Solid-Phase Chemical Synthesis of Stable Isotope-Labeled RNA to Aid Structure and Dynamics Studies by NMR Spectroscopy

RNA structure and dynamic studies by NMR spectroscopy suffer from chemical shift overlap and line broadening, both of which become worse as RNA size increases. Incorporation of stable isotope labels into RNA has provided several solutions to these limitations. Nevertheless, the only method to circumvent the problem of spectral overlap completely is the solid-phase chemical synthesis of RNA with labeled RNA phosphoramidites. In this review, we summarize the practical aspects of this methodology for NMR spectroscopy studies of RNA. These types of investigations lie at the intersection of chemistry and biophysics and highlight the need for collaborative efforts to tackle the integrative structural biology problems that exist in the RNA world. Finally, examples of RNA structure and dynamic studies using labeled phosphoramidites are highlighted.

Isotope labeled RNAs are predominantly prepared by in vitro transcription with T7 RNA polymerase (T7 RNAP) using well established protocols [30,31]. This approach is applicable to large RNA sequences and allows for uniform or nucleotide-specific labeling by mixing labeled with unlabeled ribonucleoside-5 -triphosphates (rNTPs). Although this method produces pure material in quantities amenable to NMR analysis (~0.1-1 µmol), signal overlap presents a significant problem for large RNAs. Moreover, T7 RNAP requires one or more guanosine (G) or adenosine (A) residues at the 5 -end for efficient in vitro RNA transcription [30][31][32]. In some cases, the addition of the 5 -G overhang(s) can alter the function of the RNA [33]. Additionally, the first 3 nucleotides (nts) at care must be taken when choosing these sequences [34]. To overcome the non-specificity of T7 RNAP-based in vitro transcription, three methods to prepare position-specifically labeled RNAs have emerged. In the first, Keyhani et al. developed a chemo-enzymatic synthesis that involves a single incorporation of a modified nucleoside-3′,5′-bisphosphate to the 3′-end of an RNA fragment followed by DNA-splinted ligation to complete the desired internally labeled RNA sequence [19]. This method was used to introduce photocaged, photoswitchable, and isotopically labeled nucleosides into RNAs up to 392 nts. Furthermore, this method uses standard laboratory equipment and commercially available enzymes T4 RNA ligase 1, shrimp alkaline phosphatase, and T4 RNA ligase 2, making it readily accessible to most biochemical/biophysical research groups. However, the relatively low yields from both bis-phosphorylation of the modified nucleosides (6%-22%) and DNA-splinted ligation of the RNA fragments (9%-49%) are a major drawback of this approach.
Secondly, Liu et al. developed a hybrid solid-liquid phase transcription technique using an automated robotic platform known as PLOR (position selective labeling of RNA) to prepare isotopically labeled RNAs up to 104 nts and fluorescently and heavy atom labeled RNAs up to 71 nts [35][36][37]. In this approach, the DNA template is attached to a bead and transcription is initiated by the addition of T7 RNAP and a mixture missing one (or more) of the four rNTPs, causing RNA elongation to stall at the first position where the missing rNTP would be incorporated. The beads are then washed to remove unincorporated rNTPs and T7 RNAP. Elongation is then resumed by the addition of a new mixture containing the previously omitted rNTP. Repetition of the elongation, stalling and reinitiation steps enables synthesis of position-selectively labeled RNA. The main drawbacks of this method are the inaccessibility of the automated synthesizer and the need for large, stoichiometric amounts of T7 RNAP and DNA template. Additionally, a nucleotide within a stretch of the same residues (e.g., an A in a GAAA tetraloop) cannot be individually labeled with PLOR. Lastly, at least the first ~10 nts of the RNA sequence must be composed of only 3 of the 4 residues to ensure proper formation of the processive elongation complex and subsequently stall it. Figure 1. Schematic of the utility of using isotope labeled RNA phosphoramidites and solid-phase synthesis to investigate RNA structure and dynamics. This methodology incorporates labels site-specifically into RNA, resolving NMR spectral overlap and ambiguities in NMR spectra interpretation. Magenta circle indicates a 13 C atom, D indicates a 2 H atom, and PG refers to a 2′-OH protecting group.
The final method, solid-phase chemical synthesis of RNAs with nucleoside phosphoramidites, is the main focus of this review. This approach allows for position-specific labeling that greatly reduces NMR spectral complexity and simplifies resonance assignment ( Figure 1). Furthermore, solid-phase synthesis has no sequence requirements, unlike T7 RNAP and PLOR. Despite these advantages, solid-phase synthesis is currently limited to labeling RNAs less than 76 nts [38]. Fortunately, many biologically relevant RNAs fall within this size limit [6]. For RNAs larger than 76  Figure 1. Schematic of the utility of using isotope labeled RNA phosphoramidites and solid-phase synthesis to investigate RNA structure and dynamics. This methodology incorporates labels site-specifically into RNA, resolving NMR spectral overlap and ambiguities in NMR spectra interpretation. Magenta circle indicates a 13 C atom, D indicates a 2 H atom, and PG refers to a 2 -OH protecting group.

The Phosphoramidite Method
The solid-phase chemical synthesis of RNA via the phosphoramidite method is a four-step process that is carried out in an automated synthesizer ( Figure 2A) [40,41]. Although originally developed for DNA synthesis by Beaucage and Caruthers, the phosphoramidite method has since been adapted for RNA synthesis [42]. This strategy utilizes the reaction between an activated nucleoside phosphoramidite and a solid-support bound nucleoside to synthesize the desired RNA sequence in the 3 to 5 direction ( Figure 2B). During the first step, the 4,4 -dimethoxytrityl (DMTr) protecting group at the 5 -hydroxyl (OH) of the tethered 3 -nucleoside is removed with trichloroacetic acid or dichloroacetic acid. The nucleophilic 5 -OH then attacks and removes the activated phosphoramidite to couple the two nucleosides with a phosphite triester bond. To minimize the amount of undesired RNA sequences, the unreacted tethered 3 -nucleosides are capped by acetylation of the 5 -OH groups. The final step is the oxidation of the phosphite triester to a phosphotriester with either iodine (I 2 ) or tert-butylhydroperoxide. After the entire sequence has been synthesized, the RNA is deprotected and cleaved from the solid-support. This strategy provides a powerful alternative to other methods of producing synthetic RNAs such as enzymatic ligation [43][44][45][46]. labeling strategies require a combination of solid-phase and chemo-enzymatic synthesis, RNA ligation, and T7-RNAP-based transcription.

The Phosphoramidite Method
The solid-phase chemical synthesis of RNA via the phosphoramidite method is a four-step process that is carried out in an automated synthesizer ( Figure 2A) [40,41]. Although originally developed for DNA synthesis by Beaucage and Caruthers, the phosphoramidite method has since been adapted for RNA synthesis [42]. This strategy utilizes the reaction between an activated nucleoside phosphoramidite and a solid-support bound nucleoside to synthesize the desired RNA sequence in the 3′ to 5′ direction ( Figure 2B). During the first step, the 4,4′-dimethoxytrityl (DMTr) protecting group at the 5′-hydroxyl (OH) of the tethered 3′-nucleoside is removed with trichloroacetic acid or dichloroacetic acid. The nucleophilic 5′-OH then attacks and removes the activated phosphoramidite to couple the two nucleosides with a phosphite triester bond. To minimize the amount of undesired RNA sequences, the unreacted tethered 3′-nucleosides are capped by acetylation of the 5′-OH groups. The final step is the oxidation of the phosphite triester to a phosphotriester with either iodine (I2) or tert-butylhydroperoxide. After the entire sequence has been synthesized, the RNA is deprotected and cleaved from the solid-support. This strategy provides a powerful alternative to other methods of producing synthetic RNAs such as enzymatic ligation [43][44][45][46]. coupling of residue n-1 and n shown. PG refers to the 5′-OH protecting group, which is generally DMTr. PG-2 refers to the 2′-OH protecting group, which comes in many varieties, as discussed in this review. Additional cycles can be added to the synthetic cycle to grow the RNA polymer from 3′-5′ (B).
The efficiency of the chemical synthesis of RNA by the phosphoramidite method depends on the protecting groups used, specifically at the exocyclic amino groups of the nucleobase (A, C, and G), and the 2′-and 5′-OH functionalities of the ribose sugar ( Figure 3) [40]. Typically, the 5′-OH group is protected with DMTr and the nucleobase exocyclic amino groups are protected with acetyl (Ac), phenoxyacetyl (Pac), benzoyl (Bz), or isobutyryl (iBu) groups ( Figure 3). In contrast to the 5'-OH and exocyclic amino protecting groups originally developed for DNA, the 2′-OH protecting  The efficiency of the chemical synthesis of RNA by the phosphoramidite method depends on the protecting groups used, specifically at the exocyclic amino groups of the nucleobase (A, C, and G), and the 2 -and 5 -OH functionalities of the ribose sugar ( Figure 3) [40]. Typically, the 5 -OH group is protected with DMTr and the nucleobase exocyclic amino groups are protected with acetyl (Ac), phenoxyacetyl (Pac), benzoyl (Bz), or isobutyryl (iBu) groups ( Figure 3). In contrast to the 5'-OH and exocyclic amino protecting groups originally developed for DNA, the 2 -OH protecting groups are unique to RNA. The 2 -OH protecting groups must: (1) be stable under all reaction conditions (acidic conditions of de-tritylation, basic conditions of deprotection of the nucleobase and cleavage from the solid-support), (2) be quantitatively removed under mild conditions that do not degrade the RNA, and (3) not interfere with the phosphoramidite coupling reaction [40,[47][48][49]. In the next section, we briefly summarize the commonly used 2 -OH protecting groups along with their comparative advantages and disadvantages. groups are unique to RNA. The 2′-OH protecting groups must: (1) be stable under all reaction conditions (acidic conditions of de-tritylation, basic conditions of deprotection of the nucleobase and cleavage from the solid-support), (2) be quantitatively removed under mild conditions that do not degrade the RNA, and (3) not interfere with the phosphoramidite coupling reaction [40,[47][48][49]. In the next section, we briefly summarize the commonly used 2′-OH protecting groups along with their comparative advantages and disadvantages.

Choice of RNA Phosphoramidite 2'-OH Protecting Groups
There is currently no consensus RNA 2′-OH protecting group, but the available groups can be broadly categorized into acid-, photo-, and fluoride-labile groups [40,[50][51][52][53][54]. For generating isotope labeled RNAs, the fluoride-labile 2′-OH protecting groups tert-butyldimethylsilyl (tBDMS), [(triisopropylsilyl)oxy]methyl (TOM), and 2′-cyanoethoxymethyl (CEM) group are preferred. Despite the stability under acidic conditions and ease of addition and removal, tBDMS suffers from a low coupling efficiency, long coupling time, and migration from the 2′-to 3′-OH during RNA synthesis [40]. Despite its limitations, 2′-O-tBDMS phosphoramidites have been successfully used to synthesize labeled RNAs up to 53 nts [55]. Compared to tBDMS, TOM has shorter coupling times (~6 min) and does not migrate from 2′-to 3′-OH [40]. With TOM, labeled RNAs as large as 55 nts have been successfully prepared [56]. The CEM group has a high coupling efficiency (~99%), short coupling time (2-4 min), and minimal steric hindrance that allows synthesis of RNAs as large as 170 nts [57,58]. In terms of labeled RNA, sequences up to 76 nts have been successfully prepared for NMR analysis with 2′-O-CEM phosphoramidites [38]. Two other 2′-OH protecting groups that have not yet been applied to RNA labeling are the photolabile 2-nitrobenzyloxymethyl (NBOM) group and the acid-labile bis(acetoxyethoxy)methyl ether (ACE) group [47,53,54]. The NBOM group is stable under acidic and basic conditions, has a short coupling time (2 min), and high coupling efficiency. However, its removal is not quantitative [40]. While the ACE group has a short coupling time (1 min) and is capable of generating RNAs in high yield and purity, it is acid labile and therefore not compatible with 5′-DMTr deprotection. Thus, fluoride-labile 5′-OH protecting groups are required and the synthesizer has to be modified to handle triethylamine/trihydrofluoride (TEA-3HF) as a deprotection agent [40]. Nevertheless, the ACE group is commercially successful at

Choice of RNA Phosphoramidite 2'-OH Protecting Groups
There is currently no consensus RNA 2 -OH protecting group, but the available groups can be broadly categorized into acid-, photo-, and fluoride-labile groups [40,[50][51][52][53][54]. For generating isotope labeled RNAs, the fluoride-labile 2 -OH protecting groups tert-butyldimethylsilyl (tBDMS), [(triisopropylsilyl)oxy]methyl (TOM), and 2 -cyanoethoxymethyl (CEM) group are preferred. Despite the stability under acidic conditions and ease of addition and removal, tBDMS suffers from a low coupling efficiency, long coupling time, and migration from the 2 -to 3 -OH during RNA synthesis [40]. Despite its limitations, 2 -O-tBDMS phosphoramidites have been successfully used to synthesize labeled RNAs up to 53 nts [55]. Compared to tBDMS, TOM has shorter coupling times (~6 min) and does not migrate from 2 -to 3 -OH [40]. With TOM, labeled RNAs as large as 55 nts have been successfully prepared [56]. The CEM group has a high coupling efficiency (~99%), short coupling time (2-4 min), and minimal steric hindrance that allows synthesis of RNAs as large as 170 nts [57,58]. In terms of labeled RNA, sequences up to 76 nts have been successfully prepared for NMR analysis with 2 -O-CEM phosphoramidites [38]. Two other 2 -OH protecting groups that have not yet been applied to RNA labeling are the photolabile 2-nitrobenzyloxymethyl (NBOM) group and the acid-labile bis(acetoxyethoxy)methyl ether (ACE) group [47,53,54]. The NBOM group is stable under acidic and basic conditions, has a short coupling time (2 min), and high coupling efficiency. However, its removal is not quantitative [40]. While the ACE group has a short coupling time (1 min) and is capable of generating RNAs in high yield and purity, it is acid labile and therefore not compatible with 5 -DMTr deprotection. Thus, fluoride-labile 5 -OH protecting groups are required and the synthesizer has to be modified to handle triethylamine/trihydrofluoride (TEA-3HF) as a deprotection agent [40]. Nevertheless, the ACE group is commercially successful at preparing RNAs in large quantities even with modified nucleosides (e.g., inosine, N 6 -methyladenosine, pseudouridine, 2'-fluoro-A,U,G,C) [47,53].
Given the advantages and disadvantages of each 2 -OH protecting group, the choice of which to use typically depends on the downstream application. However, few 2 -OH protecting groups are capable of making RNA in high enough quantity and purity for NMR spectroscopy. As will be discussed, 2 -O-tBDMS and 2 -O-TOM phosphoramidites have been used to great success to study RNA structure and dynamics by NMR spectroscopy [56,[59][60][61][62]. However, phosphoramidite synthesis with these protecting groups is limited to RNAs less than 60 nts. Nevertheless, unlabeled 2 -O-tBDMS and 2 -O-TOM phosphoramidites are commercially available, so researchers only need to synthesize isotope labeled phosphoramidites in-house, dramatically simplifying sample preparation. The commercially unavailable 2 -O-CEM is the only protecting group that can accommodate larger RNAs, but both unlabeled and labeled phosphoramidites must be synthesized in-house. In practice, 2 -O-tBDMS phosphoramidites are used for NMR structure and dynamics studies of RNAs prepared by solid-phase synthesis, unless the RNA under investigation is greater than 60 nts. Still, very few research groups have the resources (organic laboratory and solid-phase synthesizer) and expertise (NMR spectroscopy) necessary to fully exploit these technologies. One goal of this review is to motivate the collaboration of organic chemists with biophysicists to unlock the power of this method to study RNA structure, dynamics, and function.

Stable Isotope Labeling of RNA Phosphoramidites
In principle, any method to incorporate isotope labels into nucleobases, nucleosides, or rNTPs can be converted into phosphoramidites, using one or more of the synthetic routes described herein. The diverse methodologies to generate such labeled molecules were recently reviewed [20]. Once synthesized, phosphoramidites are characterized by 1 H/ 13 C/ 31 P NMR and mass spectrometry. The latter technique is complicated by the acid-labile nature of the phosphoramidites, which requires specialized electro-spray ionization techniques in non-aqueous buffers with lithium chloride for accurate mass determination [63,64]. Several groups have developed synthetic strategies to obtain 13 C/ 15 N-labeled RNA phosphoramidites with a variety of 2 -OH protecting groups [38,39,55,56,[59][60][61][62][65][66][67][68]. This section will highlight the many efforts employed to synthesize labeled RNA phosphoramidites.
From isotope labeled nucleobases [25][26][27]38,55,60,77], the groups of Kreutz and Micura prepared 2 -O-tBDMS and 2 -O-TOM phosphoramidites for NMR studies [59][60][61][62]. For example, uracil was synthesized, following procedures developed by Santa Lucia and Tinoco [77]. First, 13  TOM labeling protocols. Taken together, many stable isotope labeled RNA phosphoramidites have been synthesized with diverse labeling patterns. The next section is devoted to highlighting a few examples of how these labels can be exploited to study RNA structure and dynamics by NMR spectroscopy.

Facile RNA Resonance Assignments and Base Pair Interactions
An unparalleled advantage of phosphoramidite solid-phase chemical synthesis over in vitro transcription is the ability to make RNA with position-specific isotope labels that dramatically simplifies unambiguous resonance assignments (Figure 1). For example, a 40 nt RNA can be assigned with 40 samples, each containing one of the 40 residues labeled. While assigning all RNA resonances with this methodology is too cumbersome for most groups, phosphoramidites are a great tool to confirm assignments from orthogonal NMR experiments or to resolve overlapped peaks in a given NMR spectra. In addition to assigning RNA resonances, various research groups have utilized 13 C/ 15 N-labeled phosphoramidites to monitor both Watson-Crick and non-canonical base pairs in RNA [56,62].
Incorporation of 15 N-labels into RNA provides information on both canonical Watson-Crick and non-canonical base pairs in RNA [56,62,[78][79][80]. [1,NH 2 -15 N 2 ]-and [2-13 C-1,NH 2 -15 N 2 ]-guanosine were position-specifically incorporated into short (8-12 nts) RNA fragments to compare spectral properties of GU to GC base pairs [79]. Similarly, [2-13 C-1,NH 2 -15 N 2 ]-guanosine and [7,NH 2 -15 N 2 ]-adenosine were position-specifically incorporated into short (8 nts) RNAs to study face-to-face and sheared GA base pairs [78]. Additionally, [2-13 C-1,7,NH 2 -15 N 3 ]-guanosine and [8-13 C-1,7,NH 2 -15 N 3 ]-adenosine were incorporated into loop A and B domains (14-24 nts) from the hairpin ribozyme found in the satellite RNA of tobacco ringspot virus [80]. In these examples, the 13 C isotopes were used as tags to distinguish between the residues in doubly labeled samples since 1 J H2C2 or 1 J H8C8 splits the 1 H resonances for residues that contain 13 C directly coupled to 15 N. Wenter and Pitsch incorporated [1-15 N]-adenosine and -guanosine and [3-15 N]-uridine and -cytidine labels into 5 sites of the 32 nt bistable RNA [65]. An HNN correlation spectroscopy (COSY) experiment was used to correlate the hydrogen bond donor and acceptor N-atoms within a base pair formed by two 15 N-labeled residues [65,81,82]. With this approach, Wenter and Pitsch were able to unambiguously assign each HNN correlation to a particular base pair within one of the two co-existing conformers [65]. Neuner  To date, two high-resolution NMR structures have been solved using position-specific labeled RNAs by solid-phase synthesis [83,84]. Traditional NMR assignment strategies are optimized for largely helical RNAs and often fail for non-canonical tertiary structural regions. Wolter et al. solved the structure of a 34-nt GTP-binding RNA aptamer that forms exotic structural features including a trans A:A base pair, a novel GACG quartet, a protonated adenine, and a GCA base triplet [83].

Incorporating Isolated Spin-Pairs for Artifact Free RNA Dynamic Probing
In addition to NMR structural experiments, isotope labeled phosphoramidites have been used with great success to study RNA dynamics [39,55,[59][60][61]. The isolated 1 H-13 C spin-pair of the  19 51

Incorporating Isolated Spin-Pairs for Artifact Free RNA Dynamic Probing
In addition to NMR structural experiments, isotope labeled phosphoramidites have been used with great success to study RNA dynamics [39,55,[59][60][61]. The isolated 1 H-13 C spin-pair of the [6-13 C]-pyrimidine and [8-13 C]-purine phosphoramidites remove the 13 C-13 C scalar and dipolar coupling interactions and thus facilitate straightforward probing of functional dynamics in RNA. Strebitzer and Nußbaumer et al. demonstrated the potential of atom-specific 13 C-labeling to probe conformational dynamics in the ε-element of the duck Hepatitis B virus RNA (dHBVε) [55]. Wijmenga and co-workers previously showed that an RNA construct that represented the upper stem undergoes conformational fluctuations from the fast (pico-and nanosecond) to intermediate (microsecond) timescale [86]. To cross-validate this previous study, a dHBVε construct was made with [2,8-13 C 2 ]-adenosine incorporated at residues A20, A22, and A41. The kinetics of the unfolding process (k ex = 1522 ± 128 s -1 ) were quantified via a 13 C-CPMG relaxation dispersion (RD) experiment utilizing the 13 C-adenosine labels ( Figure 5A). In addition to 13 C RD experiments, the absence of nearby protons makes purine 1 H8-13 C8 and pyrimidine 1 H6-13 C6 spin-pairs well suited for 1 H RD experiments. In the latter case, pyrimidine 1 H5 must be selectively deuterated. With this in mind, Juen et al. probed the milli-to microsecond dynamics in the 27 nt A-site RNA with 1 H RD experiments, owing to the incorporation of [6-13 C-5-2 H]-uridine and -cytidine and [8-13 C]-adenosine and -guanosine labels at various sites ( Figure 5B) [61]. The overall exchange rate constant (k ex = 1880 ± 140 s −1 ) between the major and minor RNA states was obtained from the 1 H RD profiles of C7, C9, A10, A93, G94, and U95. Moreover, these data agreed with the previously reported 13 C RD experiments, validating the 1 H RD approach to study milli-to microsecond dynamics in RNA. These approaches are limited to RNAs less 9 of 15 than 70 nts and only labeled nucleobase is readily coupled to unlabeled ribose. The introduction of labeled ribose using this method remains a challenge.
experiments. In the latter case, pyrimidine 1 H5 must be selectively deuterated. With this in mind, Juen et al. probed the milli-to microsecond dynamics in the 27 nt A-site RNA with 1 H RD experiments, owing to the incorporation of [6-13 C-5-2 H]-uridine and -cytidine and [8-13 C]-adenosine and -guanosine labels at various sites ( Figure 5B) [61]. The overall exchange rate constant (kex = 1880 ± 140 s −1 ) between the major and minor RNA states was obtained from the 1 H RD profiles of C7, C9, A10, A93, G94, and U95. Moreover, these data agreed with the previously reported 13 C RD experiments, validating the 1 H RD approach to study milli-to microsecond dynamics in RNA. These approaches are limited to RNAs less than 70 nts and only labeled nucleobase is readily coupled to unlabeled ribose. The introduction of labeled ribose using this method remains a challenge.  [2,[8][9][10][11][12][13] C2]-adenosine into dHBVε RNA enabled 13 C CPMG investigation of residues A20, A22, and A41 and the overall kinetic rate of unfolding to be quantified [55]. (B) Incorporation of [8-13 C]-adenosine and -guanosine and [6-13 C-5-2 H]-uridine and -cytidine into A-site RNA allowed for the use of 1 H CPMG studies and revealed the overall exchange rate with a minor populated state [61]. Magenta colors indicate labeled residues and magenta circles refer to 13 C atoms. Two strategies to overcome the inherent size limitations of solid-phase RNA synthesis involve ligation of small chemically synthesized fragments to a larger in vitro transcribed segment and use of chemo-enzymatic methodology to incorporate both isotope labeled nucleobase and ribose moieties. This approach was recently applied to interrogate the dynamics of 96 nt CCR5 RNA that stimulates -1 programmed ribosomal frameshifting upon binding to a microRNA ( Figure 6A) [39]. Chen et al. divided the 96 nt construct into a 72 nt acceptor fragment and a 24 nt donor fragment ( Figure 6B). Eight donor fragments, each containing a different [1′, [8][9][10][11][12][13] C2]-adenosine-labeled residue were prepared by solid-phase synthesis and ligated to an unlabeled acceptor fragment with T4 DNA ligase ( Figure 6B,C). Not only did this approach allow for the unambiguous resonance assignment of each of the 8 A residues, it also facilitated dynamics measurements by a 1 H1′-13 C1′ CMPG RD experiment. Of the 7 A residues measured, 5 showed evidence of chemical exchange on the millisecond time-scale attributed to base pair opening and closing. Chen et al.  3'   [2,[8][9][10][11][12][13] C 2 ]-adenosine into dHBVε RNA enabled 13 C CPMG investigation of residues A20, A22, and A41 and the overall kinetic rate of unfolding to be quantified [55]. (B) Incorporation of [8-13 C]-adenosine and -guanosine and [6-13 C-5-2 H]-uridine and -cytidine into A-site RNA allowed for the use of 1 H CPMG studies and revealed the overall exchange rate with a minor populated state [61]. Magenta colors indicate labeled residues and magenta circles refer to 13 C atoms. Two strategies to overcome the inherent size limitations of solid-phase RNA synthesis involve ligation of small chemically synthesized fragments to a larger in vitro transcribed segment and use of chemo-enzymatic methodology to incorporate both isotope labeled nucleobase and ribose moieties. This approach was recently applied to interrogate the dynamics of 96 nt CCR5 RNA that stimulates -1 programmed ribosomal frameshifting upon binding to a microRNA ( Figure 6A) [39]. Chen et al. divided the 96 nt construct into a 72 nt acceptor fragment and a 24 nt donor fragment ( Figure 6B). Eight donor fragments, each containing a different [1 ,8-13 C 2 ]-adenosine-labeled residue were prepared by solid-phase synthesis and ligated to an unlabeled acceptor fragment with T4 DNA ligase ( Figure 6B,C). Not only did this approach allow for the unambiguous resonance assignment of each of the 8 A residues, it also facilitated dynamics measurements by a 1 H1 -13 C1 CMPG RD experiment. Of the 7 A residues measured, 5 showed evidence of chemical exchange on the millisecond time-scale attributed to base pair opening and closing. Chen et al. identified a sparsely populated (~10%) excited state with a chemical shift matching that of the CCR5:microRNA complex. Chen et al. concluded that the CCR5 samples free and bound conformations in solution and the addition of cognate microRNA stabilizes the bound conformation.

Conclusions
In summary, we present an overview of past and current RNA phosphoramidite isotope labeling methodologies and their applications in RNA structure and dynamic analysis by NMR spectroscopy. The biggest practical considerations in the solid-phase chemical synthesis of labeled phosphoramidites is 2′-OH protection and stable isotope labeling pattern, both of which are specified by the RNA of interest. Most NMR studies discussed herein were with RNAs prepared with 2′-O-tBDMS and 2′-O-TOM phosphoramidites, owing to the commercial availability of the unlabeled material. From the developments made by Kreutz and co-workers, Silantes has made the following labeled phosphoramidites commercially available: [3][4][5][6][7][8][9][10][11][12][13][14][15] (Figure 7). The availability of 13 C/ 15 N-labeled 2′-O-tBDMS phosphoramidites will increase the accessibility of these methods. Thus far, analysis of RNAs greater than 60 nts is only possible with 2′-O-CEM phosphoramidites or enzymatic ligation of smaller RNA fragments. However, RNA structural biology is moving toward larger and larger RNAs-especially as cryo-EM gains popularity. Therefore, attention must center on: (1) synthetic approaches to incorporate isotope labels into the ribose and (2) solid-phase synthetic conditions that accommodate RNAs larger than 100 nts.

Conclusions
In summary, we present an overview of past and current RNA phosphoramidite isotope labeling methodologies and their applications in RNA structure and dynamic analysis by NMR spectroscopy. The biggest practical considerations in the solid-phase chemical synthesis of labeled phosphoramidites is 2 -OH protection and stable isotope labeling pattern, both of which are specified by the RNA of interest. Most NMR studies discussed herein were with RNAs prepared with 2 -O-tBDMS and 2 -O-TOM phosphoramidites, owing to the commercial availability of the unlabeled material. From the developments made by Kreutz and co-workers, Silantes has made the following labeled phosphoramidites commercially available: [3][4][5][6][7][8][9][10][11][12][13][14][15] (Figure 7). The availability of 13 C/ 15 N-labeled 2 -O-tBDMS phosphoramidites will increase the accessibility of these methods. Thus far, analysis of RNAs greater than 60 nts is only possible with 2 -O-CEM phosphoramidites or enzymatic ligation of smaller RNA fragments. However, RNA structural biology is moving toward larger and larger RNAs-especially as cryo-EM gains popularity. Therefore, attention must center on: (1) synthetic approaches to incorporate isotope labels into the ribose and (2) solid-phase synthetic conditions that accommodate RNAs larger than 100 nts.