. M. et al. Sustained expression of microRNA-155 in hematopoietic stem cells causes a myeloproliferative disorder. J. Exp. Med. 205, 585–594 (2008). Miska, E. A. et al. Most Caenorhabditis elegans microRNAs are individually not essential for development or viability. PLoS Genet. 3, e215 (2007). Klein, U. et al. The DLEU2/miR-15a/16-1 cluster controls B cell proliferation and its deletion leads to chronic lymphocytic leukemia. Cancer Cell 17, 28–40 (2010). Medina, P. P., Nolde, M. & Slack, F. J. OncomiR addiction in an in vivo model of microRNA-21-induced pre-B-cell lymphoma. Nature 467, 86–90 (2010). Chan, J. A., Krichevsky, A. M. & Kosik, K. S. MicroRNA-21 is an antiapoptotic factor in human glioblastoma cells. Cancer Res. 65, 6029–6033 (2005). Prosser, H. M., Koike-Yusa, H., Cooper, J. D., Law, F. C. & Bradley, A. A resource of vectors and ES cells for targeted deletion of microRNAs in mice. Nature Biotechnol. 29, 840–845 (2011). Loya, C. M., Lu, C. S., Van Vactor, D. & Fulga, T. A. Transgenic microRNA inhibition with spatiotemporal specificity in intact organisms. Nature Methods 6, 897–903 (2009). Zhu, Q. et al. A sponge transgenic mouse model reveals important roles for the miRNA-183/96/182 cluster in post-mitotic photoreceptors of the retina.
J. Biol. Chem. 2865, 31749–31760 (2011). This article reports the development of the first sponge transgenic mouse that allows in vivo inhibition of one or several miRNAs. 75. Kota, J. et al. Therapeutic microRNA delivery suppresses tumorigenesis in a murine liver cancer model. Cell 137, 1005–1017 (2009). This article uses adenovirus-associated vectors to deliver miRNAs to the liver and treat cancer. 76. Czech, B. & Hannon, G. J. Small RNA sorting: matchmaking for Argonautes. Nature Rev. Genet. 12, 19–31 (2011). 77. Chicas, A. et al. Dissecting the unique role of the retinoblastoma tumor suppressor during cellular senescence. Cancer Cell 17, 376–387 (2010). 78. Stegmeier, F., Hu, G., Rickles, R. J., Hannon, G. J. & Elledge, S. J. A lentiviral microRNA-based system for single-copy polymerase II-regulated RNA interference in mammalian cells. Proc. Natl Acad. Sci. USA 102, 13212–13217 (2005). 79. Zuber, J. et al. Toolkit for evaluating genes required for proliferation and survival using tetracycline-regulated RNAi. Nature Biotechnol. 29, 79–83 (2010). 80. Fellmann, C. et al. Functional identification of optimized RNAi triggers using a massively parallel sensor assay. Mol. Cell 41, 733–746 (2011). 81. Premsrirut, P. K. et al. A rapid and scalable system for studying gene function in mice using conditional RNA interference. Cell 145, 145–158 (2011). 82. Seibler, J. et al. Reversible gene knockdown in mice using a tight, inducible shRNA expression system. Nucleic. Acids Res. 35, e54 (2007). 83. Hemann, M. T. et al. An epi-allelic series of p53 hypomorphs created by stable RNAi produces distinct tumor phenotypes in vivo. Nature Genet. 33, 396–400 (2003). 84. Zender, L. et al. An oncogenomics-based in vivo RNAi screen identifies tumor suppressors in liver cancer. Cell 135, 852–864 (2008). 85. Westbrook, T. F. et al. A genetic screen for candidate tumor suppressors identifies REST. Cell 121, 837–848 (2005). 86. Xue, W. et al. Senescence and tumour clearance is triggered by p53 restoration in murine liver carcinomas. Nature 445, 656–660 (2007). 87. Luo, J. et al. A genome-wide RNAi screen identifies multiple synthetic lethal interactions with the Ras oncogene. Cell 137, 835–848 (2009). 88. Scholl, C. et al. Synthetic lethal interaction between oncogenic KRAS dependency and STK33 suppression in human cancer cells. Cell 137, 821–834 (2009). 89. Barbie, D. A. et al. Systematic RNA interference reveals that oncogenic KRASdriven cancers require TBK1. Nature 462, 108–112 (2009). 90. Zuber, J. et al. RNAi screen identifies Brd4 as a therapeutic target in acute myeloid leukaemia. Nature 478, 524-528 (2011). 91. Gumireddy, K. et al. Small-molecule inhibitors of microRNA miR-21 function. Angew. Chem. Int. Ed. Engl. 47, 7482–7484 (2008). 92. Melo, S. et al. Small molecule enoxacin is a cancer-specific growth inhibitor that acts by enhancing TAR RNA-binding protein 2-mediated microRNA processing. Proc. Natl. Acad. Sci. USA 108, 4394–4399 (2011). 93. Garzon, R., Marcucci, G. & Croce, C. M. Targeting microRNAs in cancer: rationale, strategies and challenges. Nature Rev. Drug Discov. 9, 775–789 (2010). 94. Lanford, R. E. et al. Therapeutic silencing of microRNA-122 in primates with chronic hepatitis C virus infection. Science 327, 198–201 (2009). 95. Obad, S. et al. Silencing of microRNA families by seed-targeting tiny LNAs. Nature Genet. 43, 371–378 (2011). 96. Bonci, D. et al. The miR-15a-miR-16-1 cluster controls prostate cancer by targeting multiple oncogenic activities. Nature Med. 14, 1271–1277 (2008). 97. Kumar, M. S. et al. Suppression of non-small cell lung tumor development by the let-7 microRNA family. Proc. Natl Acad. Sci. USA 105, 3903–3908 (2008). 98. Poliseno, L. et al. A coding-independent function of gene and pseudogene mRNAs regulates tumour biology. Nature 465, 1033–1038 (2010). This elegant study shows how mRNA from genes and pseudogenes can compete for the binding of miRNAs, unveiling the complexity of miRNA regulatory networks. 99. Ruby, J. G., Jan, C. H. & Bartel, D. P. Intronic microRNA precursors that bypass Drosha processing. Nature 448, 83–86 (2007). 100. Cheloufi, S., Dos Santos, C. O., Chong, M. M. & Hannon, G. J. A dicerindependent miRNA biogenesis pathway that requires Ago catalysis. Nature 465, 584–589 (2010). Acknowledgements We apologize to all colleagues whose work could not be cited owing to space restrictions. We thank L. Dow, A. Ventura, A. Saborowski and V. Aranda for their comments on the manuscript, and G. Hannon and L. He for the many discussions. A.L. is supported by an EMBO Long-Term Fellowship. S.W.L. is a Howard Hughes Medical Institute investigator. Author Information Reprints and permissions information is available at www.nature.com/reprints. The authors declare no competing financial interests. Readers are welcome to comment on the online version of this article at www.nature.com/nature. Correspondence should be addressed to S.W.L. (firstname.lastname@example.org).
1 6 F E B R UA RY 2 0 1 2 | VO L 4 8 2 | N AT U R E | 3 5 5