Folds and Functions of Domains in RNA Modification

Folds and Functions of Domains in RNA Modification Enzymes Anna Czerwoniec, Joanna M. Kasprzak, Katarzyna H. Kaminska, Kristian Rother, Elzbieta Purta and Janusz M. Bujnicki*

Abstract

T

his chapter provides a short review of the structural biology of RNA modification enzymes, focused on comparative aspects. All modifications are introduced by protein enzymes, from relatively simple standalone catalytic domains to subunits of protein complexes or ribonucleoprotein particles. These enzymes typically comprise domains that are often specialized in the catalysis of the modification reaction or in recognition and binding of the macromolecular RNA substrate, underscoring structural and functional modularization of proteins and the interplay of these modules in complex macromolecular systems. We provide a catalog of three‑dimensional folds and molecular functions of domains implicated in RNA modifications and discuss evolution‑ ary pathways that connect different modes used by enzymes to interact with their substrates. We highlight cases of convergent evolution for unrelated enzymatic domains catalyzing very similar reactions and ‘promiscuous’ domains used for substrate recognition.

Introduction

The first RNA modification enzyme, m5U methyltransferase specific for position 54 in tRNA, now designated TrmA, has been discovered almost 50 years ago independently by two groups: Borek’s1 and Svensson’s.2 Since then, numerous new RNA modification activities have been characterized biochemically and genetically (many of them in the 1970s and 1980s). For most of them, the respective open reading frames have also been identified, prompting studies of sequence‑function relationships. Cloning of genes encoding RNA modification enzymes and their characterization at the molecular level has been particularly facilitated by whole‑genome sequencing during the last decade. However, the most solid link between sequence and function has been provided by structural studies. The first structures of RNA modification enzymes have been reported in 1996: for the tRNA‑guanine transglycosylase (TGT) from Zymomonas mobili, by the Suck group3 and for the vaccinia virus protein VP39 that acts as both an mRNA cap‑specific RNA 2′‑O‑methyltransferase and a poly(A) polymerase processivity factor, by the Quiocho group.4 Since then, many other structures of enzymes representing different activities and specificities have been determined. Most recently, structural genomics efforts have resulted in accumulation of structures for functionally uncharacterized proteins, some of which correspond to putative RNA modification enzymes. Bioinformatics analyses have helped to combine these different types of experimental data, leading to discoveries of new RNA modification enzymes based on sequence analyses as well as structural considerations.5‑7 *Corresponding Author: Janusz M. Bujnicki—Laboratory of Bioinformatics and Protein Engineering, International Institute of Molecular and Cell Biology, Ks. Trojdena 4, PL‑02‑190 Warsaw, Poland. Email: [email protected]

DNA and RNA Modification Enzymes: Structure, Mechanism, Function and Evolution edited by Henri Grosjean. ©2009 Landes Bioscience.

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Chapter 21

DNA and RNA Modification Enzymes

Detailed experimental information exists for a small fraction of all enzymes implicated in known RNA modification reactions. Only about half of  >120 reactions listed in the MODOMICS database8,9(see also Appendix 1 in this volume) have the respective enzymes characterized experimen‑ tally (genetically and/or biochemically). This means that for the other reactions that are predicted to take place, the corresponding enzymes are yet to be discovered. Furthermore, for less than 40 of the reactions, crystal structures of the respective enzymes are available and only a handful of these have the RNA substrate bound. Thus, in most cases protein‑RNA interactions have been inferred only by means of theoretical methods, such as macromolecular docking. However, even this limited number of structures has revealed many aspects of the molecular basis for recognition of RNA substrates and catalysis of the various RNA modification reactions (see other chapters of this book).

The Diversity of 3D‑Folds in RNA Modification Enzymes

Thus far, the largest number of structures has been determined experimentally for enzymes that introduce the most common modifications, namely methyltransferases (MTases) and pseudouri‑ dine synthases (PSIases), the other enzymatic activities have only one or a few representatives. Thus, in parallel with experimental structure determination for selected RNA modification enzymes, bioinformatics methods have been used to detect evolutionary relationships and make structural predictions. In particular, the fold‑recognition technique can identify a suitable structural template (a protein with known structure) for a given sequence and then a structural model for this sequence can be built by comparative modeling. Theoretical models are typically significantly less reliable than those derived from crystallographic or NMR experiments, therefore they must be interpreted with caution. In particular the potential accuracy of each theoretical model should be always carefully assessed (i.e., reliable and unreliable parts must be identified) before any functional inferences are made. We estimate that reasonably accurate models can be built for ∼60% of those currently known RNA modification enzymes, for which experimental structures are not available (A. Czerwoniec, unpublished data). Such models can be used for identification and characterization of function‑ ally important regions, e.g., active sites and RNA‑binding sites. Several published bioinformatics predictions have been validated by independent crystallographic analyses, proving the relative ac‑ curacy of theoretical models. Such examples include isopentenyltransferase MiaA and hydroxylase MiaE involved in the biosynthesis of the hypermodified nucleoside ms2io6A37 in tRNA.10 Our group has specialized in building theoretical models for methyltransferases (see e.g., refs. 7,11‑13). Theoretical models published in the literature and not yet superseded by crystallographic analyses are collected and stored in the MODOMICS database.8 Table 1 shows a list of currently known folds implicated in the catalysis of RNA modifications—here we list only single examples per fold per function, regardless of the number of structurally characterized representatives.

Catalytic Domains in RNA Methyltransferases

Thus far, 67 different methylated ribonucleosides have been discovered, including methyla‑ tions of ribose 2′‑hydroxyls and various atoms of bases and functional groups introduced by other RNA‑modification enzymes.8,14 Twenty one of these modified nucleosides possess multiple methylations. This variety at the level of the reaction substrates is reflected also at the protein level: catalytic domains of RNA methyltransferases have been found to belong to 4 unrelated superfamilies characterized by different three‑dimensional folds (Fig. 1), namely: Rossmann—fold methyltransferases (RFM), SPOUT, Radical‑SAM and the FAD/NAD(P)‑binding protein (reviews:refs. 6,7,15‑17). Among these proteins, only the SPOUT fold has not yet been found to catalyze reactions other than methylation, while other “methyltransferase folds” are known to pos‑ sess members catalyzing also other reactions. Enzymes from the RFM, SPOUT and Radical‑SAM superfamilies utilize AdoMet as the methyl group donor, while the methyltransferase with the FAD/NAD(P)‑binding domain utilizes 5,10‑methylenetetrahydrofolate (5,10‑CH2‑THF). On the other hand, there are AdoMet‑dependent methyltransferases that belong to four other folds (SET, MetH reactivation domain, tetrapyrrole methylase and ICMT), but thus far they have not been found to act on RNA.

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274

Tgt (1wke) TM0096 (1vhn) MiaB Tyw1 (2yx0) Cfr TrmB (2fca) Tyw2 TrmD (1p9p) TrmFO MnmC GidA (2cul) MnmE (1xzq) MiaA (3crm) MiaE (2itb) TruA (1dj0) ThiI (2c5s) TilS (1ni5) YadB (1nzj) MnmA (2der) Tad2p QueA (1vky) TmcA GTase (1cko) TPTase (1i9t) Cet1p (1d8i)

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transglycosylase dihydrouridine synthase methylthiotransferase ring closure in yW synthesis methyltransferase methyltransferase aminopropyltransferase methyltransferase methyltransferase demodification of cmnm5U to nm5U modification of U to cmnm5U (with MnmE) modification of U to cmnm5U (with GidA) dimethylallyltransferase hydroxylase pseudouridine synthase thiouridine synthase lysidine synthase glutamylates the modified base Q replaces oxygen by sulphur to create s2U deaminase AdoMet:tRNA ribosyltransferase‑isomerase N4‑cytidine acetyltransferase mRNA capping enzyme mRNA triphosphatase mRNA triphosphatase

TIM beta/alpha‑barrel TIM beta/alpha‑barrel TIM beta/alpha‑barrel TIM beta/alpha‑barrel TIM beta/alpha‑barrel Rossmann‑fold methyltransferase (RFM) Rossmann‑fold methyltransferase (RFM) alpha/beta knot (SPOUT) FAD/NAD(P)‑binding domain FAD/NAD(P)‑binding domain FAD/NAD(P)‑binding domain P‑loop NTPases P‑loop NTPases ferritin‑like pseudouridine synthase adenine nucleotide alpha hydrolase‑like adenine nucleotide alpha hydrolase‑like adenine nucleotide alpha hydrolase‑like adenine nucleotide alpha hydrolase‑like cytidine deaminase‑like QueA‑like GNAT ATP‑grasp phosphotyrosine protein phosphatase II CYTH‑like phosphatase

tRNA‑guanine transglycosylase DUS Radical‑SAM Radical‑SAM Radical‑SAM RFM RFM SPOUT FAD/NAD‑linked reductases D‑aminoacid oxidase GidA‑like G proteins nucleotide and nucleoside kinases ferritin‑like PUS ThiI‑like PP‑loop ATPase class I aminoacyl‑tRNA synthetase N‑Type ATP pyrophosphatase cytidine deaminase‑like QueA‑like N‑acetyltransferase guanylyltransferase dual specificity phosphatase‑like CYTH‑like phosphatase

Representative Protein and Pdb Code (if available)

Domain Fold Specific Superfamily or Family Enzyme Catalytic Function

Table 1. Folds of catalytic domains involved in RNA modification

Folds and Functions of Domains in RNA Modification Enzymes 275

DNA and RNA Modification Enzymes

Figure 1. Different folds for similar modifications: the case of RNA methyltransferases. Protein molecules are shown in the cartoon representation. Ligands and RNA (if available) are shown as black sticks. The conserved core of the catalytic domain is shown in red, additional elements are shown in grey. A) mRNA cap 1 methyltransferase VP39 with the RFM catalytic domain (monomer, 1av6), B) Cfr methyltransferase with the Radical‑SAM catalytic domain (theoretical model; K.H.K. and J.M.B. unpublished data); C) rRNA methyltransferase RlmH with the SPOUT catalytic domain (dimer; crystal structure, 1ns5: here only the single subunit is shown), D) The FAD/NAD(P)‑binding domain of the TrmFO protein (theoretical model; J.M.B., unpublished data). A color version of this image is available at www.landesbioscience.com/curie

Most of the RNA methyltransferases characterized so far belong to the RFM superfamily, with SPOUT being the second most “popular” fold among these enzymes.17,18 Among Radical‑SAM enzymes only two RNA methyltransferases have been identified to date, Cfr19 and RlmN.20 FAD/ NAD(P)‑binding proteins have only one representative among RNA methyltransferases, that of TrmFO.21,22 There are no crystal structures available for Cfr, RlmN or TrmFO, however molecular models can be built based on their homology to other proteins with known structures. Cfr/RlmN can be modeled based on the structure of e.g., molybdenum cofactor biosynthesis protein MoaA,23 while TrmFO can be modeled based on the structure of another RNA‑modification enzyme GidA, which is involved in the addition of a carboxymethylaminomethyl group to U34 in tRNA.24 All four folds of RNA methyltransferases are characterized by the “Rossmanoidal” α/β ar‑ chitecture (a parallel β‑sheet surrounded on one or both sides by α‑helices), but with different topologies. FAD/NAD(P)‑utilizing enzymes exhibit a typical Rossmann fold with six parallel strands arranged in the following order: ↓6‑↓5‑↓4‑↓1‑↓2‑↓3 and the cofactor‑binding site formed by loops following strands 1, 2 and 3.25 The topology of the RFM fold is (as the name implies) very similar to the Rossmann fold, with an additional, 7th b‑strand inserted into the sheet in an antiparallel manner (↓6‑↑7‑↓5‑↓4‑↓1‑↓2‑↓3).26 The nucleotide moiety of the co‑ factor is bound in a very similar way as in the classical Rossmann‑fold enzymes. Radical‑SAM

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Folds and Functions of Domains in RNA Modification Enzymes

277

Plethora of Functions in Radical‑SAM Enzymes

Among catalytic domains involved in RNA modification, the largest diversity of functions is seen in enzymes with the ubiquitous Rossmann‑like folds, including TIM barrel and FAD/ NAD(P)‑binding domains. Among superfamilies (within folds) special attention should be paid to the Radical‑SAM enzymes that are known for catalyzing very diverse reactions30 (see examples in a chapter by Atta, Fontecave and Mulliez in this book). Thus far, they have been implicated in RNA modification reactions as methyltransferases (Cfr and RlmN),19,20 methylthiotransferases (MiaB31 and enzymes catalyzing the tri‑ring‑formation step in Wye‑base biosynthesis (Tyw1), with the substrate tRNA bearing m1G37.32 Radical AdoMet—dependent mechanism is often used to catalyze the reactions in which the substrate activation is difficult (e.g., methylation of intracyclic carbon or ring formation).

The RFM Domain

The RFM superfamily comprises methyltransferases acting on various substrates, including RNA, DNA, proteins, lipids and a plethora of small molecules. A small fraction of RFM domains have been found to use AdoMet to catalyze different reactions (review: 17,33). In the case of RNA‑modifying enzymes such an unusual activity has been found for the Tyw2 enzyme involved in formation of the hypermodified base wybutosine (yW). Tyw2 acts similarly to MTases, but it transfers the α‑amino‑α‑carboxypropyl group of AdoMet (instead of the methyl group) to the C‑7 position of the imG‑14 intermediate to form yW‑86.34 Besides, some members of the RFM superfamily have lost the catalytic activity, but retained a function related to substrate binding or formation of an oligomeric protein complex. A number of such cases have been described in the context of RNA modification, including the Gcd10p subunit of the tRNA:m1A58 methyltrans‑ ferase Trm6,35,36 the RNA‑binding domain of the 16S rRNA:m2G1207 methyltransferase RsmC,37 the Kar4p protein, a paralog of the catalytic subunit of mRNA methyltransferase Ime4p38 and the D12 subunit of the vaccinia virus mRNA:m7G (cap 0) methyltransferase.39

Functional Convergence: The Same Function Provided by Different Folds

Interestingly, a number of cases of convergent evolution have been observed, with unrelated catalytic domains introducing the same type of methylation at the same site of homologous RNAs in different organisms. Examples include the aforementioned tRNA:m5U54 methyltransferase TrmA in Escherichia coli (an AdoMet‑dependent member of the RFM superfamily) and its un‑ related functional analog TrmFO in e.g., Bacillus subtilis (a 5,10‑CH2‑THF‑dependent FAD/ NAD(P)‑binding protein).21,22 The largest functional overlap is between enzymes from the RFM and SPOUT superfamilies, which both have been found to include the 2′‑O‑ribose and m1G methyltransferases. For instance, the 2′‑O‑ribose methylation at position 32 in tRNA is carried out by a RFM superfamily member Trm7p in Eukaryota40 and by a SPOUT superfamily member TrmJ in Bacteria.41 Likewise, m1G methylation at position 37 in tRNA is carried out by a RFM superfamily member Trm5p in Eukaryota and Archaea42,43 and by a SPOUT superfamily member TrmD in Bacteria.29,44 Interestingly, despite acting on the same substrate, Trm5p and TrmD exhibit distinct modes of tRNA recognition, with Trm5p recognizing the integrity of the L shape of the tRNA molecule and TrmD requiring just the anticodon loop capped with a stem of minimally 9 base pairs.45

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enzymes belong to the TIM‑barrel fold, which comprises eight tandemly arranged α/β ele‑ ments (‑↓1‑↓2‑↓3‑↓4‑↓5‑↓6‑↓7‑↓8‑); most of Radical‑SAM enzymes, however, lack two of these α/β elements and exhibit a ‘3/4 barrel’ architecture.27,28 Despite structural similarity to the Rossmann‑fold, Radical‑SAM enzymes bind the cofactor in a completely different way, by means of coordination by a Fe‑S cluster. Finally, the SPOUT fold has a β‑sheet with a different topology (↓5‑↓3‑↓4‑↓1‑↓2) and is characterized by the presence of a very deep knot, in which the C‑terminal a‑helix is threaded through a loop between strands 3 and 4.29 This knot forms a cofactor‑binding site.

278

DNA and RNA Modification Enzymes

A number of enzymes possess a combination of several catalytic domains. In some cases these domains catalyze consequent reactions in one pathway, as exemplified by the MnmC protein, which catalyzes the last two steps in the biosynthesis of 5‑methylaminomethyl‑2‑thiouridine (mnm5s2U) in tRNA. The C‑terminal FAD‑binding domain from the DAAO superfamily catalyzes demodifica‑ tion of cmnm5(s2)U to nm5(s2)U, while the N‑terminal domain from the RFM superfamily inde‑ pendently methylates nm5(s2)U to form the final product mnm5(s2)U.46,47 An even more complex fusion is found in plant homologs of enzymes involved in the wybutosine (yW) biosynthesis, where a single polypeptide contains orthologs of enzymes Tyw2p and Tyw3p, both with catalytic domain from the RFM superfamily, but encoding different transferase activities involved in transformation of imG‑14 to yW‑86 to yW‑72 and a C‑terminal (noncatalytic) domain homologous to Tyw4p, another enzyme from the same pathway involved in transformation of yW‑72 to yW‑58 and finally to yW (see also chapter by Urbonavicius et al in this book).34 In other cases, fused domains may be involved in completely different processes, e.g., as in the case of the Rib2/Pus8p protein, whose N‑terminal domain carries the cytoplasmic tRNA:Ψ32‑synthase activity, while the C‑terminal domain has a DRAP‑deaminase activity required for riboflavin biogenesis.48

Domains Involved in RNA‑Binding: Three Major Modes of Substrate Recognition

The catalysis of reactions involving RNA is similar to the catalysis of analogous reactions on DNA and many catalytic domains of RNA‑modification enzymes have close homologs among DNA‑modifying enzymes (examples include methyltransferases and deaminases, as exemplified in several other chapters in this book). Crystal structures of RNA modification enzymes have began to unravel how these proteins recognize the correct target nucleoside(s) in large folded or partially folded RNAs. Thus far, three major modes of interaction with the substrate have been found for RNA modification enzymes, distinguished by the character of the structural elements used for specific substrate recognition (Fig. 2). In the first mode, the binding and catalytic func‑ tions are shared by the same domain. In the second mode, catalytic domains are fused to or form a tight complex with specialized RNA‑binding domains. In the third mode, the catalytic domain forms a complex with a guide RNA that specifically recognizes the substrate (usually with help of several accessory proteins).

Figure 2. Modes of RNA recognition in RNA modification enzymes—exemplified by pseudou‑ ridine synthases. In the upper left corner of each panel, a schematic picture is shown. The common catalytic domain (in the same orientation in all three panels) is indicated in red, PUA domain is in blue, guide RNA is in green, RNA substrate is in black, other elements are in gray. A) Mode 1, catalytic domain alone, but as a homodimer: TruA (1dj0); B) Mode 2, catalytic domain with an RNA‑binding domain (PUA), TruB (1k8w); C) Mode 3, protein complex with an RNA guide, H/ACA snoRNP (2rfk). A color version of this image is available at www.landesbioscience.com/curie

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Multifunctional RNA Modification Enzymes

Folds and Functions of Domains in RNA Modification Enzymes

279

In mode 1, ‘minimalist’ enzymes comprise essentially only the catalytic domain (Fig. 1A,B). They may recognize large, well‑structured RNAs, as exemplified by the recently characterized 23S rRNA m3Ψ1915 methyltransferase RlmH (formerly YbeA) from the SPOUT superfamily that acts on a fully assembled ribosome49 and, according to a docking model, forms extensive contacts with the interfaces of both the 30S and 50S subunit.50 Another example is provided by the cap‑specific RNA 2′‑O‑methyltransferase VP39,51 which binds conformationally flexible mRNA and therefore must recognize very specific structural features. The enzymes utilizing mode 1 for binding often increase the size of the surface available to interact with the macromolecular RNA substrate by forming homooligomers comprising several identical copies of the catalytic domain (as in the case of RlmH) or by developing ‘decorations’ in the form of various protrusions that are used to recognize particular features of the substrate (as in the case of cap‑recognition by VP39).

Mode 2

In mode 2, the catalytic domain is assisted by an additional RNA‑binding domain. The most common type of RNA recognition involves fusions of the catalytic domains with unrelated domains that often belong to large promiscuous superfamilies implicated not only in RNA modification, but in RNA metabolism in general or in even broader functional context such as oligosaccharide/oligonucleotide binding (OB‑fold domains). The RNA‑binding domain can be also noncovalently bound to form a complex. Many putative RNA‑binding domains have been initially identified by bioinformatics methods in the course of large‑scale comparative analyses of proteins known for their involvement in RNA modification. Here, the group of Koonin and Aravind excelled in identifying (and naming) putative RNA‑binding domains present in pro‑ teins with different enzymatic activities. The most notable domains include: PUA (named after PseudoUridine synthase and Archaeosine transglycosylase)52 (discussed in more detail below), THUMP (named after 4‑THioUridine, Methyltransferases and Pseudouridine synthases)53 and

Figure 3. Examples of RNA‑binding domains of RNA modification enzymes. Catalytic do‑ mains are indicated in red, RNA‑binding domains are indicated in blue, other domains (if present) are indicated in cyan or gray. RNA (if present) is indicated in black. A) S4 domain in pseudouridine synthase RsuA (1vio); B) L30‑like domain in rRNA 2′‑O‑ribose methyltrans‑ ferase RlmB (1gz0); C) A degenerated copy of the RFM domain used for RNA binding in rRNA m2G methyltransferase RsmC (2pjd); D) Pre‑THUMP (cyan) and THUMP (blue) domains in thiouridine synthase ThiI (2c5s); E) TRAM (blue) and EEHEE (cyan) domains in rRNA m5U methyltransferase RumA (2bh2); C) Pre‑PUA domain, cystatin‑like fold (cyan) and PUA do‑ main (blue) in guanosine transglycolase TGT (1iq8). A color version of this image is available at www.landesbioscience.com/curie

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Mode 1

280

DNA and RNA Modification Enzymes

Domain Fold

Enzyme Catalytic Function

Representative Protein and PDB Code (if available)

THUMP

methyltransferase

Trm11p, YcbY

THUMP

thiouridine synthase

ThiI (2c5s)

THUMP

pseudouridine synthase

Pus10p (2v9k)

PUA

methyltransferase

RsmE (1nxz)

PUA

pseudouridine synthase

TruB (1k8w)

PUA

archaeosine transglycolase

ArcTGT (1iq8)

cystatin‑like (pre-PUA)

archaeosine transglycolase

ArcTGT (1iq8)

DSRDB

dihydrouridine synthase

Smm1p

DSRDB

deaminase

ADAR

DSRDB

methyltransferase

CORYMBOSA2

OB fold

guanylyltransferase

GTase (1cko)

OB fold (TRAM)

methylthiotransferase

MiaB

OB fold (TRAM)

methyltransferase

RlmD (2bh2)

EEHEE

methyltransferase

RlmD (2bh2)

LA

methyltransferase

CORYMBOSA2

KH

thiouridine synthase

MTH271

S4

pseudouridine synthase

RsuA (1vio)

RAGNYA

wybutosine biosynthesis

Tyw3p

NusB‑like

methyltransferase

RsmB (1sqf)

flavodoxin‑like

methyltransferase

TrmI (2yvl)

L30

Methyltransferase

RlmB (1gz0)

TRAM (named after tRNA methyltransferase Trm2p and methylthiotransferase MiaB)54 (Fig. 3). Some other putative RNA‑binding domains of RNA modification enzymes have been found based on detection of homology to ribosomal proteins, e.g., S4 or L30. Notably, crystallographic analyses allowed the unification of theoretically predicted domain families into broader super‑ families, e.g., the structure of 23S rRNA:m5U1939 methyltransferase RlmD (formerly RumA) revealed that TRAM is a variant of the aforementioned OB‑fold.55 Interestingly, a homolog of RlmD, the TrmA enzyme, which acts on tRNA, lacks the TRAM domain,56 demonstrating how related enzymes may differ in their ability to recognize the substrate due to gains or losses of auxiliary domains (see below). It is important to remember, however, that for many of the afore‑mentioned domains direct experimental evidence of autonomous RNA‑binding ability (in the absence of the catalytic domain) is not available.

PUA: Example of an RNA‑Binding Domain Present in Many Enzymes Involved (Not Only) in RNA‑Modification

The PUA domain was initially identified through sequence analyses in many different RNA‑binding proteins, in particular in RNA modification enzymes, including pseudouridine synthases, tRNA‑guanine transglycosylases (ArcTGTs) and RNA methyltransferases. 52 The

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Table 2. Common folds of RNA‑binding domains found in RNA modifying‑enzymes, with examples

281

three‑dimensional structure of the PUA domain (a pseudobarrel with a sheet of 5 strands; order ↓1‑↓3‑↑4‑↓5‑↑2) has been revealed by crystallographic analysis of the ArcTGT enzyme.57 Figures 2B/2C and 3C show examples of PUA domains associated with pseudouridine synthases and Arc‑TGT, respectively. Other processes that involve PUA domains include ribosome biogenesis and translation initiation. The PUA fold has been recently identified in the YTH domain58 found in e.g., RNA‑binding protein YT521‑B involved in regulation of alternative splicing (Protein Data Bank, unpublished structure 2yud, DOI: 10.2210/pdb2yud/pdb). Interestingly, PUA domains have been also identified in a functional context that has little to do with RNA metabolism, e.g., in sulfate reductases and glutamate kinases, where they are believed to fulfill regulatory roles.59 PUA domains are related to the ASCH superfamily, which is found in proteins implicated mostly in regulation of pre-mRNA processing and splicing rather than RNA modification.60 For a com‑ prehensive review of structures and functions of PUA domains see reference 61.

Mode 3

In mode 3, the oligomeric complex further binds a special guide RNAs, which is used to rec‑ ognize the target through transient base pair interactions with the substrate RNA. This mode of action is used by RNA modification systems called small nucleolar ribonucleoproteins (snoRNPs) in Eukaryota and small ribonucleoproteins (sRNPs) in Archaea. So far, these systems have not been detected in any Bacteria. The first system, the 2′‑O‑ribose methyltransferase complex comprises the catalytic subunit Nop1p/fibrillarin (a member of the RFM superfamily of methyltransferases), auxiliary proteins Nop56p, Nop58p and Snu13p and binds a guide RNA from the C/D box family62 (reviews: ref. 63 see also the chapter by Gagnon, Qu and Maxwell in this book). The second system, pseudouridine synthase complex comprises the catalytic subunit Cbf5p/dyskerin, auxiliary proteins Gar1, Nop10p and Nhp2p and binds a guide RNA from the H/ACA box fam‑ ily (reviews: refs. 63,64, see also the chapter by Grozdanov and Meier in this book). Other types of RNA‑guided systems are RNA editing in mitochondria of kinetoplastid protists, where three different editosome complexes excise or insert nucleotides into the substrate RNA (review: ref. 65) or RNA interference.66

Evolution of RNA Binding Modes

Structural and bioinformatics analyses revealed that there are evolutionary pathways connect‑ ing the different quaternary structures and the associated modes of RNA binding. The “easiest” modification is to evolve oligomerization: there are families of close homologs, which display completely different quaternary structures, as for instance tRNA:m7G46 methyltransferases, which include a monomer (TrmB) in E. coli,67 a homodimer (TrmB) in B. subtilis68 and a heterodimer of unrelated proteins (Trm8‑Trm82) in S. cerevisiae69. A number of RNA modification enzymes with known crystal structures are known to form symmetrical homooligomers. Because the substrate RNA is asymmetric, a protein subunit in the symmetric protein homooligomer must often evolve two different sites of interaction—e.g., one to perform the catalysis and another to recognize and bind another part of the substrate, adjacent to the one being modified. A very common evolutionary mechanism, namely gene duplication, can generate an additional (initially identical) copy of the catalytic domain and provide an opportunity for the two copies to undergo divergent evolution. The diverging paralogous copies (e.g., α and β) of the ancestral oligomeric protein αx may form independent homooligomers αx and βx of which one may e.g., develop a completely new specificity. This mechanism accounts for the largest fraction of new functions, in particular new specificities as well as new enzymatic activities, developed in the course of evolution. For instance most RFM superfamily methyltransferases that modify the C5 atom of pyrimidine in DNA or RNA are believed to have evolved from one ancestral enzyme by duplication and divergence.70,71 However, another possibility is for the paralogous proteins to form a new heterooligomer αyβz (often with the same stoichiometry as in the ancestral protein, in which subunits may undergo specialization and subfunctionalization. This mechanism leads to the development of heterooligomeric complex comprising homologous, but not identical sub‑ units, of which one often seems “degenerated” in the active site, while the other is unable to bind

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Folds and Functions of Domains in RNA Modification Enzymes

DNA and RNA Modification Enzymes

the substrate on its own. Such examples include deaminases that form inosine at position 34 in tRNAs (homodimeric TadA in bacteria72 and heterodimeric Tad2p/Tad3p in Eukaryota73), as well as tRNA:m1A58 methyltransferases (a homotetramer TrmI in bacteria and α2β2 heterotetramer composed of Trm6p and Trm61p subunits in Eukaryota35,36). A similar situation can occur with intragenic domain duplication, leading to a monomeric protein with two homologous domains, one being specialized towards catalysis, another towards substrate RNA binding, as exemplified by the 16S rRNA:m2G1207 methyltransferase RsmC,37 a “pseudodimeric” monomer which has structurally characterized homodimeric homologs.74 Gene duplication, combined with inter‑ and intragenic recombination, is the main mechanism contributing to the diversity of proteins using mode 2 to recognize the substrate. In particular, the acquisition of a substrate‑binding domain (e.g., a copy of a pre‑existing substrate‑binding domain of another protein) by a catalytic domain can lead to a rapid change of substrate specificity in the resulting protein. Although to our knowledge this has not yet been demonstrated for RNA modi‑ fication enzymes, experimental evidence exists for related DNA modification enzymes (e.g., DNA methyltransferases) that shuffling of specificity‑determining domains between catalytic domains results in proteins with predictably altered substrate preferences.75 Likewise, addition of generic DNA‑binding specificity domains (e.g., Zn‑fingers) to relatively nonspecific catalytic domains may result in DNA‑modifying enzymes with very stringent substrate specificity.76 In mode 3 of RNA recognition, RNA‑binding domains that are bound as subunits of the complex are often relatives of RNA‑binding domains fused to the catalytic domain by proteins using mode 2. For instance the PUA domain in the H/ACA pseudouridine synthase ribonucleo‑ protein complex (e.g., the Cbf5 protein) is a direct counterpart of a PUA domain present in the single‑subunit pseudouridine synthase TruB. In addition, Nhp2p and Snu13p components of otherwise unrelated H/ACA and C/D box snoRNPs are homologs—further, in their Archaeal counterparts they are replaced by the same ribosomal protein L7Ae (review: ref. 63). Interestingly, the Archaeal ortholog of a protein Cbf5 that introduces Ψ into multiple sites by a mechanism depending on H/ACA guide RNA and a set of essential accessory proteins was found to specifi‑ cally introduce Ψ55 in tRNA in vitro in the absence of any guide RNA or accessory proteins. Further, the activity of archaeal Cbf5 ortholog towards tRNA U55 is enhanced in the presence of the accessory proteins that form the H/ACA sRNP complex (also without the guide RNA).77,78 This indicates that acquisition of additional components can lead to evolution of a very complex system (e.g., exhibiting mode 3 of RNA recognition) from a much simpler ancestor.

General Features of Domains in RNA‑Modifying Enzymes and their Relationship to DNA‑Modifying Enzymes

RNA‑modifying enzymes share only very few generic features. Both catalytic and sub‑ strate‑binding domains are typically of α/β folds, i.e., the most typical architecture implicated in RNA binding.79 They also tend to exhibit positively charged surface patches, characteristic for pro‑ teins that bind negatively charged ligands such as the phosphate backbone. Substrate recognition by RNA‑modifying enzymes differs from DNA‑modifying enzymes because in many substrates the complex tertiary structures of RNA prevent direct recognition of the target sequence.80 In other RNA substrates, their flexibility allows for easy access to Watson‑Crick edges of the bases that are typically protected in DNA bases due to base‑pairing. Because of such flexibility, RNA‑modifying enzymes have developed a number of different strategies for substrate recognition and binding. One common feature shared by some RNA‑ and DNA‑modifying enzymes is the so‑called “base‑flipping” mechanism initially discovered for DNA methyltransferases81 and later found also for pseudouridine synthases82 and RNA methyltransferases83 (see also several chapters in this book). Interestingly, clear evolutionary links have been identified for several DNA and RNA modification enzymes, the most notable being probably DNMT2, a member of the DNA:m5C methyltransferase family, whose name is actually an abbreviation for “DNa MethylTransferase 2” and which has been found to specifically and efficiently methylate C‑5 atom of C38 in tRNAAsp  84(review: ref. 85). The cytidine deaminase‑like superfamily also groups together related DNA and RNA modifica‑

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tion/editing enzymes (see also chapters by Smith and by Forterre and Grosjean in this book). For instance, a recently characterized enzyme from Trypanosoma can mediate A‑to‑I editing of tRNA and C‑to‑U deamination of ssDNA but not both in either substrate.86 Besides, different substrate specificities are observed among closely related enzymes from the AID/APOBEC family: AID and APOBEC3 deaminate DNA to trigger pathways in adaptive and innate immunity, while APOBEC1 mediates apolipoprotein B RNA editing.87 Experimental structural analyses have provided milestones for our understanding of RNA modification pathways. The whole roadmap, however, has been charted to a large extend by bioinformatics analyses, which provided links between different domains, explaining (or predict‑ ing) their functional similarities based on detection of homology. Currently, we begin to unravel the static structures of large RNP complexes involving RNA modification enzymes.63 However, experimental and theoretical analyses have to be combined more efficiently to infer the full picture of “structural systems biology” of RNA modification in terms of pathways and networks of interactions.88 The next challenge is to rationally (re)design the domains and complexes and their protein‑RNA interactions and modify the behavior of systems (i.e., synthetic biology).89,90 In order to achieve this goal, we need to introduce a time component to the three‑dimensional pictures.91 Thus, we need to develop new approaches to and integrate data from many different sources, including high‑resolution 3D structures, theoretical structural models, biochemical and biophysical experiments involving both high‑throughput and single molecule studies, as well as computer simulations. Fulfilling such an ambitious objective will permit the visualization of RNA metabolism in four dimensions.

Acknowledgements

We thank Henri Grosjean as well as present and former members of the Bujnicki laboratory in IIMCB and at the UAM for stimulating discussions and contribution of ideas and information to this article. Our work on RNA modification enzymes has been supported by Polish Ministry of Science and Higher Education. J.M.B. acknowledges the Polish Ministry of Science for supporting his research on RNA modification (grant N301 2396 33) A.C. was supported by the Polish Ministry of Science (grant number N301 010 31/0219). K.R. was supported by the 6th FP Marie‑Curie Research EU Training Network “DNA Enzymes” (grant no. MRTNCT‑2005‑019566). J.M.B., J.K. and K.H.K. were also supported by the 6th FP Network of Excellence “EURASNET” (grant no. LSHG‑CT‑2005‑518238).

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