SPP 1784 – Excerpt from application:
"Chemical Biology of native Nucleic Acid Modifications"
Coordinator
Prof. Dr. Mark Helm Johannes Gutenberg-Universität Mainz Institut für Pharmazie und Biochemie Staudinger Weg 5 D-55128 Mainz1. Summary
The sequence genetic code based on the four canonical Watson-Crick nucleosides has been an accepted paradigm for decades, however, an important aspect of it has recently been overturned. We learned that additional nucleosides exist in DNA, which establish a second layer of information that is important e.g. to differentiation processes. Along the same line, we now start to understand that the numerous known modified RNA nucleosides are part of a similar “hidden” program that we are just beginning to understand. Indeed, deciphering the modified “genetic” code that is embodied by these modified DNA and RNA bases is currently one of the hottest topics in (Chemical) Biology. Unveiling the new hidden layer of information in the genetic code beyond the classical four letter alphabet is the aim of this application. In order to keep Germany in a leading position in this exciting field, we plan to connect the expertise present in Germany in a nationwide research program and To link the existing excellence. To this end, we have to integrate Germany’s outstanding nucleic acid expertise with scientists that are able to develop new analytical tools in order to detect and quantify the new DNA and RNA modifications. We will define the functions of these modified nucleosides by studying their protein interaction partners and by localizing the modification in specific gene regions or in special RNA species. Most of the current cutting edge challenges lie in the domain of Chemical Biology, where synthetic chemistry must develop new chemical tools to detect modifications, to synthesize modifications, and to inhibit or tag modification enzymes. New sequencing methods need to be developed that can detect and localize nucleoside modifications. Finally, we need structural biology to understand the mechanisms that allow proteins to bind specifically to the relevant parts of nucleic acids, which establish the unknown second information level beyond the sequence code. Germany has internationally leading expertise in all these key areas. Bundling this expertise is the only way to stay in a leading position in this highly competitive field. Creating a thriving network that will gain deeper inside into where, how and why native nucleic acid modifications occur and how they influence cellular processes is the central goal of this SPP.
2. Scientific background
Our view of the genetic code has recently been overthrown. It is now clear that the genetic code is more complex than though so far.[1,2] Until 2009, researchers believed that encoding life requires the four Watson-Crick bases plus a fifth base 5-methylcyctosine, which is able to selectively silence genes in higher eukaryotes. In 2009, we learned that 5-hydroxymethylcytosine is a base that is highly prevalent in stem cells and in brain tissue.[3] Over the last 3 to 4 years we then learned that these bases are needed to control silencing and reactivation of genes, which provides our genome with so far unknown plasticity.[4,5] In 2011, a German group discovered 5-formylcytosine [6] which is now called the 7th base of the genome and shortly thereafter groups in the USA and China reported the existence of 5-carboxycytosine.[7,8][1]So far, the function of these new bases is unclear, but we all believe that they function as switches able to regulate gene function, and thus establishing a completely new coding level that has escaped notice so far. These new discoveries fall into a time where new base modifications in RNA are discovered as well and where association with epigenetics has reignited the field of RNA modifications.[9,10] We have known for some time that RNA contains more than 150 modified bases, but only in very few cases was it possible to analyze their function. Together with the newly discovered DNA nucleosides, these base modifications are now thought to establish a completely novel second layer of information needed to regulate transcription and translation.
These new discoveries prompt a need to analyze nucleoside modification with higher precision. To this end new mass spec based technologies are currently being developed, that give detailed insight into the nucleoside modification content. In addition, we have to decipher the sequence context, but conventional sequencing technologies fail with modified nucleoside. The world wide efforts to develop sequencing methods that can localize modifications are just beginning to become fruitful. The first proteins are currently discovered that interact specifically with the DNA modifications and crystal structure of a few complexes start to provide insight of how the modified DNA and RNA bases are recognized. Proteomics data that can uncover the regulatory networks behind the new nucleosides are available only in a few cases. The synthesis of phosphoramidites of the modified bases needed for the synthesis of DNA and RNA with defined modifications at defined sites is a pressing problem for many modifications. Most important is that modifications are affecting the three dimensional structure of particular RNA structures. We are just at the beginning to learn how structural equilibria are affected by modified nucleosides and how less populated conformations with alternate biological function are stabilized or destabilized. We do not understand the mechanisms that regulate modification machineries and in many cases we have no information of how a cell knows where and when a certain modification has to be incorporated. More importantly, it is unclear in most cases how modifying enzymes locate a certain position for modification in the large DNA and RNA pool. Finally, the chemistry that allows proteins to change the chemical structure of a given nucleoside has been elucidated only in a very limited number of cases.[11] DNA and RNA nucleoside modifications clearly comprise a new layer of information in this sense and new modifications are still identified.[12–14] We therefore expect that many others are still to be uncovered. Even the most recent compilations[15] cannot adequately reflect the complexity of the situation: Of the over 160 modification types known in RNA and DNA, only a handful can be detected by current –“Omics” protocols.
The previously segregated fields of RNA and DNA modification (Figure 1) are now merging to create a brand new interdisciplinary research field. This SPP is designed to keep German Science into a worldwide competitive situation. The new field links the chemical alterations of the nucleoside structures to aspects of stress response, developmental biology, and epigenetics. It is becoming increasingly clear that the concerted action of nucleic acid modifying enzymes triggers and controls gene expression. The vast new field is defined by the chemistry and biology of the chemical nucleoside modifications [15] that occur in literally thousands of RNA species [2] and in different DNA regions. Among the most remarkable features of recent discoveries is the apparently reversible nature of many modifications, which is directly associated with the potential to regulate gene expression.
Structure and location of nucleic acid modifications
The current surge in the field is closely related to the recent discovery of naturally occurring oxidation products of 5-methylcytidine in DNA, which has started an avalanche of modification related research throughout the life sciences.[3,5–8] The high profile of 5-hydroxymethyl-, 5-formyl-, and 5-carboxycytosines (Figure 1A) in current literature has prompted many new developments in detection technology.[16] Consequently, new search strategies have produced novel approaches to discover and detect unusual modification structures. These new methods have recently led to the discovery that DNA phosphorothioates [17] are naturally occurring modifications. So far, they were considered artificial modifications that chemists used to stabilize oligonucleotides for in vivo applications. Significant crosstalk from the DNA field, has also promoted renewed interest in RNA modification. Figure 1A shows some modified DNA and RNA cytidines, illustrating the obvious relation between the fields. Indeed, 5-methylcytidine, 5-hydroxymethylcytidine and 5-formylcytidine are long-known modifications of ribonucleosides.[15] Compared to RNA, a smaller number of modified nucleoside are known in DNA. These mainly act as epigenetic or bacterial identity markers. In contrast, the large variety of native modifications in RNA exerts a variety of functions in the different RNA species that are part of complex regulatory cellular networks. Figure 1B shows some secondary metabolites of modified nucleosides and thus illustrates the relation of modification to basic metabolic pathways, which use ubiquitous metabolites and coenzymes to transfer methyl groups, acetyl groups, aminoacids, isoprenoids, sugars, phosphate groups and the like. Many such metabolite–RNA conjugates are known or likely to enhance the chemical functionality of their targets. A majority of modifications are chemically simple, but some are fairly sophisticated, requiring multistep biosynthesis cascades. For example, among the most complex modifications are the nearly a dozen wybutosine derivates (Figure 1C).[2]
Their biosynthesis involves an orchestrated series of about a dozen chemical transformations featuring e.g. radical chemistry and the formation of a new heterocycle.[18] More exotic chemical features (Figure 1C), such as an epoxide in epoxyqueuosine [19] or a sulfonate in 5-taurinomethyluridine [20] underpin the complex chemistry that nature developed to modify bases. Despite this, calling RNA modification “natural product chemistry on a biopolymer scaffold” has been met with some skepticism, but new discoveries have re-emphasized this principle. This applies in particular to the discoveries of covalent RNA adducts of coenzyme A [13] and NAD [21], as well as the isolation and identification of geranylated RNA (Figure 1C).[14]Altogether, the list of more than 150 chemically distinct modified ribonucleotides known to date seems not to be exhaustive, since new modified residues keep surfacing in DNA and RNA of bacteria, archaea and eukarya.[12,15]
We only have very limited insight into how and where the modifications are introduced into the different RNA species. In eukaryotes, maturation of cellular RNAs starts in the nucleus, immediately after DNA transcription by RNA polymerase. The primary RNA transcript may have 5'- and 3'-extensions, interrupting sequences (introns) and be composed of only the canonical. During the complex process of RNA maturation, the 5'- and 3'-ends are trimmed, introns are removed and initially incorporated nucleotides are chemically modified, most frequently by addition of simple chemical groups on the base or the 2'-OH of the ribose. It is clear that somewhere during the complex RNA maturation process further modifications are introduced. This happens partially in the nucleolus but some steps require transport of the RNA into the cytosol and back into the nucleus. This regulated transport is certainly another biochemical problem that needs to be investigated.
Although RNA modifications such as pseudouridine (Ψ) were first discovered in the early 60-ies [22] and extensively studied in the 70-ies/80-ies, the modified positions and the exact chemical nature of modification are known only for a limited number of the more abundant cellular RNAs, typically tRNAs and rRNA from some well-studied model species from bacteria, archaea and eukaryotes, including in particular E. coli and yeasts.[23] The knowledge of snRNA modifications is more limited, even if their modification profile is known for some model organisms.[24] Other classes of cellular RNAs were not studied at all, or only fragmentary data exist on the presence of modified residues in these molecules. The main reason for these information gaps is the absence of modern and efficient tools allowing the detection, characterization and localization of RNA modifications in rare or low abundant cellular RNAs. This important limitation also makes characterization of RNA modification enzymes difficult, since only a limited subset of their cellular substrates is really known. The discovery of 3'-terminal methylation in miRNAs and piRNAs[25–27] clearly demonstrates that modified nucleotides are also present and functionally important in these small regulatory RNA molecules. From the above follows, that RNA modification is likely to be found in abundance in all RNA species amenable to an analysis of their modification content. It has also become clear that modifications in less abundant species such as mRNA[28–30] and regulatory RNAs[26] have a huge regulatory potential. Studying this potential requires unveiling the presence of myriads of suspected modifications, which could never quite be pinpointed by current methods. [3]
Detection of DNA and RNA modifications
Of the 160 or so currently characterized modification types in DNA and RNA[15], only very few can be detected by biochemical sequencing techniques, while a vast majority is best identified and quantified by their physico-chemical properties, typically chromatography, mass-spectrometry, or a combination thereof. As these methods typically include a degradation step during sample preparation, sequence information is severely limited or non-existent. This obviously presents the most important bottleneck in studies of the biological effects of any modification, and it has been circumvented only for a handful of modifications, including 5mC, 5hoC, and 5caC in DNA[31], as well as inosine (I), 5-methylcytidine (m5C), 2’-O methylation, pseudouridine (Ψ), and, to a limited extend, 6-methyladenosine (m6A) in RNA.[32,33] For the aforementioned modifications, analysis on the genome/transcriptome scale seems possible in principle, or it has been actually conducted. Inosine, better known as the product of A-to-I editing, changes the reverse transcription (RT) profile from that of the unmodified A to one equal to G; identification is therefore conceptually easy, albeit technically challenging on a transcriptome-wide scale, by comparison of cDNA to genomic DNA sequences. The relative ease of application has resulted in several high-ranking publications on transcriptome-wide analyses[34,35], and an explosion of exciting data in the field, which now carries the firmly established branded name A-to-I editing.[36] More recently, bisulfite sequencing, a long standing technique for detection of 5mC in DNA[37], has been adapted to RNA by the Lyko group at the DKFZ in Heidelberg.[38] This chemical treatment, which has permitted transcriptome-wide studies as well[39], embodies a key concepts for identification of modifications in sequencing data: it changes the RT-profile of the analyte RNA by means of a chemical treatment, i.e. the modification is revealed by differences in RNA-seq data from before and after the treatment. One more modification, 2'-O methylation, can be detected by differential RT signatures: if the RT is performed at low dNTP concentrations, the enzyme is arrested by the modification, whereas it passes the same site at high concentrations of dNTP[40,41]. Although we are unaware of published transcriptome-wide data, we known that such work is under way from personal communications. The detection of pseudouridine by chemical treatment with CMCT and subsequent RT analysis has been reported in the mid-90ies[42], and while entire ribosomal subunits have been mapped[43], the reaction has not (yet) been adapted to transcriptome-wide detection. Finally, two top ranking papers from 2012 report on mapping m6A sites in mRNA.[28,29] However, since these data originate from RNA enriched by antibodies directed against m6A, they do not report precise mapping to the nucleotide position, neither any direct confirmation of the presence by e.g. LC-MS. The data is, however, in part consistent with previously identified consensus motifs for the corresponding methyltransferase complex.[44] Meanwhile, although there is room for technical improvement and more precise mapping of such methylation sites, these findings gave a large impetus to a newly emerging field, epigenetic RNA modifications.[9] Considering how important it is to know the exact position of a modification in the respective DNA or RNA, it is clear that we need to develop new methods that finally allow sequencing of all modifications. Next to new chemistry, the development antibodies against the modifications and of single molecule sequencing technologies might be avenues to solve this pressing problem.
A particular challenge is that modifications need to be analyzed in DNA and in all kinds of different RNA species within cells. It is hence not enough to perform one single sequencing event for a certain organism but we need multiple sequencing of different oligonucleotide species. One might need to perform sequencing of different cell types in different cellular states. Next to sequencing the determination of the modification content by LC-MS methods needs to be optimized. This requires also the development of high throughput. All methods and techniques require the synthesis of authentic standards[45], either for incorporation into oligonucleotides by solid phase synthesis, or for validation by “hard”, i.e. physicochemical characterisation such as LC-MS. [4] Another major bottleneck is the extraction of a sufficient amount of pure or highly enriched RNA sample. Various methods are based on differences in the size of different RNA molecules and use electrophoresis or chromatography.[46] Other widely used approaches use affinity chromatography with antibodies, specific for some features of a given RNA class. The use of anti-cap (m7GpppG) antibodies for isolation of snRNAs, snoRNAs and some mRNAs is described in the literature.[47,48] These methods allow rough pre-fractionation of RNAs, but are generally unable to provide RNA species of sufficient purify for LC-MS/MS analysis. An alternative general approach for isolation of specific RNA is based on the annealing of the complementary DNA oligo followed by elution of the bound RNA under selective temperature and salt conditions. A particularly interesting multiplexed application was developed in the format of column chromatography (“chaplet columns”) and also in robotized form allowing the parallel extraction[49] of multiple RNA species.
For a global analysis of the full modification content of a given small ncRNA by LC-MS, an RNA sample is required in pico- to-femtomol quantities. For a quantification, digestion of an RNA sample to nucleosides (by nuclease P1 followed by alkaline phosphatase treatment) is followed by separation on a C18 reverse-phase chromatography, which is coupled to an ESI-MS/MS triple quad instrument.[32,50] This data identifies the chemical nature of a given RNA modification as well as its abundance, but not the exact positions in the parent RNA molecule. A more cumbersome method that conserves some amount of sequence information uses a digestion of the RNA sample by RNAse T1, which faithfully cleaves after each guanosine. The resulting subset of T1 RNA fragments is then analyzed for modifications. While this method is, in principle, suitable for the determination of the exact position of a given modification, its application to routine analysis suffers from requirements for relative large amounts of sample as well as strong expertise in RNA mass-spectrometry for precise interpretation of the resulting MS spectra.[51]Although significant recent advances in mass spectrometry and nucleoside chemistry have allowed more sensitive quantification[52,53] and detection of modifications in a sequence and position specific context[54], these technical bottlenecks still throttle progress in the modification field. The mass spec approaches still remain irreplaceable for confirmation of localization data obtained by other methods.
Target recognition and catalysis
The interactions of target sites for modifications in nucleic acids are of particular interest both before and after the modification event itself. Target recognition and catalytic turnover by modification enzymes offer a plethora of aspects, as, for example they determine the extent of any downstream effect exerted by the modification. The combination of chemical biology and structural biology has produced outstanding insights, e.g. via the use of substrate analogues[55–57], which may be used to tag and detect modification sites or to biochemically elucidate reaction mechanisms. Significant progress has thus been made in the elucidation of catalytic mechanisms.[58,59] Given the multitude of different modifications and nucleic acid species, the field is obviously wide open and at the same time can rely on a large body of work, of which only the most outstanding features can be mentioned here. [5]
Arguably the most influential DNA-related discovery was the mode of the action the Ten-Eleven-Translocation proteins (TET1, TET2 and TET3), which specifically oxidize the methyl groups of 5mC to 5-hydroxymethylcytosine (5hmC).[3] These enzymes belong to the alpha-ketoglutarate/Fe2+ dependent dioxygenase family. Soon after, it was shown that the TET enzymes can further oxidize the 5hmC to 5-formylcytosine (5fC) and 5-carboxylcytosine (5caC), implying a stepwise, oxidative DNA demethylation process.[5,8,60] The currently most investigated RNA modifications are m5C[61], m6A, and pseudouridine: this is due, in large part, to the advances in transcriptome-wide detection, which have revealed many unexpected locations in mRNA and ncRNAs.[28–30] While many m5C:MTases are well characterized and their catalytic mechanism largely elucidated[61], the MTase responsible for the formation of m6A in mRNA is a poorly characterized multi-component complex[44], and currently a hot topic in the field. The catalysis of pseudouridine formation has been the object of research on many different pseudouridine synthase enzymes for some time, but still awaits ultimate clarification.
Recent insights into the function of nucleic acid modifications
As mentioned, the most intriguing latest developments come from the areas of DNA modification, mRNA modification, and tRNA modification, with other ncRNAs rapidly moving into the focus. Common to all these areas is their dynamic nature, i.e. nucleic acids modifications are removed as well as added, which implies a general capacity for the regulation of gene expression. This has been specifically noted in the scientific communities interested in epigenetics and development, but physiology and pharmacology is not lagging far behind, and a spreading to neighboring communities is highly probable.
Once modified, the mode of action by which the modification transmits its effect is of central interest to its function. Examples of direct recognition of a modification include 5mC in DNA which is specifically bound e.g. by MBD proteins and SRA proteins.[62,63] Downstream of 5mC formation, the 3 TET-dependent modified nucleosides 5hmC, 5fC and 5caC (Figure 1A) are not only intermediates in DNA demethylation, but most likely constitute new epigenetic signals on their own, as is suggested by the fact that numerous proteins in the cells can specifically recognize and bind them.[16,63] For DNA modifications, the study of these interacting enzymes constitutes the current cutting edge of the field.
For mRNA modification, the current focus is on addition as well as oxidative removal of methylgroups to the N6 of adenines. Two highly visible transcriptome wide studies have identified clusters of m6A sites in critical regions of mRNAs such as miRNA seed regions, stop codons, and splicing sites.[28,29] This modification exhibits tissue-specific regulation, and dynamics including oxidative demethylation similar to DNA via the Fat mass and obesity-associated (FTO) gene product.[64]
Modification enzymes are subject to regulation themselves. In particular for tRNA modification, recent data implies that modification enzymes play pivotal roles in the mediation of cellular responses to outside stress stimuli such as starvation or oxidative species.[65,66] The static view of tRNA modification as “concrete-cast” functional equipment has given way to viewing RNA modification as an instrument of gene regulation It is now becoming clear, that the dynamic nature of tRNA modification as a whole or of single modifications impact codon usage, and thus can switch the cell’s metabolism by selective translation of certain mRNAs. Concomitantly, interest has resurfaced and connections are being made between tRNA modifications, a cell’s overall stress response, cell development and its protein synthesis capacity.[66–68] [6]
3. Scientific goals
Understanding at the molecular and atomic resolution level
An integral understanding of nucleic acid modifications encompasses a variety of aspects, of which the molecular level is the central focus of this network. In a field that is currently wide open, urgent questions do not focus on a particular system of a defined modification or nucleic acid species. Rather, the basics of the field are currently being defined in general terms: where, how and why do modifications occur? It is along these three basic questions on detection, recognition and function, that we have identified a number of bottlenecks that currently present the biggest obstacles to faster progress in the field. These bottlenecks have obviously become so important because, as opposed to related problems in the field, they could not be solved by standard methods of molecular biology. The common motif of these bottlenecks is that their solutions involve manipulations at the atomic resolution level. This is the classical domain of chemical biology, with the impact of organic synthetic chemistry in synergy with structural biology and biochemistry. The German scientific landscape is in a privileged position to make important contributions to the solution of exactly these major bottlenecks and therefore can potentially have an enormous impact on the international level. German Science is traditionally strong in organic chemistry and that holds especially true for nucleic acid chemistry. In order to stay in a competitive situation in one of the hottest research areas right now, the participants of this funding initiative will form a network among young and established scientists from synthetic organic chemistry, chemical biology, biochemistry and structural biology including also some selected biologist from cell biology, bioinformatics, developmental biology, and epigenetics.
3.1 Detection
Detection and localization of modifications in nucleic acids are among the most important bottlenecks and therefore are a prime concern of the SPP. To collect a complete picture for a given modification, its occurrence in a particular nucleic acid species, its position in the latter, and the localization inside the cell or tissue are important must be determined as a basis for sustained research. Lower detection limits and the possibility to conduct genome-wide and transcriptome-wide analyses are eagerly awaited by the international community, and approaches to visualization of modifications or modification events inside a cell are virtually unknown.
Therefore, one particularly important set of goals of the SPP concerns analytics of modifications. Method developments that improve sensitivity, specificity, or the possibility to determine a sequence context in a genome or transcriptome wide context will be given preference. The SPP aims at establishing an overview of the distribution of modifications in the different classes of RNA, such as mRNA, tRNA and rRNA and other ncRNAs. Because detection efficiency varies with chemical structure, analytical detection must be specifically tailored to every newly discovered or investigated modification with an attention to chemical reactivity. In addition to the detection of new modifications, the SPP will focus method development onto a number of modifications currently considered particularly important. These include 5hmC, 5hmU, 5fC and 5caC on the DNA level and m6A, pseudouridine, thiouridine, inosine, queuosine, 2’-OMe, and m5C in RNA. Another important goal is the development of methods for visualization of modifications or modification events inside a cell.
3.2. Recognition
Recognition events by cognate proteins of nucleic acids and cofactors offer an intriguing possibility to investigate and manipulate modification systems. Based on efficient detection, recognition studies are essential to interpret molecular and atomic details in the bigger picture of the biological function of a given modification. Current demands on chemists that are important in the network include e.g. the development of cofactor analogues for the inhibition of modification reactions, for crystallographic studies, or as reporter moieties. The interaction of such small molecules, e.g. as inhibitors, must be characterized by enzymology. The development and the application of such chemistry are most efficient when accompanied by structural biology, which may provide a basis for the design of small molecules binding to catalytic sites. Although a number of modification enzymes are structurally characterized, the most relevant structures, i.e. those of enzymes in complex with target nucleic acids, are extremely rare. However, these are the most important structures for an understanding of recognition processes.
Therefore, one goal of the SPP is the resolution of structure and dynamics of modification enzymes and their target nucleic acids before, during, and after the catalytic modification event, by methods of structural biology including in particular NMR and crystallography. Another goal is the development and application of substrate and cofactor analogues. These may be used in enzymatic characterization of modification events, which constitutes another important goal.
3.3. Function
Molecular interactions of modified nucleic acids with cognate macromolecules mediate their downstream regulatory effects and thus are key to an understanding of regulatory aspects of modifications. A central question that needs to be addressed for every given modification is whether its function is mediated via direct recognition of e.g. a nucleoside methylation, or if the modification alters the nucleic acid’s global structure, and the altered structure is then the basis for recognition by downstream factors. Identification of the latter is also an area of high current activity and a key objective of the SPP. The knowledge on mediators of downstream effects is very limited, and the same is true for phenotypes of null mutants at the cellular and organismal level.
One goal of the SPP is the identification of modification-binding factors by proteomic analysis, and structural characterization of such factors. Another goal is to develop suitable readout systems for functional analysis of synthetically modified nucleic acids. In selected cases, it seems realistic to connect the atomic details of a disturbed molecular interaction to a disease phenotype.
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[1] T. Carell, C. Brandmayr, A. Hienzsch, M. Müller, D. Pearson, V. Reiter, I. Thoma, P. Thumbs, M. Wagner, A ngew Chem Int Ed Engl 2012, 51, 7110–31.
[2] Y. Fu, C. He, Curr Opin Chem Biol 2012, 16, 516–524.
[3] M. Tahiliani, K. P. Koh, Y. Shen, W. A. Pastor, H. Bandukwala, Y. Brudno, S. Agarwal, L. M. Iyer, D. R. Liu, L. Aravind, et al., Science 2009, 324, 930–935.
[4] M. Munzel, D. Globisch, T. Carell, Angew Chem Int Ed Engl 2011, 50, 6460–6468.
[5] S. Schiesser, B. Hackner, T. Pfaffeneder, M. Muller, C. Hagemeier, M. Truss, T. Carell, Angew Chem Int Ed Engl 2012, 51, 6516–6520.
[6] T. Pfaffeneder, B. Hackner, M. Truss, M. Munzel, M. Muller, C. A. Deiml, C. Hagemeier, T. Carell, Angew Chem Int Ed Engl 2011, 50, 7008–7012.
[7] Y. F. He, B. Z. Li, Z. Li, P. Liu, Y. Wang, Q. Tang, J. Ding, Y. Jia, Z. Chen, L. Li, et al., Science 2011, 333, 1303–1307.
[8] S. Ito, L. Shen, Q. Dai, S. C. Wu, L. B. Collins, J. A. Swenberg, C. He, Y. Zhang, Science 2011, 333, 1300–1303.
[9] Y. Saletore, K. Meyer, J. Korlach, I. D. Vilfan, S. Jaffrey, C. E. Mason, Genome Biol 2012, 13, 175.
[10] J. Kiani, V. Grandjean, R. Liebers, F. Tuorto, H. Ghanbarian, F. Lyko, F. Cuzin, M. Rassoulzadegan, PLoS Genet 2013, 9, e1003498.
[11] B. El Yacoubi, M. Bailly, V. de Crecy-Lagard, Annu Rev Genet 2012, 46, 69–95.
[12] K. Miyauchi, S. Kimura, T. Suzuki, Nat Chem Biol 2013, 9, 105–111.
[13] W. E. Kowtoniuk, Y. Shen, J. M. Heemstra, I. Agarwal, D. R. Liu, Proc Natl Acad Sci U S A 2009, 106, 7768–7773.
[14] C. E. Dumelin, Y. Chen, A. M. Leconte, Y. G. Chen, D. R. Liu, Nat Chem Biol 2012, 8, 913–919.
[15] M. Machnicka, K. Milanowska, O. Osman Oglou, E. Purta, M. Kurkowska, A. Olchowik, W. Januszewski, S. Kalinowski, S. Dunin-Horkawicz, K. M. Rother, et al., Nucleic Acids Research 2013, 41, D262–7.
[2][16] C. G. Spruijt, F. Gnerlich, A. H. Smits, T. Pfaffeneder, P. W. Jansen, C. Bauer, M. Munzel, M. Wagner, M. Muller, F. Khan, et al., Cell 2013, 152, 1146–1159.
[17] L. Wang, S. Chen, K. L. Vergin, S. J. Giovannoni, S. W. Chan, M. S. DeMott, K. Taghizadeh, O. X. Cordero, M. Cutler, S. Timberlake, et al., Proc Natl Acad Sci U S A 2011, 108, 2963–2968.
[18] M. Funamizu, A. Terahara, A. M. Feinberg, K. Nakanishi, J Am Chem Soc 1971, 93, 6706–6708.
[19] D. W. Phillipson, C. G. Edmonds, P. F. Crain, D. L. Smith, D. R. Davis, J. A. McCloskey, J Biol Chem 1987, 262, 3462–3471.
[20] T. Suzuki, T. Wada, K. Saigo, K. Watanabe, EMBO J 2002, 21, 6581–6589.
[3][21] Y. G. Chen, W. E. Kowtoniuk, I. Agarwal, Y. Shen, D. R. Liu, Nat Chem Biol 2009, 5, 879–881.
[22] W. E. Cohn, J Biol Chem 1960, 235, 1488–1498.
[23] M. Charette, M. W. Gray, IUBMB Life 2000, 49, 341–351.
[24] S. Massenet, C. Branlant, RNA 1999, 5, 1495–1503.
[25] J. Li, Z. Yang, B. Yu, J. Liu, X. Chen, Curr Biol 2005, 15, 1501–1507.
[26] B. Yu, Z. Yang, J. Li, S. Minakhina, M. Yang, R. W. Padgett, R. Steward, X. Chen, Science 2005, 307, 932–935.
[27] T. Ohara, Y. Sakaguchi, T. Suzuki, H. Ueda, K. Miyauchi, Nat Struct Mol Biol 2007, 14, 349–350.
[28] D. Dominissini, S. Moshitch-Moshkovitz, S. Schwartz, M. Salmon-Divon, L. Ungar, S. Osenberg, K. Cesarkas, J. Jacob-Hirsch, N. Amariglio, M. Kupiec, et al., Nature 2012, 485, 201–206.
[29] K. D. Meyer, Y. Saletore, P. Zumbo, O. Elemento, C. E. Mason, S. R. Jaffrey, Cell 2012, 149, 1635–1646.
[30] J. Karijolich, Y.-T. Yu, Nature 2011, 474, 395–398.
[31] H. Wu, A. C. D’Alessio, S. Ito, Z. Wang, K. Cui, K. Zhao, Y. E. Sun, Y. Zhang, Genes Dev 2011, 25, 679–684.
[4][32] S. Kellner, J. Burhenne, M. Helm, RNA Biol 2010, 7, 237–247.
[33] I. Behm-Ansmant, M. Helm, Y. Motorin, J Nucleic Acids 2011, 2011, 408053.
[34] E. Y. Levanon, E. Eisenberg, R. Yelin, S. Nemzer, M. Hallegger, R. Shemesh, Z. Y. Fligelman, A. Shoshan, S. R. Pollock, D. Sztybel, et al., Nat Biotechnol 2004, 22, 1001–1005.
[35] M. Sakurai, T. Yano, H. Kawabata, H. Ueda, T. Suzuki, Nat Chem Biol 2010, 6, 733–740.
[36] B. E. Wulff, M. Sakurai, K. Nishikura, Nat Rev Genet 2011, 12, 81–85.
[37] J. Korlach, S. W. Turner, Curr Opin Struct Biol 2012, 22, 251–261.
[38] M. Schaefer, T. Pollex, K. Hanna, F. Lyko, Nucleic Acids Res 2009, 37, e12.
[39] J. E. Squires, H. R. Patel, M. Nousch, T. Sibbritt, D. T. Humphreys, B. J. Parker, C. M. Suter, T. Preiss, Nucleic Acids Res 2012, 40, 5023–5033.
[40] B. E. Maden, M. E. Corbett, P. A. Heeney, K. Pugh, P. M. Ajuh, Biochimie 1995, 77, 22–29.
[41] B. E. Maden, Methods 2001, 25, 374–382.
[42] A. Bakin, J. Ofengand, Biochemistry 1993, 32, 9754–9762.
[43] A. Bakin, J. Ofengand, Nucleic Acids Res 1995, 23, 3290–3294.
[44] F. M. Rottman, J. A. Bokar, P. Narayan, M. E. Shambaugh, R. Ludwiczak, Biochimie 1994, 76, 1109–1114.
[45] T. Bruckl, D. Globisch, M. Wagner, M. Muller, T. Carell, Angew Chem Int Ed Engl 2009, 48, 7932–7934.
[5][46] E. Cayama, A. Yepez, F. Rotondo, E. Bandeira, A. C. Ferreras, F. J. Triana-Alonso, Nucleic Acids Res 2000, 28, E64.
[47] R. Luhrmann, B. Appel, P. Bringmann, J. Rinke, R. Reuter, S. Rothe, R. Bald, Nucleic Acids Res 1982, 10, 7103–7113.
[48] T. W. Munns, M. K. Liszewski, J. T. Tellam, H. F. Sims, R. E. Rhoads, Biochemistry 1982, 21, 2922–2928.
[49] K. Miyauchi, T. Ohara, T. Suzuki, Nucleic Acids Res 2007, 35, e24.
[50] S. Kellner, L. B. Kollar, A. Ochel, M. Ghate, M. Helm, PLoS One 2013, 8, e67945.
[51] A. M. Giessing, F. Kirpekar, J Proteomics 2012, 75, 3434–3449.
[52] S. P. Russell, P. A. Limbach, J Chromatogr B Analyt Technol Biomed Life Sci 2013, 923-924, 74–82.
[53] S. Li, P. A. Limbach, Anal Chem 2012, 84, 8607–8613.
[54] M. Taucher, K. Breuker, Angew Chem Int Ed Engl 2012, 51, 11289–11292.
[55] W. Peters, S. Willnow, M. Duisken, H. Kleine, T. Macherey, K. E. Duncan, D. W. Litchfield, B. Luscher, E. Weinhold, Angew Chem Int Ed Engl 2010, 49, 5170–5173.
[56] T. Ritschel, P. C. Kohler, G. Neudert, A. Heine, F. Diederich, G. Klebe, ChemMedChem 2009, 4, 2012–2023.
[57] M. K. McDonald, E. J. Miracco, J. Chen, Y. Xie, E. G. Mueller, Biochemistry 2011, 50, 426–36.
[58] C. J. Spedaliere, J. M. Ginter, M. V Johnston, E. G. Mueller, J Am Chem Soc 2004, 126, 12758–12759.
[59] E. J. Miracco, E. G. Mueller, J Am Chem Soc 2011, 133, 11826–11829.
[6] [60] C. S. Nabel, R. M. Kohli, Science 2011, 333, 1229–1230.
[61] Y. Motorin, F. Lyko, M. Helm, Nucleic Acids Res 2010, 38, 1415–1430.
[62] N. Sasai, M. Nakao, P. A. Defossez, Nucleic Acids Res 2010, 38, 5015–5022.
[63] L. G. Acevedo, A. Sanz, M. A. Jelinek, Epigenomics 2011, 3, 93–101.
[64] Y. Fu, G. Jia, X. Pang, R. N. Wang, X. Wang, C. J. Li, S. Smemo, Q. Dai, K. A. Bailey, M. A. Nobrega, et al., Nat Commun 2013, 4, 1798.
[65] A. Tovy, R. Siman Tov, R. Gaentzsch, M. Helm, S. Ankri, PLoS Pathog 2010, 6, e1000775.
[66] C. T. Chan, Y. L. Pang, W. Deng, I. R. Babu, M. Dyavaiah, T. J. Begley, P. C. Dedon, Nat Commun 2012, 3, 937.
[67] C. Brandmayr, M. Wagner, T. Bruckl, D. Globisch, D. Pearson, A. C. Kneuttinger, V. Reiter, A. Hienzsch, S. Koch, I. Thoma, et al., Angew Chem Int Ed Engl 2012, 51, 11162–11165.
[68] F. Tuorto, R. Liebers, T. Musch, M. Schaefer, S. Hofmann, S. Kellner, M. Frye, M. Helm, G. Stoecklin, F. Lyko, Nat Struct Mol Biol 2012, 19, 900–905.