1Corresponding author: Mirko Hennig, Department of Biochemistry and Molecular Biology, Medical University of South Carolina, 173 Ashley Ave, PO Box 250509, Charleston, SC 29425; Email: hennig@musc.edu

RNA Structure Determination by NMR: Combining Labeling and Pulse Techniques

Braden M. Roth and Mirko Hennig1 Department of Biochemistry and Molecular Biology, Medical University of South

Carolina, Charleston, SC 29425

Abstract. RNA exhibits considerable structural and functional diversity beyond well established roles of ribosomal, transfer and messenger RNAs, as illustrated by the discovery of ever increasing numbers of diverse RNA structures involved in gene processing and regulation. RNA molecules are often quite flexible; they can function in a genuinely unfolded form and adapt for recognition of both the shape and the charge distribution on a potential ligand with exquisite specificity. Liquid state NMR spectroscopy is uniquely suited to answer important questions concerning biophysical properties of RNA molecules including their three-dimensional shape, secondary structure distribution, and flexibility by looking at dynamic ensembles of structures. Here we review the fields of RNA sample preparation and NMR methodology that facilitate the determination of RNA structure in solution.

Keywords. RNA synthesis, Isotope labeling, RNA purification, NMR, resonance assignment, structure determination.

Introduction

The increasing awareness of the essential role of RNA in biological processes, including its involvement in translation, gene regulation, and viral infections, make RNA an interesting target for structural studies. Structure is the link between sequence and function and thus knowledge of the three dimensional structure of RNA is central to understanding its biology. It is indispensable for describing the underlying determinants of catalytic mechanisms, ligand binding, and molecular recognition in macromolecular assemblies.

In this review, after a brief survey of the current status, we will provide the workflow employed by our laboratory emphasizing the factors essential for the determination of RNA structures in solution using high-resolution Nuclear Magnetic Resonance (NMR) methodology and the progress that has been made so far in these areas. The key issues to determine the structure of RNA at the atomic level are: how to produce sufficient amounts of isotopically labeled RNA, and once produced, how to purify the RNA; then, how to obtain unambiguous NMR resonance assignments of the RNA; and finally, how to utilize collected restraints in NMR structure determination. The review will conclude with a brief look at the future of high-resolution NMR in the study of the structural biology of RNA in solution.

1. Current Status of RNA Structures Solved by NMR

Despite the biological importance of RNA, knowledge of their structures lags considerably behind that of proteins. Nearly 98% of the transcriptional output of the human genome is non-(protein)-coding RNA (ncRNA) [1; 2]. This estimate is based upon the fact that intronic RNA constitutes 95% of primary protein-coding transcripts [3; 4]. Yet less than 2% of Protein Data Bank (PDB) structures represent RNA. This deficit lies largely in the difficulties in applying the most productive techniques in protein structure determination to RNA. Rugged energy landscapes and multistate RNA folding kinetics pose serious obstacles to crystallization for structure determination by X-ray crystallography. Coinciding with methodological developments in the early 1990s that allowed for stable isotope labeling, the rate of deposition into the PDB of RNA structures solved using NMR methodology increased more than threefold after 1995 to an average of 23 per year. Indeed, nearly half of the RNA-only structures in the PDB have been determined by NMR. Among the 337 unique NMR structures that have been deposited between 1992 and 2009, the average length is 23 nucleotides (nt). Only three RNA NMR structures contain more than 50 nt, of which one is a homodimeric, GAAA tetraloop-receptor RNA complex totaling 86 nt (see 4.5). However, recent years have seen fewer RNA solution structures deposited in the PDB (14 in 2008 and 15 in 2009, respectively), and the two largest NMR structures were solved several years ago, in 2003 and 2004 (PDB accession codes 1P5P and 1S9S, respectively). The pace of RNA structural biology in solution remains modest because severe spectral overlap and rapid relaxation in larger RNA continue to complicate structural studies by NMR. Nevertheless, NMR offers great potential not only with respect to structural characterization of RNA but also provides the unique ability to study the details of dynamics which appear integral to understanding RNA function. The use of deuteration and of TROSY (Transverse Relaxation-Optimized Spectroscopy)-based experiments have already played an important role in extending the size of RNA molecules amenable to solution NMR studies.

2. Sample Preparation

Given the inherent insensitivity of NMR, the preparation of RNA molecules for structure determination by NMR can be quite challenging. Commercially available oligonucleotides on the millimolar scale required by NMR are often cost-prohibitive, so investigators will generally employ large-scale in vitro transcription strategies to produce the molecules in-house. In addition to scale, several custom-labeled transcripts are often required to generate a single structure, making the process expensive and labor-intensive. Therefore, careful consideration must be taken to develop a research plan that maximizes the production of multiple large-scale transcripts while minimizing effort and expense. In the following sections, strategies for nucleotide preparation, in vitro transcription, and transcript purification are presented.

2.1. Nucleotide Synthesis

The enrichment of 13C and 15N-labeled nucleotides represents a critical step in the development of multidimensional heteronuclear NMR experiments for structure determination of RNAs (reviewed in [5; 6]). Assignment of larger RNAs; however,

requires alternative labeling strategies to combat spectral crowding [7; 8]. Site-specific and random deuteration in conjunction with 13C and 15N labeling has proven essential for the solution of RNAs larger than 30 kDa [9; 10]. At considerable expense, a variety of uniformly, isotopically labeled rNTPs are commercially available (e.g. Cassia LLC/Cambridge Isotope Labs and Isotec/Sigma-Aldrich). Most recently, site-specifically deuterated rNTPs became commercially available (Cassia LLC/Cambridge Isotope Labs). The ability to produce labeled rNTPs in-house is essential for the productivity of research groups with limited financial resources and/or an interest in developing new NMR methodologies. In the case of uniform 13C, 15N, and 2H labeling, the preparation of rNTPs follows protocols pioneered by the Pardi and Williamson laboratories [11-13]. Briefly, total RNA is extracted from high-density E. coli cultures grown in media with strictly defined carbon and nitrogen sources. The RNA is hydrolyzed with Nuclease P1 and applied to a boronate resin column to separate rNMPs from dNMPs. Enzymatic phosphorylation of purified rNMPs follows a two-step process in which rNMPs are converted to rNDPs via nucleotide-specific kinases. The reaction is coupled to an ATP-regeneration step that powers the production of triphosphates. Fully-charged rNTPs are re-purified by a boronate affinity column, removing salts and high molecular weight impurities that could interfere with efficient in vitro transcription.

Myokinase, guanylate kinase and nucleoside monophosphate kinase enzymes responsible for NMP-NDP conversion are purified commercially from varying sources, but can be expensive and unstable. Alternatively, nucleotide-specific monophosphate-charging enzymes can be cloned directly from bacteria. E. coli adk, cmk, gmk and pyrH genes fused to a His6-tagged, solubility-enhancing GB1 partner [14] are easily expressed in large quantities using standard expression and purification methods. In addition, the bacterial monophosphate kinases are more efficient than their commercial counterparts and are stably stored at −80 °C (Roth and Hennig, unpublished). Finally, costs can be further reduced by modifying the ATP regeneration/triphosphate charging reaction to instead utilize phosphocreatine and creatine kinase [15].

More exotic nucleotide labeling strategies (e.g. site-specific deuteration or 19F incorporation) require additional steps including the chemical synthesis of custom-labeled ribose [7; 16; 17] and/or coupling of an isotopically-labeled base to the sugar moiety [18-20], possibly requiring an additional UTP→CTP conversion as the final step. While some specifically-deuterated NTPs are commercially available, fluorinated rNTPs are not. The details of alternative rNTP labeling are discussed in section 3.

2.2. DNA-Template Directed Synthesis of RNA Using T7 RNA Polymerase

The production of millimolar quantities of RNA required for structural analysis by NMR is generally accomplished by large-scale in vitro transcription reactions. The preferred enzyme for this reaction is bacteriophage T7 RNA polymerase because it is better characterized than other common DNA-directed RNA polymerases (T3 and SP6) [21; 22]. There are two basic strategies for oligonucleotide synthesis using T7. Short RNA (<50 nt) transcriptions can be carried out using synthetic DNA templates. The template can be comprised of complementary DNA oligonucleotides or a top strand consisting of the 17-nt T7 promoter and a complementary strand that contains the T7 promoter and the transcript of interest [23]. The second strategy utilizes a double-stranded plasmid that consists of the T7 promoter, the desired RNA product, and a

suitable restriction enzyme sequence to linearize the plasmid and terminate transcription. This strategy is preferred because it is more efficient, produces fewer abortive transcripts, and facilitates subsequent purification of the relatively small transcript from the much larger DNA template [23].

Optimization of the transcription reaction prior to large-scale production is essential since the yield of in vitro transcribed RNA is dependent on many factors which are not fully understood. The most critical factors for efficient T7 transcription; however, are the MgCl2/rNTP ratio and T7 polymerase and DNA template concentrations. Duplicate small-scale (10−50 μl) transcriptions can be designed in a rational sparse matrix [24] to find optimal conditions chosen to promote total RNA yield (e.g. when using unlabeled NTPs) or to maximize transcript yield per mole of labeled nucleotides. After optimization, a 1 mL pilot transcription is recommended to verify predicted yields and to estimate the volume required for full-scale (millimolar) transcript production.

DNA template design must take into account two disadvantages of T7-directed RNA synthesis. First, an efficient reaction requires that T7 is primed by at least two guanosines as the initiating nucleotides of the transcript [21]. Second, T7 RNA polymerase can produce “add-on” (n+1, n+2) transcripts that are suboptimal for structure determination [21]. This 3'-end heterogeneity can be resolved through purification techniques or template design that incorporates a cis-acting ribozyme.

The hammerhead ribozyme is a small (<50 nt) catalytic RNA consisting of three stems and a conserved core that efficiently cleaves substrates containing an XUN motif, where X can be any base and N must be an unpaired A, C, or U. The most efficiently cleaved sequence is the GUC triplet [25]. The hammerhead can be used to hydrolyze RNA substrates in trans [26], or engineered as part of a cis-acting, self-cleaving RNA transcript [27]. Hydrolysis of the phosphodiester bond occurs after the GUC, resulting in a population of RNA transcripts with homogeneous 3'-ends. Critical to the activity of the ribozyme is the conserved three-way junction that forms the required base pairing around the GUC triplet (Figure 1A). Proper folding of the RNA-hammerhead transcript may require modifications of nucleotides at its 3'-end in order to base pair with the desired RNA sequence and promote the formation of a third stem. Therefore, it is imperative that the potential RNA-hammerhead constructs are subjected to an RNA folding algorithm such as Quikfold [28; 29] to confirm the proper formation of the cleavage junction (Figure 1A). Additionally, purification of the desired RNA from the hammerhead requires extra steps since superior resolution is necessary to separate similarly-sized transcripts. Finally, hammerhead transcriptions introduce extra cost because a sizeable portion of the synthesized transcript is discarded following purification of the target RNA. This consideration is important when preparing NMR-scale transcripts using expensive isotope-labeled nucleotides.

Figure 1. Overview and purification of a cis-hammerhead transcript. (A) Secondary structure prediction of a 3’ hammerhead construct. Base pairing between the target transcript (black) and hammerhead (grey) forms the third stem and positions the GUC triplet in the proper context for self-cleavage, releasing a target RNA with homogeneous 3’ ends. (B) Purification of a 22mer/hammerhead RNA analyzed by 8 M urea-PAGE. (Top) FPLC anion-exchange is insufficient to resolve the hammerhead (HH) and hairpin (HP) cleavage products. Subsequent HPLC anion-exchange results in purified 22 mer RNA for NMR analysis (bottom).

2.3. RNA Purification

Traditionally, milligram quantities of in vitro-transcribed RNA were purified by denaturing (8 M urea) PAGE followed by electroelution from the gel matrix, desalting, buffer exchange, and refolding [21; 30]. The distinct advantage of this method is the ability to separate preparative quantities of RNA with single-nucleotide resolution. On the other hand, gel-purification is a labor-intensive, RNA-denaturing process that yields transcripts contaminated with water-soluble acrylamide oligomers [23]. Although the impurities can be removed through extensive washing or dialysis, the process is time-consuming and incomplete. For these reasons, new methods of purifying large-scale transcription reactions have been developed, including anion-exchange [31; 32], size-exclusion [23; 33] and DNA affinity chromatography [34]. The RNA purification protocols adapted by our laboratory utilize components of these methods to produce NMR-quality transcripts with varying size and complexity under native conditions. Briefly, large-scale (5−15 mL) T7 run-off transcriptions are centrifuged to remove pyrophosphate precipitates, then buffer-exchanged with an appropriate centrifugal filter device. Unincorporated nucleotides and short abortive transcripts are also largely removed by this step. Removal of T7 polymerase, plasmid DNA template and the remaining abortive transcripts is achieved by FPLC anion exchange chromatography (GE HiTrapQ HP). Following high-salt elution, fractions corresponding to the expected transcript are analyzed for purity by denaturing PAGE and pooled (Figure 1B). Finally, the sample is exchanged back into the low-salt buffer for NMR analysis. This protocol yields high-quality RNA samples in <2 days with minimal sample loss, but it is dependent on first-rate optimization prior to full-scale transcription. While FPLC anion exchange is well-suited to purify discrete transcripts,

its resolution is limited. Where more stringent purification is required, the sample is exchanged into a buffer suitable for HPLC anion-exchange chromatography. The Dionex DNAPac provides nucleotide resolution of pre-purified RNAs and is ideal for the separation of contaminant transcripts and hammerhead cleavage products. Again, the resolved transcripts are verified by denaturing PAGE, pooled, buffer exchanged and quantified for NMR (Figure 1B). Unlike reversed-phase chromatography, pre-purification by FPLC is a simple, robust method of “cleaning” large-scale transcripts prior to DNAPac separation in a non-denaturing environment free of potentially problematic organic solvents.

3. Labeling Approaches

NMR structure determinations of RNA are simplified by the application of multi-dimensional, heteronuclear NMR experiments. However, these experiments require milligram quantities of isotopically labeled RNA, so the production of isotopically labeled RNA remains critical to the success of these NMR-based structure studies [5; 35]. In contrast to the abundant 1H and 31P isotopes, the naturally occurring nuclei 12C and 14N cannot be readily studied with high-resolution NMR techniques. Nucleotide-specific labeling schemes are compatible with RNA synthesis using T7 RNA polymerase and relatively straightforward because the four individual rNMPs can conveniently be separated by ion exchange HPLC chromatography. Thus, all labeling schemes described can be tailored to address specific assignment problems.

3.1. Conventional Labeling of RNA with 13C and 15N

Uniformly labeled rNTPs (GCN, CCN, ACN, UCN) for in vitro transcription reactions can be readily produced by phosphorylation of nucleotides isolated from bacteria grown on 13C- and/or 15N enriched media [11-13; 36]. Through the use of 13C and 15N isotopic labeling and multidimensional heteronuclear NMR experiments, studies of 15-kDa RNAs are commonplace and methodological developments have been reviewed [37-43].

3.2. Deuteration Strategies

The specific substitution of protons with deuterium represents the most promising way to increase sensitivity while simplifying spectra. Labeling schemes involving deuteration afford two benefits: first, the spectra will be less crowded and second, the relaxation properties of carbon and the remaining proton nuclei will be favorably altered. The reduced proton spin density decreases relaxation times of protons in RNAs. In addition, any 13C atoms attached to deuterium have slower relaxation properties relative to 13C−1H moieties. Therefore, deuterium labeling has particularly aided in the study of large RNA molecules. Adopting the enzymatic synthesis strategy of the Williamson laboratory, the following differentially deuterated ribonucleotides can be prepared for transcription reactions:

Beginning with 2H,13C-uniformly labeled glucose, the ribose portion of the nucleotides are deuterated at the H3', H4', H5', H5'' positions to give [1',2',3',4',5'-13C5,3',4',5',5''-2H4]-rNTPs which greatly simplify the NMR spectra for large RNAs

[17; 44]. In addition, pyrimidine bases with deuteration at the H5 position, 5-2H(pyrimidine)-rNTPs, can be included, removing the spectral crowding from strong H5/H6 crosspeaks. This labeling pattern conserves important NOEs between base and ribose while affording all the previously described benefits of deuteration. The H2' protons, which normally reside in the most crowded region of the proton spectrum, can be identified based on their chemical shift. If faster 1H T2-relaxation caused by 13C−1H dipole-dipole interactions presents a problem, uniformly 2H labeled glucose can be used to generate [12C5,3',4',5',5''-2H4(5-2H(pyrimidine))]-NTPs for transcription reactions.

Random fractionally deuterated nucleotides, 15N,13C-(50% 2H) rNTPs or GCN50%D, CCN50%D, ACN50%D, UCN50%D), are prepared by growth of M. methylotrophus on 15N-ammonium sulfate and 13C-methanol as sole carbon and nitrogen sources [36]. Alternatively, E. coli can be grown on minimal salt media containing 15N-ammonium sulfate and 13C-glucose. The growth is carried out in 50% D2O. Perdeuterated, 15N,13C,2H rNTPs (GCND, CCND, ACND, UCND) can be synthesized by growing bacteria in 100% D2O in minimal media, optionally supplemented with 15N-ammonium sulfate and/or 13C-acetate/methanol/glucose as sole nitrogen and carbon sources [45; 46]. Random fractional deuteration as well as perdeuteration is useful for NMR of large systems due to improved relaxation properties. Apart from C5' methylene ribose carbons, random fractional deuteration of RNA does not suffer from the production of isotopomers with chemical shift heterogeneity and decreased signal intensities. A general disadvantage of deuterium labeling is the reduced 1H-1H NOE based information content essential for structure determination.

3.3. Labeling of RNA with 19F

Fluorinated ribosomal 5S-rRNA [47] and tRNA [48] isolated from E. coli grown in the presence of 5F-U revealed high levels (>80%) of incorporation of 5F-UTP in place of UTP as early as 1977. Since this pioneering work, substantial effort has been made on the study of nucleic acids using both liquid- [15; 19; 49-51] and solid-state 19F NMR [52; 53]. Advantages of using spin 19F as an NMR probe are its high sensitivity (83% of 1H) combined with 100% natural abundance and a wide chemical shift distribution (ca. 50-fold larger than that of 1H) which typically leads to well-resolved signals in one-dimensional NMR spectra. The 19F chemical shift of a covalently bound fluorine atom is extraordinarily sensitive towards changes in the local microenvironment which is primarily attributable to the anisotropic distribution of the electrons in the 2-p orbitals. The van der Waals radius of a fluorine atom, 1.35Å, is only slightly larger than that of a hydrogen (1.2Å) and smaller than a methyl group (2.0Å) providing a promising candidate to substitute for either of those without structural or functional alteration. The introduction of 19F substitutions into the heterocyclic bases is non-perturbing and provides researchers with uniquely positioned, sensitive NMR reporter groups to monitor 1) conformation, 2) molecular interactions and 3) dynamics of RNA. As a result of drastically simplified spectra, 19F-NMR spectroscopy can provide very useful information on specific aspects of the structure, interactions, and mobility of RNAs that could not otherwise be obtained.

Fluorinated RNA can routinely be prepared by either phosphoramidite-based chemical synthesis [51] or in vitro transcription reactions using RNA polymerase. We

recently established the efficient and economical enzymatic synthesis of the fluorinated nucleotide analogues 5F-UTP, 5F-CTP [18; 19], and 2F-ATP [15]. The base analog 5F-uracil is readily used as a substrate for uracil phosphoribosyl transferase, which provides a novel and efficient route to produce 5F-UMP, and ultimately 5F-UTP. The fluorinated nucleotide analogues 2F-AMP and 5F-CMP can be synthesized by enzymatic conversions of 2F-adenine using adenine phosphoribosyltransferase and by conversion of the nucleoside analog 5F-cytidine catalyzed by uridine kinase. Our laboratory subsequently demonstrated that these fluorinated rNTP analogues can be specifically incorporated into RNA and that the modified nucleotide analogue 5F-Cyt selectively base-pairs with guanine [18; 19], 5F-Ura selectively base-pairs with adenine [18; 19], and 2F-Ade selectively base-pairs with uridine in a non-perturbing way [15] in individual samples containing one of these labeling schemes. Alternative, specific labeling of RNA with 19F at the 2'-position in the ribose can be problematic especially for regions adopting non-canonical conformations because of the conformational bias imposed by the fluorine substitution. 2'-Deoxy-2'-fluorinated nucleosides are effectively locked in the C3'-endo sugar pucker normally associated with A-form RNA geometry [54-56].

4. NMR Resonance Assignments

It is the limited chemical diversity in comparison to proteins that complicate NMR (and other) studies of RNA and its complexes. RNA secondary structure is trivial by comparison with proteins, being essentially the code of Watson-Crick base pairing: Guanine (Gua) pairs with Cytosine (Cyt); Adenine (Ade) pairs with Uracyl (Ura) (Figure 2). RNA mainly adopts A-form helical geometries, which contributes to chemical shift degeneracy and makes unambiguous assignments challenging for larger RNAs. A second problem in studies of RNA with molecular weights above 25 kDa is the fast decay of the NMR signal due to relaxation. Line widths in NMR spectra are inversely proportional to the relaxation rates. Therefore the signal-to-noise in NMR spectra of larger molecules is poor. The line width problem can be overcome by (1) the use of deuteration to eliminate proton mediated relaxation pathways and (2) the NMR method called TROSY [57].

4.1. Initial Characterization of Target RNA Constructs

Characterizing the conformation of an RNA under investigation is a crucial first step before conducting a detailed and time-consuming NMR analysis. At the outset, the suitability of the system for a high-resolution structure elucidation and optimal sample conditions for acquisition of the required NMR experiments should be determined. The imino proton region of the proton NMR spectrum of an unlabeled RNA sample in H2O provides a sensitive diagnostic for this purpose. One peak should be observed for each Watson-Crick and two for each G·U wobble base pair in the molecule. Since the imino protons exchange rapidly with the bulk H2O, we typically record jump-return echo experiments that avoid presaturation, while providing the most efficient water suppression [58]. The pyrimidine base protons can provide a valuable alternative, circumventing problems related to solvent exchange or conformational heterogeneity. H5-H6 cross peaks can be conveniently monitored in WATERGATE-2D TOCSY

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4.2. Assignment Strategy for Unlabeled RNA (< 25 nt)

Imino proton resonances are assigned sequence-specifically at the earliest stage from water flip-back, WATERGATE-2D NOESY (nuclear Overhauser effect spectroscopy) [77] spectra to verify the construct integrity and secondary structure predictions. The subsequent assignment of non-exchangeable RNA resonances is commonly achieved through identification of sequential base H8 and H6 to ribose H1' nuclear Overhauser effect (NOE) patterns seen in helical regions of nucleic acid structure, in analogy to the procedure originally utilized for DNA studies in the 1980s [78] (Figure 2). Complete ribose H2', H3', H4', H5', and H5" spin system identifications are hampered by limited dispersion (ca. 1ppm) and inefficient through-bond magnetization transfer from the better resolved H1' resonances in TOCSY and COSY (correlation spectroscopy) experiments. The 3J(H1',H2') vicinal coupling is ca. 1 Hz for the C3'-endo puckers, typically found in A-form helices [79-81]. However, a number of 1H,31P-multidimensional HETCOR (heteronuclear correlation) schemes are available for sequential assignment of 31P and to extended ribose 1H resonance assignments [61; 62]. The resulting two dimensional H3'/H5'/H5'',31P-correlations can be concatenated with homonuclear 1H,1H-NOESY or TOCSY experiments to transfer magnetization to potentially better resolved resonances like H1' or aromatic H8/H6 resonances [82; 83]. Gradient-selected, sensitivity-enhanced heteronuclear single-quantum correlation experiments (HSQC) [84; 85] recorded at natural 13C abundance greatly facilitate site-specific assignments owing to distinct ribose C1', C4', C5', Ade C2, Cyt C5, and Ura C5 carbon chemical shifts. Ribose C2' and C3' as well as base Cyt/Ura C6 and Gua/Ade C8 chemical shift ranges overlap yet can be distinguished from other carbon sites in RNA (Table 2).

The 2'-hydroxyl group plays fundamental roles in both the structure and the function of RNA and is the major determinant of the conformational and thermodynamic differences between RNA and DNA. In aqueous solution the rapid exchange of the hydroxyl proton with the solvent typically prevents its observation in RNA at room temperature by NMR. Recently, our laboratory reported assignments and a conformational analysis of 2'-OH groups of a medium sized RNA by means of TOCSY and NOESY experiments at low temperature [86; 87].

4.3. Assignment Strategy for 19F-labeled RNA

Using available chemical shifts from unmodified RNA samples, we have successfully assigned the 19F resonances for each 2F-Ade, 5F-Ura and 5F-Cyt in modified, medium-sized RNA samples. The unambiguous fluorine chemical shift assignments in substituted RNA can be accomplished using an approach that combines homo- and heteronuclear through-space 19F,1H-HOESY, 1H,1H-NOESY, and 19F,19F-NOESY with through-bond 1H,19F-HMBC (heteronuclear multiple bond correlation) experiments.

4.4. Assignment Strategy for Uniformly 13C,15N-labeled RNA (< 40 nt)

The NOE-based approach described in section 4.2 relies on assumptions about structure and assignments, rendering it susceptible to errors from structural bias. A methodology that achieves sequential assignment via unambiguous through-bond correlation experiments, as is the case for proteins, would be more ideal. Such through-bond NMR experiments are usually given a name that indicates the magnetization transfer route. A possible approach for 13C labeled RNAs is HCP correlation via sequential INEPT (Insensitive Nuclei Enhancement by Polarization Transfer) transfers [88; 89] (Figure 2, Table 2). Unfortunately, complete sequential assignments of even medium-sized RNA molecules using through-bond experiments such as HCP, HCP-TOCSY and HP-HETCOR are hampered by notoriously overlapped resonances and modest sensitivity. Thus, unambiguous through-bond assignment using HCP-like experiments is always difficult and impractical for larger RNA target molecules (> 40 nt).

Out of necessity, sequential assignments are therefore achieved using through-space 1H,1H contacts derived from 3D and 4D-isotope edited NOESY experiments. Fortunately, 13C and 15N enrichment permits a suite of through-bond experiments that help to alleviate the inherent ambiguity problem of through-space contacts (Figure 2). HCCH-COSY and HCCH-TOCSY experiments are used to unambiguously assign ribose spin systems [67-71] (Figure 2). We commonly employ a hybrid version, the HCCH-COSY-TOCSY [90; 91] to unambiguously assign crowded ribose spin systems. The HCN and HCNCH experiments can elucidate intranucleotide H6/H8-H1' correlations within and between the base and ribose resonances, significantly reducing the ambiguity present in the NOESY-based assignment procedure.

Canonical base-pair hydrogen bonding of the Watson-Crick type is fundamental in all biological processes where nucleic acids are involved. NMR experiments have been introduced that allow the direct identification of donor and acceptor nitrogen atoms involved in hydrogen bonds [92; 93]. The partially covalent character of hydrogen bonds gives rise to measurable scalar spin-spin couplings of the type h2J(N,N) and h1J(H,N) that represent important additional NMR parameters for the structure determination of nucleic acids in solution [92; 93]. In addition to the unambiguous determination of donor (D) and acceptor (A) nuclei involved in hydrogen bond formation, the magnitude of the hJ(D,A) couplings reports on the hydrogen bond geometry.

4.5. Assignment Strategy for Type-Specifically Labeled RNA (> 40 nt)

Despite the use of isotope labeling with 13C and 15N, resonance assignments of larger RNAs (> 40 nt) continue to be challenging. Above 50 nucleotides, through-bond experiments start to fail because of rapid transverse relaxation. In particular, the sensitivity of through-bond experiments correlating exchangeable and non-exchangeable base protons rapidly deteriorates due to long transfer times associated with small heteronuclear 1J(C,N) and multi-bond 2,3J(C,C) couplings. The following three case studies (Table 1) were chosen based on the important state-of-the-art technologies that were applied in conjunction with the molecular weight of the targeted RNAs.

Table 1. Large RNA (greater than 50 nucleotides) NMR structures in the PDB.

PDB accession Code No # of nucleotides (nt) Description Reference 1P5P 77 HCV IRES domain II [94] 1S9S 101 MMLV Encapsidation signal [10] 2ADT 86 GAAA tetraloop-receptor [9]

Figure 3. Large RNA structures determined by NMR. A) HCV IRES Domain II (26 kDa, PDB accession code 1P5P). B) GAAA Tetraloop Receptor Complex (30 kDa, PDB accession code 2ADT). C) Moloney Murine Leukemia Virus Encapsidation Signal (34 kDa, PDB accession code 1S9S).

4.5.1. Structure of HCV IRES Domain II (26 kDa)

Larger RNAs often necessitate a “divide and conquer” strategy. This approach assumes a modular architecture of the target RNA sequence without tertiary contacts between the smaller sub-domains (e.g. distant ends of an elongated stem-loop). Accordingly, the NMR solution structure of the 77-nt domain II of the hepatitis C virus (HCV) IRES (internal ribosome entry site) RNA was determined by Puglisi and coworkers (Figure 3A). The excision of two stably folded 34- and 55-nucleotide segments representing the entire domain II allowed for high-resolution structure determinations using conventional, uniformly 13C,15N-labeled RNA samples. Subsequently, the global conformation of domain II, a distorted L-shape, could be determined using a joint refinement procedure against residual dipolar coupling (RDC) restraints from both the two smaller segments as well as 60 RDCs determined for the large domain II construct. The inclusion of RDC restraints obtained from partially oriented RNA samples enabled precise definition of the relative orientation of the distant ends of the domain II RNA. Weakly aligned RNA samples were obtained by adding Pf1-phage as a cosolute. A full-length, intact 100 kDa HCV IRES was constructed by joining a 15N-labeled domain II segment and an unlabeled segment comprising domains III and IV using T4 RNA ligase [95]. A comparison of the imino 1H and 15N chemical shift data from the 64-nucleotide domain II with the segmentally labeled, intact 314-nucleotide IRES RNA supports the notion of an independently folded subdomain and suggests an indistinguishable structure in the larger context.

4.5.2. Structure of a GAAA Tetraloop Receptor Complex (30 kDa)

In an effort to address the striking lack of RNA tertiary structures in solution, Butcher and coworkers solved the NMR structure of a homodimeric, 30 kDa GAAA tetraloop-receptor RNA complex (Figure 3B). The twofold symmetry of this homodimeric arrangement considerably reduces the complexity of the NMR spectra. A total of seven nucleotide-type-specific RNA samples, GCNCCN, ACNUCN, GCNUCN, GCN, ACN, CCN, and UCN, were prepared in addition to a uniformly labeled RNA sample, GACUCN. Adiabatic frequency-swept inversion pulses optimized to an empirically observed relationship between 1J(C,H) and carbon chemical shift are commonly used in purge-filter elements to ensure the uniform inversion of all ribose and base carbon sites. Elegant 12C-filtered/13C-edited (100% D2O) and simultaneous 12C,14N-filtered/13C,15N-edited (90% H2O/10% D2O) 3D NOESY spectra have been proposed [96] where monitored 1H magnetization that starts at either 12C- or 14N-bound protons is transferred via isotropic mixing and is subsequently detected at 13C- or 15N-bound protons. The 86-nucleotide RNA-RNA homodimer exhibits unfavorable, faster T2-relaxation of both protons and carbons when compared to the 101-nucleotide core encapsidation signal described above. Due to these limitations, the authors relied on more sensitive 2D (rather than 3D or 4D) isotope-filtered/edited NOESY experiments for unambiguously identifying NOEs between labeled and unlabeled nucleotides [97; 98]. Remarkably, the 13C-filtered experiments suffered from 1H line broadening stemming from 13C−1H dipole-dipole interactions, necessitating a selective deuteration scheme for the ribose ring to complete resonance and NOE assignments. Here, specifically deuterated [3',4',5',5''-2H4(H5-2H(pyrimidine))]-NTPs were used which offer narrower H1'- and H2'-line width with the added benefit of reduced overlap originating from H5-base proton (pyrimidine) and H3', H4', H5', and H5''-ribose resonances [7]. RDC data was essential for accurate structure determination of the distant helical ends, but the authors also had to overcome sample precipitation issues in the presence of Pf1-phage by lowering the sample concentration, permitting measurement of only 24 1H−15N RDCs.

4.5.3. Structure of the Encapsidation Signal of the Moloney Murine Leukemia Virus (34 kDa)

An alternative strategy is to resort to nucleotide-specific labeling to facilitate the NMR data accumulation and accomplish complete assignment. Summers and coworkers were able to determine the solution structure of the 101 nucleotide Moloney Murine Leukemia Virus (MMLV), both free [10] (Figure 3C) and bound to the nucleocapsid (NC) domain of Gag [99]. This represents the largest nucleic acid NMR structure to date using a total of nine differently labeled RNA samples. Resonance assignments were obtained using a new strategy that relies heavily on isotope-filtered/edited NOESY experiments in combination with nucleotide specific isotopic labeling which allows for unambiguous, nucleotide-specific identification of spin systems. Four samples were generated using nucleotide-type-specific 13C,15N-labeled RNA samples, GCN, ACN, CCN, and UCN, respectively, to help distinguish intra- from internucleotide NOE connectivities in 3D and 4D 13C-edited NOESY experiments. In addition, four nucleotide-type-specific 1H-labeled RNA samples were generated using, GH, AH, CH, and UH, respectively, with remaining nucleotides being 90% perdeuterated, XD. Finally, the exchangeable imino proton assignments were confirmed using a 15N-labeled G and U sample, GNUN. The overall structure of the RNA consists of three stem-loops, two of which coaxially stack, while a short flexible linker connects the third stem-loop. A 15N-

relaxation analysis using the GNUN-sample revealed that one stem-loop segment tumbled independently from the other two in solution, mitigating to some extent the sensitivity issues associated with slowly tumbling, large RNA molecules. RDC data could be used only for refinement of the individual, isolated stem-loop segments because the intact RNA precipitated in the presence of Pf1-phage cosolute.

Table 2. NMR experiments for RNA resonance assignment.

Approximate RNA size Isotope labeling scheme NMR experimentsa Reference

< 25 nt unlabeled: (GACU)U

1H, 1H-COSY 1H, 1H-TOCSY 1H, 1H-NOESY

31P, 1H-COSY 31P, 1H-TOCSY

31P, 1H-TOCSYNOESY 1H, 13C-HSQC

[100] [101] [77]

[61; 102] [62]

[82; 83] [84; 85]

< 40 nt uniformly 13C,15N: (GACU)CN

1H, 13C-Constant time HSQC 1H, 13C-Constant time TROSY

1H, 15N-HSQC 1H, 15N-2J-HSQC

HCCH-COSY HCCH-TOCSY

HCCH-COSYTOCSY HCP PCH HCN

HCNCH HNN-COSY

HN(C)CH-TOCSY HC(C)NH-TOCSY

3D 13C-edited NOESY 4D 13C-edited NOESY 2D 15N-edited NOESY

[103] [104] [105] [76] [64]

[65; 66] [90; 91] [59; 88]

[102; 106] [75; 107-109]

[73; 110] [92; 93]

[111-113] [114-116]

[117] [118] [119]

< 50 nt

- type-specific labeling, e.g.: (GC)CN(AU)U and (GC)U(AU)CN

- site-specific deuteration:

(GCAU)D

1H, 13C-HMQC 1H, 15N-HMQC

1H, 15N-TROSY 12C,14N-filtered/13C,15N-edited

3D NOESY

[74] [74; 120]

[121] [96; 122; 123]

> 50 nt type-specific deuteration, e.g. (G)U(CAU)D

12C,13C-filtered/edited 12C,13C-edited/filtered 2D NOESY [97; 98]

aUnderlined experiments are preferably carried out in D2O.

5. Restraint Collection for Structure Determination

After sequence specific assignments of RNAs are obtained, the structure determination is traditionally based on collecting sufficient numbers of proton-proton distance restraints utilizing NOESY experiments. Potentially, the short distance restraints between pairs of protons can be complemented with torsion angle information accessible through J-coupling constants. Vicinal 3J scalar coupling constants can provide useful structural information about the sugar pucker, the β and ε backbone torsion angle conformations, as well as the glycosidic torsion χ which defines the orientation of nucleobases with respect to the sugar moiety (Figure 2).

In general, there is a practical difficulty in defining RNA structures precisely by NMR because traditional NOE and J-coupling based structure calculation relies on either short-range distance (< 6 Å) or local torsion angle information. The structural analysis of the RNA backbone conformation is complicated by the lack of useful 1H−1H NOE distance restraints available that define the backbone torsions. RNAs often are elongated structures which are better approximated as cylindrical rather than globular shapes. Thus there is a lack of NOE information between distant ends of the molecule, and as a result, the relative orientations of helical segments at opposite ends of the molecule are poorly defined. Recent advances in methodology have aimed to alleviate or overcome this shortcoming [124; 125]. Experiments to measure orientational, rather than distance dependent dipolar couplings, and cross-correlated relaxation rates have been developed providing additional structural information. RDC data not only provide additional information for a better definition of the global orientation of the segments with respect to each other but also carries valuable information on the dynamical properties of the RNA studied.

5.1. Proton-Proton Distance Restraints

NOEs provide distance restraints for pairs of hydrogen atoms. Only short proton-proton distances in the range < 6 Å are accessible through NOESY-type experiments. The intensity of NOESY cross peaks is approximately proportional to the inverse of the averaged distance to the power of six, <1/rij

6>, assuming an isolated pair of proton spins i and j.

For RNA NMR studies, NOE derived distance restraints are often determined semi-quantitatively and placed into four categories: Strong, medium, weak and very weak NOEs. A conservative approach sets all the lower bounds to 1.8 Å (van der Waals radius) with upper bounds ranging from 3.0 Å for the most intense NOEs to 7.0 Å for the weakest NOEs found in H2O experiments. Potential problems with interpretation of obtained NOESY cross peak intensities in terms of 1H−1H distances in structure calculations arise mainly from the phenomenon of spin diffusion. Spin diffusion causes a breakdown of the isolated spin pair approximation because nearby protons provide competing indirect pathways for observing the direct NOE between the two protons. Spin diffusion effects are significant at longer NOESY mixing times (> 100 ms) leading to damped direct-pathway cross peak intensities and resulting in underestimated interproton distances. Furthermore, multistep transfer pathways can result in false NOE assignments. However, in an early stage of the assignment procedure based on NOESY correlations, spin diffusion pathways can aid the identification of spin systems. Thus, for assignments it is recommended to analyze NOESY spectra acquired with a range of mixing times (ca. 50−200 ms).

5.2. Torsion Angle Restraints

J-coupling restraints can be implemented in two different ways during structure determination. They can be introduced qualitatively by restricting a torsion angle in a loose manner (± 30°) to one of the three staggered rotamers along the phosphodiester backbone or defining the preferred ribose sugar pucker such as C2'-endo or C3'-endo. Alternatively, vicinal J-couplings can be quantitatively related to a certain torsion angle

using semi-empirical Karplus relations of the form: 3J = A cos2θ + B cosθ + C, where θ is the intervening torsion angle [63; 126].

The quantitative analysis of scalar J-couplings, especially in the case of homonuclear 3J(H,H) couplings related to the ribose sugar pucker, becomes more and more difficult with increasing molecular weight and largely fails for RNAs larger than 40 nt. In contrast, the efficiency of cross-correlated relaxation pathways scales linearly with the overall correlation time of the molecule, which is related to its size. Cross-correlated relaxation rates have been introduced to high resolution NMR as a novel parameter for structure determination [127; 128] and allow the characterization of conformations for larger RNA molecules for which J-coupling analysis is not feasible anymore.

5.2.1. Sugar Pucker

The ribose sugar geometry is defined by five alternating torsion angles (ν0 through ν4, Figure 2). Usually, the ribose sugar adopts one of the energetically preferred C2'-endo (South) or C3'-endo (North) conformations. A number of 1H,1H and 1H,13C scalar couplings are available to determine the sugar pucker qualitatively with the combination of H1'-H2' and H3'-H4' coupling constants being the most useful for smaller RNAs. The 3J(H1',H2') vicinal coupling is > 8 Hz for C2'-endo puckers and ca. 1 Hz for the C3'-endo puckers, typically found in A-form helices [79; 80; 89]. The opposite behavior is expected for the 3J(H3',H4') coupling constant with C2'-endo puckers associated with small and the C3'-endo puckers associated with relatively large coupling constant values. An alternative is the measurement of cross-correlated relaxation rates between neighboring 13C−1H dipoles within the ribose ring to define the sugar pucker. For RNAs, cross-correlated relaxation rates can be measured using an experiment that belongs to the HCCH class, and precisely determine the ribose sugar pucker without the need of any empirical Karplus parameterization [129]. The resolution of this experiment can be further enhanced by adding a CC-TOCSY transfer [130].

5.2.2. γ Torsion Angle

Measurement of the γ torsion is difficult due to the need for stereospecific assignments of the H5' and H5'' proton resonances. In principle, the two-bond C4',H5'/H5'' couplings can be used in conjunction with the vicinal H4',H5'/H5'' couplings to define γ [81; 131].

5.2.3. Glycosidic Torsion Angle χ

The preferred orientation around χ in A-form helix is anti, which makes the base accessible for commonly found hydrogen bonding interaction. Two heteronuclear vicinal 1H,13C couplings contain useful information about the glycosidic torsion angle χ. The 3J(H1',C) couplings involving the C4,C8 carbons in purines and the C2,C6 carbons in pyrimidines, all depend on the χ torsion [81; 132]. We have shown that the magnitude of a five bond scalar 5J(H1',F)-coupling observed upon 5F-Ura and 5F-Cyt substitutions also depends on the glycosidic torsion angle χ [18]. Alternative applications have been published where the cross-correlated relaxation between two

13C−1H dipoles or a dipole and the glycosidic 15N CSA is utilized to collect information about the glycosidic torsion angle χ [133-135].

5.2.4. ε and β Torsion Angles

The ε and β torsions can be determined by measuring a variety of 13C,31P and 1H,31P scalar couplings. Some of these torsions may be measured directly in 2D 1H,31P heteronuclear HETCOR experiments [61; 102] and non-refocused 1H,31P HSQCs if the phosphorous and proton resonances are sufficiently resolved. However, both the ribose proton and phosphorus resonances involved are generally overlapped for even moderate size RNAs. Accurate measurements for 13C,31P and 1H,31P couplings can be obtained from both phosphorous-fitting of doublets from singlets [136] or spin echo difference experiments [137-141]. J-HMBC techniques can be applied to determine 3J(H,P) couplings [142]. A quantitative version of the HCP experiment allows for quantitation of 3J(C4',P) [143].

5.2.5. α and ζ Torsion Angles

The α and ζ torsions are not accessible by J-coupling measurements because the involved 16O nuclei have no magnetic moment. Some groups have used 31P chemical shifts as a guide for loose constraints on these torsions [144]; however, the correlation between 31P chemical shifts and the phosphodiester backbone conformation is not well understood in RNA. Cross-correlated relaxation rates have been employed to gain information on the α and ζ torsions. The cross-correlated relaxation between a ribose 13C−1H dipole and the 31P chemical shift anisotropy (CSA) carries valuable structural information about the phosphodiester conformation [145].

5.3. RDC Restraints

Several methods have been developed to create a slightly anisotropic environment for molecules tumbling in solution. This results in a small degree of alignment of the molecule, such that the dipolar couplings no longer average to zero, while retaining the quality of high-resolution NMR spectra. The most promising systems for NMR studies of partially aligned systems are dilute liquid crystalline bicelles [146] or Pf1 bacteriophage solutions [147; 148]. RDCs depend on the average value of an orientational function and the inverse cube of the distance, 1/r3, between the coupled nuclei. Two polar angles θ and φ characterize the orientation of the internuclear vector that connects coupled nuclei with respect to the principal axis system of the molecular alignment tensor A. Thus, for a directly bonded pair of nuclei with known distance, such as 1H−13C or 1H−15N in labeled RNA, angular restraints can be extracted from dipolar coupling data and incorporated during the structure calculation. Three experiments form the basis of our strategy in regard to RNA structure determination. To measure nucleobase 1DHC residual dipolar couplings, constant-time (CT)-TROSY and CT-anti-TROSY spectra [149; 150] are acquired in the absence (isotropic) and the presence (anisotropic) of Pf1 phage. 1DHC values are obtained by subtracting the isotropic values from the anisotropic J-couplings. To measure one-bond ribose 1' through 4' 1DHC, a J-modulated CT-HSQC experiment is acquired and analyzed as described [151]. Either a J-modulated 1H-15N-HSQC or a gradient-enhanced, interleaved inphase-antiphase (IPAP)-HSQC experiment provides additional 1DHN

RDC restraints [152; 153]. Field-induced alignment studies are also feasible for RNA and the obtained RDC data can complement data measured in the presence of external aligning media to resolve redundancies [154]. The measurement of independent sets of RDCs can provide a detailed view of both RNA structure and dynamics. A minimum of five RDCs must be measured per base or rigid sub-segment to establish segment specific order tensors, although in practice more are desirable to improve the precision to which individual As can be determined. A comparison of principal values of ordering can provide valuable information about relative motional amplitudes between segments. Measurements that allow one to obtain dipolar generalized order parameters SRDC (that are identical to order parameters derived from heteronuclear spin relaxation measurements) are sensitive to a much broader motional time scale (ps-ms, thus potentially covering biologically relevant μs-ms time scales) [155-158].

6. Structure Calculation

In the early stages of a project, we typically employ qualitative NOESY NMR data reporting on RNA secondary structure in combination with thermodynamic-based folding algorithms as implemented in the NMR-assisted prediction of secondary structure (NAPSS) to obtain accurate low-resolution structures [159]. Together with the initial characterization, this is most helpful in assessing the folding state of the target RNA.

Input data for RNA structure calculations include the previously introduced experimental restraints: NOE-based 1H−1H distance, torsion angle, cross-correlated relaxation rate, and RDC restraints. Direct experimental evidence for base paring interactions can be obtained through measurement of hJ(D,A) couplings and hydrogen bond restraints in form of donor-acceptor distances can be introduced. Non-experimental constraints include planarity restraints and conformational database potentials of mean force [160] that have been introduced to reproduce planar base pairs, torsion angle correlations, and sequential and nonsequential base-base interactions observed in RNA crystal structures. Such constraints must be applied with great care since they have no experimental basis; their inclusion introduces bias and apparent improvements in precision at the cost of imposing conformations that may not be present. Small angle X-ray scattering (SAXS) data can provide useful low resolution information on the global structural features of RNA-protein complexes facilitating the determination of overall dimensions, radius of gyration and shape of biomolecules [161-163]. The mutually complementary NMR and SAXS data serve to reduce angular degrees of freedom and to confine the translational degrees of freedom, respectively. Solution X-ray scattering data was successfully employed to refine the 30 kDa RNA−RNA complex described in section 4.5.3 [164].

Once a restraint set is assembled, we typically generate an initial NOE-derived structure for refinement with the RDC data by subjecting 100 random extended structures to a simulated annealing protocol using XPLOR-NIH [165] as described [166; 167]. Starting structures are calculated from randomized RNA coordinates using solely energy terms from holonomic constraints such as geometric and non-bonded terms. Torsion angle dynamics (TAD) as implemented in XPLOR and CNS prove to be robust and have a higher convergence rate with respect to molecular dynamics in

Cartesian coordinate space [168]. The ca. 25 lowest energy structures without NOE violations ≥0.5 Å are refined via a three step RDC-based protocol designed first to refine the local structure and then the global structure [169]. A new module has been developed for fitting SAXS data via XPLOR-NIH [162]. The lowest energy structures after simulated annealing and subsequent refinement against sets of RDCs and SAXS data are minimized using the AMBER module Sander [170]. Due to more adapted force fields, AMBER yields better and more consistent results for nucleic acids [171].

A number of statistics are commonly evaluated to judge the convergence and quality of the family of calculated RNA NMR structures: Root Mean Square Deviation (RMSD), number of NOE, RDC, and torsion restraints; residual distance, dipolar coupling, and torsion violations; and the largest distance, dipolar coupling, and torsion violations. Typically, the distance restraints are further dissected into the number of interresidue, intraresidue, and intermolecular NOEs. The NMR input data have to be satisfactorily reproduced in high quality structures. Thus, the process of NMR assignment and restraint collection and subsequent structure calculation is iterative; in the process, wrong assignments will be corrected and additional restraints may be identified. Useful RMSDs to consider include only regions of interest and are usually a more accurate descriptor of the quality of the structure than the overall global RMSD. Local RMSDs are given because the overall global value can easily be in the 2.0−3.0 Å range, which might otherwise be indicative of poor convergence. Most RNA structures studied include poorly defined regions such as a disordered loop, terminal base pairs, or a nucleotide lacking internucleotide NOEs.

Finally, the obtained structures are validated by carrying out structure calculations omitting a randomly chosen subset of the RDC data while refining against the remaining RDCs [172]. The accuracy of a family of RNA NMR structures is cross-validated by the agreement between the back-calculated RDCs derived from the structures and the omitted RDC subset. Alternatively, a comparison between calculated and observed 1H chemical shifts represents another possibility for cross-validation of structures derived from NMR restraints [173].

7. Future Perspectives

7.1. Segmental Labeling of RNA.

Resonance overlap caused by the limited chemical and structural diversity presents an inherent barrier to investigate larger RNA in solution by NMR. As the number of resonances increases, even uniform or nucleotide-type specific labels are not sufficient to provide the necessary spectral dispersion. The isotopic labeling of RNA segments, facilitated by T4 RNA ligase, simplifies the spectral complexity by reducing the number of NMR-active atoms. T4 RNA ligase catalyzes the ligation of single-stranded RNA or DNA to either oligoribo- or oligodeoxyribonucleotides through the formation of a standard 3'→5' phosphodiester bond with hydrolysis of ATP to AMP and PPi [174; 175]. An elegant and economic approach using in vitro transcription and hammerhead ribozyme cleavage for generating complementary labeling schemes has recently been described [95; 176]. The incorporation of isotopically labeled RNA fragments into the middle of the full-length target RNA molecule necessitates a three-way ligation; such a

multiple segmental labeling of RNA with three segments has recently been demonstrated [177].

RNA structure determination remains a key challenge and represents the next frontier to modern structural biology. The dominance of ncRNAs in the genomic output of higher organisms suggests that they are not simply occasional transcripts with peculiar structure and function, but rather that they may constitute an extensive but hitherto incompletely characterized regulatory network within higher organisms. A decade of structural genomics dramatically increased the database of known protein structures by developing and applying methodologies to determine them as rapidly and cost-effectively as possible. To date, though conceptually conceived a decade ago [178], the primary mission of high-throughput determination of RNA structures has not been seriously undertaken. Nevertheless, a tremendous amount of exciting research is currently underway. Improvements in the fields of RNA NMR methodology and biochemistry, paired with technical advances in NMR instrumentation have paved the way for more streamlined structural efforts targeting RNA in the future.

Acknowledgments

The authors gratefully acknowledge past and present members of the Hennig laboratory for helpful discussions, Drs Christina Mozes and Brendan Duggan for critical reading of the manuscript, and Dr Brendan Duggan for PDB data base mining. This work was supported by funding from the National Institutes of Health (AI064307, AI081640 and RR024442) and by the National Science Foundation (NSF 0845512).

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