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Trends Biochem Sci. Author manuscript; available in PMC Sep 6, 2010.
Published in final edited form as:
PMCID: PMC2933830

Structural Features of Metabolite-Sensing Riboswitches


Riboswitches, metabolite-sensing RNA elements found in untranslated regions of the transcripts they regulate, possess extensive tertiary structure to couple metabolite binding to genetic control. Herein, we discuss recently published structures from four riboswitch classes and compare these natural RNA structures to those of in vitro selected RNA aptamers, which bind similar ligands to those of the riboswitches. Additionally, we examine the glmS riboswitch, the first example of a ribozyme-based riboswitch. This RNA provides the latest twist in the riboswitch field and portends exciting advances in the coming years. Our knowledge of the mechanisms of genetic regulation by riboswitches has increased mightily in recent years and will continue to grow as new riboswitch classes and ligands are discovered and structurally characterized.

Keywords: RNA structure, riboswitch, ribozyme, transcription attenuation, noncoding RNAs, posttranscriptional regulation, crystallography, NMR

Riboswitches: Remnants of an ancient mode of genetic regulation?

The “central dogma” states that information is stored as DNA, transcribed into RNA, and translated into proteins, which are traditionally thought to satisfy most cellular structural and catalytic requirements. Genetic control of these steps is believed to occur primarily through the action of regulatory proteins; however, the importance of noncoding RNAs in these processes has become increasingly appreciated in recent years [1]. In eubacterial organisms, regulatory RNAs can elicit control of gene expression through regulation of transcription attenuation, translation initiation [reviewed in 2,3], or mRNA stability [4]. These eubacterial regulatory RNAs function via diverse intermolecular (trans) or intramolecular (cis) mechanisms to regulate the target mRNA. Trans-acting regulatory RNAs interact with target mRNAs to control translation, mRNA stability, or transcription termination [reviewed in 5,6,7], whereas others regulate gene expression through specific sequestration of RNA-binding proteins [8]. Cis-acting RNAs, which are the focus of this review, are located within untranslated regions (UTRs) of the mRNA transcript that they regulate. Genetic control by the latter RNA elements can be accomplished either through the sole action of the RNA molecule or in conjunction with recruited protein factors.

A classic example of a cis-acting regulatory RNA is the transcription attenuation control of tryptophan biosynthesis genes in certain Gram-negative bacteria [9]. For this mechanism, the kinetics of translation of a short leader peptide dictates the conformational state of the overall 5′-UTR. Under tryptophan-depleted conditions the translating ribosome stalls at tryptophan codons within the leader peptide thereby allowing for formation of an antiterminator helix and ultimately, synthesis of the downstream trp operon. In contrast, during tryptophan-replete conditions the translating ribosome masks sequence elements required for antiterminator formation, instead allowing for the mutually exclusive formation of an intrinsic terminator. In other microorganisms, a tryptophan-binding protein (TRAP) associates with a different regulatory RNA in order to control transcription attenuation or translation initiation of downstream tryptophan biosynthesis genes [10]. In addition to these mechanisms, a class of regulatory RNAs termed T-box RNAs specifically associates with cognate uncharged tRNAs in order to promote expression of downstream amino acid biosynthesis and transport genes [11]. They are unable to respond to charged tRNAs, therefore the level of downstream gene expression is based upon the direct sampling of intracellular tRNA charging ratios by T-box regulatory RNAs. The fact that such distinct RNA-based mechanisms are similarly employed to ascertain the status of intracellular amino acid pools hints at the broad array of RNA elements that are likely to exist in modern biology.

A more recently characterized class of cis-acting regulatory RNAs, commonly referred to as riboswitches, regulate gene expression via direct association with small molecule metabolites [12] (Figure 1). These metabolites, which are essential to all forms of life, are bound by the RNAs with high affinity and selectivity. Riboswitches typically regulate expression of genes responsible for the synthesis and transport of the target metabolites. Whereas a few examples of these RNA motifs appear in eukaryotes [13], they are mainly found within eubacterial genomes. For example, in Bacillus subtilis, greater than 2.1% of the genome is believed to be under the regulatory influence of metabolite-sensing RNAs [3]. Several different mechanistic strategies are employed by these RNAs for genetic regulation. Generally, riboswitches exert control over translation initiation or formation of a transcription terminator helix within the 5′-UTR of the genes they regulate (transcription attenuation). While it is possible that accessory proteins influence these genetic mechanisms in vivo, they are not required for binding of metabolites to the RNAs. This observation, coupled with their widespread phylogenetic distribution, argues that riboswitches have been evolutionarily maintained for several billion years, although considerably more research is required to fully establish these claims. To date, biochemical evidence for riboswitch function has been obtained for RNAs that respond to adenosylcobalamin [14,15], thiamine pyrophosphate (TPP) [16], flavin mononucleotide (FMN) [17,18], guanine [19], adenine [20], a precursor for queuine [21], lysine [22], glycine [23], glucosamine-6-phosphate (GlcN6P) [4], and S-adenosylmethionine (SAM) [2426; reviewed in 27,12]. Given their ability to function as sensitive sentinels for intracellular metabolites in a protein-independent manner, these RNAs are likely to require exquisite structural sophistication. Therefore, riboswitches are excellent targets for structural analysis due to their overall importance in microbial genetic regulation, the possibility that they might resemble ancient RNA structures from a primordial world, and their striking ability to form efficient chemical-binding pockets. To that end, the aptamer domains (see Box 1) of several classes of riboswitches have been structurally characterized within the past three years. Specifically, structural models have been obtained for riboswitches that sense SAM, TPP, GlcN6P, and purine nucleobases [2834 reviewed in 35]. These studies have revealed the architectures that are unique to each riboswitch but have also identified common underlying mechanistic features employed by these important regulatory RNAs.

Figure 1
Secondary structure and genetic control mechanisms of structurally characterized riboswitches. The secondary structures and mechanisms of genetic control for each riboswitch class highlighted in this review are shown. Diversity in RNA-based genetic control ...

Riboswitches bind ligands with high selectivity utilizing unique architectural features

Most riboswitches have been subjected to chemical and enzymatic probing analyses in the absence and presence of their target ligand [briefly reviewed in 36]. The conclusions that can be extracted from these data vary dramatically per technique. For example, an alteration of the overall structure of riboswitches in the presence of target metabolites is supported by the differential binding of oligonucleotides between conformational states [37]. In addition to this technique, general secondary structure probing techniques such as in-line probing have produced higher resolution structural data. Specifically, in-line probing compares the relative spontaneous scission rates for backbone internucleotide linkages within RNA molecules that have been 5′-radiolabeled [38]. The products of these reactions are resolved by gel electrophoresis and analyzed by phosphor imaging. The rate of spontaneous cleavage is predominately dependent upon proper `in-line' positioning between the 2′-hydroxyl and the oxyanion leaving group, although metal ions can also affect these cleavage rates at high concentrations Therefore, regions of the RNA that exhibit greater or lesser flexibility can be visualized and quantified in this manner. This approach has proved to be remarkably successful in establishing the overall secondary structure patterns for these RNA species. Furthermore, monitoring these changes over a range of ligand concentrations has allowed for determination of apparent KD values, which for certain riboswitches have also been validated via several biophysical techniques [33,35,36,3941]. However, the most reliable secondary structures have been obtained when structural probing data have been combined with the comparative sequence analyses of riboswitches from different species. Secondary structure patterns of riboswitches that have been established in this manner have generally agreed with three-dimensional structural models as determined by X-ray crystallography. Exceptions include regions of extensive non-canonical base interactions and tertiary base contacts that are difficult to predict by sequence analysis alone.

Although most helical regions have been successfully predicted by biochemical probing and sequence analyses, recent crystallographic data reveal how they, and their intervening nucleotides, are arranged to form the tertiary conformations adopted for purine-, SAM-, TPP-, and GlcN6P-sensing riboswitches (Figure 2) [2834,42]. Similar to published structures of unrelated RNAs, there is extensive co-axial stacking of helices within riboswitches. However, most important for establishing the tertiary fold are nucleotides in interhelical regions, bulges, and terminal loops that are predominately responsible for long-range base connections. These residues stably bring together pre-formed secondary structure elements through a combination of hydrogen bonding interactions and extensive base stacking, ultimately stabilizing the tertiary architecture. Another functional consequence of these long-range base interactions is to stabilize the close packing of parallel helices.

Figure 2
Global visualization of riboswitch structure and ligand binding. (a) to (e) Representative structures are presented for each structurally characterized riboswitch class. The natural ligand and PDB accession codes are listed in parentheses along with number ...

RNA structures are sculpted in part from small stable structural motifs that rapidly form during the process of RNA folding [43]. Riboswitches are no exception to this rule as they also contain a subset of this steadily increasing catalog of motifs. For example, the kink-turn motif (K-turn), common in rRNA and noncoding RNAs [44], is also present in the SAM riboswitch (Figure 1c and and2c).2c). As in other RNAs, this motif introduces a sharp angle between two adjacent helices (P2a and P2b), ultimately assisting the formation of a long-range pseudoknot that is required for ligand binding. Another common RNA building unit, the GNRA tetraloop, is present within the glmS ribozyme and mediates direct contact between distantly located regions of the molecule (P1, P4) (Figure 1e and and2e)2e) [31]. Finally, A-minor motifs [45], which result from the docking of adenosines into the minor groove of a distantly located base pair, are common within riboswitch structures and are integral components for tertiary structure stabilization. In addition to these motifs riboswitches can also contain unique structural elements. Most notably in this regard is an interaction between terminal loops of P2 and P3 helices for guanine and adenine riboswitches [33,39,46]. This is a stable motif, comprised of two base quadruples and a noncanonical base pair, which still adopts its configuration even when removed from the full context of the purine riboswitch [46]. However, despite its ligand-independent formation, it is required for association of the purine ligand to the RNA.

Guanine and Adenine Riboswitches

For purine-sensing RNAs, residues important for ligand binding and long-range base interactions are largely located within the interhelical regions of a three-helix junction. X-ray crystallographic and NMR analyses demonstrate extensive hydrogen bonding interactions between virtually all functional groups within the nucleobase ligand and RNA (Figure 2a and b) [33,34,47]. Specifically, the sugar edge of the purine ligand pairs to the Watson-Crick face of the U51 base. Additionally, a hydrogen bond is formed between the U22 ribose 2′-hydroxyl and the Hoogsteen edge of the ligand. These ligand interactions are largely identical for both guanine- and adenine-sensing riboswitches with the exception of one position. The pyrimidine residue at the 74 position interacts with the nucleobase ligand via Watson-Crick base-pairing. For guanine riboswitches this position is conserved as a C whereas it is retained as a U for adenine-sensing RNAs. This enables the pairing interaction that occurs between the ligand and position 74 to function as the primary determinant for discrimination between intracellular purine nucleobases. The result is that the ligand is virtually engulfed within the core of the RNA structure. Specifically, the purine ligand fits into a pocket within the middle of a stacked column of base triples.

SAM-I Riboswitch

Similar to purine-sensing RNAs, almost all functional groups of the SAM ligand are recognized by RNA functional groups (Figure 2c). The adenine moiety of SAM forms a base triple with residues from the P3 helix whereas the methionine chain carboxyl group hydrogen bonds to the Watson-Crick face of a guanine in the P1–P2 interhelical region. These observations provide at least one explanation for how a highly negatively charged RNA polymer can surmount electrostatic difficulties inherent in specific recognition of a negatively charged ligand. These interactions help position the SAM ligand into a binding pocket that is situated between the minor grooves of the P1 and P3 helices. Additionally, a P3 guanosine interacts at the methionine portion. Therefore, the ligand binding pocket is ultimately created from two distantly located helices that are brought together by ligand association [28]. This compaction is likely to be assisted by a pseudoknot, which appears to be formed even in the absence of ligand interactions [26]. Both the length of the methionine chain and the partial positive charge located at the methyl-bound sulfur atom had been predicted to be important features in ligand binding [48]. Indeed, oxygen groups from P1 uridines might contribute to electrostatic interactions to the charged sulfur group. An interesting final note regarding SAM recognition is that the metabolite is bound in its stacked form, with the methionine chain directly above and parallel to the adenine base. This contrasts with recognition of SAM by proteins wherein the metabolite typically adopts its extended conformation [28].

TPP Riboswitch

The TPP-sensing RNA is the most widespread riboswitch class. It is also the only riboswitch class to have been identified in eukaryotes [13]. Therefore, recent structures from the Ban [29] and Patel [30] research groups provide a unique opportunity for comparative structural analysis of riboswitches from distantly related species (Figure 2d). Indeed, the overall architectures of this RNA class from E. coli and Arabadopsis are identical, although there are a few modest differences within the ligand pocket [49]. In contrast to the other riboswitches, not all ligand functional groups are specifically recognized. The ligand pocket is constructed largely in two halves, located within parallel helices. The pyrimidine ring of TPP is recognized by one helix whereas binding to the pyrophosphate moiety occurs within an interhelical region between coaxially stacked helices located parallel to the pyrimidine-binding helix. Surprisingly, the thiazole moiety of TPP is not specifically recognized by RNA through hydrogen bonding or base stacking interactions, accounting for the prior observation that TPP analogs containing alterations in this region were still capable of associating with the RNA [16,42,50]. Similar to the SAM sensor, the TPP riboswitch class also faces the electrostatic challenge of accommodating the negative charge of the pyrophosphate linkage. However, for this riboswitch class, this goal is accomplished through specific coordination of Mg2+.

What is the consequence of ligand binding to these different RNAs? Ligand binding precipitates long-range tertiary contacts, thereby helping to induce a compact global conformation. These tertiary contacts are of considerable importance for the purposes of genetic control. In all noncatalytic riboswitches structures, ligand binding facilitates formation of higher order structural features at or near to the P1 helix. The latter is a helix that is formed between sequences at the 5′ and 3′ termini of the RNA aptamer. For many riboswitches, the 3′ portion of the P1 helix is capable of base-pairing to an alternate portion of the molecule, which in turn includes nucleotides that are required for genetic control (e.g., intrinsic transcription terminator or Shine-Dalgarno-sequestering helix). Therefore, a most exciting feature of riboswitch structures is the structural complexity induced by ligand binding at or near the P1 helix. For example, association of the ligand with purine riboswitches induces formation of base triples with P1 residues. Similarly, binding of SAM to the riboswitch also directly involves P1 nucleotides. In fact, the P1 helix together with P3 comprises the SAM binding pocket for these RNAs. Finally, binding of TPP leads to formation of tertiary contacts juxtaposing the P1 helix. These ligand-induced tertiary contacts are also accompanied by extensive base stacking, sometimes with nucleotides from distant regions of the molecule. Together these ligand-induced structural features are likely to stabilize the P1 helix and alter the prevalence of alternate pairing routines that involve P1 nucleotides. Therefore, structural analyses of riboswitch RNAs have elegantly revealed how RNA can build a diverse range of small molecule binding pockets and couple this information to genetic control.

Riboswitches are active: glmS is a metabolite-sensing ribozyme

The discovery of the glmS riboswitch upstream of the glmS gene in Gram-positive bacteria not only added a new ligand binding class of riboswitch, but also a new mechanism of genetic regulation [4]. Namely, in response to binding GlcN6P, the glmS riboswitch is stimulated at least 100,000-fold to site-specifically self-cleave in an autocatalytic manner. As such, glmS represents the first example of a metabolite-responsive ribozyme. The mechanistic details for how self-cleavage of this ribozyme within the 5′-UTR of the glmS gene imparts genetic control in vivo have not been revealed. However, it has been generally assumed that self-cleavage by the glmS ribozyme somehow targets the downstream mRNA for rapid degradation by cellular factors. Since glmS has two primary functions, ligand binding and RNA cleavage, it is expected to share properties of both the aforementioned riboswitch classes and small self-cleaving ribozymes. Focusing on this comparison of the glmS structures to metabolite-sensing riboswitches, general rules for ligand binding remain the same whereas the additional catalytic functions of the RNA impose different characteristics on the aptamer structure. Specifically, binding of GlcN6P is mediated by extensive hydrogen bonds throughout the molecule, additional stacking to nucleobases and charge stabilization by Mg2+ for the sugar moiety and phosphoryl group, respectively (Figure 3) [31].

Figure 3
Stereo views of riboswitch aptamer ligand binding sites. The ligand binding sites from the (a) T. tencongensis SAM-I (b) E. coli TPP, and (c) T. tencongensis glmS riboswitches are shown with bases that make direct hydrogen bonding interactions to their ...

The additional functional role of glmS, namely catalysis, is likely to account for the most obvious difference between this RNA and other riboswitch aptamers: the RNA is nearly twice the size of any of the structurally characterized riboswitches. Though riboswitches with larger aptamer domains exist, the size or complexity of the ligand generally accounts for this difference. Unlike non-catalytic riboswitches whose tertiary structures are dynamic and become ordered upon ligand binding, glmS is fully folded and poised for binding of GlcN6P [31,32,51]. For this reason, multiple structures of glmS ribozymes were able to be determined in the presence or absence of natural ligands or potential inhibitors. These studies combined with subsequent fluorescence resonance energy transfer (FRET) experiments suggest that GlcN6P acts an enzyme cofactor [52], providing the final piece to a preorganized nucleobase active site. Elucidating the catalytic contribution of each component of the glmS active site is being intensely studied and future work should provide additional insight into this remarkable genetic regulatory ribozyme.

Riboswitches are sophisticated versions of in vitro selected RNA aptamers

Prior to the discovery of natural riboswitches, in vitro selected RNAs provided the first evidence that RNA was capable of binding a variety of small molecules [53,54]. A subset of these in vitro evolved RNAs have been structurally characterized using NMR or crystallographic techniques [5560]. Relevant to riboswitches, the structure of these RNA aptamers bound to AMP, GTP, FMN, and vitamin B12 have been elucidated (Figure 4). The AMP- and GTP-bound aptamers can be compared most directly to the guanine- and adenine-sensing riboswitches. In addition, examination of the FMN- and vitamin B12-bound aptamers allows speculation about the binding of these same ligands to their naturally occuring riboswitch counterparts. In vitro evolved aptamers tend to be much smaller than their natural counterparts with minimal motifs comprising ~30–50 nucleotides [61] whereas riboswitches tend to be ~70–170 nucleotides in length (compare Figure 4 and Figure 2) with the recently published preQ1-binding riboswitch minimal aptamer domain being an outlier at ~34 nucleotides [36,21].

Figure 4
Overview of ligand binding to in vitro evolved RNA aptamers. Non-natural RNA aptamer structures are shown as sticks with a ribbon drawn through the riboses of the RNA. The ligands are listed along with the PDB accession codes in parentheses and the number ...

Both riboswitch aptamer domains and RNA aptamers from in vitro evolution employ common mechanisms to bind small organic molecules, including base pairing, aromatic stacking, and metal-dependent stabilization of electronegative moieties, but they differ in the extent of direct and global interactions required to achieve molecule binding. In general, artificial aptamers do not interact with all portions of the ligand whereas natural aptamers (aptamer domains from riboswitches) typically bury the ligand in a site created by tertiary structure elements. As a result, artificial aptamer RNAs generally have lower affinities and less ligand specificity than their natural counterparts. A comparison of the structural basis for ligand binding of AMP, GTP, FMN, and vitamin B12 aptamers demonstrates this point (compare Figure 2a to e and Figure 4). The recently characterized GTP aptamer serves as the closest mimic of the binding mode of riboswitches [57]. All portions of GTP in this structure are recognized and nearly buried. Consequently the KD for this aptamer is ~75 nM, a number comparable to riboswitches and approximately two to three orders of magnitude tighter than most other in vitro selected aptamers (Figure 4b). However, this increase in affinity does not correlate with subsequent increases in specificity of the aptamer, which is far inferior to the natural counterparts [62]. As a final comparison, two aptamer structures have been solved bound to FMN and vitamin B12, ligands for riboswitches that have no structural models to date. When examining these in vitro selected aptamer structures in light of the biochemical data on ligand specificity for their natural riboswitch counterparts, it is evident that in vitro evolved aptamers partially reveal general modes of ligand binding to RNA, but will lack the sophistication of naturally evolved riboswitches. In short, in vitro evolved RNAs have been of great utility for proof of principle experiments on the diversity of RNA ligands and could be well suited for technology-based applications as sensors, but natural riboswitches are likely to have literally evolved beyond the current level of our in vitro selection capabilities.

Riboswitches are still ripe for discovery: new classes, ligands and applications

Metabolite-sensing RNAs have garnered much attention and inspired new ideas in the field of RNA biology and gene regulation since their discovery. The presence of naturally occuring metabolite-binding RNA aptamers has also bolstered the RNA world theory of evolution as it ascribes yet another role for RNA that might have existed in primordial organisms. Examination of natural RNA aptamers in the context of in vitro selected aptamers has illustrated the fundamental elements mediating ligand binding to all RNAs and accentuates the sophistication required by riboswitches to discriminate their cognate ligand in a cellular pool of metabolic intermediates. As well, riboswitches can act as ribozymes in response to metabolite as demonstrated by the glmS aptamer. While the recent explosion of structural analyses of riboswitches has enhanced our knowledge of RNA aptamers, this work is only a harbinger of exciting developments in this field as much remains unknown.

Many riboswitches have been described with intriguing metabolite ligands including adenosylcobalamin, lysine, glycine, and flavin mononucleotide that await structural analyses. Based on the current number of orphan riboswitches and the genes they regulate, the repertoire of ligands for riboswitches should expand within and beyond small organic molecules. As an example of a potential non-metabolite ligand, levels of intracellular metal ions have been proposed to regulate the expression of the Salmonella enterica mgtA gene via an RNA element located within its 5′ UTR [63]. Additionally, a broadly distributed orphan riboswitch class appears to regulate the expression of an MgtE-family magnesium transporter (ykoK) in Bacillus subtilis as well as many genes responsible for metal uptake in other organisms [64, 65], allowing one to wonder whether multiple classes of RNAs could function as metal ion sensors. The utilization of better bioinformatic search algorithms and genetic screens could uncover additional examples of structural noncoding RNAs, including new riboswitches. Lastly, riboswitches are current and future targets for technology and therapeutic applications. On the technological side, riboswitches seem primed for use as in vivo metabolite sensors or as modules for inducible control of gene expression. Furthermore, riboswitches are targets for known antimicrobials; and as riboswitches have not yet been identified in humans, they could be ideal targets for future drug development [66]. In closing, the RNA world might have benefited from riboswitches in the evolutionary process, but it is certain that present-day riboswitches have helped evolve our thinking about the seemingly unlimited possibilities for RNA.

Box 1. Apt usage of the term `aptamer'

The term aptamer originated from the Latin word, aptus, meaning `to fit' and has been traditionally used to refer to RNA and DNA species that have been artificially engineered to bind other molecules. In recent years, this term has been applied to natural RNAs that possess the ability to bind metabolites. In this review, the expression will refer to any region of nucleic acid, natural or synthetic, possessing the ability to bind a ligand.


Funding for CD is provided by the Sara and Frank McKnight Fund for Biochemical Research. Research on regulatory RNAs in the Winkler laboratory is supported by the University of Texas Southwestern Medical Center Endowed Scholars Program, the Searle Scholars Program, the National Institutes of Health, and the Welch Foundation.


1. Hannon GJ, et al. The expanding universe of noncoding RNAs. Cold Spring Harbor Symp. Quant. Biol. 2006;71:551–564. [PubMed]
2. Gollnick P, Babitzke P. Transcription attenuation. Biochim Biophys Acta. 2002;1577:240–250. [PubMed]
3. Winkler WC. Metabolic monitoring by bacterial mRNAs. Arch. Microbiol. 2005;183:151–159. [PubMed]
4. Winkler WC, et al. Control of gene expression by a natural metabolite-responsive ribozyme. Nature. 2004;428:281–286. [PubMed]
5. Brantl S. Regulatory mechanisms employed by cis-encoded antisense RNAs. Curr. Opin. Microbiol. 2007;10:1–8. [PubMed]
6. Gottesman S, et al. Small RNA regulators and the bacterial response to stress. Cold Spring Harb. Symp. Quant. Biol. 2006;71:1–11. [PMC free article] [PubMed]
7. Storz G, et al. Regulating bacterial transcription with small RNAs. Cold Spring Harb. Symp. Quant. Biol. 2006;71:269–273. [PubMed]
8. Babitzke P, Romeo T. CsrB sRNA family: sequestration of RNA-binding regulatory proteins. Curr Opin Microbiol. 2007;10:1–8. [PubMed]
9. Yanofsky C. The different roles of tryptophan transfer RNA in regulating trp operon expression in E. coli versus B. subtilis. Trends Genet. 2004;20:367–374. [PubMed]
10. Gollnick P, et al. Complexity in regulation of tryptophan biosynthesis in Bacillus subtilis. Annu. Rev. Genet. 2005;39:47–68. [PubMed]
11. Henkin TM, Grundy FJ. Sensing metabolic signals with nascent RNA transcripts: the T box and S box riboswitches as paradigms. Cold Spring Harb. Symp. Quant. Biol. 2006;71:231–237. [PubMed]
12. Winkler WC, Breaker RR. Regulation of bacterial gene expression by riboswitches. Annu. Rev. Microbiol. 2005;59:487–517. [PubMed]
13. Sudarsan N, et al. Metabolite-binding RNA domains are present in the genes of eukaryotes. RNA. 2003;9:644–647. [PMC free article] [PubMed]
14. Nou X, Kadner RJ. Adenosylcobalamin inhibits ribosome binding to btuB RNA. Proc. Natl. Acad. Sci. 2000;97:7190–7195. [PMC free article] [PubMed]
15. Nahvi A, et al. Genetic control by a metabolite binding mRNA. Chem Biol. 2002;9:1043–1049. [PubMed]
16. Winkler WC, et al. Thiamine derivatives bind messenger RNAs directly to regulate bacterial gene expression. Nature. 2002;419:952–956. [PubMed]
17. Mironov AS, et al. Sensing small molecules by nascent RNA: a mechanism to control transcription in bacteria. Cell. 2002;111:747–756. [PubMed]
18. Winkler WC, et al. An mRNA structure that controls gene expression by binding FMN. Proc. Natl. Acad. Sci. U S A. 2002;99:15908–15913. [PMC free article] [PubMed]
19. Mandal M, et al. Riboswitches control fundamental biochemical pathways in Bacillus subtilis and other bacteria. Cell. 2003;113:577–586. [PubMed]
20. Mandal M, Breaker RR. Adenine riboswitches and gene activation by disruption of a transcription terminator. Nat. Struct. Mol. Biol. 2004;1:29–35. [PubMed]
21. Roth A, et al. A riboswitch selective for the queuosine precursor preQ1 contains an unusually small aptamer domain. Nat. Struct. Mol. Biol. 2007;14:308–317. [PubMed]
22. Sudarsan N, et al. An mRNA structure in bacteria that controls gene expression by binding lysine. Genes Dev. 2003;17:2688–2697. [PMC free article] [PubMed]
23. Mandal M, et al. A glycine-dependent riboswitch that uses cooperative binding to control gene expression. Science. 2004;306:275–279. [PubMed]
24. Epshtein V, et al. The riboswitch-mediated control of sulfur metabolism in bacteria. Proc. Natl. Acad. Sci. U S A. 2003;100:5052–5056. [PMC free article] [PubMed]
25. McDaniel BA, et al. Transcription termination control of the S box system: direct measurement of S-adenosylmethionine by the leader RNA. Proc. Natl. Acad. Sci. USA. 2003;100:3083–3088. [PMC free article] [PubMed]
26. Winkler WC, et al. An mRNA structure that controls gene expression by binding S-adenosylmethionine. Nat. Struct. Biol. 2003;9:701–707. [PubMed]
27. Nudler E, Mironov AS. The riboswitch control of bacterial metabolism. Trends Biochem. Sci. 2004;29:11–17. [PubMed]
28. Montange RK, Batey RT. Structure of the S-adenosylmethionine riboswitch regulatory mRNA element. Nature. 2006;441:1172–1175. [PubMed]
29. Thore S, et al. Structure of the eukaryotic thiamine pyrophosphate riboswitch with its regulatory ligand. Science. 2006;312:1208–1211. [PubMed]
30. Serganov A, et al. Structural basis for gene regulation by a thiamine pyrophosphate-sensing riboswitch. Nature. 2006;441:1167–1171. [PubMed]
31. Klein DJ, Ferre-D'Amare AR. Structural Basis of glmS ribozyme activation by glucosamine-6-phosphate. Science. 2006;313:1752–1756. [PubMed]
32. Cochrane JC, et al. Structural investigation of the glmS ribozyme bound to its catalytic cofactor. Chem. Biol. 2007;14:97–105. [PMC free article] [PubMed]
33. Batey RT, et al. Structure of a natural guanine-responsive riboswitch complexed with the metabolite hypoxanthine. Nature. 2004;432:411–415. [PubMed]
34. Serganov A, et al. Structural basis for discriminative regulation of gene expression by adenine- and guanine-sensing mRNAs. Chem. Biol. 2004;11:1729–1741. [PubMed]
35. Gilbert SD, et al. Structural studies of the purine and SAM binding riboswitches. Cold Spring Harb. Symp. Quant. Biol. 2006;71:259–268. [PubMed]
36. Mandal M, Breaker RR. Gene regulation by riboswitches. Nat. Rev. Mol. Cell Biol. 2004;5:451–463. [PubMed]
37. Mironov AS, et al. Sensing small molecules by nascent RNA: A mechanism to control transcription in bacteria. Cell. 2002;111:747–756. [PubMed]
38. Soukup GA, Breaker RR. Relationship between internucleotide linkage geometry and the stability of RNA. RNA. 1999;5:1308–1325. [PMC free article] [PubMed]
39. Lemay JF, et al. Folding of the adenine riboswitch. Chem. Biol. 2006;8:857–868. [PubMed]
40. Wickiser JK, et al. The speed of RNA transcription and metabolite binding kinetics operate an FMN riboswitch. Mol. Cell. 2005;1:49–60. [PubMed]
41. Wickiser JK, et al. The kinetics of ligand binding by an adenine-sensing riboswitch. Biochemistry. 2005;44:13404–13014. [PubMed]
42. Edwards TE, Ferre-D'Amare AR. Crystal structures of the Thi-box riboswitch bound to thiamine pyrophosphate analogs reveal adaptive RNA-small molecule recognition. Structure. 2006;14:1459–1468. [PubMed]
43. Leontis NB, Westhof E. Analysis of RNA motifs. Curr. Opin. Struct. Biol. 2003;13:300–308. [PubMed]
44. Klein DJ, et al. The kink-turn: a new RNA secondary structure motif. EMBO J. 2001;20:4212–4221. [PMC free article] [PubMed]
45. Nissen P, et al. RNA tertiary interactions to the large ribosomal subunit: the A-minor motif. Proc. Natl. Acad. Sci. USA. 2001;98:4899–4903. [PMC free article] [PubMed]
46. Noeske J, et al. Interplay of `induced fit' and preorganization in the ligand induced folding of the aptamer domain of the guanine binding riboswitch. Nucleic Acids Res. 2007;35:572–83. [PMC free article] [PubMed]
47. Noeske J, et al. An intermolecular base triple as the basis of ligand specificity and affinity in the guanine- and adenine-sensing riboswitch RNAs. Proc. Natl. Acad. Sci. USA. 2005;102:1372–1377. [PMC free article] [PubMed]
48. Lim J, et al. Molecular recognition characteristics of SAM-binding riboswitches. Angew. Chem. Int. Edn. Engl. 2006;45:964–968. [PubMed]
49. Winker WC, Dann CE., III RNA allostery glimpsed. Nat. Struct. Mol. Biol. 2006;7:569–571. [PubMed]
50. Sudarsan N, et al. Thiamine pyrophosphate riboswitches are targets for the antimicrobial compound pyrithiamine. Chem. Biol. 2005;12:1325–1335. [PubMed]
51. Hampel KJ, Tinsley MM. Evidence for preorganization of the glmS ribozyme ligand binding pocket. Biochemistry. 2006;25:7861–7871. [PubMed]
52. Tinsley, et al. Trans-acting glmS catalytic riboswitch: Locked and loaded. RNA. 2007;13:468–477. [PMC free article] [PubMed]
53. Ellington AD, Szostak JW. In vitro selection of RNA molecules that bind specific ligands. Nature. 1990;346:818–822. [PubMed]
54. Szostak JW. In vitro genetics. Trends Biochem Sci. 1992;17:89–93. [PubMed]
55. Jiang F, et al. Structural basis of RNA folding and recognition in an AMP-RNA aptamer complex. Nature. 1996;382:183–186. [PubMed]
56. Dieckmann T, et al. Solution structure of an ATP-binding RNA aptamer reveals a novel fold. RNA. 1996;2:628–640. [PMC free article] [PubMed]
57. Carothers JM, et al. Solution structure of an informationally complex high-affinity RNA aptamer to GTP. RNA. 2006;12:567–579. [PMC free article] [PubMed]
58. Fan P, et al. Molecular recognition in the FMN-RNA aptamer complex. J. Mol. Biol. 1996;258:480–500. [PubMed]
59. Sussman D, et al. The structural basis for molecular recognition by the vitamin B 12 RNA aptamer. Nat Struct Biol. 2000;7:53–57. [PubMed]
60. Sussman D, Wilson C. A water channel in the core of the vitamin B(12) RNA aptamer. Structure. 2000;8:719–727. [PubMed]
61. Patel DJ, et al. Structure, recognition, and adaptive binding in RNA aptamer complexes. J. Mol. Biol. 1997;272:645–664. [PubMed]
62. Carothers, et al. Aptamers selected for higher-affinity binding are not more specific for the target ligand. J Am Chem Soc. 2006;128:7929–79237. [PubMed]
63. Cromie MJ, et al. An RNA sensor for intracellular Mg2+ Cell. 2006;125:71–84. [PubMed]
64. Barrick JE, et al. New RNA motifs suggest an expanded scope for riboswitches in bacterial genetic control. Proc. Natl. Acad. Sci. U.S.A. 2004;101:6421–6426. [PMC free article] [PubMed]
65. Griffiths-Jones S, et al. Rfam: annotating noncoding RNAs in complete genomes. Nucleic Acids Res. 2005;33:D121–124. [PMC free article] [PubMed]
66. Blount, Breaker RR. Riboswitches as antibacterial drug targets. Nat Biotechnol. 2006;24:1558–1564. [PubMed]
67. Shi H, Moore PB. The crystal structure of yeast phenylalanine tRNA at 1.93 A resolution: a classic structure revisited. RNA. 2000;6:1091–1105. [PMC free article] [PubMed]
68. Lamour V, Darst SA. Crystal structure of Thermus aquaticus Gfh1, a Gre-factor paralog that inhibits rather than stimulates transcript cleavage. J Mol Biol. 2006;356:179–188. [PubMed]
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