key: cord-0032230-20uiyjd5
authors: Dayie, Theodore K.; Olenginski, Lukasz T.; Taiwo, Kehinde M.
title: Isotope Labels Combined with Solution NMR Spectroscopy Make Visible the Invisible Conformations of Small-to-Large RNAs
date: 2022-04-20
journal: Chem Rev
DOI: 10.1021/acs.chemrev.1c00845
sha: 2eaf60958a2c4195b070297f37549817ebc3b2d9
doc_id: 32230
cord_uid: 20uiyjd5

[Image: see text] RNA is central to the proper function of cellular processes important for life on earth and implicated in various medical dysfunctions. Yet, RNA structural biology lags significantly behind that of proteins, limiting mechanistic understanding of RNA chemical biology. Fortunately, solution NMR spectroscopy can probe the structural dynamics of RNA in solution at atomic resolution, opening the door to their functional understanding. However, NMR analysis of RNA, with only four unique ribonucleotide building blocks, suffers from spectral crowding and broad linewidths, especially as RNAs grow in size. One effective strategy to overcome these challenges is to introduce NMR-active stable isotopes into RNA. However, traditional uniform labeling methods introduce scalar and dipolar couplings that complicate the implementation and analysis of NMR measurements. This challenge can be circumvented with selective isotope labeling. In this review, we outline the development of labeling technologies and their application to study biologically relevant RNAs and their complexes ranging in size from 5 to 300 kDa by NMR spectroscopy.

RNA is central to medicine, chemical and structural biology, and basic research. For more than a half-century, it has been known that the code of life is imprinted in DNA sequences, following the so-called "sequence hypothesis", usually wrongly labeled as the "central dogma" in the popular parlance. 1 In the last several decades, it has become increasingly clear that the functions of cells are also transacted by DNA's lesser-known relative, RNA. 2 Indeed, the varied roles that RNAs play in both normal and dysfunctional cells have motivated RNA-based therapeutic development, as highlighted by the recent SARS COV-2 mRNA vaccines. 3−9 Additionally, RNAs are central to the workings of molecular nanomachines such as the ribosome 10−12 and the spliceosome 13−15 to name a few. Moreover, thanks to the advent of genomic sequencing efforts, we now understand that the amount of RNA sequence transcribed in humans exceeds the number of protein sequences translated by at least 50-fold ( Figure 1A ). 16 Paradoxically, the number of RNA-only structures deposited in the Protein Data Bank (PDB) remains below 1%, whereas the number of protein-only structures is a staggering 87% ( Figure 1B) . This paucity undercuts current understanding of RNA structure−function relationships. Nuclear magnetic resonance (NMR) spectroscopy accounts for ∼35% of the RNA structures deposited in the PDB and ∼7% of the protein structures, making it competitive with other biophysical tools such as X-ray crystallography and more recently cryo-electron microscopy (cryo-EM) ( Figure 1C ). 17 Moreover, NMR spectroscopy provides high-resolution structural dynamic information in solution, rendering it an ideal tool to study RNA and its interactions with macromolecules or small drug-like compounds or both. 18−25 However, unlike proteins, which are made up of 20 unique amino acid building blocks, RNAs are composed of only four aromatic nucleotides [i.e., adenosine (Ade or A), guanosine (Gua or G), cytidine (Cyt or C), and uridine (Uri or U)] that resonate over a very narrow chemical shift region. This poor chemical shift dispersion is further exacerbated with increasing RNA size. To overcome these limitations, novel isotope labeling strategies that incorporate atom-specific labels (e.g., uridine 13 C6) or expand the number of NMR probes beyond the traditional 1 H− 15 N and 1 H− 13 C spin pairs (e.g., 13 C− 19 F) have been developed.

In this review, we will outline the development of isotope labeling technologies for RNA NMR and some of the exciting new applications enabled by these labels to study small-to-large RNAs. Specifically, we will begin by detailing the benefits afforded by each common NMR-active isotope (Section 2). Next, we outline the various technologies that incorporate such labels into RNA building blocks and eventually into RNA (Section 3). This discussion will center around chemoenzymatic labeling, a method that our group has extensively developed for the past near-decade. Next, we will examine how these labels benefit dynamics measurements (Section 4) and can be leveraged to study interactions involving large RNA systems (Section 5). Finally, to conclude, we will comment on how isotope labeling can advance the field of RNA chemical and structural biology (Section 6).

Frederick Soddy is credited with coining the word "isotope" from the Greek isos (iσος) and topos (τoπος) meaning "same place", 26 with the idea that stable isotopes are chemical elements that occupy the same position in the periodic table but differ in mass due to a different number of neutrons within the atomic nucleus. Stable isotopes have been used in a wide range of applications in industry, academia, and medicine. 26 In particular, stable isotopes have significantly impacted methods such as NMR and mass spectrometry (MS) . For this work, we will focus on how these probes impact RNA NMR spectroscopy, with special emphasis on proton (hydrogen-1 or 1 H), deuterium (hydrogen-2 or 2 H), carbon-13 ( 13 C), nitrogen-15 ( 15 N), fluorine-19 ( 19 F), and phosphorus-31 ( 31 P) ( Table 1 ).

The proton isotope has high natural abundance (∼100%) and the highest sensitivity of NMR receptive and stable nuclei (Table 1) . Therefore, homonuclear two-dimensional (2D) 1 H− 1 H NMR methods were attractive in the early days of NMR analysis. However, the very limited resolution of ribose and aromatic nucleobase resonances in the RNA 1 H spectra restricted such studies to small RNAs (<5 kDa). Within the ribose, all protons with the exception of H1′ (i.e., H2′, H3′, H4′, H5′, and H5′′) are clustered within a narrow ∼0.6−0.8 ppm range (Figure 2A) . 29 Within the nucleobase, the chemical shift distribution of all protons is limited to 1 ppm or less, except for imino protons with a dispersion of ∼4 ppm ( Figure  2A ). 30, 31 Taken together, the distribution of proton resonances leads to severe chemical shift overlap that worsens as RNAs grow in size due to increased line broadening ( Figure 2B ). This, in part, explains the paucity of NMR structures of large RNAs (e.g., > 60 nt) ( Figure 2C ).

Unlike protons, with a chemical shift span of 2−15 ppm, 15 N and 13 C nuclei in nucleic acids have larger chemical shift distributions among the various atomic sites. For example, 13 C nuclei in RNA have chemical shifts from 61 (C5′-ribose) to 170 (nonprotonated pyrimidine nucleobase C4) ppm, and 15 N nuclei from 70 (amino nitrogen) to 240 (nonprotonated purine nucleobase N7) ppm. 29−31 Introduction of the 15 N isotope (0.37%) into RNA nucleobases circumvents the extensive line broadening arising from the electric quadrupole moment of the naturally abundant 14 N isotope (99.63%) ( Table 1) . Incorporation of the 15 N isotope has several additional advantages. As a spin 1/2 nucleus with low gyromagnetic ratio (γ) ( Table 1) , the 15 N isotope provides very narrow spectral lines. Nitrogen atoms, like protons and carbon, are distributed in nucleic acid major and minor grooves, and both grooves serve as important sites for metal, drug, or macromolecule interactions. However, given the wider chemical shift dispersion of 15 N over the 1 H nucleus and its narrower linewidths over 13 C and 1 H nuclei, 15 N is more suited to monitor those grooves, especially in larger RNAs. However, nitrogen's low-γ is also an "Achilles heel". In the absence of appropriate NMR cryogenic probes and the availability of high magnetic fields, detecting low-γ nuclei such as 15 N has been very unattractive. Increasing the availability of such probes is expected to reverse this trend. Nevertheless, these considerations suggest that the shortcomings of proton NMR can be overcome by heteronuclear NMR methods. 32 Beginning in the 1980s, several groups introduced 15 N, 2 H, and 13 C labels to facilitate NMR studies of RNAs and proteins. 33−42 Depending on the scientific question, these labels were introduced uniformly or selectively using bacteria in vivo or enzyme catalyzed synthesis in vitro. Selective enrichment was achieved by growing auxotrophs on obligate chemically synthesized compounds. 13 C-labeling of bacterial tRNAs 33−35 and 15 N-labeling of tRNA and 5S rRNA enabled various atomic sites in these RNAs to be monitored by NMR. Uniform 15 N-labeling was also applied to 5S rRNA in vivo. 36−39 To extend this labeling to additional RNAs, several research groups developed in vitro methods to convert ribonucleoside 5′-monophosphates isolated from bacteria grown on 15 N-, 2 H-, and 13 C-sources into the corresponding triphosphates for in vitro transcription. 43−47 These uniform 15 N-and 13 C-labeling technologies did extend the use of NMR to medium-sized RNAs (MW < 20 kDa). However, two perennial challenges of low signal-to-noise and decreased spectral resolution remained. The latter problem arises from the reintroduction of spectral overlap along the heteronuclear dimension as the RNA grows in size, and the former arises from increased relaxation that results from the slower overall tumbling of large biomolecules. The next section will describe recent labeling methods to overcome both problems.

Deuteration (i.e., replacement of protons with deuterons) simplifies the multiplicity of spin−spin interactions, eliminates nonessential resonance lines, reduces spectral crowding, helps to identify coupling patterns, and improves calculation of coupling constants with precision. 48 Given the smaller γ of the deuterium spin relative to proton (γ D ≈ γ H /6.5) (Table 1) , the relaxation rates for deuterated nuclei are scaled proportionally by 2% [(γ D /γ H ) 2 ≈ 0.02]. By eliminating competing relaxation pathways of dipolar coupled protons, deuteration suppresses spin diffusion within a relaxation network, leading to smaller linewidths and higher signal-to-noise for the remaining protons and directly attached 13 In addition to 2 H, magnetically active nuclei such as 19 F have valuable spectroscopic properties that confer clear advantages in the study of macromolecular structure and conformational changes. 56 These benefits include the 100% natural abundance of 19 F (Table 1) , a comparably large γ (94% of 1 H) (Table 1) , and a superior chemical shift dispersion that is ∼6-fold that of 1 H. 18, 57 Furthermore, 19 F is sensitive to changes in its local chemical environment, making it a useful probe of conformational changes. 18, 56, 57 Finally, fluorine has an atomic radius (1.35 Å) slightly larger than that of a hydrogen (1.20 Å) but slightly smaller than that of a methyl group (2.00 Å). The 19 F nuclei is therefore expected to substitute for either group without serious structural perturbations, 58 making it a valuable tool for the in vitro study of medically important RNAs. 59 Finally, 19 F is virtually absent in biological systems and therefore offers 19 F NMR a biorthogonal advantage of background-free drug screening. 60 Taken together, 19 F is an attractive probe for studying RNAs in solution. Details of new technologies developed to incorporate 19 F into nucleobases 16 Organismal complexity increases with RNA coding but decreases with protein coding capacity as a percentage of the DNA genomic output. (B) Percentage of RNA-only and protein-only structures deposited in the PDB. Given that this analysis excluded DNA-only structures and structures of protein−DNA/RNA complexes, the percentages do not sum to 100%. (C) Percentage of RNA-only and protein-only structures deposited in the Nucleic Acid Database (NDB) and PDB, sorted by structure determination technique. Given that this analysis is self-contained within categories, the percentages sum to 100%. NMR accounts for a larger fraction of RNA structures as compared to proteins. PDB and NDB statistics were accessed from https://www.rcsb.org/ and http://ndbserver.rutgers.edu/ in January 2022. 13 C, and 19 F Labels. The uracil nucleobase is easily assembled using a method initially devised by Roberts and Poulter, 81 later streamlined by SantaLucia and Tinoco and co-workers, 71 and further improved by Kreutz and coworkers. 82 In the original synthetic eight-step pathway described by Roberts and Poulter, the 13 C label can be placed in any position of the six-membered ring simply by changing the 13 C-source. 81 SantaLucia and Tinoco and co-workers streamlined this to a three-step reaction scheme to make 13 Clabeled cyanoacetyl urea from inexpensive commercially available 13 C-labeled precursors. 71 A slightly modified approach from Kreutz and co-workers uses bromoacetic rather than chloroacetic acid. Bromoacetic acid is the preferred starting material due to the lower costs and better handling of the cyanide reagent. 74, 82 Other methods with fewer steps exist such as condensation of malic or propiolic acid and urea. 83, 84 Even though these are straightforward two-step reactions, execution is not as convenient or cost-effective.

Using the Poulter-SantaLucia-Kreutz approach, 71,74,81,82 [1-13 C]-and [2-13 C]-bromoacetic acid selectively incorporate 13 C at uracil C4 and C5, respectively. Use of 13 C-urea, on the other hand, delivers 13 C at the C2 site, and that of 13 Cpotassium cyanide ( 13 C-KCN) labels the C6 site. Finally, 15 Nurea installs 15 N at N1 and N3. All possible uracil heteroatom positions can therefore be labeled in good yields, and these reactions can be easily scaled to gram quantities. 74, 82 An example of a synthetic scheme using the Poulter-SantaLucia-Kreutz approach 71,74,81,82 is shown for uracil C6 labeling Here, bp and nc refer to canonical Watson−Crick base pair and noncanonical base pairs, respectively. A schematic of RNA ribose and nucleobase structures and numbering are shown above the spectrum. (B) Nucleobase region of 1 H NMR spectra for RNAs of increasing size. Both signal overlap and broad linewidths worsen as RNAs grow in size. In fact, for the best visual representation, the signals corresponding to the 61 and 232 nt RNAs were increased to display them on a similar scale to that of the 14 nt RNA. (C) Histogram of RNA NMR structures in the NDB, sorted by RNA size (in nt, bin = 10 nt). Given the challenges faced by RNA NMR, there are only 23 NMR structures corresponding to RNAs > 60 nt. NDB statistics were accessed from http://ndbserver.rutgers.edu/ in January 2022.

(Scheme 1). 82, 85 In brief, bromoacetic acid 1 reacts with 13 C-KCN and sodium carbonate (Na 2 CO 3 ) in a Kolbe nitrile reaction to form 2-[cyano-13 C]acetic acid 2. Treatment of 2 with urea in the presence of acetic anhydride (Ac 2 O) then yields a urea intermediate 3 that can be readily converted to [6-13 C]-uracil 4 using a palladium catalyst (e.g., Pd/BaSO 4 ) under hydrogen atmosphere (H 2 ). Given that pyrimidine H5/ H6 protons have three-bond scalar coupling ( 3 J H5/H6 ≈ 8 Hz 29 ) and strong dipolar coupling (H5−H6 distance of 2 Å) that complicate NMR experiments, selective and quantitative deuteration can be achieved by reacting 4 with triethylamine (TEA) to form the desired [6-13 C, 5-2 H]-uracil 5. 85 Taken together, 5 was synthesized with four-steps in 63% overall yield (Scheme 1). 82, 85 Given the valuable spectroscopic properties of 19 F (Section 2.4), uracil can be fluorinated with the commercially available Selectfluor, as recently reported. 18, 57, 86 This synthetic scheme is similar to that described for uracil C6 labeling (Scheme 1), 82 18, 86, 87 In summary, 11 was synthesized in five-steps with a total yield of 38% (Scheme 2). 18, 57, 82, 85, 86 Finally, thymine C6 can be selectively labeled with a threestep synthesis in a manner similar to uracil labeling (Schemes 1 and 2). 18, 57, 82, 85, 86 In brief, bromopropionic acid 12 is used in a Kolbe nitrile reaction followed by addition of urea and Ac 2 O to form intermediates 13 and 14. 89, 90 Then reaction of 14 with Pd/BaSO 4 in H 2 forms the desired [6-13 C]-thymine 15 in 45% overall yield (Scheme 3). 89 3.1.1.2. Purine Synthesis with C8 Specific Labeling. As with pyrimidines, purine nucleobases can be selectively labeled with 13 C and 15 N isotopes using commercially available precursor compounds. In the early 1990s, SantaLucia and Tinoco and co-workers described an effective purine synthesis using 13 C-formic acid to label purine C8. 71 More recently, Kreutz and co-workers streamlined and improved the efficiency of such labeling in one-step reactions. 75, 85, 91 Here, the condensation of 13 The 13 C isotope can be incorporated at the purine C2 site starting with 5-aminoimidazole-4-carboxamide (AICA) and ethylsodium 13 C-xanthate to form [2-13 C]-hypoxanthine, [2-13 C]-adenine, or [2-13 C]-guanine. 98 A preferred alternative for purine C2 labeling uses the method of Battaglia and Ouwerkerk and co-workers, wherein sodium ethoxide (C 2 H 5 ONa) mediates cyclization of ethyl cyanoacetate 21 with 13 C-thiourea 22 to give [2-13 C]-6-amino-2-thiouracil 23. 99,100 Unlabeled sodium nitrite (NaNO 2 ) is then used for nitrosylation (the 15 N-labeled form can also be used to introduce a second isotope label) to form 24. Then sodium dithionite (Na 2 S 2 O 4 ) mediates the reduction of the nitroso group to yield 25 followed by desulfurization over Raney-Nickel to form the diaminopyrimidine 26. 101 Treatment of the product with sulfuric (H 2 SO 4 ) and formic (HCOOH) acids yields [2-13 C]-hypoxanthine 27. 102 Subsequent reaction with phosphorus oxychloride (POCl 3 ) and N,N-dimethylaniline (N,N-DMA) yields [2-13 C]-6-chloropurine 28. 103 In the final step, reaction with methanolic NH 3 in a microwave reactor yields the desired [2-13 C]-adenine 29 (Scheme 6). 100 Alternative purine synthesis pathways have been devised to enable specific labeling of adenine C2 or any purine nitrogen position. 98 99, 100 and demonstrated its utility in NMR analysis of RNA structure and dynamics (Scheme 6). 104 3.1.2. Specific 15 N Labeling. Several approaches have been reported for the synthesis of atom-specific 15 N-labeled nucleobases and nucleosides as well as their incorporation into the corresponding rNTPs and amidites for RNA synthesis. 98 74, 115 or by chemical synthesis from a transiently protected uridine amidite. 85 In this way, all uracil isotope labeling patterns will be retained in CTP and cytidine amidites. Moreover, additional 15 N-labeling of the cytidine N4 amino group can be achieved using 15 NH 4 Cl in the enzymatic 74 or chemical 85 reaction, as will be described in Sections 3.2 and 3.3.

3.1.2.2. Purine N1, N3, N7, and N9 Labeling. Synthesis of adenine N1 labeling occurs in two-steps. 101 Here, commercially available 5-aminoimidazole-4-carbonitrile 30 reacts with diethoxymethyl acetate (DEMA) to yield intermediate 31.

Subsequent reaction of 31 with aqueous ammonia (NH 3 ) readily forms the desired product [1-15 N]-adenine 32 with a total yield of 60% (Scheme 7) 101 Adenine labeled at N3, on the other hand, can be synthesized in six steps. 108 In addition, purine N7 labeling is readily achieved and has been widely adapted. 99 103 In the final step, reaction with methanolic NH 3 in a microwave reactor gives the desired [7-15 N]-adenine 51 with a total yield of 18% (Scheme 10). 100, 104, 106, 111 As mentioned above, we recently showcased the same synthetic scheme while also incorporating selective 13 C2 labeling. 104 Finally, in the synthesis of N9-labeled adenine, 5-amino-4,6dichloropyrimidine 52 is converted to a [9-15 N]-6-chloropurine 53 using aqueous 15 NH 3 and DEMA. 118 Then a reaction with aqueous NH 3 yields the desired [9-15 N]-adenine 54. This simple three-step reaction proceeds with an overall yield of 79% (Scheme 11). 118 3.1.3. Nucleobase Labels: Summary and Outlook. As described in Sections 3.1.1 and 3.1.2, and shown in Schemes 1−11, a wide range of isotope-labeled nucleobases ( Table 2) are now available to the scientific community. Of all synthetic procedures, purine C8 sites are most readily labeled in one chemical step in a single day and with high yield (64−94%) ( Table 2) . Conversely, adenine N3 is the least readily labeled, taking 11 days (Figure 2 ). Adenine C2 and N7 have the lowest Scheme 8. Synthetic Route to [3] [4] [5] [6] [7] [8] [9] [10] [11] [12] [13] [14] [15] N]-Adenine 108a a Adenine N3 and its amino group can also be labeled at by 15 Adapted with permission from Dayie and co-workers. Copyright 2020 Springer Nature. 104 Adenine C2 can also be labeled if 13 C-labeled thiourea is used as the starting material. overall yields of 18% (Table 2 ). In future work, it would be advantageous to focus on improving yields and reducing the number of chemical steps. Nevertheless, these RNA labeling patterns are commonly chosen based on the experimental information required and less often dictated by the relative time and yield of the building blocks.

With chemically synthesized isotope-labeled nucleobases inhand, this section outlines the various enzymatic methods that can be used to build them into isotope-labeled rNTPs (and dNTPs). Alternatively, this can be accomplished using Escherichia coli 45 126 However, D-ribose is a more cost-effective labeled precursor than D-glucose for the selective 13 C-or 2 H-ribose labeling of rNTPs. 128 On the basis of earlier work by Whitesides and coworkers, 131−133 our group truncated the relatively complex Gilles-Schramm-Williamson method 65,122−125 to use 10 enzymes instead of 18, and two cofactor regeneration systems (dATP and creatine phosphate) ( Table 3 ). This chemoenzymatic labeling 74,75,129 is a versatile technology to couple nucleobase to ribose followed by subsequent phosphorylation to the rNTP in a one-pot enzymatic reaction. 74, 75, 129 The nucleobase and ribose building blocks can be unlabeled, isotope-labeled, chemically synthesized, or commercially available. This method therefore permits a diverse set of labeling patterns. Moreover, this approach has many advantages over previously reported de novo 72,73 or chemical 134−138 synthesis methods including fewer enzymes, fewer synthetic steps, and greater yields. This method affords the facile coupling of chemically synthesized uniformly 15 N-and 13 C/ 15 N-labeled uracil (Scheme 1) 82,85 to commercially available unlabeled D-ribose and 13 C-labeled D-ribose. The resulting uniformly 15 N-labeled and uniformly 13 C/ 15 N-labeled UTP provided 338-and 14-fold savings over the commercially available material from CIL, respectively. However, the main advantage of chemo-enzymatic synthesis is the ability to generate noncommercially available atom-specific labeling patterns.

We showcased the power of this method with the synthesis of [1′,5′,6-13 C 3 , 1,3-15 N 2 ]-pyrimidine rNTPs using six enzymes (Table 3 ). 74 We also used this method to synthesize [1′,8-13 C 2 ]-, or [2′,8-13 C 2 ]-, or [1′,5′,8-13 C 3 ]-ATPs and -GTPs with five enzymes (Table 3) . 75 First, 13 Total reaction time was based on the time required for all chemical steps. In addition, 16 h were added for any explicit mention of overnight procedures, and 24 h were added for any chromatographic purifications. b Number in parentheses represents the number of chromatographic purification steps. c All data for [2-13 C]-adenine and [7-15 N]-adenine labeling came from the same doubly labeled [2-13 C, 7-15 N]-adenine labeling scheme. 104 d This synthetic procedure is from Dayie and co-workers. 116 the final phosphorylation to afford the 5′-triphosphates 80−82 (Scheme 13). 74, 75, 129 Similar to the Gilles-Schramm-Williamson method, 65,122−125 a final 15 N label can be introduced at the CTP 83 amino group if 15 NH 4 Cl is used alongside CTPS in the final enzymatic step (Scheme 13). 74, 75, 129 These atomspecifically labeled rNTPs can then be used with in vitro transcription to make RNAs without any size limit. Importantly, these labeling patterns reduced spectral crowding, increased signal-to-noise ratios, facilitated direct carbon detection experiments, and eliminated 13 C− 13 C scalar and dipolar couplings. 63, 74, 75, 86, 104 As with the Gilles-Schramm-Williamson method, 65 18, 86 ) . It is worth noting that Serianni and co-workers have also developed a complementary approach to enzymatically couple nucleobase and ribose sources using four enzymes (Table 3) . 130 Their method uses hypoxanthine 84 and 1-O-acetyl-2,3,5-tri-O-benzoyl-α-D-ribofuranoside (ATBR) 85 in a Vorbruggen reaction (detailed in Scheme 15) to yield inosine 86. Then purine nucleoside phosphorylase (PNPase) (EC 2.4.2.1) replaces the hypoxanthine moiety on the C1 position of 86 with a phosphate group to give α-D-ribofuranosyl-1-phosphate sodium salt (αR1P) 87 (Table 3) . 130 Then 87 is glycosylated by PNPase with adenine or guanine or by UPase (EC 2.4.3.2) with uracil to form nucleosides 88−90, respectively (Scheme 14). 130 Products 88−90 can then be converted to the desired rNTP or amidite with further enzymatic or chemical synthesis.

Labeling. While these chemo-enzymatic methods enable straightforward atom-specific labeling, they rely solely on DNA templatedirected T7 RNA polymerase-based in vitro transcription and are therefore unable to incorporate these labels positionspecifically (e.g., nucleotide 5). Fortunately, there are two alternative enzymatic methods capable of such position-specific labeling, both of which are compatible with the isotope-labeled rNTP building blocks described above. Wang and co-workers developed a hybrid solid−liquid phase transcription technique that employs an automated robotic platform known as position-selective labeling of RNA (PLOR). 139 In PLOR, the DNA template is attached to beads and RNA synthesis is initiated by the addition of T7 RNA polymerase and a mixture of three of the four rNTP building blocks (e.g., ATP, GTP, and CTP). The beads are then washed and a new rNTP mixture is added, this time containing the previously omitted building block. Thus, PLOR can incorporate any isotope-labeled rNTP (e.g., [6-13 C, 5-2 H]-UTP) position-specifically, assuming the desired labeling site (e.g., uridine 10) does not coincide with a stretch of identical nucleotides (e.g., UUU). While isotope labeling by PLOR has aided NMR studies of RNA, 139−141 its widespread use is still limited due to the requisite equipment needed and its laborious nature.

Schwalbe and co-workers developed an alternative chemoenzymatic approach for position-specific labeling. 142 Impor- Given that there is overlap in the enzymes used in the methods of Schramm-Williamson and co-workers 65,122−125 and Dayie and coworkers, 74, 75, 129 only the unique enzymes are listed for the latter. All enzymes are commercially available except APRT, UPRT, XGPRT, CTPS, and RK. 129 These are currently only available in a few academic laboratories. At some point, these plasmids would be available at Addgene. Chemical Reviews pubs.acs.org/CR Review 22%) and ligation (9−49%) reactions are a major drawback. 142 More recent efforts by Schwalbe and co-workers to improve this technology include the addition of magnetic streptavidin beads as a solid-support and 5′-biotinylated RNA. 143 3.2.3. rNTP Labels: Summary and Outlook. As described in Section 3.2.1 and shown in Scheme 13, the chemo-enzymatic labeling method developed by Dayie and coworkers 74, 75, 129 permits the synthesis of a versatile assortment of rNTPs with atom-specific isotope labels (Table 4 ). While there are other enzymatic methods to generate both atomspecific (e.g., the Gilles-Schramm-Williamson 65,122−125 or Serriani 130 methods shown in Schemes 12 and 14, respectively) and position-specific (e.g., PLOR 139 and the Schwalbe method 142, 143 ) labels, no other technique offers the versatility and simplicity that is afforded by the Dayie method. Our one-pot chemo-enzymatic approach can produce isotopelabeled purine and pyrimidine rNTPs in a few days and with high yield (75−95%) ( Table 4 ). The main disadvantage of this method is the need to express and purify five noncommercial enzymes in-house (Table 3) . However, providing these plasmids to Addgene will make our method widely accessible to the field.

While the enzymatic production of RNA with isotope-labeled rNTPs 44,45,69−75 is the most widely used approach to obtain labeled RNA, an attractive alternative is to use isotope-labeled amidites and solid-phase synthesis. Like PLOR introduced by Wang and co-workers 139 and the chemo-enzymatic approach developed by Schwalbe and co-workers, 142,143 the amidite method offers the advantage of position-specific RNA labeling. However, even though amidite labeling is currently the most effective and widely used method for position-specific labeling, its utility for NMR studies is limited to RNAs ≈ 60 nt.

3.3.1. 15 N and 13 C Labeling. The Kreutz and Micura groups have used isotope-labeled nucleobases to prepare 2′-Otert-butyldimethylsilyl (tBDMS) and 2′-O-[(triisopropylsilyl)oxy]methyl (TOM) phosphoramidites for NMR studies, 57, 82, 85, 89, 110, 111, 144, 145 as recently reviewed. 61 A representative example of [6-13 C, 5-2 H]-pyrimidine 2′-O-TOM amidite syntheses is shown in Schemes 15 and 16. 85 In brief, [6-13 C, 5-2 H]-uracil 5 is coupled to ATBR under Vorbruggen conditions 138 to give the 2′,3′,5′-O-benzoyl (Bz)-protected 91, which is then fully deprotected to nucleoside 92 after treatment with methylamine (CH 3 NH 2 ) in ethanol (C 2 H 5 OH). Addition of 4,4′-dimethoxytrityl chloride (DMT-Cl) and TOM-Cl protects the 5′-and 2′-hydroxyl (OH) to form 93 and 94, respectively. Finally, phosphitylation of the 3′-OH of 94 with 2-cyanoethyl N , N -diisopropylchlorophosphoramidite (CEP-Cl) and N,N-diisopropylethylamine (DiPEA) yields the desired [6-13 C, 5-2 H]-uridine 2′-O-TOM amidite 95 with five-steps in 22% total yield (Scheme 15). 85 The corresponding cytidine derivative is obtained from 94 in four additional steps (Scheme 16). 85 75 The [6-13 C, 1,3-15 N 2 ]-uracil and -cytosine nucleobases, on the other hand, were coupled to [1′,5′-13 C 2 ]-D-ribose only. 74 Nevertheless, the reported times, enzymatic steps, and yields are representative of all ATP, GTP, CTP, and UTP reactions made with this method. b Total reaction time was based on the time required for all chemical steps. In addition, 24 h were added for any chromatographic purification. c Number in parentheses represents the number of chromatographic purification steps. Since the time of our original publication, 74 pyrimidine rNTP synthesis now only requires one chromatographic purification. 18, 86 Ac-cytidine 2′-O-TOM amidite 99. Starting from uracil 5, this cytidine synthesis has an overall yield of 14% (Scheme 16). 85 In contrast to pyrimidines, the starting purine is protected before beginning the nucleosidation reaction 85 Guanosine synthesis, on the other hand, proceeds from a N 2 -isobutyryl (iBu) protected guanine 106 made from [8-13 C]guanine 20 with a yield of 77%. From there, however, synthesis proceed as with adenine. That is, 106 is reacted under Vorbruggen conditions 138 to form 107, which is then 2′,3′,5′-O-deprotected to nucleoside 108. Again, 5′-OH tritylation, 2′-OH TOM protection, and 3′-OH phosphitylation yields 109, 110, and 111, respectively. In summary, [8-13 C]-N 2 -iBuguanosine 2′-O-TOM amidite 111 was synthesized with an overall yield of 18% (Scheme 18). 85 Importantly, Schemes 15−18 can be adapted to prepare 2′-O-tBDMS amidites simply by altering the 2′-OH protection reaction steps.

However, these 2′-O-tBDMS or 2′-O-TOM amidites are not suitable for producing RNAs > 60 nts. Instead, amidites with 2cyanoethoxymethyl (CEM) as the 2′-OH protecting group 146, 147 are used, due to its increased coupling efficiency, which rivals that in DNA synthesis. 80 Using a protocol developed by Yano and co-workers, 146 91 While the benefits of the CEM amidite method are attractive for obvious reasons, it has not gained widespread use due to the commercial unavailability of both unlabeled and isotope-labeled CEM amidites.

3.3.2. 19 F Labeling and Post-transcriptional Modifications. Another benefit of labeling with amidites is the position-specific incorporation of modified building blocks. Indeed, many epigenetic and post-transcriptional modifications modulate the structure, dynamics, and folding of RNAs, and NMR is providing new insights into their functions. 148 These studies have been greatly aided by the synthesis of 13 C-or 15 Nlabeled amidites bearing modifications such as uridine 5oxyacetic acid (cmo 5 U) 14 9 and N 6 -methyladenine (m 6 A). 150, 151 In collaboration with the Al-Hashimi group, Kreutz and co-workers synthesized a 15 N-labeled cmo 5 U amidite. 149 Their synthetic route begins from bromoacetic acid 1 and through intermediates 112 and 113 to assemble [1, [3] [4] [5] [6] [7] [8] [9] [10] [11] [12] [13] [14] [15] 149 Another example from the Al-Hashimi and Kreutz groups showcases the synthesis of a 13 C-labeled m 6 A amidite. 150 Their synthetic route begins with ethyl cyanoacetate 21 and 13 Cthiourea 22 and through intermediates 23−25 to assemble [2-13 C]-5,6-diamino-4-pyrimidinone 26, as in Scheme 6. 104 In contrast to Scheme 6, however, H 13 COOH was used with H 2 SO4 to introduce a second 13 . 150 Commercially, INNotope has 13 C-labeled N 1 -methyladenine, m 6 A, and N 3 -methylcytidine 2′-O-tBDMS amidites available. Finally, [1, [3] [4] [5] [6] [7] [8] [9] [10] [11] [12] [13] [14] [15] N 2 ]-pseudouridine (Ψ) amidites can be made from 15 N-labeled uracil with 11 steps in 6% total yield. 152 Additionally, building on the work shown in Scheme 2, 18, 57, 82, 85, 86 Kreutz and co-workers showcased new methods to incorporate 19 F− 13 C into the pyrimidine nucleobase of amidites. 18 57 These labeling topologies not only capitalize on the beneficial spectroscopic properties of the 19 F nuclei (Section 2.4) but also open the door to NMR studies of large RNAs, as will be discussed in greater detail in Section 5.

3.3.3. Synergy between Phosphoramidites and Chemo-enzymatic Labeling. In principle, any nucleobase labeling scheme described in Section 3.1 can be coupled to any commercially available 13 C-or 2 H-labeled D-ribose (from Omicron Biochemicals or CIL) with the chemo-enzymatic method (Section 3.2) and built into an amidite with a variety of 2′-OH protecting groups (Section 3. (Table 5 ) are becoming available to the scientific community. For all synthetic protocols, pyrimidine C6/C5 and purine C8 sites are most readily labeled. The production of these 2′-O-TOM amidites is streamlined 85 and proceeds quickly (∼1 week) and with adequate yields (14−18%) ( Table 5 ). The introduction of 19 F labels and post-transcriptional modifications, on the other hand, dramatically increases the time of synthesis (i.e., up to 10 days) and reduces the overall reaction yields (i.e., as low as 1%) (Table 5) . Nevertheless, the benefits afforded by the position-specific incorporation of these labels into RNA more than offsets these shortcomings. As with nucleobase labeling, researchers are typically motivated by the scientific question they are pursuing rather than the relative yields of each labeling reaction. Still, improvements in reaction yields and reduction in chemical steps would be advantageous for future work. Despite the synergy between the synthesis of nucleobases (Section 3.1), rNTPs (Section 3.2), and amidites (Section 3.3), and their contribution to RNA labeling for applications with solution NMR spectroscopy, a number of insurmountable limitations remain for RNAs prepared enzymatically (using, e.g., T7 RNA polymerase) and chemically (i.e., solid-phase synthesis). The former is incapable of position-specific labeling and the latter is size limited, even though both methods can install isolated 1 H− 13 C spin pairs into RNA that remove the 13 C− 13 C scalar and dipolar couplings that are normally present in uniformly labeled RNA, as will be detailed in Section 4.

Again, unlike DNA template-directed in vitro transcription, a tremendous advantage to the field is that amidite labeling and solid-phase synthesis can provide direct read-outs of the biophysical consequences of post-transcriptional modifications. This will be discussed in greater detail in Section 4.2.2.3. However, despite this strength, the "size problem" of solidphase synthesis limits the production of RNAs to ∼60 nt, beyond which it is exceedingly difficult to prepare RNA in high yield and sufficient purity for NMR studies. Even though the 2′-O-CEM 91, 146, 147 protecting group initially held promise for synthesizing larger RNAs, it has not gained widespread use. Conversely, while much larger RNAs can be transcribed enzymatically, larger RNAs always carry with them more extensive signal overlap and broader linewidths. These complications make NMR analysis of RNAs > 60 nt extremely difficult, even when atom-specific labeling is used. However, introducing 13 C− 19 F spin pairs into RNA, 18, 57, 86 leveraging the spectral properties of the 15 N nuclei, 53, 154 or combining selective deuteration with 1 H NMR 17,53−55 all hold promise to lessen the burden imposed by overlap and broad lines. This will be discussed in detail in Section 5.

It is clear that elucidating the structure, interactions, and dynamics of large RNAs and their complexes (e.g., those implicated in viral transcription, splicing, nuclear export, translation, packaging, and particle assembly) requires developing breakthrough technologies and new experimental strategies to solve the structures of such large RNAs rapidly and accurately. While the advances in the synthesis of atomspecific isotope-labeled rNTPs and amidites are essential first steps in this direction, the ability to incorporate these labels position-specifically will be a game changer for RNA structural and chemical biology. Overnight, it would transform our ability to perform position-specific readouts in vitro and in vivo. Moreover, it would enable scientists to peer directly into the active site of RNA enzymes, visualize the binding pockets of RNA−drug complexes, and exquisitely map out the interfaces of RNA−protein, RNA−RNA, or RNA−DNA−RNA hybrids. . List of possible atom-specifically isotope-labeled nucleobase and ribose labeling patterns. These can be coupled to form rNTPs via chemo-enzymatic synthesis but also converted into amidites with further chemical synthesis. Nucleobase labeling patterns (unmodified and modified) are based on the synthetic schemes described in Sections 3.1 and 3.3. These need not be mutually exclusive, and some labeled sites can be incorporated simultaneously. Labeled ribose, on the other hand, is available from commercial sources (Omicron Biochemicals and CIL).

At least that is the dream. While we await these technological advances, the availability of these isotope-labeled RNA building blocks with diverse labeling topologies ( Figure 3 ) still bodes well to address structural dynamic features of RNAs with NMR spectroscopy as well as MS or small angle neutron/ X-ray scattering. The remaining sections highlight how the labels described in Section 3 can be exploited to study RNA structure, interactions, and dynamics by NMR spectroscopy.

Originating more than 45 years ago, early investigations of RNA dynamics were limited to the study of bacterial tRNAs using one-dimensional (1D) NMR methods. 155 More than a decade later, development of 1D and 2D heteronuclear polarization transfer schemes to measure heteronuclear relaxation rates 156 (Figure 4 ). For these low populated states, we can extract chemical shifts (structure), rates (kinetics), and populations (thermodynamics) under various physiological conditions of temperature, salt, pH, and cellular environment. Finally, we can examine how the cellular milieu modulates the structure, dynamics, and interactions of RNA in real time.

On the picosecond (ps)-to-nanosecond (ns) (ps-ns) time scales, spin relaxation provides information about the amplitude and time scale of motions powered by the bond vectors (e.g., 15 N− 1 H, 13 C− 1 H, 13 C− 19 F, 1 H− 1 H) reorienting relative to the external applied magnetic field ( Figure   4 ). 169,175−177 Longitudinal relaxation describes the return to the equilibrium distribution of spins along the z-axis, with a characteristic exponential time constant T 1 (or rate constant R 1 = 1/T 1 ). Transverse relaxation, on the other hand, describes the decay of magnetization in the transverse xy-plane, with a characteristic decay time constant T 2 (or rate constant R 2 = 1/ T 2 ). Larger R 2 values produce broader peaks and lower peak heights in an NMR experiment. The linewidth, defined as fullwidth at half-height (given in Hz), is Δν 1/2 = R 2 /π. The heteronuclear Overhauser effect (hNOE) measures the enhancement of the heteroatom magnetization that arises from saturating the proton magnetization, and is mediated by their dipolar interaction.

For an isolated pair of spin-1/2 nuclei S and I (here, S is 15 N, 13 C, 31 P, 19 F; and I is 1 H), R 1 , R 2 , and the hNOE of nucleus S are related to the rotational diffusion tensor of the molecule according to well-known relations: 178, 179 ω , σ x = σ 33 − σ 11 , σ y = σ 33 − σ 22 , σ 11 , σ 22 , and σ 33 are the principal components of the chemical shielding anisotropy (CSA) tensor, 180,181 J(ω) is a spectral Chemical Reviews pubs.acs.org/CR Review density function, which is assumed to be a Lorentzian (e.g., simplest form is ω = τ ωτ + J( ) 1 ( ) C C 2 ), γ i is the gyromagnetic ratio of spin i, r SI is the distance between spins I and S, h is Planck's constant, and R ex is the exchange contribution to R 2 due to slow (i.e., microsecond-to-millisecond, μs-ms) motions. The raw data represented by the three relaxation parameters (R 1 , R 2 , and hNOE) reveal the nucleotide level variation of the dynamic motions encoded in the RNA primary sequence. Additional motional variables such as the overall correlation time (τ C ) and generalized order parameter (S) can be fit within a Model Free formalism 182, 183 to describe fast (i.e., ps-ns) motions. Though, for reasons enumerated below, this becomes problematic for large uniformly labeled RNAs. 184 The RNA motions reported by R 1 , R 2 , and hNOE are easily probed by 13 C 163−166,184−189 and 15 N 163,167,190 nuclei. 15 N sites are present in the four nucleobases at the following sites: adenosine (Ade)-H2-N1, Ade-H2-N3, Ade-H8-N7, and Ade-H8-N9, guanosine (Gua)-H1-N1, Gua-H8-N7, and Gua-H8-N9, uridine (Uri)-H3-N3, and Uri-H6-N1, and cytidine (Cyt)-H6-N1 (Figures 2 and 4) . These are suitable reporters of hydrogen-bonding and non-hydrogen-bond dynamics that occur in base-paired and nonbase-paired regions. However, solvent exposed imino regions are usually broadened beyond detection. Nonprotonated nitrogen sites such as Ade-N1 and Ade-N3, purine (Pur)-N7 and Pur-N9, and pyrimidine (Pyr)-N1 remain underutilized. The limited availability of directly protonated imino nitrogen probes has made protonated carbons an attractive alternative for probing RNA relaxation. These sites are found in both the ribose (C1′−C5′) and nucleobase (Ade-C2, Pur-C8, Pyr-C5, and Pyr-C6) moieties (Figures 2 and 4) .

Despite the greater number of detectable 13 C nuclei in RNA, complications arise for measurements and analysis of 13 C relaxation. First, the carbon sites are linked by intricate multibond couplings (i.e., to 15 N, 13 C, and 1 H nuclei) that are proximally positioned within 3 Å or less. Therefore, 13 C spins do not approximate an isolated two-spin system. In uniformly labeled samples, these extensive dipolar couplings complicate 1 3 C R 1 r a t e m e a s u r e m e n t s a n d a n a l ysis 120,121,185,186,188,189,191−195 in biopolymers of large size (τ C > 7 ns). Given this fact, our group has developed pulse schemes (based on the isolated 1 H− 15 N backbone amide spin pair in proteins 196 ) to leverage the isolated 1 H− 13 C spin pairs afforded by our atom-specifically labeled RNA samples ( Figure  5 ). Pulse scheme for transverse relaxation optimized spectroscopy (TROSY)-detected experiments for measuring (A) rotating-frame (R 1ρ ) (from which R 2 can be calculated 186, 196 ) and (B) 13 C R 1 rates in selectively labeled RNA, adapted from previous reports. 196 Quadrature detection and sensitivity-enhanced/gradient-selection is implemented using the Rance- Kay 197, 198 echo/antiecho scheme with the polarity of G 1 inverted and phase Φ 4 and Φ 5 incremented 180°for each second FID of the quadrature pair. Simulated R 1 rates and R 1 difference (defined as above) for the nuclei highlighted in panel A. R 1 simulations were carried out for 800 MHz field and R 1 difference simulations were run at multiple magnetic fields. All simulations were carried out at various τ C values, and additional details can be found in the original works. 188, 189 Chemical Reviews pubs.acs.org/CR Review Theoretical simulations of R 1 rates for Pyr-C5 and Pyr-C6, ribose C1′, Ade-C2, and Pur-C8 in uniformly and selectively labeled RNAs suggest that the various 1 H− 13 C, 13 C− 13 C, and 13 C− 15 N dipolar couplings ( Figure 6A ) present in uniformly labeled samples lead to overestimated R 1 rates ( Figure 6B ). Moreover, this discrepancy, measured by the R 1 difference (where R 1 difference = [100 × (R 1,uni − R 1,sel )/R 1,uni )]), increases with higher molecular weights and magnetic field strengths ( Figure 6B ). Experimental measurements with our customized pulse sequence (for selectively labeled RNA) ( Figure 5 ) and those of others 186 (for uniformly labeled RNA), corroborated our simulations, suggesting that these discrepancies in R 1 cannot be wholly ignored, even for fairly isolated Ade-C2 and Pur-C8 sites. 188, 189 Taken together, the contribution of 13 C− 13 C dipolar interactions needs to be explicitly taken into consideration in data analysis of uniformly labeled RNA. Spin relaxation measurements on uniformly labeled RNA from Al-Hashimi and co-workers 186 demonstrate that this is not an insurmountable hurdle. Nevertheless, the focus of our discussion on RNA dynamics will center on slower conformational exchange motions, which will be discussed in Section 4.2.

Spin-1/2 nuclei with a positive gyromagnetic ratio either align parallel (α, high-populated, favorable energetic state) to the static NMR magnetic field (B 0 ) or antiparallel (β, lowpopulated, unfavorable state). The net bulk magnetization, oriented parallel to B 0 , can be realigned with radiofrequency (RF) pulses along a direction perpendicular to B 0 . The magnetization then precesses about B 0 at a resonant Larmor frequency (ω) characteristic of the nucleus. When Fourier transformed, this detectable oscillating time-domain signal yields a frequency-domain NMR spectrum with signals at characteristic frequencies for each nucleus. When referenced against a standard frequency (e.g., sodium-3-(trimethylsilyl)-1propanesulfonate (DSS) for 1 H), we obtain a fieldindependent chemical shift that is directly proportional to the energy difference between the α and β states.

For RNA exchanging between two states A and B, the chemical shift difference (Δω) between the two states and the exchange rate constant [k ex , sum of the forward (k AB ) and reverse (k BA ) rate constants] or the exchange lifetime (τ ex = 1/ k ex ) determine if two distinct NMR peaks are observed and what signal intensity and linewidth are obtained for a given nucleus. 199, 200 In the slow exchange regime, two distinct peaks are detected at the chemical shifts of the individual states, and the peak intensities are proportional to the populations of each state. In the fast exchange regime, k ex is much larger than Δω, and therefore, a single peak is observed at the populationweighted average chemical shift. In the intermediate exchange regime, which, as its name implies, lies between the fast and slow time scales, k ex ≈ Δω.

Regardless of the exchange regime, if chemical exchange is present, R 2 increases by R ex , which depends on k ex and Δω and can therefore be modulated by magnetic field strength. 199 (Figure 4) . Moreover, even processes slower than seconds can be studied with real-time NMR (Figure 4 ). 209 For two-site exchange, a general expression for the R 2 rate constant (R CPMG (τ cp )) for state A (where p A > p B ), that encompasses all conformational exchange time scales, is given by the Carver-Richards equation: 199, 210 i k j j j j j j 

where R 2 A/B and p A/B are the R 2 rate and relative populations of the A/B state, respectively. A main disadvantage of the CPMG experiment is that only the magnitude (and not the sign) of Δω is obtained. Still, this disadvantage of the CPMG experiment is offset by the relative ease of its implementation and data analysis. That is, conformational exchange is easily detected by a nonflat CPMG curve when plotting R 2,eff versus v CPMG ( Figure 7A ). Nonexchanging nuclei, on the other hand, have no dependence of R 2,eff on v CPMG and therefore appear as flat curves ( Figure 7A ). R 1ρ and CEST experiments provide more robust information regarding the chemical shifts of state B. For a two-site model, Δω, k ex , and p B can be extracted from CEST profiles using the Bloch-McConnell 7 × 7 matrix (including the equilibrium magnetization terms). 213−215 By combining all data sets, global k ex and p B values can be fit numerically for all the CEST profiles, plotted as I/I 0 versus spin-lock offset (in Hz) ( Figure  7B ). The 7 × 7 two-site Bloch-McConnell equation is derived from the relaxation matrix and the kinetic rate matrix for an exchanging two-site system: 208, 212, 214, 215 (11) where R 1 A/B , ω A/B , and ω 1 are the R 1 rate of the A/B state, the offset of the B 1 spin-lock field from the peaks in the A/B state (in rad s −1 ), and the B 1 field strength (in rad s −1 ), respectively. The evolution of magnetization for the peak in state A during the CEST spinlock period is given by

Similarly, under the R 1ρ model for two-site exchange, the R 1ρ value for state A magnetization is given by 216 (13) and (15, 16) where Ω = ω rf − Ω obs is the difference between the resonance frequency of the observed nucleus (Ω obs ) and the spinlock transmitter frequency (ω rf ). For R 1ρ experiments, conformational exchange can be detected by plotting R 2,eff versus Ω/2π ( Figure 7C ). The expression for CEST and R 1ρ (eqs 11−16) provide insight into the parameters that are important for acquiring useful data. For example, higher B 1 fields decrease chemical shift resolution between states and also broadens linewidths ( Figure 7B,C) .

While almost all RD studies involve two-site systems, expressions for CPMG, R 1ρ , and CEST models for characterizing N-site exchange have been described by Arthur Palmer III and co-workers. 199 Indeed, work from Al-Hashimi and coworkers on Watson−Crick mismatches and base pair reshuffling in RNA feature R 1ρ and CEST data that described three-site exchange. 217 4.2.1. Slow Motions: Are Selective Labels Needed? As with spin relaxation, the scalar and dipolar couplings present in uniformly labeled samples can lead to complications in RD and CEST experiments. As we have discussed elsewhere, 75 numerous spectroscopic solutions have been proposed to circumvent the problems that arise from 13 C− 13 C couplings that exist in uniformly labeled RNA. These advances include constant time evolution, 218−221 adiabatic band selective decoupling, 222−224 and selective cross-polarization with weak RF fields. 225−227 These solutions have benefited RD and CEST experiments to varying degrees in RNA. Specifically, 13 C− 13 C scalar couplings (e.g., C1′−C2′ or C5−C6) complicate CPMG Chemical Reviews pubs.acs.org/CR Review experiments 228,229 to a much larger degree than both CEST and R 1ρ. However, these couplings still pose a problem to CEST 230,231 and R 1ρ 212 and oscillations are sometimes observed in the decay profiles of C1′ and C6 nuclei. Moreover, as with spin relaxation, these couplings must be explicitly taken into consideration in data analysis. The number of coupled homogeneous differential equations (n) is equal to (2 × 4 m ) − 1, where m is the number of weakly coupled nuclear spins in an m-spin system. Therefore, for 1-, 2-, and 3-spin systems, n = 7, 31, and 127, respectively. 214, 215, 231 This transforms the CEST matrix (eq 11) from 7 × 7 to 31 × 31 for 13 C− 13 C scalar coupled spin pairs found in the nucleobase and ribose moieties. Atom-specific labeling (Section 3), on the other hand, circumvents this problem entirely, and dramatically simplifies NMR spectra, especially when incorporated position-specifically via solid-phase synthesis (Section 3.3). However, a drawback for selective labels is the obvious reduction of probe sites.

Nevertheless, using both selective and uniformly labeled RNA, CEST and R 1ρ experiments have now been applied to the protonated nucleobase (Pyr-C5 and Pyr-C6, Pur-C8, and Ade-C2) and ribose (C1′-C5′) carbons, the nucleobase imino (Gua-N1 and Thy/Uri-N3) and amino (Gua-N2) nitrogen, nucleobase (Uri-H3, Gua-H1, Ade-H2, Pur-H8, Pyr-H5, and Pyr-H6) and ribose H1′ protons, as well as nonprotonated (Gua-N7, Ade-N1, and Pur-N7) and amino (Cyt-N4) nitrogen sites (Figure 4) . 75 

Experiments in Selectively Labeled RNA. As highlighted above, implementation of RD experiments on selectively labeled RNA circumvents all complications from strong 13 C− 13 C scalar couplings and permits straightforward data analysis. The following sections will be devoted to showcasing examples of CPMG, CEST, and R 1ρ experiments performed on selectively labeled RNAs. Specifically, we will highlight recent work from our group 228,234 using isotope-labeled rNTPs and from Kreutz and Al-Hashimi and co-workers 151 using isotope-labeled amidites with post-transcriptional modifications. Figures 8A,B) . 228 The SQ 1 H CPMG experiment was amenable to Pur-H8 sites, detecting exchange in G19 and A21. The extracted exchange rate (k ex = 4000 ± 100 s −1 ) from a global fit was consistent with that determined from a standard 1 H− 13 C TROSY CPMG experiment (k ex = 3000 ± 800 s −1 ), demonstrating that these new experiments are feasible for RNA ( Figure 8B ). 228 Moreover, these data agree with R 1ρ measurements on uniformly labeled RNA from Al-Hashimi and co-workers, which suggests that each measurement, using various methods and labeling techniques, is picking up fundamental motions within this RNA. 256 In addition, these SQ experiments could provide important data on 1 H chemical shifts, which are currently lacking, such as ribose H1′ and Pyr-H6. In the latter case, however, the presence of Pyr-H5 can cause dispersive CPMG patterns for the H6 site. 228 Fortunately, Pyr-H5 deuteration is easily achieved (Scheme 1), 85 and therefore, this experiment can be readily implemented to obtain data for Pyr-H6 sites.

Our group also designed a CH 2 1 H− 13 C TROSY-detected CPMG pulse sequence ( Figure 8C) 228, 257 to leverage the isolated 13 C spin at the ribose C5′ position (Figure 4 ) afforded by our chemo-enzymatic labeling (Sections 3.1 and 3.2). 74, 75, 129 This new CPMG experiment was implemented using the selectively labeled ([1′,5′,6-13 C 3 , 5-2 H]-CTP) ironresponsive element (IRE) RNA and detected exchange in C18−C5′ ( Figure 8D ). 228 These data were then globally fit with additional CPMG data from other nuclei to obtain chemical shift (Δω = 2.5 ± 0.2 ppm), population (p B = 1.7 ± 0.2%), and exchange rate (k ex = 3600 ± 300 s −1 ) information that suggests a significant structural rearrangement in the IRE triloop ( Figure 8D ). 228 4.2.2.2. CEST in Atom-and Position-Specifically Labeled RNA. In addition to using selective labels to benefit CPMG experiments, they can also be used to simplify CEST experiments. Specifically, our group combined enzymatic ligation, chemo-enzymatic labeling, and newly developed CEST experiments ( Figure 9A ) to study the conformational equilibria of the SAM-II riboswitch in the apo (ligand-free) state. 234 To understand the formation of the SAM metabolitebinding pocket, a SAM-II RNA was constructed via DNA splinted ligation with T4 DNA ligase (EC 6.5.1.1) of two RNA fragments: an unlabeled 31 nt acceptor fragment and a [1′,6-13 C 2 , 5-2 H]-CTP labeled 21 nt donor fragment. This strategy enabled position-specific labeling, given that there was only one cytidine (C43) in the donor sequence and therefore permitted direct monitoring of the G22−C43 base pair interaction in the SAM binding pocket. Moreover, the isolated spin pair labeling topology enabled the design of a 1 H CEST experiment, and simplified setup and analysis of 1 H and 13 C CEST experiments without complications from 13 C− 13 C couplings to Cyt-C1′ and Cyt-C6 sites. 230 To leverage the labeling scheme, our group designed a new 13 C CEST experiment based on previous pulse schemes 212, 214 and used it on the apo SAM-II riboswitch ( Figure 9A ). The CEST profiles of C43−C1′ and C43−C6 indicated two states of the free SAM-II riboswitch: one that matched the resonance of the ligand-free, highly populated conformation (i.e., state A) and another that matched the ligand-bound, transient conformation (i.e., state B) ( Figure 9B ). 234 We then used our new 1 H CEST experiment ( Figure 9A) to indirectly obtain the C43−H1′ chemical shift of state A and B. 234 In agreement with the 13 C data, the 1 H chemical shift of state B matched the ligand-bound SAM-II ( Figure 9B ). 234 Taken together, these results suggest that the apo SAM-II exists in a dynamic equilibrium (k ex = 36 ± 3 s −1 ) between an open (highly populated, p A = 90.5 ± 0.5%) and a partially closed (transient, Chemical Reviews pubs.acs.org/CR Review p B = 9.5 ± 0.5%) state ( Figure 9B ). 234 Moreover, these results underscore the emerging consensus that transient, low populated states likely enhance rapid ligand recognition and therefore play a potentially ubiquitous role in RNA recognition and signaling.

Perhaps the greatest benefit of selective labeling is the ability to monitor the structural dynamic consequences of epigenetic and post-transcriptional modifications. Using labels created by Kreutz and co-workers, the Al-Hashimi group has been at the forefront of exploring how these modifications alter the dynamic ensembles of nucleic acids. 149 150 The effect of m 6 A on hybridization kinetics stands in contrast to the effect of mismatches. Mismatches also slow the rate of duplex annealing but dramatically increase the rate of duplex melting. 276−278 Of critical importance, the methylamino group of the m 6 A nucleobase can form two rotational isomers that interconvert on the millisecond time scale 279, 280 (Figure 10A ). The preferred syn isomer (i.e., high-populated, state A) cannot form a canonical Watson−Crick base pair with uridine due to a steric clash between the uridine keto group and the methylamino 279−281 and is therefore mismatch-like ( Figure  10A ). Instead, when base-paired with uridine, the methylamino rotates into the anti isomer (i.e., transient, state B) to form a canonical Watson−Crick m 6 A:U base pair ( Figure 10A ).

Kinetic mechanisms that involve binding and conformational change can occur via pathways wherein the conformational change occurs prior to (conformational selection, CS) or post (induced fit, IF) binding. Al-Hashimi and co-workers employed their recently developed RD-based and CEST experiments 31,148,168,237−246 to measure hybridization kinetics of single-and double-stranded RNA (ssRNA and dsRNA, respectively) harboring atom-and position-specifically labeled m 6 A probes (i.e., [2, [8] [9] [10] [11] [12] [13] C 2 ]-m 6 A or [ 13 CH 3 ]-m 6 A) ( Figure  10B ) to determine how m 6 A modulates hybridization. 151 In this way, they had direct readouts of the effects of the m 6 A isomers on Watson-Crick or mismatch-like hybridizations. They showed that m 6 A with the methylamino group in the anti conformation forms a Watson−Crick base pair with uridine that transiently isomerizes on the millisecond time scale to a singly hydrogen-bonded (p B ≈ 1%) mismatch-like conformation, with the methylamino group in the syn conformation. 151 This rapid interconversion between Watson−Crick and mismatch forms, combined with different syn:anti preferences in ssRNA and dsRNA states, hints at how m 6 A slows duplex annealing without affecting melting via two pathways in which isomerization occurs before (CS) or after (IF) duplex annealing ( Figure 10C ). 151 4.2.3. Examples of Relaxation Dispersion Experiments without Selectively Labeled RNA. While RD experiments work well with selective labels, it is not a prerequisite, as long as care is taken to either minimize strong 13 C− 13 C scalar couplings (i.e., probe nuclei where these are Figure 11A ), which has been characterized extensively by Zhang and co-workers. 250 Here, they showed that the riboswitch aptamer adopts a nearidentical solution structure 250 with (holo) and without (apo) the fluoride ligand, in agreement with X-ray crystal structures In solution, the aptamer adopts near-identical structures in the apo and holo forms, in agreement with crystallography. 250, 282 (C) Schematic of the equilibrium between the highly populated apo state (i.e., State A) and the transient "holo-like" conformation of the apo state (i.e., State B). Exchange parameters were extracted from a global fit of the CEST data. The transient "holo-like" conformation of the apo state (i.e., State B) occludes the formation of a reverse Hoogsteen base pair in the highly populated conformation of the apo state (i.e., State A) to signal transcription termination. Additional details can be found in the original work. 250 Figure 12 . (A) Secondary structure of the mir-34a−mSirt1 duplex. 247 Nucleotides that were found to be in exchange are circled. (B) Schematic of the equilibrium between the highly populated 7-mer-A1 and transient 8-mer-GU mir-34a−mSirt1 duplex. Exchange parameters were extracted from a global fit of the R 1ρ data (i.e., gG8-H1, gG8-N1, gG8-C8, tC17-C1′, tA19-C8, tU20-C1′, tU21-C6, and tA22-C8). The boxed nucleotides represent the critical switch from the gG8:tC17 to gG8:tU21 base pair. (C) Replotted functional data 247 showing the percentage of target repression for each miR-34a duplex. The transient 8-mer-GU reduces target mRNA levels ∼2-fold compared to the highly populated 7-mer-A1. The 8-mer-GU duplex therefore represents a "catalytically competent RISC". Additional details can be found in the original work. 247 ( Figure 11B ). 282 Moreover, these states also undergo very similar dynamic motions across a wide range of time scales, as determined from 13 C spin relaxation rates and residual dipolar couplings (RDCs). 250 However, functional assays indicate that transcription activation is fluoride-dependent and kinetically driven. 282, 283 What is more, mutational studies suggest that a prefolded "holo-like" apo state lowers the kinetic barrier for ligand binding, enabling efficient fluoride sensing to activate transcription below or near the toxicity threshold. Until recently, the mechanism by which this holo-like apo state achieves the "transcription−off" state remained unknown. 250 To shed light on this mechanism, 13 C CEST experiments were implemented on uniformly 13 C/ 15 N-GTP-and uniformly 13 C/ 15 N -ATP/UTP labeled aptamer RNA. For the holo state, CEST profiles consistently showed a single, highly populated conformation (i.e., state A). 250 A subset of CEST profiles of the apo state, on the other hand, revealed the presence of conformational exchange to a transient state (i.e., state B). 250 The nucleotides that undergo chemical exchange were localized to the junction of P3, J13, J23, and the 3′-tail, suggesting a concerted transition ( Figure 11A,B) . A global fit of the CEST data determined the population (p B = 1.4 ± 0.1%) and lifetime (τ B = 3.2 ± 0.3 ms) of the holo-like conformation of the apo state. This fleeting process differentiates the apo and holo states. Rapid transition to the hololike conformation of the apo state, which unlocks the highly conserved reverse Hoogsteen base pair located at the interface between the aptamer domain and the expression platform, promotes strand invasion and provides a path to transcription termination ( Figure 11C ). 250 Conversely, fluoride binding allosterically suppresses access to the holo-like conformation of the apo state, ensuring continued gene transcription. 250 RNA can also regulate the initial steps of translational silencing. This process begins when a mature miRNA binds to the human Argonaute (Ago2) protein to form the RNAinduced silencing complex (RISC). 284 Here, translational silencing is predominantly controlled by base pair complementarity between the "seed" region of the miRNA and the target mRNA. 284−289 Interestingly, data from bioinformatics, 290 structural, 291 and mutational 292 studies all suggest that RNA dynamics within the central bulge of miRNA− mRNA duplex likely controls mRNA fate. To test this hypothesis, Petzold and co-workers used R 1ρ experiments coupled with molecular dynamics simulations to investigate the structural dynamics of the interaction between miR-34a and its miRNA recognition element in the 3′-UTR of silent information regulator 1 mRNA (mSirt1) ( Figure 12A ). 247 Using these experiments, the authors detected chemical exchange in nucleotides surrounding the central bulge of the miR-34a−mSirt1 duplex ( Figure 12A ). 247 In this structural rearrangement, the gG8:tC17 base pair ('g' refers to the guide miRNA and 't' refers to the target mRNA) interconverts from a highly populated (i.e., state A) to a transient (i.e., state B) conformation. A global fit of the R 1ρ data determined the exchange rate (k ex = 1008 ± 12 s −1 ) and population (p B = 0.9 ± 0.2%) of the unfavorable state ( Figure 12B ), 247 and the chemical shift data 247 from 1 H (Δω −2.20 ± 0.02 ppm) and 15 N (Δω −3.8 ± 0.1 ppm) R 1ρ experiments suggest formation of a gG8:tU21 wobble pair ( Figure 12B ), 247 a motif seen in other miRNAs. 293, 294 Taken together, the miR-34a−mSirt1 binding site is in equilibrium between a highly populated 7mer-A1 and a transient 8-mer-GU ( Figure 12B ).

Next, Petzold and co-workers sought to investigate the functional relevance of the 8-mer-GU unfavorable state using a functional assay and simulated complexes of human Ago with 7-mer-A1 and 8-mer-GU 34a−mSirt1 duplexes. Interestingly, the switch to the 8-mer-GU state causes coaxial stacking of the seed and supplementary helix fitting into Ago2, reminiscent of an active state in prokaryotic Ago. 295, 296 Moreover, this state enhances repression of the target mRNA, revealing the importance of this dynamic miRNA−mRNA structure ( Figure  12C ). 4.2.3.2. CEST and R 1ρ Experiments in Unlabeled RNA. After highlighting RD experiments in selectively and uniformly labeled RNA, we will conclude this section with a brief description of two pulse schemes that permit R 1ρ 252 and CEST 253 experiments in unlabeled RNA. In the first, Petzold and co-workers developed a SELOPE homonuclear NMR method by combining the selective excitation of specific groups of protons and reduction of spectral crowding using coherence transfer among scalar coupled protons. These coherence transfers take advantage of uniform homonuclear three bond Chemical Reviews pubs.acs.org/CR Review scalar coupling between H5 and H6 for pyrimidine bases ( 3 J H5H6 ∼ 8−10 Hz) or between H1' and H2' for ribose in C2'endo conformation ( 3 J H1'H2' ∼ 8Hz). Taken together, SELOPE permits well-resolved 1D and 2D spectra of unlabeled RNA.

To demonstrate the utility of this method to probe RNA transient states, Petzold and co-workers adapted the SELOPE pulse scheme to include a spinlock ( Figure 13A ). 252 As proofof-concept, this new 1 H R 1ρ SELOPE experiment was used to detect chemical exchange in the central bulge region of the GUG RNA ( Figure 13B ). 252 Importantly, this method enables the use of lower spinlock strengths to measure slower exchange time scales. 252 Building on this work, Al-Hashimi and co-workers introduced a high-power 1 H CEST SELOPE experiment to target imino protons ( Figure 13C ). 253 To showcase the utility of this method, Watson−Crick to Hoogsteen exchange of G:C and A:T base pairs in DNA were monitored ( Figure 13D ). 253 Importantly, Al-Hashimi and co-workers showed that short relaxation delays could be used to characterize fast exchange events that effectively minimize NOE effects that complicate 1 H RD experiments. 214,253,297−302 Moreover, their approach also takes advantage of high-power RF fields recently shown to extend the time scale sensitivity of CEST to include faster exchange processes that were traditionally only detectable by R 1ρ . 253, 303 While both of these exciting new advancements hold promise, they are inherently limited to small RNAs. However, RNA biology is increasingly moving toward larger and larger RNAs. This important topic will be the focus of the next section.

NUCLEIC ACIDS Until now, most studies of RNA dynamics have focused on relatively small systems. However, RNA structural biology is increasingly moving toward larger RNAs, especially as cryo-EM advances in resolution and popularity. 304−306 Solution NMR spectroscopy, unlike X-ray crystallography and cryo-EM, is the only biophysical technique capable of probing nucleic acid conformational dynamics on a wide range of time scales in a physiologically relevant environment. Moreover, four technological advances have expanded the types of problems that NMR can tackle in studies of molecular nanomachines on the order of 1 MDa: (1) commercial availability of high-field magnets, up to 1.2 GHz 1 H Larmor frequency (28.2 T), 307 (2) specialized probes (e.g., cryo-probes) that minimize noise associated with the NMR signals, 308 (3) new isotope labeling technologies (described in Section 3), and (4) the design of new NMR experiments that are tailored to the isotope labeling used (described in Section 4). Our final section will describe how new labeling efforts can be leveraged to study large RNAs by NMR.

Taking inspiration from protein labeling, 309 our group installed 19 F directly next to a 13 C spin in UTP (Scheme 2) 18, 86 and showed that, compared to the 13 C− 1 H spin pair, 13 C− 19 F had better sensitivity, ∼6-times wider chemical shift Figure 14 . (A) Simulated 13 C R 2 rates (linewidths) at various magnetic fields in RNAs of various molecular weights (as measured by τ C ) to compare the relative TROSY effects of 13 C− 1 H or 13 C− 19 F spin pairs, which are shown on the left. (B) Same simulated rates as in A for each spin pair but only at the magnetic fields corresponding to the narrowest linewidths (smallest R 2 ) (600 and 950 MHz for 13 C− 19 F or 13 C− 1 H, respectively, as shown by the gray lines in panel A). 18 (B) Representative 19 F− 13 C TROSY spectrum to highlight the dispersion of resonances based on secondary structure (i.e., G:U wobble base pairs, nonhelical nucleotides, and helical A:U base pairs). Spectral regions are colored to match the respective uridines on the corresponding RNA. Additional details can be found in the original works. 18, 57 dispersion and ∼2-times more favorable relaxation properties in 2D TROSY experiments ( Figure 14A ,B). 18 Importantly, the high sensitivity of the 19 F nucleus enabled clear delineation of helical and nonhelical regions as well as G:U wobble and Watson−Crick base pairs ( Figure 14C ). 18, 57 In parallel, the Kreutz group incorporated 13 C− 19 F into both cytidine and uridine 2′-O-tBDMS amidites (Schemes 21 and 22) to show the same effect in RNAs made by solid-phase synthesis. 57 These findings suggest that structural insights are possible even in the absence of complete resonance assignment, which is a substantial bottleneck for large RNAs. Moreover, these labeling schemes can be readily adapted to exploit 19 F CEST and R 1ρ experiments, which have been described for proteins up to 360 kDa. 310−315 An alternative approach to heteronuclear correlation experiments that include nuclei with large CSAs such as 13 C and 19 F, which broaden the lines of nearby protons, was recently described by Bax and Summers and co-workers. 53 This approach capitalizes on the favorable relaxation properties of 15 N nuclei within RNA nucleobases. Here, they employed 1 H− 15 N heteronuclear multiple quantum coherence (HMQC) experiments to measure 15 N R 1ρ rates and RDCs in a large 232 nt (∼78 kDa) RNA by selectively transferring magnetization from Ade-H2 to Ade-N1/N3 via the two-bond scalar coupling ( 2 J HN ≈ 15 Hz 29 ) ( Figure 15 ). Extending this method in the same 232 nt RNA, Marchant and Tjandra and co-workers measured pseudocontact shifts using the two-bond scalar coupling of Ade-H8-N7 ( 2 J H8N7 ≈ 11 Hz 29 ) and Ade-H8-N9 ( 2 J H8N9 ≈ 8 Hz) for coherence transfer. 316 Importantly, both experiments would benefit by atom-specific labeling. That is, selective 15 N labeling of Ade-N1 or Ade-N3 (described in Section 3.1.2.2) (Schemes 7 and 8) would reduce crowding considerably and direct magnetization transfer uniquely from Ade-H2 rather than splitting it between both sites, as in uniformly 15 N-labeled RNA (Figure 15 ). In the same way, selective 15 N labeling of Pur-N7 or Pur-N9 (described in Section 3.1.2.2) (Schemes 9−11) would again reduce crowding and direct coherence transfer uniquely from Pur-H8 ( Figure 15 ). However, selective pulses can be deployed to affect the same decrowding and directed transfer. These labeling topologies could then be leveraged to probe two-bond 15 N CEST in large RNAs, as recently described by Zhang and co-workers. 232 Our final example of harnessing the versatility of the 15 N nuclei is one that exploits the narrow linewidths in 1 H− 15 N TROSY experiments compared to its 1 H− 13 C counterpart ( Figure 16A ). Here, Furtig and Schwalbe and co-workers investigated several reconstituted complexes between an adenine-sensing riboswitch and the 30S ribosome by NMR spectroscopy. 154 In particular, they implemented the 1 H− 15 N BEST-TROSY pulse scheme 317, 318 to obtain incredible spectra for a massive-sized complex (>800 kDa) ( Figure 16B ). Taken 154 Additional details can be found in the original work. 154 Chemical Reviews pubs.acs.org/CR Review together, Furtig and Schwalbe and co-workers succeed in illuminating the dynamic network that links the riboswitch RNA regulator, adenine ligand inducer, and ribosome protein S1 modulator during translation initiation. 154 

In humans, RNA transcripts exceed the number of proteins decoded by more than 50-fold, and yet the number of RNA structures remains below 1%, preventing a detailed understanding of RNA function (Figure 1 ). It is therefore essential to characterize RNA structural dynamics and interactions at atomic resolution to fill this critical knowledge gap. Over the past two decades, NMR spectroscopy has assumed a central role in RNA structure determination and probing dynamics on functionally relevant time scales in solution. In this review, we have summarized some of the many contributions of solution NMR studies to our knowledge of RNA structure, dynamics, and interactions, as facilitated by isotope labeling. We have presented a detailed overview of the prominent role stable isotopes continue to play in NMR analysis of nucleic acids (Section 2), how to synthesize these labels and introduce them into RNA (Section 3), and how these labels benefit NMR analysis. Of great interest, selective isotope labeling alleviates spectral crowding and removes dipolar and scalar couplings to simplify NMR dynamics measurements and data interpretation (Section 4). Moreover, recent advances in labeling open the door to study large RNA systems in a manner previously thought impossible (Section 5). As new orthogonal technologies are developed to better characterize the functional relevance of RNA, their structural dynamics will become increasingly important to better understand the cellular basis of RNA-based dysfunction that leads to various diseases. We anticipate that several imminent breakthrough technologies, some described herein, will enable NMR spectroscopy to continue to play a pivotal role in shining light on the structure, dynamics, and function of the important "dark matter of the genome", RNA in vitro, in cellulo, and in vivo. carbodiimide GK = guanylate kinase GND = phosphogluconate dehydrogenase GTP = guanosine 5′-triphosphate Gua = guanine H 2 SO 4 = sulfuric acid HC(OC 2 H 5 ) 3 = triethyl orthoformate HCOOH = formic acid HCOONH 2 = formamide HF = hydrogen fluoride HMQC = heteronuclear multiple quantum coherence hNOE = heteronuclear Overhauser effect HXK = hexokinase iBu = isobutyryl IF = induced fit KCN = potassium cyanide k AB = forward reaction rate k BA = reverse reaction rate k ex = exchange rate MK = myokinase mRNA = messenger RNA MS = mass spectrometry ms = millisecond mSirt1 = silent information regulator 1 mRNA m 6 A = N 6 -methyladenine Na 2 CO 3 = sodium carbonate NaNO 2 = sodium nitrite Na 2 S 2 O 4 = sodium dithionite NaOD = sodium deuteroxide NaOH = sodium hydroxide NDB = nucleic acid database NH 3 = ammonia NH 4 Cl = ammonium chloride NH 4 15 NO 2 = 15 N-labeled ammonium nitrate NH 4 OH = ammonium hydroxide NMPK = nucleoside-monophosphate kinase NMR = nuclear magnetic resonance N,N-DMA = N,N-dimethylaniline NPE = nitrophenyl ethanol ns = nanosecond nt = nucleotide OH = hydroxyl p A = population of state A p B = population of state B PDB = protein data bank Pd/BaSO 4 = palladium catalyst PNPase = purine nucleoside phosphorylase POCl 3 = phosphorus oxychloride PRPPS = phosphoribosylpyrophosphate synthetase ps = picosecond pTSA = para toluene sulfonic acid Pur = purine PYKF = pyruvate kinase Pyr = pyrimidine RD = relaxation dispersion RDC = residual dipolar coupling R ex = chemical exchange contribution to R 2 RF = radiofrequency RISC = RNA-induced silencing complex RK = ribokinase rNTPs = ribonucleoside 5′-triphosphates RPI1 = ribose-5-phosphate isomerase 1 rSAP = recombinant shrimp alkaline phosphatase R 1 = longitudinal relaxation rate R 1ρ = rotating-frame transverse relaxation rate R 2 = transverse relaxation rate S = order parameter SELOPE = selective optimized proton experiment SO 2 Cl 2 = sulfuryl chloride SQ = single quantum ssRNA = single-stranded RNA tBDMS = 2′-O-tert-butyldimethylsilyl TC = temperature compensation TEA = triethylamine TiBSC = 2,4,6-triisopropylbenzenesulfonyl chloride TiPCEP = 2-cyanoethyl N,N,N′,N′-tetraisopropylphosphorodiamidite TOM = 2′-O-[(triisopropylsilyl)oxy]methyl TROSY = transverse relaxation optimized spectroscopy T 1 = longitudinal relaxation time T 2 = transverse relaxation time UMPK = uridine monophosphate kinase UPRT = uracil phosphoribosyltransferase Ura = uracil UTP = uridine 5′-triphosphate XGPRT = xanthine-guanine phosphoribosyltransferase ZWF = glucose-6-phosphate dehydrogenase αR1P = α-D-ribofuranose 1-phosphate Δω = chemical shift difference ω = Larmor frequency τ C = overall correlation time τ ex = exchange lifetime μs = microsecond Chemical Reviews pubs.acs.org/CR Review 1D = one-dimensional 2D = two-dimensional 5FU = [5-13 C, 5-19 F, 6-2 H]-uracil

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Syntheses of All Singly Labeled [ 15 N]Adenines: Mass Spectral Fragmentation of Adenine

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NMR Chemical Exchange Measurements Reveal That N 6 -Methyladenosine Slows RNA Annealing

A Quantitative Model Predicts How m 6 A Reshapes the Kinetic Landscape of Nucleic Acid Hybridization and Conformational Transitions

Pseudouridines in RRNA Helix 69 Play a Role in Loop Stacking Interactions

Synthesis of Atom-Specific Nucleobase and Ribose Labeled Uridine Phosphoramidite for NMR Analysis of Large RNAs

Switching at the Ribosome: Riboswitches Need rProteins as Modulators to Regulate Translation

Observation of Resonances from Some Minor Bases in the Natural-Abundance Carbon

Nuclear Magnetic Resonance Spectrum of Unfractionated Yeast Transfer Ribonucleic Acid. Evidence for Fast Internal Motion of the Dihydrouracil Rings

Measurement of 13 C Relaxation Times in Proteins by Two-Dimensional Heteronuclear 1 H-13 C Correlation Spectroscopy

Measurement of Carbon-13 Longitudinal Relaxation Using 1 H Detection

Sensitivity Enhancement of 13 C T1Measurements via Polarization Transfer

Backbone Dynamics of a Free and a Phosphopeptide-Complexed Src Homology 2 Domain Studied by 15 N NMR Relaxation

Relaxation-Rate Measurements for 15 N-1 H Groups with Pulsed-Field Gradients and Preservation of Coherence Pathways

Mapping of Spectral Density Functions Using Heteronuclear NMR Relaxation Measurements

Backbone Dynamics of Proteins as Studied by 15 N Inverse Detected Heteronuclear NMR Spectroscopy: Application to Staphylococcal Nuclease

Base Flexibility in HIV-2 TAR RNA Mapped by Solution 15 N, 13 C NMR Relaxation

Active Site Dynamics in the Lead-Dependent Ribozyme

13 C Spin Relaxation Measurements in RNA: Sensitivity and Resolution Improvement Using Spin-State Selective Correlation Experiments

13 C Relaxation and Dynamics of the Purine Bases in the Iron Element RNA Hairpin

Base Dynamics in a UUCG Tetraloop RNA Hairpin Characterized by 15 N Spin Relaxation: Correlations with Structure and Stability

Characterizing Micro-to-Millisecond Chemical Exchange in Nucleic Acids Using Off-Resonance R 1ρ Relaxation Dispersion

Quantitative 2D and 3D Γ-HCP Experiments for the Determination of the Angles α and ζ in the Phosphodiester Backbone of Oligonucleotides

RNA Phosphodiester Backbone Dynamics of a Perdeuterated CUUCGg Tetraloop RNA from Phosphorus-31 NMR Relaxation Analysis

A Suite of 2 H NMR Spin Relaxation Experiments for the Measurement of RNA Dynamics

Characterizing RNA Dynamics at Atomic Resolution Using Solution-State NMR Spectroscopy

RNA Dynamics: It Is about Time

Cross-Correlated 19 F Relaxation Measurements for the Study of Fluorinated Ligand-Receptor Interactions

NMR Characterization of the Dynamics of Biomacromolecules

Protein Dynamics from NMR

Nuclear Magnetic Resonance Relaxation in Determination of Residue-Specific 15 N Chemical Shift Tensors in Proteins in Solution: Protein Dynamics, Structure, and Applications of Transverse Relaxation Optimized Spectroscopy

Theory and Practice of Nuclear Spin Relaxation In Proteins

Measurement of 15 N Chemical Shift Anisotropy in Solution

Chemical Shift Tensors of Protonated Base Carbons in Helical RNA and DNA from NMR Relaxation and Liquid Crystal Measurements

Model-Free Approach to the Interpretation of Nuclear Magnetic Resonance Relaxation in Macromolecules. 1. Theory and Range of Validity

Model-free approach to the interpretation of nuclear magnetic resonance relaxation in macromolecules. 2. Analysis of experimental results

13 C NMR Relaxation Studies of RNA Base and Ribose Nuclei Reveal a Complex Pattern of Motions in the RNA Binding Site for Human U1A

Binding of U1A Protein Changes RNA Dynamics as Observed by 13 C NMR Relaxation Studies

Dynamics of Large Elongated RNA by NMR Carbon Relaxation

Dynamics and Metal Ion Binding in the U6 RNA Intramolecular Stem-Loop as Analyzed by NMR

Deleterious Effects of Carbon-Carbon Dipolar Coupling on RNA NMR Dynamics

Quantifying the Effects of Long-Range 13 C-13 C Dipolar Coupling on Measured Relaxation Rates in RNA

Resolving the Motional Modes That Code for RNA Adaptation

Rotational Diffusion Tensor of Nucleic Acids from 13 C NMR Relaxation

Alternate-Site Isotopic Labeling of Ribonucleotides for NMR Studies of Ribose Chemical Reviews pubs.acs.org/CR Review

Conformational Dynamics in RNA

Experiments for the Measurement of Carbon Relaxation Properties in Highly Enriched, Uniformly 13 C, 15 N-Labeled Proteins: Application to 13 C α Carbons

Extensive Backbone Dynamics in the GCAA RNA Tetraloop Analyzed Using 13 C NMR Spin Relaxation and Specific Isotope Labeling

Carbonyl Carbon Probe of Local Mobility in 13 C, 15 N-Enriched Proteins Using High-Resolution Nuclear Magnetic Resonance

Measurement of 15 N relaxation rates in perdeuterated proteins by TROSY-based methods

Sensitivity Improvement in Proton-Detected Two-Dimensional Heteronuclear Correlation NMR Spectroscopy

Pure Absorption Gradient Enhanced Heteronuclear Single Quantum Correlation Spectroscopy with Improved Sensitivity

Nuclear Magnetic Resonance Methods for Quantifying Microsecond-to-Millisecond Motions in Biological Macromolecules

Visualizing Transient Dark States by NMR Spectroscopy

Solution NMR Spin Relaxation Methods for Characterizing Chemical Exchange in High-Molecular-Weight Systems

Characterization of the Dynamics of Biomacromolecules Using Rotating-Frame Spin Relaxation NMR Spectroscopy

2D Heteronuclear NMR Measurements of Spin-Lattice Relaxation Times in the Rotating Frame of X Nuclei in Heteronuclear HX Spin Systems

Nuclear Magnetic Resonance Study of the Protolysis of Trimethylammonium Ion in Aqueous Solution Order of the Reaction with Respect to Solvent

Modified Spin-Echo Method for Measuring Nuclear Relaxation Times

Effects of Diffusion on Free Precession in Nuclear Magnetic Resonance Experiments

Invisible" Excited Protein States in Slow Exchange with a Major State Conformation

Very Fast Two-Dimensional NMR Spectroscopy for Real-Time Investigation of Dynamic Events in Proteins on the Time Scale of Seconds

A General Two-Site Solution for the Chemical Exchange Produced Dependence of T 2 upon the Carr-Purcell Pulse Separation

Studying Sparsely Populated Conformational States in RNA Combining Chemical Synthesis and Solution NMR Spectroscopy

Characterizing Slow Chemical Exchange in Nucleic Acids by Carbon CEST and Low Spin-Lock Field R 1ρ NMR Spectroscopy

Reaction Rates by Nuclear Magnetic Resonance

Measurement of Proton Chemical Shifts in Invisible States of Slowly Exchanging Protein Systems by Chemical Exchange Saturation Transfer

Simulations of NMR Pulse Sequences during Equilibrium and Non-Equilibrium Chemical Exchange

Off-Resonance R 1ρ NMR Studies of Exchange Dynamics in Proteins with Low Spin-Lock Fields: An Application to a Fyn SH3 Domain

Dynamic Basis for dG-dT Misincorporation via Tautomerization and Ionization

Optimization of Constant-Time Evolution in Multidimensional NMR Experiments

Improved 3D Triple-Resonance NMR Techniques Applied to a 31 KDa Protein

Investigation of Complex Networks of Spin-Spin Coupling by Two-Dimensional NMR

Homonuclear Broadband-Decoupled Absorption Spectra, with Linewidths Which Are Independent of the Transverse Relaxation Rate

Resolution Enhanced Homonuclear Carbon Decoupled Triple Resonance Experiments for Unambiguous RNA Structural Characterization

Base-Type-Selective High-Resolution 13 C Edited NOESY for Sequential Assignment of Large RNAs

Multisite Band-Selective Decoupling in Proteins

Frequency-Switched Single-Transition Cross-Polarization: A Tool for Selective Experiments in Biomolecular NMR

Excitation of Selected Proton Signals in NMR of Isotopically Labeled Macromolecules

Selective Cross-Polarization in Solution State NMR

NMR Probing of Invisible Excited States Using Selectively Labeled RNAs

Strong Coupling Effects during X-Pulse CPMG Experiments Recorded on Heteronuclear ABX Spin Systems: Artifacts and a Simple Solution

13 C α CEST Experiment on Uniformly 13 C-Labeled Proteins

Effects of J Couplings and Unobservable Minor States on Kinetics Parameters Extracted from CEST Data

Probing Excited Conformational States of Nucleic Acids by Nitrogen CEST NMR Spectroscopy

Direct Evidence for (G)O6···H 2-N4(C) + Hydrogen Bonding in Transient G(Syn)-C + and G(Syn)-m 5 C + Hoogsteen Base Pairs in Duplex DNA from Cytosine Amino Nitrogen off-Resonance R 1ρ

Relaxation Dispersion Measurements

Riboswitch Samples at Least Two Conformations in Solution in the Absence of Ligand: Implications for Recognition

Probing RNA Dynamics via Longitudinal Exchange and CPMG Relaxation Dispersion NMR Spectroscopy Using a Sensitive 13 C-Methyl Label

Extending the Range of Microsecond-to-Millisecond Chemical Exchange Detected in Labeled and Unlabeled Nucleic Acids by Selective Carbon R 1ρ NMR Spectroscopy

Widespread Transient Hoogsteen Base Pairs in Canonical Duplex DNA with Variable Energetics

Visualizing Transient Watson-Crick-like Mispairs in DNA and RNA Duplexes

Visualizing the Formation of an RNA Folding Intermediate Through a Fast Highly Modular Secondary Structure Switch

Probing Conformational Transitions Towards Mutagenic Watson-Crick-like G·T Mismatches Using Off-Resonance Sugar Carbon R 1ρ Relaxation Dispersion

Shortening the HIV-1 TAR RNA Bulge by a Single Nucleotide Preserves Motional Modes Over a Broad Range of Time Scales

Direct NMR Evidence That Transient Tautomeric and Anionic States in dG· dT Form Watson-Crick-like Base Pairs

Resolving Sugar Puckers in RNA Excited States Exposes Slow Modes of Repuckering Dynamics

Atomic Structures of Excited State A-T Hoogsteen Base Pairs in Duplex DNA by Combining NMR Relaxation Dispersion, Mutagenesis, and Chemical Shift Calculations

Invisible RNA State Dynamically Couples Distant Motifs

Evaluating the Uncertainty in Exchange Parameters Determined from Off-Resonance R 1ρ Relaxation Dispersion for Systems in Fast Exchange

Base-Pair Conformational Switch Modulates miR-34a Targeting of Sirt1 mRNA

Capturing Excited States in the Fast-Intermediate Exchange Limit in Biological Systems Using 1 H NMR Spectroscopy

Zhang, Q. Visualizing a Protonated RNA State That Modulates MicroRNA

An Excited State Underlies Gene Regulation of a Transcriptional Riboswitch

Measuring Residual Dipolar Couplings in Excited Conformational States of Nucleic Acids by CEST NMR Spectroscopy

Efficient Detection of Structure and Dynamics in Unlabeled RNAs: The SELOPE Approach

Rapid Assessment of Watson-Crick to Hoogsteen Exchange in Unlabeled DNA Duplexes Using High-Power SELOPE Imino 1 H CEST. Magn

Probing Microsecond Time Scale Dynamics in Proteins by Methyl 1 H Carr-Purcell-Meiboom-Gill Relaxation Dispersion NMR Measurements. Application to Activation of the Signaling Protein NtrC r

Quantifying Millisecond Exchange Dynamics in Proteins by CPMG Relaxation Dispersion NMR Using Side-Chain 1 H Probes

Visualizing Transient Low-Populated Structures of RNA

Relaxation-Optimized NMR Spectroscopy of Methylene Groups in Proteins and Nucleic Acids

2′-O-Methylation Can Increase the Abundance and Lifetime of Alternative RNA Conformational States

Pseudouridine and N 6 -Methyladenosine Modifications Weaken PUF Protein/RNA Interactions

A Potentially Abundant Junctional RNA Motif Stabilized by m 6 A and Mg 2+

m 6 A Minimally Impacts the Structure, Dynamics, and Rev ARM Binding Properties of HIV-1 RRE Stem IIB

Writing and Erasing mRNA Methylation

Dynamic RNA Modifications in Gene Expression Regulation

Gene Expression Regulation Mediated through Reversible m 6 A RNA Methylation

N 6 -Methyladenosine Modification of HCV RNA Genome Regulates Cap-Independent IRES-Mediated Translation via YTHDC2 Recognition

Hepatitis B Virus X Protein Recruits Methyltransferases to Affect Cotranscriptional N 6 -Methyladenosine Modification of Viral/Host RNAs

N 6 -Methyladenosine Modification of Hepatitis B Virus RNA Differentially Regulates the Viral Life Cycle

N 6 -Methyladenosine Modification and the YTHDF2 Reader Protein Play Cell Type Specific Roles in Lytic Chemical Reviews pubs

Viral Gene Expression During Kaposi's Sarcoma-Associated Herpesvirus Infection

N 6 -Methyladenosine of HIV-1 RNA Regulates Viral Infection

Dynamics of Human and Viral RNA Methylation During Zika Virus Infection

N6-Methyladenosine in Flaviviridae Viral RNA Genomes Regulates Infection

FTO-Dependent Demethylation of N6-Methyladenosine Regulates mRNA Splicing and is Required for Adipogenesis

N 6 -Methyladenosine-Dependent Regulation of Messenger RNA Stability

Nuclear m 6 A Reader YTHDC1 Regulates mRNA Splicing

Epitranscriptomic m 6 A Regulation of Axon Regeneration in the Adult Mammalian Nervous System

Mismatching Base-pair Dependence of the Kinetics of DNA-DNA Hybridization Studied by Surface Plasmon Fluorescence Spectroscopy

Real-Time Reliable Determination of Binding Kinetics of DNA Hybridization Using a Multi-Channel Graphene Biosensor

A Rule of Seven in Watson-Crick Base Pairing of Mismatched Sequences

Effects of Methylation on the Stability of Nucleic Acid Conformations. Studies at the Polymer Level

Effects of Methylation on the Stability of Nucleic Acid Conformations: Studies at the Monomer Level

Structure and Thermodynamics of N 6 -Methyladenosine in RNA: A Spring-Loaded Base Modification

Fluoride Ion Encapsulation by Mg 2+ Ions and Phosphates in a Fluoride Riboswitch

Widespread Genetic Switches and Toxicity Resistance Proteins for Fluoride

The Crystal Structure of Human Argonaute2

Structure of Yeast Argonaute with Guide RNA

MicroRNA Targeting Specificity in Mammals: Determinants beyond Seed Pairing

Determinants of Targeting by Endogenous and Exogenous MicroRNAs and siRNAs

The Structure of Human Argonaute-2 in Complex with miR-20a

Mechanisms of Post-Transcriptional Regulation by MicroRNAs: Are the Answers in Sight?

Beyond the Seed: Structural Basis for Supplementary MicroRNA Targeting by Human Argonaute2

Argonaute Divides Its RNA Guide into Domains with Distinct Functions and RNA-Binding Properties

An Interplay of miRNA Abundance and Target Site Architecture Determines miRNA Activity and Specificity

Pairing Beyond the Seed Supports MicroRNA Targeting Specificity

Structure-Based Cleavage Mechanism of Thermus Thermophilus Argonaute DNA Guide Strand-Mediated DNA Target Cleavage

Propagation and Cleavage of Target RNAs in Ago Silencing Complexes

Probing Slow Timescale Dynamics in Proteins Using Methyl 1 H CEST

Separating Dipolar and Chemical Exchange Magnetization Transfer Processes in 1

Hsp70 Biases the Folding Pathways of Client Proteins

Off-Resonance Rotating-Frame Amide Proton Spin Relaxation Experiments Measuring Microsecond Chemical Exchange in Proteins

A New Amide Proton R 1ρ Experiment Permits Accurate Characterization of Microsecond Time-Scale Conformational Exchange

Using Amide 1 H and 15 N Transverse Relaxation To Detect Millisecond Time-Scale Motions in Perdeuterated Proteins: Application to HIV-1 Protease

Extending the Sensitivity of CEST NMR Spectroscopy to Micro-to-Millisecond Dynamics in Nucleic Acids Using High-Power Radio-Frequency Fields

Cryo-EM Structures of Full-Length Tetrahymena Ribozyme at 3.1 Å Resolution

Accelerated Cryo-EM-Guided Determination of Three-Dimensional RNA-Only Structures

A Viral RNA Hijacks Host Machinery Using Dynamic Conformational Changes of a tRNA-like Structure

Protein In-Cell NMR Spectroscopy at 1.2 GHz

Cryogenically Cooled ProbesA Leap in NMR Technology

Aromatic 19 F-13 C TROSY: A Background-Free Approach to Probe Biomolecular Structure, Function, and Dynamics

19 F NMR Studies of a Desolvated Near-Native Protein Folding Intermediate

The Role of Dimer Asymmetry and Protomer Dynamics in Enzyme Catalysis

Conformational Selection and Functional Dynamics of Calmodulin: A 19 F Nuclear Magnetic Resonance Study

19 F NMR Reveals Multiple Conformations at the Dimer Interface of the Nonstructural Protein 1 Effector Domain from Influenza A Virus

A Suite of 19 F Based Relaxation Dispersion Experiments to Assess Biomolecular Motions

Unveiling the Activation Dynamics of a Fold-Switch Bacterial Glycosyltransferase by 19 F NMR

Long-Range RNA Structural Information via a Paramagnetically Tagged Reporter Protein

Recovering Lost Magnetization: Polarization Enhancement in Biomolecular NMR

BEST-TROSY Experiments for Time-Efficient Sequential Resonance Assignment of Large Disordered Proteins