RNA Damage and Repair 3030765709, 9783030765705

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Ioly Kotta-Loizou   Editor

RNA Damage and Repair

RNA Damage and Repair

Ioly Kotta-Loizou Editor

RNA Damage and Repair

Editor Ioly Kotta-Loizou Imperial College London London, UK

ISBN 978-3-030-76570-5 ISBN 978-3-030-76571-2 https://doi.org/10.1007/978-3-030-76571-2

(eBook)

© Springer Nature Switzerland AG 2021 This work is subject to copyright. All rights are reserved by the Publisher, whether the whole or part of the material is concerned, specifically the rights of translation, reprinting, reuse of illustrations, recitation, broadcasting, reproduction on microfilms or in any other physical way, and transmission or information storage and retrieval, electronic adaptation, computer software, or by similar or dissimilar methodology now known or hereafter developed. The use of general descriptive names, registered names, trademarks, service marks, etc. in this publication does not imply, even in the absence of a specific statement, that such names are exempt from the relevant protective laws and regulations and therefore free for general use. The publisher, the authors, and the editors are safe to assume that the advice and information in this book are believed to be true and accurate at the date of publication. Neither the publisher nor the authors or the editors give a warranty, expressed or implied, with respect to the material contained herein or for any errors or omissions that may have been made. The publisher remains neutral with regard to jurisdictional claims in published maps and institutional affiliations. This Springer imprint is published by the registered company Springer Nature Switzerland AG. The registered company address is: Gewerbestrasse 11, 6330 Cham, Switzerland

Preface

Ribonucleic acid (RNA) is a macromolecule that plays a central role in cell physiology: RNA molecules act as intermediates between deoxyribonucleic acid (DNA), where genetic information is stored, and proteins, which perform the necessary functions within cells. Traditionally, the structural and functional properties of RNA are closely linked to gene expression; however, RNA-based enzymes called ribozymes are also involved in catalysis and small RNAs regulate key cellular processes, such as cell growth, division, differentiation, aging, and death. RNA is a sensitive macromolecule that can be easily damaged, by radiation, alkylating agents and oxidative stress, similar to the widely studied DNA and protein macromolecules, but also by ribonucleases, ribotoxins, and clustered regularly interspaced short palindromic repeats (CRISPR)-Cas systems which specifically target RNA. Therefore, cells have developed mechanisms to protect and/or repair RNA molecules. “RNA damage and repair,” to my knowledge the first book on this topic, presents an overview of the biology of RNA damage and repair in prokaryotic and eukaryotic cells. Individual chapters cover expression regulation, enzymology, and the physiological role of such systems, together with their links to important human diseases. The first three chapters focus on RNA damage and repair in prokaryotic organisms, mainly bacteria. Chapter “Endoribonucleases of the Toxin–Antitoxin Systems Induce Abortive Infection,” describes how bacteria utilize enzymes that damage RNA to protect themselves against bacteriophage infection and the response of bacteriophage in this arms race. In this context, the toxin–antitoxin systems MazF-MazE and RnlARnlB in the model organism Escherichia coli and ToxIN in the plant pathogen Pectobacterium atrosepticum are discussed in detail. Chapter “The Lifecycle of Ribosomal RNA in Bacteria,” focuses on ribosomal (r)RNA: non-coding, ubiquitous, stable, catalytic RNA molecules that play a crucial structural and functional role in protein production. The complex cellular processes regulating expression, maturation and modification of rRNA molecules and their

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subsequent assembly in functional ribosomes are described together with quality control mechanisms, potentially damaging factors and repair systems. Chapter “The Rtc RNA End Healing and Sealing System,” focuses on two enzymes conserved in all three kingdoms of life: RNA ligase RtcB and RNA cyclase RtcA. The biochemistry and structure of these enzymes are discussed, together with their biological role in prokaryotes and eukaryotes, and their expression regulation in bacteria under the control of the enhancer-binding protein RtcR and its CRISPR associated Rossmann fold domain. The next three chapters focus on RNA damage and repair in eukaryotic organisms, including plants. Chapter “Oxidative and Nitrative RNA Modifications in Plants,” illustrates how chemical damage to RNA caused by reactive oxidative and nitrative species in plant cells plays a regulatory role in both physiological and stress induced processes. Modified RNA transcripts containing 8-hydroxyguanine and 8-nitroguanine are associated with seed dormancy and germination, microbe infection and plant defense responses, and heavy metal induced stress. Chapter “The Role of Ribonucleases in RNA Damage, Inactivation, and Degradation,” describes how cells utilize enzymes that target RNA as defense mechanisms to protect themselves from damaged self and foreign RNA. These mechanisms act against defective messenger (m)RNA molecules that stall ribosomes and interfere with translation or viral RNA molecules and immunity-related transcripts during infection. Chapter “Cytoplasmic mRNA Recapping: An Unexpected Form of RNA Repair,” focuses on the 7-methylguanosine cap at the 5’ terminus of eukaryotic mRNAs, whose presence affects translation and mRNA decay. In addition to the established nuclear co-transcriptional capping, mRNAs may be decapped and recapped in the cytoplasm, a process that potentially has significant implications for the cell proteome. The last two chapters focus on the links between RNA damage and physiological human diseases. Chapter “Adenosine-to-Inosine RNA Editing: A Key RNA Processing Step Rewriting the Transcriptome in Normal Physiology and Diseases,” illustrates how damage to RNA molecules in the form of mutations caused by adenosine-to-inosine editing is regulated, together with its effects on numerous cellular processes including RNA silencing, splicing and polyadenylation, circadian rhythm, innate immunity, apoptosis, and cancer. Chapter “RNA-Mediated Metabolic Defects in Microsatellite Expansion Diseases,” focuses on the role of RNA metabolism in neurodegenerative disorders, such as Huntington’s disease, various forms of dystrophy, ataxia, epilepsy, and dementia. The molecular mechanisms underpinning RNA toxicity, including protein loss-of-function, protein gain-of-function, and RNA gain-of-function, are discussed in detail.

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Thanks to the high-quality contributions from all the authors involved, I hope that “RNA damage and repair” will be a book useful to all life scientists interested in the biology and homeostasis of RNA, coding or non-coding, intact, modified, or cleaved, in bacteria, plants, or mammals, under physiological conditions or during infection and disease. London, UK

Ioly Kotta-Loizou

Contents

Part I

RNA Damage and Repair in Prokaryotes

Endoribonucleases of the Toxin-Antitoxin Systems Induce Abortive Infection . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Yuichi Otsuka

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The Lifecycle of Ribosomal RNA in Bacteria . . . . . . . . . . . . . . . . . . . . . Maria Grazia Giuliano and Christoph Engl

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The Rtc RNA End Healing and Sealing System . . . . . . . . . . . . . . . . . . . Danai Athina Irakleidi, Harry Beaven, Martin Buck, and Ioly Kotta-Loizou

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Part II

RNA Damage and Repair in Eukaryotes

Oxidative and Nitrative RNA Modifications in Plants . . . . . . . . . . . . . . . Jagna Chmielowska-Bąk, Karolina Izbiańska-Jankowska, Magdalena Arasimowicz-Jelonek, Joanna Deckert, and Jolanta Floryszak-Wieczorek The Role of Ribonucleases in RNA Damage, Inactivation and Degradation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Fabian Hia and Osamu Takeuchi

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Cytoplasmic mRNA Recapping: An Unexpected Form of RNA Repair . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 109 Daniel R. Schoenberg Part III

RNA Damage in Human Diseases

Adenosine-to-Inosine RNA Editing: A Key RNA Processing Step Rewriting Transcriptome in Normal Physiology and Diseases . . . . . . . . 133 Priyankaa Pitcheshwar, Haoqing Shen, Jian Han, and Sze Jing Tang RNA-Mediated Metabolic Defects in Microsatellite Expansion Diseases . . 153 Nan Zhang ix

Part I

RNA Damage and Repair in Prokaryotes

Endoribonucleases of the Toxin-Antitoxin Systems Induce Abortive Infection Yuichi Otsuka

Abstract A competitive bacteria-phage coevolution is often referred to as an “arms race.” On the one hand, bacteria have evolved a variety of defense mechanisms against phages such as inhibition of phage adsorption and cleavage of phage nucleic acids through the restriction-modification system or the CRISPR-Cas system. On the other hand, phages have also devised many means to protect themselves from bacterial defense mechanisms. The abortive infection (Abi) system, one of the bacterial defense mechanisms, inhibits phage propagation by killing phage-infected cells before the phage life cycle is completed. Abi can be achieved through the toxinantitoxin (TA) system, a widespread genetic module in prokaryotes composed of a toxin and the corresponding antitoxin. In this system, the toxin inhibits an essential cellular process to arrest cell growth, while the antitoxin neutralizes the toxicity of a cognate toxin. MazF and RnlA toxins of the Escherichia coli TA systems and ToxN toxin of the plant pathogen Pectobacterium atrosepticum TA system have been well studied and demonstrated to possess endoribonuclease activities required for Abi. In this chapter, I will discuss the functions, structures, and regulations of their endoribonucleases. I will also discuss how these endoribonucleases induce Abi and how phages counteract Abi. Keywords Bacteria · Bacteriophages (phages) · The abortive infection (Abi) · The toxin-antitoxin (TA) systems · Endoribonucleases · MazF · RnlA · ToxN

Y. Otsuka (*) Department of Biochemistry and Molecular Biology, Graduate School of Science and Engineering, Saitama University, Saitama City, Saitama, Japan e-mail: [email protected] © Springer Nature Switzerland AG 2021 I. Kotta-Loizou (ed.), RNA Damage and Repair, https://doi.org/10.1007/978-3-030-76571-2_1

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1 Introduction 1.1

The Arms Race Between Bacteria and Phages

Bacteriophages or phages, which are viruses that specifically infect bacteria, are the most abundant and diverse biological entities on earth. Phages inhabit essentially everywhere and repeatedly infect bacteria at a high rate in the global ecosystem (Chibani-Chennoufi et al. 2004). They are estimated to outnumber bacteria by approximately 10 times; for example, there are ~1030 phages in the ocean, and phage infection occurs at a rate of ~1023 per second (Suttle 2007). Since phages constantly predate bacteria, bacteria have evolved a variety of defense mechanisms for survival and prosperity. The diversity and complexity of the bacterial defense mechanisms against phages are impressive; bacteria prevent phage adsorption by altering or concealing the phage receptors, inhibit phage propagation by cleaving the phage nucleic acids through the restriction-modification or the CRISPR-Cas systems, or commit suicide via abortive infection (Abi) before the production of phage progeny (Dy et al. 2014; Rostøl and Marraffini 2019). Similarly, phages have evolved to protect themselves from the bacterial defense mechanisms (Samson et al. 2013b). For example, some phages have inhibitory molecules that neutralize the CRISPR-Cas (Bondy-Denomy et al. 2013, 2015; Athukoralage et al. 2020) and Abi systems (Kaufmann 2000; Blower et al. 2012; Otsuka and Yonesaki 2012) or form a nucleus-like structure as a barrier to DNA-targeting CRISPR-Cas and restriction-modification systems (Malone et al. 2020; Mendoza et al. 2020). This competitive bacteria-phage coevolution is often referred to as an “arms race” (Hampton et al. 2020).

1.2

The Abortive Infection (Abi) System

While many of the bacterial defense mechanisms including the restrictionmodification and the CRISPR-Cas systems protect a single bacterial cell from getting infected by the phage, the Abi system triggers the premature death of a phage-infected cell before the phage completes its life cycle. Therefore, few phage progenies propagate, protecting the surrounding clonal bacteria from subsequent phage infection. The Abi system can be viewed as “altruistic suicide.” Although the Abi systems are becoming more widespread and diversified, the components and mechanisms leading to Abi remain elusive. E. coli PrrC, an endoribonuclease for the anticodon loop of tRNALys, is involved in Abi. After phage infection, PrrC is activated and inhibits both E. coli and phage translation, which kills the infected cells before phage progenies are produced (Kaufmann 2000). In Lactococcus sp., approximately 20 Abi-related genes have been identified (Chopin et al. 2005). For example, AbiZ interacts with Holin, a phage protein that forms a pore on the bacterial inner membrane, and induces premature

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cell lysis (Durmaz and Klaenhammer 2007). AbiB leads to nonspecific degradation of mRNAs to inhibit host and phage translation (Parreira et al. 1996). In Staphylococcus sp., a serine-threonine kinase, STK2, is activated by the phage protein PacK and phosphorylates the proteins involved in cellular metabolism, which results in the death of premature infected cells (Depardieu et al. 2016). To achieve Abi, bacteria need a toxic molecule to commit suicide upon phage infection. At the same time, bacteria also need an antitoxic molecule to repress the expression of the toxic molecule or counteract its toxicity; otherwise, they cannot grow normally. This is reminiscent of the toxin-antitoxin (TA) system (Gerdes et al. 2005). The TA system is a genetic module composed of a toxin and its cognate antitoxin and is abundantly found in prokaryotic genomes (Pandey and Gerdes 2005). A TA toxin inhibits an essential cellular process and arrests cell growth, while the antitoxin neutralizes the toxicity of a cognate toxin (Harms et al. 2018). As expected, many studies have reported a direct link between the Abi and the TA systems. The Hok-sok system encoded in an E. coli plasmid R1 was discovered as the first TA system that participates in Abi, in which the Hok toxin forms pores on the inner membrane and induces cell lysis before producing phage progenies (Pecota and Wood 1996). Later on, some TA toxins with an endoribonuclease activity have been revealed to protect against phage propagation.

1.3

The Toxin-Antitoxin (TA) System

The TA genetic module was originally discovered in the plasmid of E. coli in 1983 (Karoui et al. 1983; Ogura and Hiraga 1983). Since then, TA systems have been abundantly found not only in plasmids but also on bacterial chromosomes (Fozo et al. 2010; Leplae et al. 2011; Goeders et al. 2016). In 2005, it was also discovered in an archaeal genome (Pandey and Gerdes 2005). To date, many researchers have extensively investigated the function, regulation, and role of TA systems. The TA system is composed of two factors: a toxin and a cognate antitoxin. While a toxin arrests cell growth when it is expressed in a cell, the antitoxin neutralizes the toxicity of the cognate toxin (Harms et al. 2018). The genes encoding the toxin and the antitoxin are generally contiguous and have been discovered in almost all sequenced bacterial genomes. Notably, most of the bacteria, especially free-living bacteria, tend to have multiple TA loci (Pandey and Gerdes 2005). In fact, E. coli K-12 strain has 36 TA loci, and Mycobacterium tuberculosis has more than 88. The benefit of such a high number of TA systems is still unknown. In addition, how bacteria obtained that many TA loci on chromosomes remain unclear. Toxins in the TA system are stable proteins that inhibit one of the essential cellular processes, such as maintenance of the plasma membrane integrity (Kim et al. 2018; Wilmaerts et al. 2019; Otsuka et al. 2019), DNA replication (Jiang et al. 2002; Harms et al. 2015; Jankevicius et al. 2016), translation (Zhang and Inouye 2011; Castro-Roa et al. 2013; Germain et al. 2013; Cheverton et al. 2016; Masuda and Inouye 2017), peptidoglycan synthesis (Mutschler et al. 2011), and cell division

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(Tan et al. 2011; Heller et al. 2017). In contrast, antitoxins are generally labile (Tsuchimoto et al. 1992; Lehnherr and Yarmolinsky 1995; Koga et al. 2011) and are constantly expressed to compensate for degradation in order to successfully inhibit their cognate toxins (Li et al. 2014); the TA system is therefore referred to as an “addiction module.” If gene expression from TA loci is impaired, labile antitoxins rapidly diminish, which increases the level of free toxins and causes cell growth arrest. To date, TA systems are classified into six types according to the nature and function of antitoxins. In type I and III TA systems, antitoxins are small noncoding RNAs. Type I antitoxins bind to the cognate toxin mRNAs to inhibit their translation initiation or to induce their degradation (Gerdes and Wagner 2007; Fozo et al. 2008), while type III antitoxins inactivate their cognate toxins through protein-RNA interactions (Goeders et al. 2016). Type II, IV, V, and VI antitoxins are proteins. Type II is the best-characterized TA system where antitoxins bind the cognate toxins directly (Leplae et al. 2011; Fraikin et al. 2020). Type IV antitoxins do not interact with the cognate toxins but function on the same target. For example, E. coli CbeA antitoxin neutralizes the toxicity of the CbtA toxin by interfering with the binding of the toxin to its target protein, FtsZ and MreB (Masuda et al. 2012; Wen et al. 2017). In type V or VI TA systems, only one example has been identified. The GhoS antitoxin of type V E. coli TA system is an endoribonuclease that specifically cleaves the GhoT toxin mRNA to block its expression (Wang et al. 2012). The SocA antitoxin of a type VI Caulobacter TA system promotes degradation of the SocB toxin mediated by ClpXP protease and consequently neutralizes the toxicity of SocB (Aakre et al. 2013). Elucidating the biological roles of the TA systems is an active research topic, as these systems are involved in a wide range of bacterial events. TA loci located on plasmids promote plasmid maintenance in growing bacterial populations through the mechanism known as “post-segregational killing” (Karoui et al. 1983; Ogura and Hiraga 1983; Thisted and Gerdes 1992). The roles of TA loci on bacterial chromosomes are still elusive although the following roles are suggested: biofilm formation (Kim et al. 2009; Mandell et al. 2019), persistent cell formation (Germain et al. 2013; Harms et al. 2016; Page and Peti 2016; Semanjski et al. 2018), stress response (Wang et al. 2011; Hu et al. 2012), and phage defense (named, Abi) (Emond et al. 1998; Hazan and Engelberg-Kulka 2004; Fineran et al. 2009; Koga et al. 2011; Alawneh et al. 2016). Many TA systems, including Hok-sok of the type I TA system, are known to participate in Abi (Pecota and Wood 1996). Among them, the MazF toxin of MazEMazF and the RnlA toxin of RnlA-RnlB in E. coli type II TA systems and the ToxN toxin of ToxIN in the plant pathogen P. atrosepticum type III TA system have been well studied (Otsuka 2016; Goeders et al. 2016; Fraikin et al. 2020; Jurėnas and Van Melderen 2020). Interestingly, all of them have endoribonuclease activities. In this chapter, three TA toxins, MazF, RnlA, and ToxN, are highlighted. The functions, structures, and regulations of their endoribonucleases, Abi mechanisms induced by them, and phage mechanisms to overcome Abi are discussed.

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2 Functions, Structures, and Regulation of the MazF Endoribonuclease E. coli MazF toxin is a sequence-specific endoribonuclease that recognizes the ACA sequence and cleaves RNA at the 50 side of the first A residue (denoted as #ACA, the arrow indicates the cleavage site of RNA) (Zhang et al. 2003). MazF homologs exhibit different sequence specificities. MazF homologs from Gram-positive bacteria (Staphylococcus aureus, Bacillus subtilis, and Clostridium difficile) cleave RNA at the U#ACAU sequence (Zhu et al. 2009; Park et al. 2011; Rothenbacher et al. 2012). In contrast, archaeal MazF homologs from Haloquadratum walsbyi and Methanohalobium evestigatum cleave RNA at the UU#ACUCA sequence and the C#UGGU or U#UGGU sequence, respectively (Yamaguchi et al. 2012; Ishida et al. 2019). MazF mainly targets single-stranded RNAs (Zhang et al. 2003). Its cleavage generates a 5´ RNA fragment with the 20 ,30 -cyclic phosphate at the 30 -end and the 30 fragment with the 50 -hydroxyl group at the 50 -end (Zhang et al. 2005; Mets et al. 2017), while most E. coli endoribonucleases (such as RNase E, RNase P, and RNase III) that participate in RNA processing and decay yield a hydroxyl group at the 30 -end and a phosphate at the 50 -end (Cannistraro and Kennell 1993). E. coli MazF exhibits nuclease activity in the absence of Mg2+, and the addition of Mg2+ inhibits its activity (Zhang et al. 2005). Structures of bacterial MazF toxins alone and their complex with an RNA substrate have been unraveled (Kamada et al. 2003; Simanshu et al. 2013; Zorzini et al. 2014; Chen et al. 2017b; Hoffer et al. 2017). The E. coli MazF monomer consists of a seven-stranded, twisted antiparallel β-sheet linked by three α-helices (Kamada et al. 2003). MazF forms a homodimer, and a positively charged concave at the interface between the two MazF subunits binds to one RNA molecule (Simanshu et al. 2013; Zorzini et al. 2016). Structural comparisons among TA toxins revealed that several TA toxins (Kid and ToxN) with endoribonuclease activity are homologous to MazF (Kamada et al. 2003; Blower et al. 2011). Surprisingly, MazF is structurally similar to the CcdB toxin (Kamada et al. 2003), despite their completely different functional activities. The ccdB gene is encoded in E. coli F-plasmid, and its product obstructs DNA replication by binding to DNA gyrase (Dao-Thi et al. 2005). In addition to their structural similarities, the inhibitory mechanisms of cognate antitoxins to their toxins are analogous (Zorzini et al. 2016). There are several possible mechanisms underlying MazF toxicity. E. coli MazF cleaves the ACA sequence in the coding and untranslated regions of mRNAs in vivo, which causes the reduction of translatable mRNAs (Zhang et al. 2003; Culviner and Laub 2018; Mets et al. 2019). In addition, MazF cleaves rRNAs at multiple sites (Mets et al. 2017; Culviner and Laub 2018). Upon MazF expression, cleavages of rRNAs would lead to the reduction of intact ribosomal particles and/or the accumulation of aberrant ribosomes with fragmented rRNAs; consequently, the activity of protein synthesis would dramatically decrease. Notably, E. coli MazF cleaves 16S rRNA to generate specialized ribosomes that translate specific mRNAs. MazF cleaves 16S rRNA at nucleotide positions 1396 and

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1500 located in the decoding center and removes the anti-Shine-Dalgarno (SD) sequence (Vesper et al. 2011), which generates a specialized ribosome with a truncated 16S rRNA. In addition, MazF generates leaderless mRNAs through its preferential cleavages at or closely upstream of the AUG start codon. Unlike normal ribosomes, specialized ribosomes appear to specifically translate the leaderless mRNAs lacking the SD sequence. Since the leaderless mRNAs mostly encode stress-responsive proteins (Moll and Engelberg-Kulka 2012; Sauert et al. 2016; Nigam et al. 2019), this is considered to be a mechanism to cope with stresses. However, other studies have reported contradictory data showing no enrichment of any stress-responsive proteins in the MazF-expressing cells (Culviner and Laub 2018; Mets et al. 2019; Wade and Laub 2019; Kaldalu et al. 2019). Therefore, this concept is still under debate. Intriguingly, M. tuberculosis encodes the highest number of mazF genes (10 mazF genes) (Sala et al. 2014). Each MazF toxin exhibits different specificities to the recognition sequence and the RNA substrate (tRNA, mRNA, or rRNA). MazF-mt3 and MazF-mt6 toxins cleave mRNAs at the U#CCUU and UU#CCU sequences, respectively (Schifano et al. 2014; Hoffer et al. 2017). In addition, MazFmt3 cleaves the anti-SD sequence of the 16S rRNA and loop 70 of the 23S rRNA (Schifano et al. 2014). MazF-mt6 also cleaves loop 70 of the 23S rRNA (Schifano et al. 2013). These toxins impair translatable mRNAs and intact ribosomes, ultimately leading to bacterial growth arrest. MazF-mt9 cleaves the anticodon sequence of tRNALys43-UUU (Schifano et al. 2016; Chen et al. 2017b). It is unlikely to inhibit the translation globally because a proteome analysis detected hundreds of newly synthesized proteins (Barth et al. 2019). Instead, the loss of tRNALys43-UUU mediated by MazF-mt9 leads to ribosome stalling on mRNAs containing AAA Lys codon. mRNAs harboring stalled ribosomes are recognized as aberrant and are positively digested by cellular RNases. Consequently, mRNAs containing the AAA Lys codon are specifically untranslated causing cell growth arrest. MazE antitoxin inhibits the MazF endoribonuclease via direct binding. The structure of their complex has been unraveled (Kamada et al. 2003; Simanshu et al. 2013; Ahn et al. 2017). One MazE dimer binds two MazF dimers, forming a heterohexamer with MazE2:MazF4 stoichiometry. The unstructured C-terminal extension of MazE binds to the dimer of MazF. The strong negative charge of its C-terminal extension mimics an RNA molecule and traps MazF to suppress its activity. Intriguingly, the binding of one MazE to one of the two RNA binding sites within a MazF dimer efficiently inhibits the catalytic activity of both sites. MazF activity is suggested to be enhanced by a peptide called extracellular death factor (EDF) in E. coli, B. subtilis, and Pseudomonas aeruginosa. EDFs are secreted from bacteria and induce interspecies cell death (Kolodkin-Gal et al. 2007; Kumar et al. 2013). EDFs of E. coli and B. subtilis are the NNWNN pentapeptide and the RGQQNE hexapeptide, respectively. P. aeruginosa secretes three different EDFs; the INEQTVVTK, VEVSDDGSGGNTSLSQ, and APKLSDGAAAGYVTKA peptides. EDFs appear to enhance the endoribonuclease activities of MazF toxins directly or indirectly by preventing the formation of a MazE-MazF complex (Belitsky et al. 2011; Erental et al. 2012; Oron-Gottesman et al. 2016; Nigam et al.

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2018). Although the use of the EDF peptides as antibiotics has been proposed, their impact on cell growth and MazF activity has not been established yet (Jurėnas and Van Melderen 2020).

2.1

The Abortive Infection Induced by the MazF Endoribonuclease

In 2004, the Engelberg-Kulka group showed that E. coli MazF-mediated growth arrest led to the inhibition of the lysogenic phage P1 propagation (Hazan and Engelberg-Kulka 2004). The number of P1 phage progenies induced from lysogenic E. coli by high temperatures was significantly increased by the disruption of the mazE-mazF locus. Furthermore, the transduction of P1 phage was lethal to ΔmazEmazF non-lysogenic E. coli, but not to the wild-type cells. Thus, MazE-MazF plays an inhibitory role in P1 phage propagation, suggesting the MazF-induced Abi. It remains unknown how MazF is activated after heat induction or P1 phage infection or how it leads to Abi. Our paper published in 2016 demonstrated the direct link between Abi and MazF (Alawneh et al. 2016) (Fig. 1). Virulent phage T4 progenies were significantly

Fig. 1 MazF-induced Abi and the inactivation of MazF by T4 phage Alt protein. (a) In the absence of phage infection, MazF endoribonuclease activity is repressed by MazE via direct binding. (b) If a T4 phage lacking Alt infects a bacterium, host gene expression is immediately shut off, and labile MazE is degraded. Consequently, MazF is activated and degrades host and phage RNAs, which causes Abi. (c) If a T4 phage harboring Alt infects a bacterium, Alt is injected with phage genomic DNA into a host cell and immediately ADP-ribosylates MazF. Consequently, MazF is inactivated, which allows T4 phage to propagate

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increased by the disruption of mazE-mazF and decreased by ectopically expressed MazF. Upon MazF expression, E. coli and T4 phage mRNAs were mostly degraded, and protein synthesis was inhibited. Therefore, MazF endoribonuclease is a factor leading to Abi against the T4 phage. To understand the mechanism of MazF activation after phage infection, the changes in MazE antitoxin and MazF toxin were examined. While MazF was stable during infection, MazE disappeared immediately after infection. The different stabilities of MazE and MazF likely contribute to MazF activation. T4 phage infection results in the immediate blockage of E. coli gene expression in several ways such as DNA cleavage and modifications of RNA polymerase (Miller et al. 2003). The blockage of E. coli gene expression probably promotes the disappearance of labile MazE, which in turn activates a more stable MazF that triggers Abi.

2.2

Phage Mechanism to Overcome MazF-Induced Abi

The T4 phage can grow on E. coli cells expressing MazF, albeit less efficiently than ΔmazE-mazF cells. This result suggests that the T4 phage may possess an inhibitory factor against MazF. A line of experimental results supports this hypothesis (Alawneh et al. 2016). Firstly, MazF was ADP-ribosylated immediately after T4 phage infection. Secondly, ADP-ribosylation was not observed in cells infected with the T4 phage lacking Alt, one of the three ADP-ribosyltransferases encoded by the T4 phage that is injected with phage DNA upon infection (Tiemann et al. 2004; Depping et al. 2005). An ADP-ribosyltransferase catalyzes the transfer of an ADP-ribosyl group from the substrate, nicotinamide adenine dinucleotide (β-NAD+), to a specific amino acid (frequently arginine or histidine) of the acceptor protein. Alt ADP-ribosylated the arginine residue of MazF at position 4. Importantly, the ADP-ribosylation of MazF by Alt resulted in the partial reduction of the MazF RNA cleavage activity in vitro. This is the first example showing that chemical modification regulates the activity of a bacterial toxin in the TA systems. Taken together, T4 Alt ADP-ribosylates MazF immediately after infection and renders this toxin inactive, leading to efficient growth of the T4 phage (Fig. 1).

3 Functions, Structures, and Regulation of RnlA Endoribonuclease E. coli RnlA (initially called RNase LS) is an endoribonuclease without strict sequence specificity (Kai et al. 1996; Otsuka and Yonesaki 2005; Otsuka et al. 2007). It preferentially cleaves RNA at the 50 side of pyrimidines in vivo, but not frequently (Kai et al. 1996). Interestingly, RnlA appears to cleave mRNA when the target is translatable (Kai and Yonesaki 2002; Yamanishi and Yonesaki 2005). The

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RnlA-mediated cleavage occurred specifically when a stop codon (or a rare codon) was introduced into the coding region of target mRNAs. However, the disruption of the SD sequence or introduction of stop-codon-suppressing tRNA eliminated RnlAmediated cleavages. RnlA likely cleaves mRNA when the translocation of ribosomes stops or slows down. The linkage between the ribosome and the TA toxin with endoribonuclease activity was first reported for a RelE toxin of E. coli RelBRelE TA system. The RelE-family toxins (RelE, YoeB, YafQ, HigB) associate with ribosomes and cleave mRNAs in the ribosomal A site between the second and the third bases of the codon (Neubauer et al. 2009; Prysak et al. 2009; Hurley and Woychik 2009; Feng et al. 2013). RnlA and RelE-family toxins appear to have different modes of RNA cleavage mechanisms. It remains unclear what kind of end structures RNA fragments have after RnlA cleavage or whether RnlA cleaves only single-stranded RNA. Although in vitro RNA cleavage assay with ribosomal fractions prepared from E. coli cell extracts expressing RnlA demonstrated that some of the RnlA-mediated cleavages were dependent on magnesium ions (Otsuka et al. 2007), the metal dependency of RnlA activity needs to be further investigated. The structure of the E. coli RnlA toxin has been unraveled (Wei et al. 2013). RnlA is composed of three independent domains: NTD (N-terminal domain; 1–90), NRD (N-repeated domain; 91–197), and DBD (Dmd-binding domain; 198–356). NTD is composed of a four-stranded twisted antiparallel β-sheet (β1–β4) and two helices (α1 and α2). NRD has a five-stranded twisted antiparallel β-sheet (β5–β9) and two helices (α4 and α5). NTD and NRD are connected by a short loop. Surprisingly, the topology of NRD is highly homologous to NTD except for the β5 strand, albeit their low similarities in amino acid sequence. Although NTD and NRD share structural similarity with other known proteins, their functions remain unclear. The C-terminal DBD is primarily composed of eight helices (α7–α14) in addition to two short β-strands. Since the expression of DBD alone exhibited cell growth arrest, DBD is responsible for the toxicity and the endoribonuclease activity of RnlA. Residues in or around the α14 helix form a continuous positively charged surface patch and are thought to play a role in RNA substrate recognition. Moreover, DBD is responsible for RnlA dimerization and the interaction with a cognate antitoxin RnlB or a phage antitoxin Dmd (Wei et al. 2013; Wan et al. 2016; Naka et al. 2017). A total of 20 RnlA homologs were found in different bacterial species ranging from Proteobacteria to Firmicutes (Naka et al. 2014). Half of these homologs are similar in size to E. coli K-12 RnlA (357 amino acids). The other half is larger than RnlA by 270–330 amino acids, and their C-terminal moieties are homologous to RnlA. Interestingly, each of these N-terminal sequences encodes an RNase HI domain, raising the possibility that RNase HI and RnlA are functionally coupled. E. coli RNase HI cleaves RNA in a DNA-RNA duplex (Miller et al. 1973) and is responsible for DNA replication, transcription, and DNA repair (Itoh and Tomizawa 1980; Drolet 2006; Tadokoro and Kanaya 2009). Supporting this idea, our group reported the involvement of RNase HI in the regulation of RnlA endoribonuclease activity (described below) (Naka et al. 2014, 2017). DBD of RnlA with an endoribonuclease activity belongs to the high eukaryotes and prokaryotes nucleotide-binding (HEPN) domain superfamily (Anantharaman

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et al. 2013). The HEPN superfamily is composed of all-α-helical catalytic domains. Proteins containing the HEPN domain typically have metal-independent endonuclease activities. Interestingly, the components involved in bacterial defense systems including the TA, the restriction-modification, and the CRISPR-Cas systems often share the HEPN domain (Anantharaman et al. 2013; Yao et al. 2015; Abudayyeh et al. 2016; Jia et al. 2018). This agrees with the data showing that RnlA participates in Abi (Koga et al. 2011; Otsuka and Yonesaki 2012). The expression of RnlA induces the cleavage of bulk E. coli mRNAs, leading to the inhibition of protein synthesis and the arrest of cell growth (Koga et al. 2011). Additionally, RnlA cleaves T4 phage mRNAs at the late stage after infection and consequently blocks the propagation of the T4 phage (Kai et al. 1996). Whether RnlA targets tRNAs and rRNAs remains to be determined. The disruption of RnlA in E. coli led to the accumulation to a high level of a 307-nucleotide fragment with an internal sequence of 23S rRNA (Otsuka and Yonesaki 2005). Therefore, RnlA appears to play a role in the digestion of this short fragment. Interestingly, this fragment seems to contain a putative SD sequence and an open reading frame encoding a polypeptide of 27 amino acids. Because the SD-like sequence is properly located upstream of the AUG start codon, this short RNA may encode the functional polypeptide. An analogous mechanism has been reported where a part of the rRNA may function as an mRNA; overexpression of a functional peptide comprising five amino acids that may be encoded in the 23S rRNA renders cells resistant to erythromycin (Tenson et al. 1996). The RnlB antitoxin is supposed to bind RnlA directly to inhibit the RnlA endoribonuclease activity (Koga et al. 2011; Naka et al. 2017). The structure of the RnlA-RnlB complex has not been elucidated yet because RnlB is unstable and degraded before and/or during purification. Recently, native mass spectrometry (MS) and small-angle X-ray scattering (SAXS) experiments demonstrated that RnlA and RnlB formed a heterotetramer with RnlA2: RnlB2 stoichiometry (Garcia-Rodriguez et al. 2020). This complex was crystallized, but its structure could not be unraveled. In 2016, the complex structure of the LsoA toxin, a RnlA homolog from enterohemorrhagic E. coli O157:H7, and the Dmd antitoxin from the T4 phage was unraveled (Wan et al. 2016). Dmd is an inhibitor of LsoA and is inserted into the deep groove between NRD and DBD of LsoA. Upon Dmd binding, NRD shifts significantly from a closed to an open conformation. Additionally, three residues (R243, E246, and R305) located in DBD are highly conserved among the six RnlA homologs (E. coli K-12 RnlA, E. coli O157:H7 LsoA, Citrobacter braakii LsoA, Vibrio cholerae RNase, Klebsiella pneumoniae RNase, and Arsukibacterium ikkense RNase) and are responsible for the endoribonuclease activity of LsoA. Considering that these residues are located in the Dmd-binding groove, Dmd would occupy the active site of LsoA via substrate RNA mimicry to block its RNase activity.

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The Abortive Infection Induced by RnlA Endoribonuclease

Two type II TA systems, rnlA-rnlB, encoded by the E. coli K-12 chromosome, and lsoA-lsoB of E. coli O157:H7, encoded by a plasmid pOSAK1, antagonize the T4 phage (Koga et al. 2011; Otsuka and Yonesaki 2012) (Fig. 2). Endoribonuclease activities of both toxins (RnlA and LsoA) increase after T4 phage infection, because T4 infection shuts off E. coli gene expression, and consequently the labile RnlB and LsoB diminish. When a dmd mutant of the T4 phage infects E. coli, free RnlA or LsoA degrades most of the E. coli and T4 phage mRNAs, rendering this mutant phage incapable of propagation (Kai et al. 1996; Otsuka and Yonesaki 2012). Thus, RnlA and LsoA function as phage defense agents, leading to Abi. Interestingly, RnlA-induced Abi was not observed in ΔrnhA (a gene encoding RNase HI) cells (Naka et al. 2014). In vivo and in vitro analyses have demonstrated that RNase HI is an activating factor for RnlA activity. Therefore, RNase HI may activate RnlA after T4 phage infection, although its molecular mechanism remains to be determined. Recently, we reported that RNase HI recruits RnlB antitoxin to RnlA through the NRD of RnlA for inhibiting RnlA toxicity (Naka et al. 2017). Taken together, RNase HI is an essential component of the RnlA-RnlB TA system, where it plays two contrary roles in controlling RnlA activity.

Fig. 2 RnlA-induced Abi and the inactivation of RnlA by T4 phage Dmd protein. (a) In the absence of phage infection, RnlA endoribonuclease activity is repressed by RnlB. (b) If a T4 phage lacking Dmd infects a bacterium, host gene expression is immediately shut off, and labile RnlB is degraded. Consequently, RnlA is activated with the aid of RNase HI and degrades host and phage RNAs, which causes Abi. (c) If a T4 phage encoding the dmd gene infects a bacterium, Dmd is expressed immediately after infection and binds to RnlA for inactivation, which allows T4 phage to propagate

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Phage Mechanisms to Overcome RnlA-Induced Abi

When Dmd is expressed immediately after T4 phage infection, as found in wild-type T4, the phage can grow normally. During infection, T4 Dmd, rather than RnlB or LsoB, suppresses the endoribonuclease activity of RnlA or LsoA through direct binding (Otsuka and Yonesaki 2012; Wan et al. 2016) (Fig. 2). The T4 phage is considered to have evolutionally obtained its antitoxin against bacterial toxins for survival. Dmd antitoxin exhibits a unique property; in general, an antitoxin only inhibits the cognate toxin. Indeed, RnlB and LsoB are not interchangeable in the inhibition of RnlA or LsoA activity. Dmd, in contrast, can inhibit both RnlA and LsoA toxins. Dmd does not share any sequence similarity with RnlB or LsoB. Therefore, Dmd is a novel antitoxin with broad specificity and is likely to function differently from canonical bacterial antitoxins. Some T4 phage mutants that are able to partially escape RnlA-induced Abi have been isolated (Kai et al. 1998). Among them, two independent mutants contained a nonsense or missense mutation in motA or asiA, respectively (Ueno and Yonesaki 2001) (Unpublished data). Both encode transcriptional activators for the middle genes of the T4 phage. When a motA dmd or asiA dmd double mutant of the T4 phage infects, activation of RnlA is delayed. The T4 phage may control RnlA activity, that is, a middle gene product may participate in RnlA activation after phage infection (Miller et al. 2003).

4 Functions, Structures, and Regulations of ToxN Endoribonuclease ToxNPa encoded in a cryptic plasmid (pECA1039) of the plant pathogen P. atrosepticum is a sequence-specific endoribonuclease and cleaves RNAs at the 50 side of the third A-residue (AA#AU or AA#AG) (Short et al. 2013). ToxNBt encoded in a cryptic plasmid (pAW63) of Bacillus thuringiensis with 30% amino acid identity to ToxNPa, or AbiQ encoded in a cryptic plasmid (pSRQ900) of Lactococcus lactis with 31% identity, cleaves RNAs at A#AAAA or A#AAA, respectively (Samson et al. 2013c; Short et al. 2013). The ToxN homologs belong to the type III TA system and specifically target their cognate antitoxin RNAs as well as cellular mRNAs (Goeders et al. 2016). Similar to MazF, the ToxN cleavage yields a 5´ RNA fragment with a 20 -30 cyclic phosphate at the 30 -end (Blower et al. 2011). The same functionality of MazF and ToxN is supported by their overall structural similarity (Kamada et al. 2003; Blower et al. 2011). Contrary to RnlA, both ToxN and MazF do not require the ribosome for their cleavages. ToxN is proposed to cleave RNAs in a metal-independent manner. It is still unclear whether ToxN cleaves only single-stranded RNA. The structures of the three type III TA toxins (ToxNPa, ToxNBt, and AbiQ) have been unraveled (Blower et al. 2011; Samson et al. 2013c; Short et al. 2013). All the

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toxins are globular with a β-sheet core-fold region surrounded by helices and loops. For example, AbiQ has a core of six antiparallel β-sheets surrounded by six α-helices. ToxN homologs cleave a variety of mRNAs, leading to bacterial growth arrest (Fineran et al. 2009; Samson et al. 2013c). Whether they target tRNAs and rRNAs remains unclear. As a characteristic of the type III TA systems, a toxin specifically cleaves its cognate antitoxin precursor transcript to generate functional antitoxin RNAs (Goeders et al. 2016). A ToxIPa precursor transcript is composed of 5.5 repeats of a 36-nucleotide. ToxIBt or antiQ precursor transcripts contain 2.9 repeats of a 34-nt or 2.8 repeats of a 35-nt, respectively. An appropriate number of repeats is likely to be important for their antitoxic activities in vivo (Blower et al. 2009; Bélanger and Moineau 2015). The specific cleavage of these precursor transcripts by cognate toxins generates monomeric fragments, each of which is a highly structured RNA with a pseudoknot structure essential for its antitoxic function. In the case of ToxINPa, each mature 36-nt RNA fragment interacts directly with a ToxN molecule and ultimately forms a triangular heterohexamer with ToxI3:ToxN3 stoichiometry (Blower et al. 2011). In this complex, toxins occupy the apices and are held together by mature antitoxin monomers. Each end of ToxIPa RNA interacts with a different ToxNPa monomer via electropositive grooves. Hydrogen bonds are formed between the RNA bases of ToxIPa and the amino acids of ToxNPa. ToxINBt and AbiQ-antiQ exhibit a similar structure to ToxINPa (Samson et al. 2013c; Short et al. 2013). Another type III TA system CptINEr from Eubacterium rectale, which is distantly related to the ToxIN family, also forms the overall similar structure, although it is composed of two toxin monomers assembled with two antitoxin RNAs (Rao et al. 2015). Therefore, the alternating interaction between a toxin protein and an antitoxin RNA is a common feature of the type III TA system. Since an antitoxin RNA becomes both an inhibitor and a substrate of the cognate toxin, the regulation of the toxin activity by the antitoxin RNA, especially the process of a complex formation and dissociation, is considered to be organized. ToxINPa complex can be automatically formed if the precursor ToxI RNA or a mature form of the ToxI RNA is mixed with ToxN in a solution without any cellular proteins (Short et al. 2013). It remains unclear when a precursor antitoxin RNA is cleaved by a toxin (before, during, or after their complex formation) and how toxins are released from their stable complex for activation.

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The Abortive Infection Induced by ToxN Endoribonuclease

ToxIN is known to play a role in Abi (Fineran et al. 2009) (Fig. 3). L. lactis AbiQantiQ of the ToxIN family effectively inhibits some Lactococcal phages (Samson et al. 2013a). When AbiQ-antiQ were expressed in E. coli cells, an Abi phenotype against several coliphages including the T4 phage was observed, indicating that

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Fig. 3 ToxN-induced Abi and the inactivation of ToxN by phage small RNAs. (a) In the absence of phage infection, ToxN processes toxI precursor RNAs to generate mature ToxI RNAs. Subsequently, ToxN endoribonuclease activity is repressed by mature ToxI RNAs via direct binding. (b) If a phage infects a bacterium, the complex is dissociated, and mature ToxI RNAs is degraded. Consequently, ToxN is activated and degrades host RNAs, which causes Abi. (c) If a phage encoding pseudo-toxI RNA or toxI RNA infects a bacterium, pseudo-toxI RNAs or toxI RNAs are expressed from the phage genome during infection and bind to ToxN for inactivation, which allows the phage to propagate

AbiQ-antiQ is effective against a wide range of phages, and their effect is not limited to L. lactis. ToxINPa exhibits a strong Abi phenotype against a variety of phages, while ToxINBt does not exhibit an Abi phenotype against any phages tested (Short et al. 2013). As for the molecular mechanism of ToxIN-induced Abi, the major question is how the complex is dissociated, and the toxin is activated after phage infection. Like MazF and RnlA (Otsuka and Yonesaki 2012; Alawneh et al. 2016), the toxins could be released if antitoxin levels are reduced following the blocking of host transcription immediately after phage infection. Another possibility is that a specific phage protein could trigger the release of the toxin from the complex and/or the degradation of antitoxin RNAs. A gene product of the ΦM1 phage, M1–23, may be a candidate for a phage protein which activates ToxNPa, because mutations in the M1–23 gene allowed the ΦM1 phage to escape ToxNPa-induced Abi (Blower et al. 2017). The M1–23 mutant phages could also grow in cells harboring a different type III TA system (TenpIN), while wild-type phage could not. The function of M1–23 against the ToxINPa and TenpIN TA systems are still unknown. Besides, there are two interesting results. Several ToxN mutant proteins with the endoribonuclease activity did not exhibit Abi phenotype (Blower et al. 2009). Similarly, the AbiQ mutant, in which the serine at position 51 was replaced by threonine, retained the endoribonuclease activity but lost the Abi activity (Samson et al. 2013c). It is

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considered that these mutant proteins could not be adequately activated after phage infection.

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Phage Mechanisms to Overcome ToxN-Induced Abi

Phages escape ToxN-induced Abi using various means. Although ToxNPa prevents ΦTE phage from propagation, this phage has obtained spontaneous mutations at a low frequency to avoid ToxNPa-induced Abi (Blower et al. 2012). Most of the mutants extended a short region of their genome to produce small RNA repeats termed “pseudo-ToxI” (Fig. 3). This “pseudo-ToxI” was similar, but not identical, to ToxIPa in sequence. This RNA was produced during phage infection and suppressed the ToxNPa activity, thus allowing these mutants to avoid ToxNPa-induced Abi. Besides, an Abi-escaped mutant acquired the DNA region corresponding to ToxIPa into the phage genome from the plasmid encoding the original toxINPa locus through homologous recombination (Fig. 3). Serratia mutant phages escaped from ToxNPa-induced Abi using three different means: a mutation of asiA encoding a putative phage transcriptional activator, a mutation of an unknown gene, and a deletion of a large region of the phage genome (Chen et al. 2017a). Interestingly, a mutation of asiA in the T4 phage genome also allows the phage to escape RnlA-induced Abi (unpublished data). AsiA-controlled genes may be involved in ToxN- or RnlA-induced Abi against Serratia phages or the T4 phage, respectively.

5 Conclusion In this chapter, MazF, RnlA, and ToxN toxins with endoribonuclease activities that participate in Abi are highlighted. An open question in TA toxin-induced Abi is how a toxin is activated after phage infection. An activation model is proposed; phage infection immediately stops the transcription of TA genes, which leads to the loss of antitoxins, and consequently, toxins are released and activated. Thus, blocking the transcription of the TA gene loci is a trigger for toxin activation. However, this model cannot fully explain several experimental data. Firstly, an antitoxin bound to a toxin is protected from protease attack, thereby a toxin is unlikely to be released and activated simply after transcription from its TA locus is stopped (LeRoux et al. 2020). Secondly, RnlA digests almost all mRNAs present in the late stage of infection in the absence of T4 phage antitoxin Dmd; however, it does not cleave any mRNAs in the early and middle stages (Kai et al. 1996). Similarly, AbiQ also appears to be activated at the late stage (Emond et al. 1998). Such drastic and instantaneous activation of RnlA and AbiQ is unlikely to be accomplished by merely blocking the transcription of the TA gene loci after infection. A phage protein may be involved in the activation of RnlA or ToxN. In support of this idea, mutations in

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phage genes encoding transcriptional activators (AsiA of Serratia phages ΦCHI14 and ΦCBH8 or AsiA and MotA of T4 phage) allow phages to partially escape ToxNor RnlA-induced Abi, indicating that phages may control toxin activity (Ueno and Yonesaki 2001; Chen et al. 2017a) (unpublished data). Phages appear to escape from TA toxin-induced Abi using a variety of ways such as the acquisition of an inhibitor against a toxin, inactivation of a toxin by chemical modification, and mutations of phage genes that participate in toxin activation. All 36 TA toxins of the E. coli K-12 strain can be activated after T4 phage infection probably because the infection shuts off E. coli gene expression and consequently the labile antitoxins diminish. However, the T4 phage can efficiently grow in E. coli, suggesting that the T4 phage may be equipped with inhibitory factors against activating toxins. As described above, the T4 phage has the Dmd antitoxin that can neutralize two toxins, although an antitoxin generally inhibits the cognate toxin only (Otsuka and Yonesaki 2012). Phages might harbor such multifunctional antitoxins to deal with many activating toxins. An interesting candidate is a T4 phage protein, Pin, which is an inhibitor of E. coli Lon cellular protease that degrades many antitoxins (Skorupski et al. 1988). The sana TA system of Shewanella protected E. coli from the T7 phage, and the disruption of T7 Gp4.5, a Lon-interacting protein, allowed the T7 phage to grow (Sberro et al. 2013). This implies that Gp4.5 inhibits Lon protease, which consequently causes the prevention of antitoxin degradation and toxin activation. Besides the three TA toxins featured in this chapter, other toxins with endoribonuclease activities have been found and characterized. In addition to basic research, research applying these endoribonucleases in biotechnology are progressing. For example, MazF that specifically cleaves the ACA RNA sequence is commercially available as “mRNA interferase”. This can be utilized for the analysis of the RNA structure. Endoribonucleases with strict sequence specificities will contribute to RNA research like restriction enzymes have greatly promoted DNA research. The restriction-modification and the CRISPR-Cas systems that function as phage defense have been currently applied in technologies for gene manipulation and genome editing and have greatly contributed to the development of various fields of biology. The TA system also has the potential to be a powerful tool for combatting pathogenic bacteria and antimicrobial-resistant bacteria. For example, the TA loci are naturally found in the genomes of these bacteria. If a TA system specific to these bacteria is identified and a compound that disrupts the complex between its toxin and antitoxin is discovered, we might be able to eliminate these bacteria specifically. Although it has been over 40 years since a TA system was discovered, there are still many basic and practical issues that need to be addressed. Acknowledgments We cordially thank Prof. Toshimitsu Kawate at Cornell University and Prof. Tetsuro Yonesaki at Osaka University for the invaluable help with the manuscript. This work was supported in grants from the program Grants-in-Aid for Scientific Research (C)(17K08837 and 20K07493) from the Ministry of Education, Culture, Sports, Science and Technology of Japan.

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The Lifecycle of Ribosomal RNA in Bacteria Maria Grazia Giuliano and Christoph Engl

Abstract Ribosomal RNA is the main constituent of ribosomes, where it not only provides a structural scaffold for r-proteins but also a decoding mechanism for the translation of mRNAs into proteins. Modulation of the cellular rRNA content is at the heart of the strictly regulated biosynthesis of ribosomes and allows the cells to adapt their translation machinery to environmental fluctuations. In bacteria, the genes encoding rRNA molecules are typically organised in operons. The operons encode large primary transcripts that are processed and chemically modified. The resulting rRNAs are assembled into ribosomes together with ribosomal proteins in a multistep process. The activity of the assembled ribosome is constantly monitored, and a large set of enzymes is responsible for rRNA quality control, repair and degradation in case of ribosomal damage. The synchronisation of these processes creates an elaborate network of reactions in charge of the synthesis of the translation apparatus, its modification in response to environmental changes and its protection against stress factors. This review illustrates the current knowledge of the key events that occur in the lifecycle of rRNA molecules in bacteria, and it highlights the importance of a fine regulation of rRNA content for the maintenance of ribosome homeostasis. Keywords Ribosome biogenesis · rDNA operons · rRNA maturation · rRNA quality control · rRNA repair

1 Introduction RNA molecules in cells have functional significance (Bellacosa and Moss 2003), as they assume essential roles in the expression of genetic information retained by the DNA. The non-coding ribosomal RNA (rRNA) is the primary constituent (~60%) of

M. G. Giuliano · C. Engl (*) School of Biological and Chemical Sciences, Queen Mary University of London, London, UK e-mail: [email protected] © Springer Nature Switzerland AG 2021 I. Kotta-Loizou (ed.), RNA Damage and Repair, https://doi.org/10.1007/978-3-030-76571-2_2

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ribosomes. The precise folding of rRNA molecules generates the catalytic centres within the ribosome, providing not only a structural scaffold for ribosomal proteins (r-proteins) but also a mechanism for decoding messenger RNAs (mRNAs) into growing amino acid chains during the translation process. A significant amount of cellular resources and energy are used during protein synthesis, with more than 2/3 of the ATP requirements for biomass spent on protein translation in bacteria (Stouthamer and Bettenhaussen 1973; Tempest and Neijssel 1984; Russell and Cook 1995; Maitra and Dill 2015; Hu et al. 2019); it is therefore essential for the cells to strictly regulate this process. One way of controlling protein synthesis is by altering rRNA content. In all three domains of life, the biosynthesis of rRNA molecules comprises several steps that are highly regulated and monitored, allowing the cells to adapt their translational machinery throughout the life cycle and in accordance to changing environmental conditions. In bacteria, the genes encoding rRNA molecules (rDNAs) are typically organised in operons, whose number per chromosome can differ markedly from species to species. The operons encode large primary transcripts that undergo post-transcriptional processing and chemical modifications. The resulting rRNAs are assembled into ribosomes together with ribosomal proteins in a multistep process aided by assembly factors. The activity of the assembled ribosome is constantly monitored, and a large set of enzymes is responsible for rRNA quality control, repair and degradation in case of ribosomal damage under stress conditions. All these processes together create an elaborate network of reactions in charge of the synthesis of the translation apparatus, its modification in response to environmental changes and its protection against stress factors. In this chapter, we summarise the key events in the lifecycle of rRNA in bacteria to highlight the fine-tuning of rRNA content in order to maintain ribosome homeostasis.

2 Localisation and Copy Number of rDNA Operons in Bacteria In bacterial genomes, the genes for rRNAs—rrf (5S rDNA), rrs (16S rDNA) and rrl (23S rDNA)—are typically encoded in a single operon (rrn), which can be present in multiple copies distributed on one or more chromosomes (Suwanto and Kaplan 1989; Michaux et al. 1993; Rodley et al. 1995; Yamaichi et al. 1999) and on plasmids (Kunnimalaiyaan et al. 2001; Battermann et al. 2003). In Enterobacteria, most of the rrn operons are located in the half of the chromosome which is proximal to the origin of replication (oriC), and even though they are up to 180
apart on the E. coli genetic map, they come to close proximity in the folded 3D structure of the chromosome (Gaal et al. 2016). To our knowledge, the only bacteria known to have the rrn operon exclusively on small high copy number plasmids (9.4 and 6.6 kb, respectively) (referred to as rrn-plasmids) rather than on the chromosome are a clade within the genus Aureimonas (Anda et al. 2015) and three novel Oecophyllibacter

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saccharovorans strains (Ha5T, Ta1 and Jb2) (Chua et al. 2020, 2021). The number of rrn operons differs markedly among bacterial genomes, ranging from 1 to 17; E. coli, for example, contains 7 rrn operons (rrnA-E, rrnG and rrnH) (Stoddard et al. 2015; Espejo and Plaza 2018). The copy number variation of rrn operons has been correlated with the speed with which cells can synthesise ribosomes and therefore with their ability to rapidly respond to fluctuating growth conditions and resource availability (Klappenbach et al. 2000). Bacteria with few (1–2) rrn operons tend to be slow growing and oligotrophic organisms (for example, aquatic bacteria isolated from oligotrophic environments) (Fegatella et al. 1998; Strehl et al. 1999; Lauro et al. 2009), while those with many (around 9) rrn operons are copiotrophic and grow more rapidly during resource abundance (Klappenbach et al. 2000; Stevenson and Schmidt 2004; Dethlefsen and Schmidt 2007; Lauro et al. 2009; Roller et al. 2016). Thus, the average number and distribution of rrn copies per cell can provide important information on the ecological strategy of bacterial communities (Gao and Wu 2018). The rrn operon copy number variation has also been associated with biofilm formation under fluctuating environments (Niederdorfer et al. 2017) and ecological microbial succession (Nemergut et al. 2016). Bacteria with low rrn copy number (LCN) are more abundant in laboratory-grown oligotrophic biofilms than in eutrophic biofilms, and during the initial phase of biofilm formation (up to 25 days) high rrn copy number (HCN) species are significantly more abundant in biofilms from eutrophic than from oligotrophic systems (Niederdorfer et al. 2017). Furthermore, biofilms grown in oligotrophic systems have extended lag phases and lower growth rates than biofilms from eutrophic systems. However, a prevalence of HCN is observed in biofilms from oligotrophic glacier-fed streams, a natural highly unsteady environment due to snow and ice melts, and it was suggested that a short lag phase combined with a high growth rate is beneficial for biofilms to thrive under fluctuating environments (Niederdorfer et al. 2017). The discovery of rrn-plasmids in Aureimonas and O. saccharovorans reinforces the need to further investigate the role and modulation of multiple rrn operons in bacteria. The presence of rrn operons on high copy number plasmids could offer a selective advantage to the host under changing environmental conditions, ensuring a high rate of rRNA and protein synthesis (Anda et al. 2015; Chua et al. 2021). Another role for rrn-plasmids might be the recruitment of RNA polymerase (RNAP) to specific locations inside the cells, thus changing the pattern of RNAP foci formation and the bacterial nucleoid structure (see the aforementioned connection between the 3D structure of the chromosome and rrn co-localisation), and consequently a potential role in reprogramming global transcription (Anda et al. 2015). Indeed, it has been shown that in E. coli recombinant plasmids harbouring rrn operons impact the distribution of RNAP as well as cell growth (Cabrera and Jin 2006). Furthermore, E. coli strains with six of the seven rrn operons deleted are marked by redistribution of RNAP and decreased growth rate (Jin et al. 2016). A recent study further demonstrated that active transcription of rDNA is a driving force for nucleoid compaction (Martin et al. 2018). Hence, the localisation and copy number of rrn operons have a significant impact on bacterial physiology and reflect the copiotrophic/oligotrophic lifestyle of bacteria.

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Recently, the existence of widely distributed operon-unlinked 5S, 16S and 23S rDNAs has emerged (Brewer et al. 2020), and it seems to be widespread in bacteria and archaea (Ahn et al. 2020). In this peculiar genomic arrangement, the rRNAs are encoded by two separate transcription units, one for the 16S and one for the 23S–5S rRNAs. The 16S rDNA is separated from the 23S–5S rDNAs by internal transcribed spacer (ITS) regions greater than 1500 bp in length, differing from the canonical rrn operons that have an average ITS length of 419 bp. These operon-unlinked rDNAs are mainly associated with symbiotic bacteria (Ahn et al. 2020). However, the reasons and possible advantages of the rrn operon disruption are still unclear.

3 Organisation of rDNA Operons in Bacteria Bacterial rrn operons generally contain genes for 16S, 23S and 5S rRNA (Lafontaine and Tollervey 2001). Located at the 50 and 30 end of the operon are conserved external transcribed spacers (50 -ETS and 30 -ETS), and an internal transcribed spacer (ITS) separates the 16S gene from the 23S and 5S genes. A transfer RNA (tRNA) gene is usually located in the ITS, and additional tRNAs may also be encoded downstream of the 5S rDNA (Kaczanowska and Rydén-Aulin 2007). In E. coli six of the seven rrn operons show this general structure (Neidhardt et al. 1996), with the exception of the rrnD operon which contains two genes for 5S rRNA.

4 Transcription Regulation of rDNA Genes Along the Microbial Growth Curve Bacteria are able to adjust their rRNA content and thus the biosynthesis of the translation apparatus to the growth condition encountered (Klappenbach et al. 2000; Roller et al. 2016). Along the growth curve, bacteria change rRNA expression according to nutrient availability. In E. coli the rRNA fraction of the total transcription increases with increasing growth rate, reaching around 70% during exponential growth (Bremer and Dennis 1987; Dennis and Bremer 2008). Furthermore, recent studies suggest that sequence variations between multiple rrn operons within the same genome could yield subpopulations of ribosomes with different mRNA substrates and thus a targeted protein expression that increases the ability to adapt to new conditions (Chen et al. 2020; Culviner and Laub 2018; Kurylo et al. 2018; Maeda et al. 2015; Song et al. 2019). Moreover, in E. coli cells exposed to nutrient limitations, the upregulation of the rrnH operon—encoding a 16S rRNA characterised by ten variant nucleotides, nine of them clustered within helix 33 (h33) (forming the small ribosomal subunit head domain and A-site)—generates a specific pool of active ribosomes that selectively affects the expression of genes involved in stress responses (Kurylo et al. 2018). It was further shown that the rrsH

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sequence variants cause an alteration in the kinetics of binding interactions between AdhE, RelE, RelA and other translation factors and the ribosome, influencing both initiation and elongation phases of protein synthesis (Kurylo et al. 2018). The idea that multiple rrn copies in a cell might generate subpopulations of ribosomes with different mRNA substrates is also supported by findings in Vibrio vulnificus (Song et al. 2019). Here, ribosomes containing rRNAs encoded by the most divergent rrnI operon specifically translate mRNAs involved in temperature adaptation and nutrient shifts (Song et al. 2019).

4.1

Ribosomal DNA Promoters and Cis-Regulatory Elements

Bacteria implement an elaborate mechanism of transcription regulation to control the cellular rRNA content along the growth curve. The expression of rRNA genes is regulated by a combination of strong promoters (P1 and P2) and regulatory regions upstream of P1 that allow the recruitment of multiple transcription factors (Fis, H-NS and Lrp) (Fig. 1). Additional control is exerted via transcription initiating nucleotides (iNTPs) and the second messenger (p)ppGpp (Paul et al. 2004b; Gralla 2005). In E. coli, P1 and P2 are located 120 base pairs apart. The core promoter element of both P1 and P2 contains (i) the 10 and 35 region for binding σ70, (ii) an A-T rich binding site (UP element) for the α subunits of RNAP upstream of the 35 region, (iii) a G-C-rich region (discriminator) between 10 and the transcription start site and (iv) a consensus sequence recognised by σ32 (Newlands et al. 1993), the

Fig. 1 Regulation of rrn operons. Schematic representation of core elements of P1 and P2 promoters and regulatory regions upstream of P1 present in each E. coli rrn operon. The number of repeats of the transcription binding sites is indicated if known and (✔) if unknown

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sigma factor controlling the heat shock response during log-phase growth (Straus et al. 1987). The discriminator causes the formation of short-lived open complexes (Murray and Gourse 2004) enabling a rapid promoter response to the second messenger (p)ppGpp (Gafny et al. 1994; Paul et al. 2004b; Schneider and Gourse 2003; Haugen et al. 2006). The upstream regulatory regions can differ significantly between rrn operons containing varying numbers of binding sites and affinities for the transcription factors Fis, H-NS and Lrp (Fig. 1). This suggests a differential regulation of individual rrn operons (Hirvonen et al. 2001; Hillebrand et al. 2005). Indeed, Fis has the highest affinity for rrnH and weakest affinity for rrnE while H-NS binds weakly to rrnH and strongly to rrnB and rrnD (Hillebrand et al. 2005). Fis and H-NS specifically regulate the P1 promoter, and the corresponding regulatory regions upstream of P1 are characterised by varying levels of A-T content between the seven rrn operons that cause distinct curvatures in the DNA. DNA curvature is known to affect the strength of bacterial promoters and to change in response to environmental perturbations. Indeed, the extent of curvature upstream of P1 plays a role in sensing and responding to environmental changes (Hillebrand et al. 2005; Pul et al. 2008). Transcription from P2 is inhibited by transcription from P1 (Gafny et al. 1994) and in addition it is regulated in response to the cellular levels of CTP and GTP (Murray et al. 2003a; Murray and Gourse 2004). Thus promoter selection is highly coordinated, whereby P1 is responsible for intensive rRNA synthesis during rapid growth, while P2 is in charge of rRNA synthesis at low growth rates, during stationary phase and during outgrowth from stationary phase (Sarmientos and Cashel 1983; Murray et al. 2003a).

4.2 4.2.1

Trans-regulatory Elements Regulation by Transcription Factors

Fis is a small nucleoid-associated protein (Dillon and Dorman 2010) involved in the organisation and maintenance of nucleoid structure (Schneider et al. 2001; Cho et al. 2008). Fis also directly modulates the transcription of around 21% of genes (Cho et al. 2008). It binds to the upstream regulatory regions of P1 and to the C-terminal domain of the RNAP alpha subunit, stabilising the RNAP-P1 preinitiation complex (presumably at the open complex step) and leading to transcription activation (Hirvonen et al. 2001; Zhi et al. 2003). In vivo the occupancy of Fis-binding sites correlates with changes in Fis levels, and in vitro the activation of rrn promoters varies with Fis concentration. The largest effect on transcription is thereby exerted when Fis binds to sites proximal to the promoter while the impact of distal Fis-binding sites located further upstream of P1 varies between the rrn operons and ranges from 8% at rrnG to 73% at rrnA (Hirvonen et al. 2001). Transcription activation by Fis is antagonised by H-NS, a nucleoid-associated protein (Dame et al. 2000) which binds to sites that overlap with the Fis binding sites. H-NS condenses

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DNA and thereby acts as a repressor of transcription (Dorman 2004; Arold et al. 2010; Grainger 2016). H-NS also aids the interaction of the transcriptional repressor Lrp with regulatory regions upstream of P1 (Pul et al. 2005) to further downregulate expression (Pul et al. 2007). Overall, expression from rrn P1 promoters is a function of the relative amounts of Fis and H-NS. Specifically, expression from P1 is activated when Fis levels are high and H-NS levels are low and vice versa. The transcription of fis is increased at high levels of its initiating nucleotide CTP (Walker et al. 2004) and repressed by (p)ppGpp during nutrient limitation (Mallik et al. 2006). Thus during exponential growth, cells contain up to 60,000 copies of Fis, while its levels dramatically decrease in stationary phase to less than 100 molecules per cell (Azam et al. 1999; Dillon and Dorman 2010). As a consequence, Fis-dependent rRNA expression decreases in the stationary phase (Hirvonen et al. 2001).

4.2.2

Regulation by NTP and (p)ppGpp

Ribosomal promoters have an unusually high Km value for initiation by the +1 (Schneider et al. 2002) and +2 initiating nucleotides (iNTPs) (Lew and Gralla 2004)—GTP for the rrnD operon and ATP for the remaining six rrn operons of E. coli. The low affinity for iNTPs causes inhibition of rrn transcription in conditions where iNTP concentrations are limited. Cellular ATP and GTP pools increase with growth rate (Gaal et al. 1997; Bagnara and Finch 1973; Moses and Sharp 1972; Murray et al. 2003a, b) and begin to decrease in mid-log phase (Buckstein et al. 2008), reaching the lowest levels in stationary phase (Buckstein et al. 2008). During outgrowth from the stationary phase, the concentration of iNTPs increases rapidly in parallel with increased activity of the rrnB P1 promoter (Murray et al. 2003b). However, rRNA transcription during the stationary phase or in the earliest times in outgrowth is mainly driven by the rrn P2 promoters, which would provide the active ribosome pool available before the P1 promoters are activated (Murray and Gourse 2004). NTP pools, however, are not the only nucleotides involved in controlling rRNA synthesis. The second messenger nucleotide (p)ppGpp triggers what is termed the stringent response under nutrient deprivation (Irving et al. 2020; Potrykus and Cashel 2008; Potrykus et al. 2011). During a stringent response, the cellular resources and energy are redirected from intensive ribosomal synthesis, nucleotide biosynthesis and DNA replication to amino acid synthesis and transport, as well as the maintenance of essential cell functions only. (p)ppGpp inhibits rRNA transcription by binding directly to RNAP near the catalytic centre in the secondary channel (Potrykus and Cashel 2008; Hauryliuk et al. 2015; Liu et al. 2015; Anderson et al. 2019; Anderson et al. 2020; Zhang et al. 2019; Mechold et al. 2013; Gourse et al. 2018; Artsimovitch et al. 2004). There are three proposed mechanisms through which (p)ppGpp binding inhibits rRNA transcription: (i) (p)ppGpp competes with the initiating NTP for the active site, (ii) (p)ppGpp pairs with a cytosine residue (s) upstream of the transcription start site, or (iii) (p)ppGpp decreases the lifetime of

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the open complex formed at all rRNA promoters (Artsimovitch et al. 2004). The binding of (p)ppGpp to RNAP is stabilised by RNAP-binding factor DskA, amplifying the magnitude of the inhibitory effects of (p)ppGpp on all P1 and P2 promoters (Paul et al. 2004a; Lemke et al. 2011). The binding of (p)ppGpp and DksA affects transcription initiation at rrnB P1 by shifting promoter DNA into an incorrect position in the active site of the RNAP, preventing transcription initiation. Given that the lifetime of rrnB P2 open complex is two- to threefold longer than in rrnB P1, rrnB P2 is less affected by (p)ppGpp compared to rrnB P1 (Murray and Gourse 2004). In the current model, (p)ppGpp binds to two sites within RNAP to regulate transcription at rrnB P1: One site is located at the β0 –ω interface causing a three to fourfold inhibition, and one site requires DksA causing an additional fivefold inhibition (Gourse et al. 2018). Although (p)ppGpp and DksA regulate all rrn promoters in E. coli, the degree of inhibition varies depending on the promoter sequences and the nature and concentration of the iNTP (Kolmsee et al. 2011; Gourse et al. 2018); transcription from rrnD P1, the only rrn operon that uses GTP as iNTP, is subject to a weaker (p)ppGpp inhibition compared to the remaining six promoters (Kolmsee et al. 2011). Notably, the direct binding of RNAP by (p)ppGpp described above is not observed in bacteria belonging to the phyla Firmicutes, Actinobacteria and Deinococcus–Thermus. Here, (p)ppGpp controls growth indirectly through decreasing GTP levels causing a downregulation of rRNA promoters that use GTP as the transcription initiating nucleotide (Irving et al. 2020). Taken together, the regulation of rrn expression is not only dependent on the levels of Fis, H-NS and Lrp but also on (p)ppGpp and NTPs (Schneider and Gourse 2003; Gralla 2005; Gourse et al. 2018).

5 Processing of Premature rRNA into Functional rRNAs In all organisms, mature rRNAs are generated by post-transcriptional processing of the primary transcripts. In bacteria, nucleolytic cleavage of the primary rRNA transcript (containing all three rRNAs separated by intervening sequences) occurs via RNase III and produces separate precursors for the three individual rRNAs (pre-5S, pre-16S and pre-23S rRNA) (Malagon 2013). These are further processed by secondary nucleolytic cleavages to yield the final mature 5S, 16S and 23S rRNA (Srivastava and Schlessinger 1990; Condon 2007; Deutscher 2015) (Fig. 2). The rRNA processing events occur co-transcriptionally and in an assembly-assisted manner (Srivastava and Schlessinger 1988; Shajani et al. 2011). They take place within the nucleoid (Bohne 2014), where 10 to 15% of ribosome particles at different maturation stages are located (Bakshi et al. 2012).

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Fig. 2 Biogenesis of rRNA in bacteria (schematic representation adapted from Weis et al. 2015). The primary transcript comprising the three rRNAs (16S, 23S and 5S), as well as external and internal transcribed spacers (ETS and ITS) is processed by nucleolytic cleavage mediated by RNase III. Intermediate premature rRNAs are released from the primary transcript and further processed at 50 and 30 ends by specific ribonucleases. The ribonucleases currently identified in E. coli are shown. During maturation rRNAs are also subject to chemical modifications. Location, number and type of chemical modifications found in E. coli are indicated

5.1

Maturation of 23S rRNA in E. coli

Compared to the mature 23S rRNA, the pre-23S rRNA contains three to seven additional nucleotides at the 50 end and eight additional nucleotides at the 30 end (Dönhöfer et al. 2009). The 30 and 50 end maturation appears to be coordinated whereby processing at the 30 end facilitates 50 end maturation (Gutgsell and Jain 2010). RNase PH initiates 30 end maturation and removes a CC dinucleotide (Gutgsell and Jain 2012) which allows RNase T to proceed with maturation (Li et al. 1999a; Gutgsell and Jain 2012). Recent findings suggest that the 30 -50 exoribonuclease RNase R in cooperation with RNA chaperone Hfq is also involved in this process (Dos Santos et al. 2020), given that pre-23S rRNA accumulates in a ΔrnrΔhfq double mutant; the precise mechanism however remains to be determined. Our understanding of 50 end maturation is also still incomplete; yet it involves the single-stranded endoribonuclease RNase G (Song et al. 2011) and the 50 -30 exonuclease RNase AM (Jain 2020).

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Maturation of 16S rRNA in E. coli

After release from the primary transcript by RNase III, the pre-16S rRNA, also referred to as 17S rRNA, contains 115 and 33 nucleotides at the 50 and 30 ends, respectively (Dönhöfer et al. 2009). In absence of 30 end maturation, 50 end processing proceeds less efficiently (Sulthana and Deutscher 2013). Thus, it appears that similar to the 23S rRNA, 50 and 30 end maturation of 16S rRNA is linked. In E. coli, RNase E and RNase G are responsible for processing the 50 end of 17S rRNA (Li et al. 1999b). RNase E cleaves 66 nucleotides upstream of the 50 end, and RNase G subsequently removes the residual extra nucleotides (Li et al. 1999b). However, it has been shown recently that the exonuclease RNase AM also contributes to the maturation at the 50 end of 17S rRNA by removing three unprocessed nucleotides of 17S rRNA after RNase G cleavage (Jain 2020). Maturation of the 30 end of 16S rRNA is less well understood (Deutscher 2009). However, the removal of extra 33 nucleotides during 30 end maturation requires endonuclease YbeY as well as exonucleases RNase II, RNase R and RNase PH (Sulthana and Deutscher 2013; Jacob et al. 2013; Ghosal et al. 2018; Smith et al. 2018). Thus, an E. coli ΔybeY mutant grown at 45
C accumulates only partially 30 end matured 16S rRNA (Jacob et al. 2013). YbeY also interacts with proteins involved in ribosome biogenesis, such as the ribosomal protein S11 and the ribosome-associated GTPases Era and Der (Vercruysse et al. 2016), and overexpression of Era partially suppresses the growth defect of the E. coli ΔybeY mutant and improves 16S rRNA maturation in presence of RNase II, RNase R and RNase PH (Ghosal et al. 2018). Similarly, B. subtilis YqfG, a homologue of E. coli YbeY, is involved in the 30 end maturation of 16S rRNA, and the absence of YqfG results in the accumulation of pre-16S rRNA bearing 30 extensions (Baumgardt et al. 2018).

5.3

Maturation of 5S rRNA in E. coli

RNase III cleavage also releases pre-5S rRNA from the primary transcript. Subsequent processing is performed by RNase E, RNase T and RNase AM (Misra and Apirion 1979; Li and Deutscher 1995; Jain 2020). RNase E cleaves the pre-5S rRNA at three nucleotides upstream from its 50 and 30 end (Roy et al. 1983). RNase T then removes two additional nucleotides at the 30 end (Li and Deutscher 1995), and RNase AM completes the maturation of the 50 end (Jain 2020).

6 Bacterial rRNA Fragmentation RNase III is further responsible for rRNA fragmentation, the post-transcriptional cleavage of rRNAs at intervening sequences (IVS) (Evguenieva-Hackenberg 2005). These sequences are removed from the primary transcript without re-ligation, and

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the resulting rRNA fragments can thus be found in assembled ribosomes (Burgin et al. 1990; Gregory et al. 1996; Pronk and Sanderson 2001). In Alphaproteobacteria rRNA fragmentation requires the nucleolytic activity of multiple enzymes, some of them are still unknown. IVS have been detected in 16S and 23S rRNAs, with a sporadic distribution among bacteria and high sequence variability (EvguenievaHackenberg 2005). It has been suggested that IVS in rRNAs are more common amongst bacteria that are symbionts or pathogens of eukaryotic hosts and that IVS are acquired by prokaryotes through lateral transfer rather than vertical inheritance (Baker et al. 2003). In line with this, IVS in Enterobacteriaceae are confined to pathogenic strains (Pronk and Sanderson 2001). Nonetheless, the prevalence of IVS and the physiological implications of rRNA fragmentation in bacteria remain largely unknown. It has however been proposed that rRNA fragmentation might create targets for degrading ribonucleases RNase E, RNase R and PNPase via the formation of monophosphorylated 50 and 30 ends (Evguenieva-Hackenberg 2005). This might increase the efficiency of rRNA decay (Cheng and Deutscher 2003) during exponential growth when the abundance of rRNAs that contain IVS is increased (Evguenieva-Hackenberg 2005). However, stationary phase fragmentation of 16S rRNA at the tip of helix 6 in 30S subunits of E. coli (which does not contain IVS) (Luidalepp et al. 2016) shows that this phenomenon is not confined to the exponential phase and it can be independent of IVS. Stationary phase-specific cleavage of 16S rRNA within the 30S subunit has been shown to be an efficient means to attenuate ribosomal translation activities in non-proliferating environments (Luidalepp et al. 2016). Another role for rRNA fragmentation in the pathogenic strains Y. enterocolitica and S. typhimurium might be the removal of IVS to protect 23S rRNA fragments from unknown bacteriocins in the gut (Skurnik and Toivanen 1991). For S. typhimurium, it was also suggested that stationary phase fragmentation allows a quick adaptation to the rapid environment fluctuations experienced in animal hosts (Hsu et al. 1994).

7 Chemical Modification of rRNA Molecules In all domains of life, rRNA molecules are subject to chemical modifications. In bacteria, they contribute to the fine-tuning of rRNA content, to the maintenance and modulation of ribosome function and to ribosome heterogeneity in response to environmental changes (Decatur and Fournier 2002; Lapeyre 2004; Baxter-Roshek et al. 2007; Chow et al. 2007; Sergiev et al. 2011; Xue and Barna 2012; Byrgazov et al. 2013; Agris 2015; Polikanov et al. 2015). They also enhance the interaction of the ribosome with ligands and can confer antibiotic resistance (Doi and Arakawa 2007; Costello et al. 2019). Some modification enzymes play a role in rRNA quality control and act as assembly factors that check the accuracy of ribosome assembly (Gutgsell et al. 2005; Siibak and Remme 2010; Arai et al. 2015; Stojković et al. 2016; Leppik et al. 2017; Abedeera et al. 2020; Jayalath et al. 2020; Wang et al.

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2020). For an extensive overview on the consequences of chemical modification of rRNA, we refer the reader to other reviews (Sergiev et al. 2011; Agris 2015). Chemical modifications occur post-transcriptionally during maturation and map to specific regions of the assembled ribosome—mostly, but not exclusively, in the 50S subunit (95% in E. coli)—in the peptidyl transferase centre, the A- and P-sites, the exit tunnel and in the intersubunit bridges (Decatur and Fournier 2002; BaxterRoshek et al. 2007). The chemical modification found in the decoding region on the small subunit can influence ribosome structure and function; however, they are not essential for cell viability or ribosome assembly, in contrast to the modifications located in the large subunit (Byrgazov et al. 2013). In bacteria, rRNA molecules are covalently modified at the riboses and nucleobases by site-specific enzymes (Lapeyre 2004). There are three main types of covalent modifications: (i) methylation of nucleotide bases, (ii) pseudouridylation and (iii) methylation of the 20 -hydroxyl group of ribose (Decatur and Fournier 2002) (Fig. 2). A recent review illustrates the enzymes responsible for rRNA nucleotides modifications and positions of modified residues in E. coli (Sergiev et al. 2018). In E. coli, 21 nucleotide bases are methylated by a set of 21 methyltransferases (Golovina et al. 2012). Although further investigation is needed to unravel the advantages of rRNA base methylation, Pletnev et al. (2020) recently showed that rRNA methylation impacts fundamental steps of ribosome biogenesis such as rRNA processing and ribosome assembly (Pletnev et al. 2020). In E. coli nearly one third of the modifications found within the ribosome are pseudouridines (the base isomerisation of uridine to pseudouridine). One is found in 16S rRNA (modified by RsuA) (Conrad et al. 1999) and ten in 23S rRNA (modified by RluA-F) (Conrad et al. 1998; Raychaudhuri et al. 1998, 1999; Del Campo et al. 2001; Del Campo and Ofengand 2004). Yet, an E. coli strain lacking pseudouridynated rRNA due to deletion of all rRNA pseudouridine synthases showed only minor defects in ribosome biogenesis and function (O’Connor et al. 2018). Thus to date, the role of pseudouridynation is not clear. Methylation of the 20 -hydroxyl group of ribose has so far been identified at four residues in E. coli: (i) 16S rRNA residue C1402 located in the P-site of the 30S subunit (methylated by RsmI) (Zhao et al. 2014, 2016), (ii) 23S rRNA residue Gm2251 located in the P-site of the 50S subunit (methylated by RlmB) (Lövgren and Wikström 2001), (iii) 23S rRNA residue C2498 located in the peptidyl transferase loop (methylated by RlmM) (Purta et al. 2009) and (iv) 23S rRNA residue U2552 in the A loop (methylated by RrmJ (FtsJ)) (Bügl et al. 2000; Caldas et al. 2000; Hager et al. 2004) or RlmE) (Arai et al. 2015). Methylation of C1402 plays a role in regulating the shape and function of the P-site and increases the decoding fidelity; 20 -O-methylation of U2552 influences the assembly of the 50S subunit (Wang et al. 2020). Absence of U2552 20 -O-methylation delays 50S subunit assembly at multiple late stages and compromises translation initiation and elongation (Wang et al. 2020). The roles of O-methylation of G2251 and C2498 remains unclear. A ΔrlmB mutant has no ribosome assembly defects (Lövgren and Wikström 2001) and a ΔrlmM mutant only shows a slight reduction in fitness (Purta et al. 2009).

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8 Assembly of rRNAs into Ribosomes In bacteria the translationally active 70S ribosome is formed by the subunits 30S (16S rRNA, 21 r-proteins) and 50S (5S and 23S rRNAs, 33 r-proteins). The assembly of rRNAs into subunits is a multistep process involving a series of rRNA folding and r-protein binding events (Williamson 2003). Aided by additional factors such as RNA chaperones, RNA helicases and ribosome-dependent GTPases, r-proteins bind rRNA co-transcriptionally, cooperatively and hierarchically (de Narvaez and Schaup 1979) in 50 -30 direction, whereby binding of early r-protein influences the binding of late r-proteins. For a detailed overview on ribosome assembly, we refer the reader to articles by Williamson and co-workers (Shajani et al. 2011; Sashital et al. 2014; Davis et al. 2016; Davis and Williamson 2017).

9 rRNA Quality Control Ribosome biosynthesis is associated with a significant energy cost to the cell, and thus it is advantageous to resolve any faults in the structure and function of the ribosomal components during rather than at the end of the synthesis process. Consequently, cells have invested in surveillance pathways responsible for the quality control of rRNA production and ribosome assembly. To limit the cost of ribosome biosynthesis, faulty particles are degraded so that their components can be recycled to build new correctly structured particles. In bacteria, several ribonucleases are involved in ribosomal quality control, rRNA turnover and stress responses. Delays in ribosome assembly increase the exposure of rRNA to nucleolytic cleavage, and mutations in factors involved in ribosome assembly enhance rRNA fragmentation. The idea that rRNA maturation and ribosome assembly are strongly interconnected, so that the correct development of one process depends on the success of the other, is supported by the evidence that the E. coli RNase R acts in collaboration with (i) two RNA remodelling factors (DEADBox proteins DeaD and SrmB) (Jain 2018), (ii) the RNA chaperone Hfq (Dos Santos et al. 2020; Andrade et al. 2018; dos Santos et al. 2019; Quendera et al. 2020; Santiago-Frangos and Woodson 2018; Vogel and Luisi 2011) and (iii) RNase YbeY (Jacob et al. 2013). Specifically, double deletion of either DeaD or SrmB with RNase R increases rRNA breakdown (Jain 2018). As mentioned in the previous section on rRNA maturation, inactivation of both RNase R and Hfq results in strong accumulation of rRNA precursors and causes a sharp reduction in the levels of 70S ribosomes, suggesting the presence of severe assembly defects (Dos Santos et al. 2020; Domingues et al. 2015). RNase R and YbeY are responsible for the degradation of defective 70S ribosomes, in a process mediated specifically by defective 30S ribosomal subunits (Jacob et al. 2013). It has been proposed that YbeY initiates the degradation of 70S ribosomes bearing defective 30S subunits by performing

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endonucleolytic cleavage in exposed rRNA single strands, and afterwards the helicase activity of RNase R unwinds the rRNA (Awano et al. 2010) and continues the degradation of rRNA exonucleolytically (Jacob et al. 2013). Many other RNases are also involved in the degradation of faulty ribosomal particles in bacteria; these include PNPase, RNase III, RNase J and RNase I (Kitahara and Miyazaki 2011; Cameron et al. 2018; Datta and Burma 1972; Jones et al. 2014; Deutscher 2015). In most gram-negative bacteria, including E. coli, PNPase performs rRNA quality control by degrading 16S and 23S fragments generated by RNase E cleavage of improperly processed, damaged or overabundant rRNAs (Cheng and Deutscher 2003; Sulthana et al. 2016; Cameron et al. 2018). In some bacteria such as the radiation-resistant D. radiodurans, PNPase degrades misfolded rRNAs in response to nutrient starvation in complex with the Ro sixty-related protein Rsr (Chen et al. 2013; Cameron et al. 2018). It has been shown that in S. venezuelae cells lacking RNase III (double-strand specific RNase) or RNase J (a RNase that has RNase E-like endoribonucleolytic and a 50 -to-30 exonuclease activity) have higher levels of inactive ribosome dimers (100S ribosomes) than wild-type cells. Thus, multiple RNases contribute to the surveillance pathways responsible for the fine control of ribosome homeostasis; a detailed understanding of their interactions however is only just emerging.

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Damage and Repair of Bacterial rRNA

Similar to DNA, RNA is subject to damage by a range of abiotic and biotic factors. These include reactive oxygen species (M. Liu et al. 2012; Willi et al. 2018), alkylating agents (Yoshizawa et al. 1999; Sedgwick 2004) and enzyme-catalysed cleavage via toxins belonging to the RelE, MazF/Kid, Doc and PIN domain superfamilies of RNA endonucleases (Burroughs and Aravind 2016; Harms et al. 2018). Bacteria have developed dedicated repair systems to mitigate RNA damage and while the biochemistry of RNA repair is quite well established (Burroughs and Aravind 2016), a precise understanding of its physiology is still elusive (Bellacosa and Moss 2003; Wurtmann and Wolin 2009; Burroughs and Aravind 2016). RNA repair involves sequential steps of modifying the ends of damaged RNA (“cleaning” or “healing”) and subsequent rejoining of those modified ends (“sealing”) by RNA ligases (Burroughs and Aravind 2016). Two main classes of ligases have been identified: the ATP-dependent RNA ligases, and the GTP-dependent RtcB-like ligases. ATP-dependent RNA ligases contain an ATP-grasp protein fold and catalyse the joining of RNA molecules via a three-step mechanism similar to ATP-dependent DNA ligases: step 1, the RNA ligase reacts with ATP to form a covalent ligase-AMP intermediate; step 2, the AMP is transferred to the 50 -phosphate RNA end to form RNA-adenylate; and step 3, the RNA-adenylate reacts with the 30 -OH RNA end to form a phosphodiester bond, releasing the AMP (Gu et al. 2016). In contrast, RtcB joins either 30 -phosphate or 20 ,30 -cyclic phosphate RNA ends to 5’-OH RNA ends, in

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a unique three-step mechanism that requires GTP and Mn2+: step 1, formation of a covalent RtcB-(histidinyl-N)-GMP intermediate via reaction with GTP; step 2, transfer of guanylate to a polynucleotide 30 -phosphate to form a polynucleotide-(30 )pp(50 ) G intermediate; and step 3, attack of a 50 -OH on the –N(30 )pp(50 )G end to form the splice junction (Tanaka et al. 2011a, b; Chakravarty and Shuman 2012; Desai and Raines 2012; Englert et al. 2012; Desai et al. 2013). The 20 ,30 -cyclic phosphate RNAs are typically produced through RNA cleavage by ribonucleases, or they can be generated de novo by a widely conserved RNA cyclase named RtcA (Shigematsu et al. 2018). RtcA is thereby the enzyme responsible for the “healing” step during the RtcAB-dependent RNA repair. In E. coli RtcA is encoded in a σ54-regulated operon together with RtcB (Genschik et al. 1998). Expression of the RtcAB system is induced under stress conditions such as tRNA cleavage (Engl et al. 2016; Hughes et al. 2020), starvation (Engl et al. 2016), oxidative stress (Engl et al. 2016; Kurasz et al. 2018) and treatment with antibiotics that target the ribosome (Engl et al. 2016). In addition, RtcB expression in S. typhimurium is induced by nucleic acid crosslinking agents and by overexpression of endonuclease YafQ (Kurasz et al. 2018). In E. coli RtcB maintains ribosome homeostasis and 16S rRNA stability under nutrient starvation (Engl et al. 2016) and increases bacterial survival during treatment with antibiotics that target the ribosome (Engl et al. 2016). It can further repair truncated 30 -ends of 16S rRNA after cleavage by the stress-induced endonuclease MazF (Temmel et al. 2017).

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Conclusion

In all living organisms, fine-tuning of rRNA content is essential for an adequate biosynthesis and functioning of ribosomes in response to the cellular needs. This is especially relevant for bacteria that colonise a wide range of habitats and are often exposed to fluctuating growth conditions. The findings summarised in this chapter illustrate the heterogeneous nature of rRNA that underpins the adaptability of the protein synthesis machinery. However, much remains to be uncovered, and thus the lifecycle of rRNA is still an exciting and active field of research. Acknowledgements This work was supported by the Wellcome Trust [213955/Z/18/Z].

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The Rtc RNA End Healing and Sealing System Danai Athina Irakleidi, Harry Beaven, Martin Buck, and Ioly Kotta-Loizou

Abstract RNA occupies a central position in living systems, as an intermediate in information flow from DNA to proteins, as structural element of the ribosomes and as a regulator of gene expression at transcriptional and translational levels. Its formation and maturation is controlled through numerous enzymatic activities. Rather less well understood are the systems that act to restore the integrity of RNA. Here we describe one such system, focusing on the bacterial Rtc RNA end modification and ligation mediated respectively by the RtcA and RtcB enzymes, whose expression relies on an enhancer binding transcription control protein RtcR. Keywords RNA repair · RtcA RNA cyclase · RtcB RNA ligase · RtcR transcriptional activator · CRISPR-Cas systems · CARF domain · Ribotoxins · RNA damage · RNA end sealing

1 Introduction Ribonucleic acid (RNA) is a biopolymer, usually single-stranded (ss), whose structure was first characterised in 1965. RNA molecules consist of ribonucleotides linked via phosphodiester bonds that form chains of varying lengths. Ribonucleotides are composed of four different nitrogenous bases (adenine, cytosine and guanine and uracil) attached to a ribose sugar. As opposed to the more stable deoxyribonucleic acid (DNA), which evolved as the preferred genetic carrier, RNA contains a chemically reactive hydroxyl group that renders it prone to hydrolysis and therefore unstable.

D. A. Irakleidi · H. Beaven · M. Buck (*) · I. Kotta-Loizou (*) Department of Life Sciences, Faculty of Natural Sciences, Imperial College London, London, UK e-mail: [email protected]; [email protected] © Springer Nature Switzerland AG 2021 I. Kotta-Loizou (ed.), RNA Damage and Repair, https://doi.org/10.1007/978-3-030-76571-2_3

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The three most widely known types of RNA are messenger (m)RNA, transfer (t)RNA and ribosomal (r)RNA, all participating in the genetic flow of information from DNA to protein. For many years the perceived roles of RNA in the cell were limited to protein synthesis, but the established functions have greatly increased during the past few decades. In a broader sense, RNAs are classified as coding and non-coding (ncRNAs); the latter is divided further into housekeeping ncRNAs (including tRNAs and rRNAs) and into regulatory, small (200 nt) ncRNAs. Regardless of their classification, ncRNAs have a wide spectrum of diverse biological roles at epigenetic, transcriptional and post-transcriptional levels, such as regulating chromatin modification, gene expression and microRNA levels and functions. In the light of the above, this chapter focuses on the universally conserved RNA 30 -terminal phosphate cyclase (Rtc) enzymes, the RNA cyclase RtcA and the RNA ligase RtcB, which act together as components of an RNA end healing and sealing system in Escherichia coli and other prokaryotes. The expression regulation of the Rtc system via its transcriptional activator RtcR is evolutionarily linked to the prokaryotic clustered regularly interspaced short palindromic repeats (CRISPR)Cas systems.

2 Biological RNA Damage Functionally important RNA molecules are very highly conserved both in terms of sequence and structure, rendering RNA one of the least evolved cellular components in nature. Due to this “lack” of evolution and its inherent instability, RNA is a common target in a range of biological conflict systems that aim at disabling a prokaryotic or eukaryotic cell: effectors can target either RNA associated with invasive, exogenous material as a means of self-preservation or endogenous, “self” RNA as a way of inducing cell apoptosis or cell dormancy to eliminate the exogenous threat (Burroughs and Aravind 2016). Such effectors include ribotoxins, enzymes that cleave essential and conserved RNA molecules such as tRNAs to endonucleotically induce cell death. Bacterial ribotoxins are expressed by the same operon as their respective labile antitoxins; the toxin and antitoxin form a stable complex to prevent the toxin from harming the cell. Under stress, the antitoxin is degraded and the toxin is freed (Zhang et al. 2005). Numerous such toxin-antitoxin (TA) systems with slightly different modes of action have been identified in E. coli, but the most well-characterised is the MazEF TA system (Zhang et al. 2005; Nariya and Inouye 2008). Endoribonuclease effectors are also an integral part of some CRISPR-Cas systems.

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CRISPR-Cas Systems

The prokaryotic CRISPR-Cas system was first discovered accidently in 1987 by a team of scientists that identified a group of common interspaced repeat elements, initially in the E. coli genome and subsequently in other archaeal and bacterial species (Ishino et al. 1987). CRISPR-Cas systems act as an intrinsic adaptive response of archaeal and bacterial immune systems against invading exogenous DNA, such as phages (Rath et al. 2015). CRISPR loci have been found in both chromosomal and plasmid DNA and are formed by short palindromic repeats with unique sequence spacers in between (Rath et al. 2015). Their length is widely conserved among species: the repeats are 21–48 bp and the spacers 26–72 bp (Jansen et al. 2002; Bolotin et al. 2005). The number of different spacers present in any bacterial cell at any given time is an evolutionary representation of the different viral infections encountered by this cell and its ancestral line, ranging from just a couple to several hundred. The spacers are responsible for the recognition and inactivation of exogenous nucleic acids, from which they themselves are probably derived (Rath et al. 2015). In many species, CRISPR-based immunity relies on a family of RNA-guided CRISPR-associated (Cas) proteins, encoded by genes adjacent to the CRISPR loci. Other species lack these adjacent genes and rely on trans-encoding factors instead (Rath et al. 2015). The Cas protein family is diverse, accommodating members different in both function and structure: most have nuclease activity, while others serve as helicases or RNA-binding proteins (Makarova et al. 2002; Rath et al. 2015). The Cas nuclease binds to a single-stranded long precursor CRISPR RNA (pre-crRNA) transcript, leading to the formation of the CRISPR-ribonucleoprotein (crRNP) complex. The Cas nuclease together with cofactors processes the pre-crRNA transcript into a shorter mature crRNA molecule corresponding to a single spacer, complementary to the DNA or RNA sequence targeted for cleavage (Rath et al. 2015). Following assembly of the complex, the crRNA targets the Cas protein to the respective target area, based on simple complementarity principles (Jinek et al. 2012). The only additional requirement for the binding event to proceed is the presence of a 2–6 nt protospacer adjacent motif (PAM) sequence at the end of the DNA region targeted for cleavage, usually 3–4 nt after the designated cleavage site. The PAM sequences help distinguish exogenous from self DNA and are used as recognition and binding sites for Cas proteins that will unwind dsDNA, allowing subsequent cleavage (Gleditzsch et al. 2019). When catalytically active, the Cas nuclease will induce an RNA-programmed double-stranded break (DSB) in the genomic DNA (Fig. 1), thereby disrupting it and diminishing the threat.

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Fig. 1 The molecular mechanism underpinning CRISPR-Cas immunity: (1) entry of viral nucleic acid into the cell and acquisition of new target spacer sequence as part of the CRISPR locus, (2) transcription of the spacer sequence from the CRISPR locus, (3) crRNA processing and maturation, (4) detection and cleavage of the viral nucleic acid as guided by the Cas nuclease and the crRNA molecule

CRISPR systems are broadly divided into two main classes; class 1, which relies on multisubunit protein complexes and class 2, which involves single multidomain protein effectors (Shmakov et al. 2017; Makarova et al. 2020). Each class is subdivided into three types (types I, III, IV and types II, V, VI respectively) and further subtypes, depending on the signature genetic localisation and orientation as well as on protein sequence and conservation (Makarova et al. 2020). Ca. 90% of CRISPR systems in bacteria and archaea (including all hyperthermophiles) are class 1 (Makarova et al. 2015), with the remaining 10% being class 2 and exclusively found in bacteria, but not in hyperthermophiles (Chylinski et al. 2014; Makarova et al. 2015; Shmakov et al. 2017). For instance, the well-characterised CRISPR-Cas9 system belongs to the class 2 type II subcategory. Although CRISPR systems predominantly use DNA as a natural substrate, recent studies suggest that some CRISPR-associated nucleases can also recognise and cleave ssRNA molecules in a mode of action independent of the PAM sequence (Makarova et al. 2020). For example, a putative class 2 type IV single-effector system relies on the C2c2 (Cas13a) nuclease, which does not share any homology with other characterised DNA-targeting nucleases but is instead formed by two ribonuclease (RNase)-activity

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higher eukaryotes and prokaryotes nucleotide-binding (HEPN) domains; therefore, C2c2 is an RNA-guided CRISPR component with RNase activity (Abudayyeh et al. 2016; East-Seletsky et al. 2016; Shmakov et al. 2017). The system is expected to function in a similar way to the typical CRISPR-Cas9 DNA-targeting system, with the only difference being that its substrate is ssRNA. Additional RNA-guided RNA-targeting systems have been identified (VI-B1 and VI-B2), which operate through Cas13b nucleases that can cleave RNA through their target-activated RNase activity. However, unlike C2c2, Cas13b nucleases are differentially regulated by additional proteins which are co-expressed by the VI-B1 and VI-B2 loci (Abudayyeh et al. 2016; East-Seletsky et al. 2016; Smargon et al. 2017).

3 RNA Repair To counteract the systems that operate against RNA in the cell, a range of RNA protection and repair systems are in place, ensuring proper maintenance and potential damage repair. Since the discovery of RNA repair effectors in vivo is only recent, their characterisation is relatively poor (Burroughs and Aravind 2016). However, some common characteristics of the known RNA repair systems have been identified, including (1) RNA ligation effectors with endoribonuclease activity, such as T4 RNA ligases; (2) RNA elongation effectors, typically polymerase-mimicking nucleotidyltransferases, such as the DNA polymerase β CCA enzymes that add a CCA trinucleotide to the 30 termini of tRNAs, in a template-independent fashion, to stabilise the CCA-lacking mature tRNAs following cleavage by the metallo-betalactamase (MBL) fold endonuclease tRNase Z (Vogel et al. 2005); (3) “cleaning” domains, structurally and functionally diverse, that modify RNA termini, either by adding or by removing phosphate groups or by performing additional modifications to prevent degradation, so that the termini can be processed by the ligase and/or elongation effectors; and (4) cofactors and additional binding protein components that increase the catalytic rate of these reactions (Burroughs and Aravind 2016).

4 The RtcA and RtcB Enzymes RtcA and RtcB are two evolutionary conserved RNA healing and sealing enzymes with homologs in species across all life kingdoms, excluding fungi and plants (Tanaka and Shuman 2011). The biochemistry and structure of both enzymes are well-studied, but their physiological roles in the cells are not well understood, with the exception of RtcB in humans and metazoans.

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The RtcA RNA Cyclase

The E. coli RtcA is a 338 aa enzyme catalysing a phosphate transfer reaction that converts RNA 30 -monophosphate termini into 20 ,30 -cyclic phosphate termini, which are subsequently ligatable and can serve as substrates for RNA end sealing (Das and Shuman 2013). Cyclisation can be divided into three nucleotidyl transfer reactions. Initially, RtcA and ATP yield an RtcA-AMP intermediate, where AMP is linked via a phosphoramide covalent bond to the NE atom of His-309 (Chakravarty et al. 2011) and coupled with the release of a PPi molecule (Filipowicz et al. 1985). Subsequently, AMP is transferred from the RtcA-AMP intermediate to the RNA 30 -monophosphate terminus yielding an activated phosphoanhydride intermediate RNA(30 )pp(50 )A (Reinberg et al. 1985). Finally, complete cyclisation occurs through the interaction of the 2’-OH ribose terminus with the 30 -phospate terminus of RNA (30 )pp(50 )A resulting in the generation of the 20 ,30 -cyclic diphosphate RNA product and the release of AMP (Billy et al. 1999). Although these steps and intermediate products strongly mimic those catalysed by typical RNA/DNA ligases, both the active-site composition and the tertiary structure of RtcA shares very few, if any, common features with those of typical polynucleotide ligases (Palm et al. 2000; Tanaka et al. 2010). Resolved crystal structures of RtcA (Fig. 2) reveal a fold of four tandem domains, each consisting of four β-sheet strands connected to two α-helices (Palm et al. 2000; Tanaka et al. 2010). Domains 1, 2 and 4 are identical and are arranged in a globular unit with a pseudo-threefold symmetry. Domain 3 has a different tertiary arrangement and is embedded into this globular unit between the β1 strand and the α1 helix of the fourth domain (Tanaka et al. 2010; Chakravarty et al. 2011). Crystal structures of the RtcAAMP intermediate provide further insight into RtcA substrate specificity. Initially the preference for ribonucleotide substrates is due to the formation of a hydrogen bond network between Asp-306 and Gln-307 and the 20 and 30 free oxygen molecules provided by the riboses present. Secondly, the preference for ATP as a cofactor, as opposed to GTP, is explained by the fact that following binding to His-328, the ATP adenine base is embedded within a hydrophobic pocket lined by Pro-150, Phe-154 and Tyr-303, with the latter further favouring the insertion of the ATP purine C2 subdomain into the pocket (Chakravarty et al. 2011). The same intermediate structures have been identified as containing both one citrate and two sulphate anions, held within the RtcA N-terminus by electrostatic interactions and hydrogen bonds with adjacent conserved residues (Chakravarty et al. 2011). Although it is still unclear how their presence aids catalytic activity, it is speculated that these anions mimic either the substrate RNA phosphate atoms or the PPi adenylation released product (Tanaka and Shuman 2009; Chakravarty et al. 2011).

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Fig. 2 Crystal structures of RtcA complexes in Escherichia coli, obtained via X-ray diffraction (1.85–1.90 Å). (a) The RtcA-ATP initial complex; (b) the RtcA hydrophobic-lined pocket, where ATP is embedded; (c) the RtcA-AMP intermediate; and (d) the RtcA four tandem domain fold structure, with domains annotated: domain 1 (aa 4–86) in red, domain 2 (aa 86–177) in light blue, domain 3 (aa 186–277) in purple and domain 4 (aa 178–185 and 278–339) in green

4.2

The RtcB RNA Ligase

E. coli RtcB is a 408 aa long RNA ligase that repairs RNA breakage by sealing 20 ,30 -cyclic phosphate termini or 30 -phosphate termini with 50 -hydroxyl RNA termini (Tanaka et al. 2011a, b; Desai and Raines 2012; Chakravarty and Shuman 2012; Maughan and Shuman 2016). RtcB is characterised as an “atypical” ligase in that, unlike most ‘typical’ ligases such as T4 ligases, which use ATP and magnesium (Mg) II as cofactors, it relies on GTP and manganese (Mn) II.

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Fig. 3 Crystal structure of the RtcB/Mn2+-GMP intermediate complex in Pyrococcus horikoshii, obtained via X-ray diffraction (2.4 Å). (a) The tetrahedral orientation of the Mn(t) divalent cation held by active-site side chains; (b) the positioning of the two divalent cations, Mn(t) and Mn(o), in the RtcB active site in the presence of the conserved Cys-98 residue side chain; (c) the octahedral orientation of the Mn(o) divalent cation held by active-site side chains; and (d) the RtcB/Mn2+GMP intermediate complex in a twofold symmetry

RtcB-mediated RNA ligation occurs through three GTP-dependent nucleotidyl transfer sub-reactions that ultimately lead to the formation of a 30 ,50 -phosphodiester bond between the substrate termini. Initially, GTP becomes covalently bound to the His-404 residue and forms an RtcB/Mn2+-GMP intermediate upon the release of PPi (Desai et al. 2013). Subsequently, the GMP moiety is attached to the 30 phosphate substrate terminus, which hydrolyses into an activated 30 -monophosphate group (Tanaka et al. 2011b; Chakravarty and Shuman 2012; Chakravarty et al. 2012). Finally, the 5’-OH substrate terminus attacks the 30 -phosphate one to form a covalent 30 -50 phosphodiester bond, releasing the GMP moiety (Desai et al. 2013). Structural analysis of the Pyrococcus horikoshii RtcB ligase (Fig. 3), which possesses an almost identical structure to its E. coli homolog, has shown that under physiological conditions, two Mn2+ cations and a GTP molecule are covalently bound to the active site of the ligase. In addition, four sulphate anions (SO42 1–4) surround the putative catalytic site (Englert et al. 2012). The first divalent cation, referred to as Mn(t) is found in a tetrahedral orientation with the active-site residues Asp-95, Cys-98 and His-203 together with a sulphate ion that replaces the

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20 -30 -cyclic phosphate RNA substrate in states of inactivity (Englert et al. 2012). The second divalent cation, Mn(o), is instead found in an octahedral coordination with the conserved residues Cys-98, His-234 and His-329 and three water molecules (Englert et al. 2012). The two different positions of the divalent ions within the active site and their interactions with the same Cys-98 residue explain why Mn is preferred over Mg as a cofactor: the distance between the sulphate anions upon interaction with Mn matches that of a typical diphosphate bond in a 50 -3’ ssRNA molecule, such as the RtcB substrate. Therefore, Mn prevents steric clashes following substrate binding and during catalysis (Englert et al. 2012). Finally, crystal structures of the guanylated RtcB/Mn2+-GMP complex, following release of PPi, suggest that the conserved His-404, which forms part of a His-404-Asp-95 dyad, is the site of RtcB guanylation. More precisely, GTP interacts with RtcB through its α-phosphate, forming a covalent phosphoramide link with the Nε2 atom of His-404, inducing a conformational change to the structure of RtcB. This conformational change leads to the optimal alignment of the His-404-Asp-95 dyad, so that Asp-95 can form a hydrogen bond with the Nδ1-H donor group of His-404 and reversibly interact with the His-404 backbone amide. The His-404-Aps-95 dyad then catalyses RtcB guanylation and gives rise to a 50 -GMP-His (GPH) intermediate (RtcB/Mn2+GMP complex) (Englert et al. 2012).

4.3

The RtcB RNA Ligase in Prokaryotes

There are numerous examples of RtcB homologs in other prokaryotic species such as the bacteria Thermobifida fusca and Thermus thermophilus and the archaeon Pyrococcus horikoshii (Desai et al. 2015), which display close structural similarities to the E. coli RtcB, with virtually identical folds, similar histidine-coated hydrophilic active-site pockets and sulphate anion binding sites. However, unlike E. coli and T. fusca, the RtcB in T. thermophilus and P. horikoshii acts in an archease-dependent fashion instead. Archease is a protein that binds to RtcB in order to accelerate the ligation process by modifying NTP specificity, therefore allowing the use of either GTP or ATP as a cofactor (Desai et al. 2014). Crystal structures (Desai et al. 2014) of archease suggest that it has an anionic surface charge, which allows it to embed into and bind to the cationic-coated active site of RtcB (Desai et al. 2014). In addition, a metal ion binding site has been identified at the surface of archease, potentially essential for optimal activity (Desai et al. 2014). Although archease is conserved across all kingdoms of life, its expression has been detected in a small percentage of bacterial taxa, suggesting that numerous bacteria perform RtcB ligation in an archease-independent manner (Desai et al. 2015).

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The RtcB RNA Ligase in Metazoans

The most well-studied homolog is the human RtcB, also known as HSP117, which modifies RNA in human cells for the purposes of regulating dynamic cellular functions. Human RtcB, similar to its bacterial homolog, is a noncanonical RNA ligase that relies on GTP and Mn(II) for enzymatic catalysis, instead of ATP and Mg (II) (Desai and Raines 2012). RtcB was first identified in humans as a tRNA ligase (Popow et al. 2011), since tRNAs are an indispensable component of protein synthesis and tRNA splicing is crucial for tRNA maturation, involving precursor tRNA intron removal followed by exon ligation (Hirata 2019). The introns are enzymatically cleaved by endonuclease A (Abelson et al. 1998), and the exons are religated by RtcB (Englert et al. 2011; Popow et al. 2011; Hirata 2019). RtcB also re-seals the X-box-binding protein 1 (XBP1) exons, after intron removal by the endoplasmic reticulum (ER) transmembrane sensor inositolrequiring enzyme 1 (IRE1) (Sidrauski et al. 1996; Chen and Brandizzi 2013; Lu et al. 2014; Jurkin et al. 2014; Kosmaczewski et al. 2014). XBP1 is a cell- and condition-specific transcriptional regulator that contributes to the maintenance of ER homeostasis. XBP1 activity has been correlated with the upregulation of ER-resident protein folding chaperones in cases of ER stress (Yoshida et al. 2001; Calfon et al. 2002; Lee et al. 2002, 2003), which is primarily characterised by the accumulation of unfolded proteins within the organelle (Cao and Kaufman 2012; Hetz 2012). XBP1 splicing, and therefore RtcB that mediates it, play a crucial role in the adaptive unfolded protein response that restores ER protein homeostasis in eukaryotic cells. The RtcB homolog in Caenorhabditis elegans is involved in regulating neuronal axon growth: it localises in neurons in an injury-dependent manner and inhibits axon regeneration, potentially assisting the more controlled neuronal development and wiring, and preserving resources needed to properly address axonal damage (Kosmaczewski et al. 2015). Unlike tRNA or XBP1 ligation, RtcB activity on axon regeneration occurs independently of known cofactors, including archease, a typical cofactor for both tRNA and XBP1 ligation (Kosmaczewski et al. 2015). Naturally, there are a large number of RtcB homologs found across species, such as in the white-tufted-ear marmoset Callithrix jacchus and in the king cobra Ophiophagus hannah. Both these RtcB homologs perform a GTP and Mn(II)dependent ligation of 20 -30 -cyclic phosphate or 30 -phosphate termini with 5’-OH RNA termini in a process almost identical to that of bacterial RtcB. Despite potential structural or minor functional discrepancies, the process of RtcB-mediated RNA repair via religation is a highly conserved pathway across all kingdoms.

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5 The Bacterial Rtc RNA Repair System In E. coli, the rtcB and the rtcA genes are genetically linked and under the control of the rtcBA operon (Fig. 4), indicating that they contribute towards the same biological process (Das and Shuman 2013). The RtcA RNA cyclase, whose enzymatic reaction does not yield a fully sealed final product, is considered to play a supportive role to the RtcB RNA ligase, instead of being the main enzyme. In addition to RNA 30 -monophosphate termini, RtcA also uses RNA 20 -monophosphate termini as substrates, in order to provide 20 ,30 -cyclic phosphate products that will be subsequently religated by RtcB (Tanaka and Shuman 2009; Tanaka et al. 2011b; Das and Shuman 2013). Moreover, the RtcA 20 ,30 -cyclic phosphate products may be involved in signalling pathways for rtcBA expression regulation (Makarova et al. 2014). In fact, 20 ,30 -cyclic phosphate-containing RNAs may be a hidden layer of the transcriptome that could play roles in a wide range of physiological processes (Shigematsu et al. 2018). The activation of the rtcBA operon and the expression of both proteins are regulated by the alternative sigma factor σ54 in cooperation with RtcR, an additional transcription factor that controls the stress-induced Rtc RNA repair system (Genschik et al. 1998). The σ54 subunit does not share homology with any other known sigma factors (Keppetipola et al. 2009) and is an indispensable component of the bacterial RNA polymerase (RNAP) complex, required for promoter recognition and transcription initiation (Studholme and Dixon 2003). RtcR is a σ54-dependent transcriptional activator which is independently expressed by a neighbouring gene, oppositely oriented to the rtcBA operon (Genschik et al. 1998). Expression regulation of the Rtc system is not well understood, although a CRISPR-Cas-associated Rossmann fold (CARF) domain is known to be involved.

Fig. 4 Schematic representation of the Rtc system in Escherichia coli. The transcriptional activator RtcR is expressed by an oppositely oriented neighbouring gene, in a σ 70-dependent manner. In a hexameric arrangement, RtcR binds the rtcBA upstream activating sequence (UAS) and induces transcription of the rtcBA operon by the RNA polymerase (RNAP), in a σ 54-dependent manner

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Diversity of the rtc Operon in Bacteria

RtcA and RtcB homologs have been identified in a range of different bacterial taxa, but do not necessarily share the same genetic arrangement as in E. coli, where the rtcA and rtcB genes are genetically linked, controlled by the same operon and clustered with rtcR (rtcR ! rtcB–rtcA). Overall, this genetic organisation is predominantly conserved across most bacterial taxa that display rtc genetic linkage; however, there are some notable exceptions. For example, in the δ-proteobacterium Pelobacter carbinolicus, the order of the rtcA and rtcB genes is reversed as compared to E. coli (rtcR ! rtcA–rtcB) (Das and Shuman 2013). Furthermore, in the γ-proteobacterium Hahella chejuensis, the rtcR gene is transcribed in the same direction as the other two and, once again, rtcA precedes rtcB (! rtcR–rtcA–rtcB). Nevertheless, despite such differences, a highly conserved feature across all bacteria expressing the Rtc proteins is the regulation of the rtcBA operon by RtcR and the σ54 factor (Das and Shuman 2013). Unlike E. coli, many of the bacterial taxa that display rtc genetic linkage carry additional protein coding genes close to the rtcBA operon, either directly preceding it or located between rtcA and rtcB. The most common example of this feature is that of the gene coding for archease, which is present in all bacterial species that display an archease-dependent RtcB ligation. For instance, the archease in Thermus thermophilus is encoded by an rtc neighbouring gene, whose expression is also regulated by the rtcBA operon (Desai et al. 2014). Another example is the presence of the gene encoding the RNA-binding protein Ro in the rtcBA operon preceding the rtcB gene (rtcR !ro–rtcB–rtcA). This arrangement is found in a few Proteobacteria including Salmonella typhimurium and Pseudomonas fluorescens, together with the planctomycete Pirellula staleyi and some chlamydiae (Das and Shuman 2013). Ro is a relatively small (ca. 60 kDa) bacterial protein that performs ribonucleotide quality control by recognising damaged or misfolded RNAs and binding to them in order to induce their removal (Chen et al. 2003). Consequently, Ro may be expressed alongside RtcA and RtcB in order to participate in RNA repair.

5.2

The RtcR Transcriptional Regulator and Its CARF Domain

The RtcR protein consists of three domains; an N-terminal regulatory domain, a NtrC-like AAA+ (ATPase associated with diverse cellular activities) domain and a helix-turn-helix (HTH) DNA-binding domain (Makarova et al. 2014; Engl et al. 2016). The regulatory domain is a divergent member of the CARF family, whose canonical members can be found in most CRISPR-Cas systems (Wiedenheft et al. 2012; Koonin and Makarova 2013; Makarova et al. 2014). The core of the CARF domain is a nucleotide-binding, Rossmann-like fold, consisting of six core strands, two of which (strands 5 and 6) form a β-hairpin

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(Makarova et al. 2014). Higher-sequence conservation is identified in strands 1 and 4, in locations associated with ligand binding sites: the one terminus of strand 1 contains a polar residue, usually Ser or Thr, whereas the sequences immediately following strand 4 have highly conserved basic residues (Lys or Arg), often as part of a set signature, [D/N]X[S/T]XXX[R/K] (Makarova et al. 2014). These conserved motifs participate in the formation of a substrate-binding pocket, and the presence of the highly basic Lys and/or Arg residues within the pocket supports the hypothesis that the primary substrate for at least the majority of CARF domains is negatively charged nucleotides or nucleotide-derived molecules (Lintner et al. 2010). However, RtcR does not have basic residues downstream of strand 4 within the active-site pocket, suggesting that it operates under the control of different ligand types (Makarova et al. 2014). CARF-containing proteins often include a bent, C-terminal helix-turn-helix domain and an intermediate domain with DNase or RNase activity (Lintner et al. 2010; Kim et al. 2013; Makarova et al. 2014). CARF domain activity is regulated by ligand binding events: following ligand binding, the CARF domain acts as a signal transductor influencing the effector domain of the protein. In the case of CRISPRCas systems, the ligand is usually a cyclic oligoadenylate (cAn, n ¼ 2–6) produced by the Palm domain of cyclic oligoadenylate synthases triggered by the presence of foreign nucleic acids in the cell (Kazlauskiene et al. 2017; Niewoehner et al. 2017). Interestingly, CARF domains are also responsible for the degradation of the cyclic oligoadenylate signals, leading to deactivation of the CRISPR nucleases when they are no longer needed (Athukoralage et al. 2020). Other CARF domains may control different effector domains such as GTPases or ATPases (Makarova et al. 2014). RtcR, the most well-characterised “CRISPRindependent” protein with a CARF domain is a transcriptional activator, controlling the activity of the RtcR AAA+ domain, and subsequently expression of the rtcBA operon. Normally, the CARF domain inhibits the AAA+ domain, and the expression of the rtcBA operon is repressed (Genschik et al. 1998). Under stress conditions that require the RNA end healing and sealing activity of the Rtc system, an as yet unknown ligand binds onto the CARF domain. The latter undergoes a conformational change that alleviates the inhibition of the AAA+ domain, allowing activation of the σ54 transcription factor and expression of the rtcBA operon. The exact ligand (s) of the RtcR CARF domain and events leading to the derepression of the AAA+ domain have not yet been determined. The ligands may be linear or cyclic oligonucleotides or damaged RNA molecules with 20 ,30 -cyclic phosphate termini generated by RtcA (Makarova et al. 2014). Furthermore, in Salmonella enterica serovar Typherium, RtcR may activate the expression of the rtcAB operon upon binding to 20 ,30 -cyclic phosphate tRNAs, in an RtcA-dependent manner (Hughes et al. 2020).

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Expression and Function of the Rtc System in Bacteria

Work on RtcB in metazoa illustrate its involvement in tRNA splicing (Popow et al. 2011), ER stress (Kosmaczewski et al. 2014) and neuron regeneration (Kosmaczewski et al. 2015); however, it does not provide any insight on the potential role of the Rtc RNA repair system in prokaryotes. The cellular and genetic conditions under which the rtcBA operon is expressed, and therefore the RtcA and RtcB enzymes are physiologically important, have been mostly studied in E. coli and in S. enterica serovar Typherium (Table 1). In E. coli, the Rtc system has been shown to help the bacterium respond to stress conditions such as defects in translation, oxidative stress and exposure to antibiotics. Lack of individual rtc genes influences cell phenotypes with ΔrtcR, ΔrtcA and ΔrtcB mutants forming weaker biofilms and showing increased expression of genes related to chemotaxis and mobility (Engl et al. 2016). The RtcA and RtcB are associated with the ribosome, where RNA is integral to both structure and function (Engl et al. 2016; Temmel et al. 2016). Protein synthesis is fundamental for survival, and in bacteria it is performed by a 70S 2-subunit ribosomal complex containing as many as 54 different proteins. Given the large number of ribosomal proteins physically interacting with RtcB (Temmel et al. 2016), the primary role of the Rtc in bacteria may be to repair RNA vital to the integrity of translation within the cell. Both ribosome generation and protein synthesis are very energy-consuming, and, therefore, the Rtc system may ensure that altered or dysfunctional ribosomal components are repaired or removed in time to allow high translational efficiency and that all the energy and resources utilised are not wasted. For instance, RtcB may be involved in a mechanism used to cope with cellular stress by reprogramming protein synthesis, following stress-induced cleavage by MazF at an ACA site of the 43 nt at the 30 terminus of 16S rRNA in the 70S ribosomal assembly (Vesper et al. 2011). The reaction generates 16S rRNA fragments with a 20 ,30 -cyclic phosphate terminus (70SΔ43) and a 50 -hydroxyl (5’-OH) terminus (RNA43), both remaining associated with the 30S ribosomal subunit, and is reversible by RtcB ligation (Temmel et al. 2016). The stress-specialised ribosomes generated lack helix 45 and the anti-Shine-Dalgarno (aSD) sequence, both crucial for normal initiation of mRNA translation (Shine and Dalgarno 1974), and are able to selectively translate exclusively MazF-processed mRNAs, leading to a form of translational reprogramming (Sauert et al. 2016; Temmel et al. 2016; Vesper et al. 2011). In S. enterica serovar Typherium, the Rtc system has been shown to be induced by genotoxic, metabolic and oxidative stress conditions (Kurasz et al. 2018) and a range of genetic lesions involved in DNA replication and repair (Hughes et al. 2020). Not all these conditions and genetic lesions overlap with those that activate the expression of the Rtc system in E. coli (Engl et al. 2016). Potentially the complexity of the S. enterica Rtc locus allows for the chaperoning of damaged RNA to converge stress pathways not possible in E. coli.

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Table 1 List of conditions that induce activation of the Rtc RNA repair system in the bacteria Escherichia coli and Salmonella enterica serovar Typhimurium Inducing agent Antibiotics Gentamicin Chloramphenicol Tetracycline Minocycline Rolitetracycline Ceftriaxone Enoxacin Chemicals

Cupric chloride Nickel chloride Potassium tellurite Cisplatin Hydrogen peroxide Bleomycin Methyl methanesulfonate Mitomycin C

Ribotoxins

VapC Colicin D MazF YafQ

Genetic lesions

gor ybaK yobF ftsK guaB ndk nhaA parA/B pdxB pnp polA recC

Description Inhibits protein synthesis and ribosome translocation Inhibits protein synthesis Inhibits protein synthesis (tetracycline derivative) (tetracycline derivative) Inhibits cell wall biosynthesis Blocks DNA gyrase and topoisomerase IV Redox-active metal ion Inhibits superoxide radicals Generates superoxide radicals

Bacterium E. coli

References Engl et al. (2016)

E. coli

Engl et al. (2016)

Induces nucleic acid crosslinking Oxidising agent

S. enterica

Kurasz et al. (2018)

Inhibits DNA replication Sulfhydryl-reactive agent Induces nucleic acid crosslinking tRNase that inhibits protein synthesis RNase and DNase Part of the MazF-MazE toxinantitoxin system Part of the YafQ-DinJ toxin-antitoxin system Glutathione oxidoreductase tRNA deacetylase Stress-induced peptide DNA translocase Inosine monophosphate dehydrogenase Nucleoside diphosphate kinase N+/H+ for pH homeostasis DNA gyrase/DNA primase Erythronate-4-phosphate dehydrogenate Polynucleotide phosphorylase DNA polymerase I ExoDNase

Hughes et al. (2020)

E. coli

S. enterica E. coli

S. enterica

Engl et al. (2016) Temmel et al. (2016) Kurasz et al. (2018) Engl et al. (2016) Hughes et al. (2020)

(continued)

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Table 1 (continued) Inducing agent rnhA rodZ ruvA/C sraG truA uvrD yebC

Description RNase HI

Bacterium

References

Transmembrane component of cytoskeleton Involved in Holliday junction resolution Transcriptional regulator of pnp tRNA pseudouridine synthase DNA helicase Transcriptional regulator

6 Conclusions The widely distributed Rtc system in bacteria affords research opportunities to work out how RNA damage is sensed through ligand binding to CARF domain targets. This presents opportunities to understand how CARF domain signalling is elaborated to include perhaps polynucleotide binding to CARF domains, as well as the established cyclic oligoadenylates. How the Rtc system enables stresses to be overcome requires a knowledge of which RNAs are recovered by the end modifying and end sealing activities, with some rRNAs presenting attractive areas for investigation. Whether the signals sensed by the CARF domains are themselves molecules for repair is unknown. They need not be, but would be stress-induced either directly or indirectly to couple stress to target repair. Advances in RNA sequencing methodologies can facilitate such studies. The use of live cell imaging methods to follow the locations of RNA and proteins should help establish how RNA damage is perceived and repaired on a temporal and spatial level, providing insights into the cell biology of an area of RNA biochemistry that serves many aspects of bacterial and phage physiology. Acknowledgements The work was supported by a Leverhulme Trust Research Project Grant [RPG-2019-092] awarded to MB and IK-L.

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Part II

RNA Damage and Repair in Eukaryotes

Oxidative and Nitrative RNA Modifications in Plants Jagna Chmielowska-Bąk, Karolina Izbiańska-Jankowska, Magdalena Arasimowicz-Jelonek, Joanna Deckert, and Jolanta Floryszak-Wieczorek

Abstract Nitro-oxidative modifications of biomolecules result from the presence of reactive oxygen species (ROS) and reactive nitrogen species (RNS) in the cellular environment. Unlike ROS/RNS-mediated modifications of proteins and fatty acids, nucleotides embedded in nucleic acids have not been a major focus of studies on nitro-oxidative metabolism in plants. However, RNA structure being more susceptible to oxidative/nitrative attack than DNA can be considered as an important functional biotarget of ROS/RNS generated under both physiological and stress conditions. The formation of 8-hydroxyguanine (8-OHG) and 8-nitroguanine (8-NG), the most common ROS- and RNS-mediated RNA modifications, respectively, could therefore affect protein synthesis to efficiently fine-tune various cellular responses. This chapter emphasizes the importance of 8-OHG and 8-NG not only as a symptom of ROS/RNS toxicity but implicates the nitro-oxidative RNA modifications in the regulation of multiple plant biological processes. Keywords Reactive oxygen species · Reactive nitrogen species · 8-Hydroxyguanine · 8-Nitroguanine

1 Introduction Reactive oxygen species (ROS) and reactive nitrogen species (RNS) are commonly found by-products of an oxygen- and nitrogen-rich environment. In higher plants they play a role as important signaling molecules involved in a plethora of physiological processes ranging from seed germination to senescence and cell death (e.g.,

J. Chmielowska-Bąk · K. Izbiańska-Jankowska · M. Arasimowicz-Jelonek (*) · J. Deckert Department of Plant Ecophysiology, Faculty of Biology, Adam Mickiewicz University in Poznan, Poznan, Poland e-mail: [email protected] J. Floryszak-Wieczorek Department of Plant Physiology, Poznan University of Life Sciences, Poznan, Poland © Springer Nature Switzerland AG 2021 I. Kotta-Loizou (ed.), RNA Damage and Repair, https://doi.org/10.1007/978-3-030-76571-2_4

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Fig. 1 ROS/RNS-dependent modifications of guanine nucleotides embedded in RNAs as potential regulators of posttranscriptional gene expression during plant development and plant stress responses

Considine et al. 2015). Various kinds of internal or external stimuli are able to disturb cellular ROS/RNS homeostasis, leading to the formation of oxidized and nitrated derivatives. In general, the accumulation of oxidized/nitrated macromolecules, i.e., proteins, lipids, and nucleotides, may be responsible for cell dysfunction, but it could also be important for plant defense and signaling. Unlike ROS/RNSmediated modifications of proteins and lipids, nucleotides have not been a major focus of studies on nitro-oxidative metabolism in plants. However, 8-nitroguanosine 30 ,50 -cyclic monophosphate (8-nitro-cGMP), a major nitrated cGMP derivative, has been characterized as a novel second messenger involved in abscisic acid and nitric oxide (NO)-induced stomatal closure (Joudoi et al. 2013). Nevertheless, nitrooxidative modifications of nucleotides embedded in nucleic acids are much less recognized in plants. In the case of DNA, oxidation/nitration is perceived as a symptom of ROS/RNS toxicity and cellular damage, while the role of oxidized and nitrated RNA is still under debate. It should be noted that RNA is more abundant within the cell than DNA, accounting for 80–90% of the total nucleic acid pool of the cell (Li et al. 2020). Moreover, RNA bases are not protected by hydrogen bonds, making RNA more susceptible to oxidative/nitrative attacks than DNA. Thus, RNA probably contains most of the oxidized and nitrated nucleotides under both physiological and nitrooxidative stress conditions (Castellani et al. 2008). As a result the incorporation of 8-hydroxyguanine (8-OHG) and 8-nitroguanine (8-NG) into RNA is the most common ROS- and RNS-mediated RNA modification, respectively, and could therefore play an important role in fundamental processes. According to the recent discussion on experimental data obtained in animal and plant models (ArasimowiczJelonek and Floryszak-Wieczorek 2019), the functional role of nucleotide oxidation/ nitration of transcripts may be connected with selective modifications of protein synthesis in response to endogenous and exogenous factors that function as a smart redox switch towards metabolism adjustment (Fig. 1). In confirmation, messenger (m)RNA harbors fivefold higher 8-OHG levels when compared with total

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RNA, which consists mainly of ribosomal (r)RNA and transfer (t)RNA (Simms et al. 2014).

2 Oxidative Modifications of RNA in Plants ROS are frequently called double-faced molecules. On the one hand, their overaccumulation leads to damage of biological molecules and affects cellular homeostasis. On the other hand, a certain ROS level is indispensable for proper cell functioning (Mittler 2017; Schieber and Chandal 2014). The role of the products of oxidation processes is also far from evident. For example, products of lipid peroxidation, such as malondialdehyde (MDA) and thiobarbituric acid reactive substances (TBARS), are considered markers of oxidative stress and membrane damage. In parallel oxylipins, which are derived from the oxygenation of membrane polyunsaturated fatty acids, are involved in the cellular signaling network. Similarly, short peptides formed as a result of protein oxidation may act as organelle-specific ROS sensors (reviewed in Chmielowska-Bąk et al. 2015). In the case of nucleic acids, DNA oxidation is perceived as a symptom of ROS toxicity (Evans et al. 2004), while the role of RNA oxidative modifications is still under discussion. Among numerous ROS-dependent RNA modifications, 8-OHG is the most studied. In animal models elevated 8-OHG levels are associated with various pathological conditions such as cancer, diabetes, amyotrophic lateral sclerosis (ALS), Parkinson’s and Alzheimer’s disease, as well as schizophrenia (Kong and Lin 2010; Poulsen et al. 2012). In plants the role of this oxidative modification is still relatively poorly examined, which is reflected in the limited number of publications on this topic, as listed in Table 1.

Table 1 Studies on RNA oxidative modifications (8-OHG and 8-hydroxyguanosine) in plants Plant species Sunflower (Helianthus annuus) Wheat (Triticum aestivum) Green alga (Chlamydomonas reinhardtii) Soybean (Glycine max)

Process Elevated 8-OHG levels in the transcripts of seeds subjected to the process of dry after-ripening Elevated 8-OHG levels in the transcripts of seeds subjected to the process of dry after-ripening Accumulation of 8-OHG enriched RNA in chloroplasts

References Bazin et al. (2011) Gao et al. (2013) Zhan et al. (2015)

Induction of 8-OHG formation in total RNA and mRNA in response to Cd stress

Arabidopsis (Arabidopsis thaliana)

Increased 8-OHG levels in total RNA in plants infected with cyst nematode Heterodera schachtii

ChmielowskaBąk et al. (2018) Labudda et al. (2018)

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In the case of plants, the occurrence of 8-OHG was studied for the first time in sunflower (Helianthus annuus) seeds. Bazin et al. (2011) discovered that the level of this modification is elevated in nondormant sunflower seeds when compared to dormant ones. The 8-OHG enrichment was predominant in mRNA, indicating that this RNA type is more susceptible to oxidation. It has been further evidenced that 8-OHG is found in a certain set of transcripts encoding proteins involved in cellular metabolism, stress response, and transport. Thus, the formation of this oxidative modification is not a random process. To elucidate the impact of 8-OHG enrichment in mRNA on the translation process, the research team carried out in vitro experiments on the rabbit reticulocyte lysate system. The results demonstrated that a high 8-OHG level in transcripts is associated with hampered translation and a decrease in the level of proteins (Bazin et al. 2011). The study indicates that selective oxidation of particular transcripts and the associated decrease in the level of encoded proteins play a role in the acquisition of germination potential by seeds (Bazin et al. 2011; ElMaarouf-Bouteau 2013). This hypothesis was confirmed by a subsequent study carried out on wheat (Triticum aestivum L.). The authors compared sets of transcripts enriched with 8-OHG in dormant seeds and seeds subjected to the process of after-ripening, which induces their germination potential. The results showed that after-ripening processes resulted in elevated 8-OHG levels in 80 transcripts, mainly associated with the nutrient reservoir (gliadin, glutenin, avenin) and the α-amylase inhibitor activity. Enrichment of 8-OHG in this particular set of mRNAs suggests that transcript oxidation constitutes a newly discovered mechanism involved in the regulation of storage material metabolism and energy supply during the process of germination. In turn, dormant seeds contained 40 highly oxidized mRNAs. The identified 8-OHG-enriched transcripts are involved in oxidative phosphorylation and ribosome biogenesis (Gao et al. 2013). The induction of 8-OHG formation in response to stress factors was demonstrated for the first time in soybean seedling (Glycine max). In the study seedlings were exposed to cadmium (Cd) at two concentrations of 10 and 25 mg/l. A short-term treatment with the metal at the lower concentration resulted in a significant increase in the 8-OHG level in total RNA and mRNA. In turn, common markers of oxidative stress such as lipid peroxidation and formation of abasic sites (AP sites) in mRNA were induced in response to more severe Cd stress – at higher metal concentrations (25 mg/l) and longer treatment times (24 h). The results indicate that an elevated 8-OHG level precedes the symptoms of metal-dependent cellular damage (Chmielowska-Bąk et al. 2018). An increase in the 8-OHG level has also been evidenced by biotic stress. Arabidopsis (Arabidopsis thaliana) plants infected with a cyst nematode Heterodera schachtii (Schmidt) showed an altered oxidative homeostasis including ROS overproduction, intensified lipid peroxidation, modulated activity of the antioxidant system, and 8-OHG enrichment in RNA. The level of 8-OHG was the highest after 3 days of infection, remained elevated after 5 days, and thereafter declined to the control levels. In contrast, lipid peroxidation was most intense at the later stages of infection (Labudda et al. 2018).

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The main sites of ROS production in plant cells include mitochondria, chloroplasts, peroxisomes, plasma membrane, cell wall, and endoplasmic reticulum (Das and Roychoudhury 2014). It may be suspected that these structures are involved in the generation of oxidative modifications in transcripts. In a unicellular alga Chlamydomonas reinhardtii, 8-OHG-enriched RNA has been detected in the chloroplast, particularly in the pyrenoid – a microcompartment associated with CO2 assimilation. Further research highlighted the role of the ribulose bisphosphate carboxylase-oxygenase (RuBisCO) large subunit (RBCL) in the regulation of 8-OHG metabolism. A knockout mutant with an attenuated RBLC expression contained higher 8-OHG levels. The authors suggested that RBCL acts as a bifunctional protein – a component of the RuBisCO holoenzyme and an RNA-binding protein involved in the regulation of 8-OHG levels (Zhan et al. 2015). Taken together the results of described studies show that in plants 8-OHG formation is associated with normal physiological processes (breakage of seed dormancy) and an early reaction to stress factors (nematode infection and metal stress). The studies on the unicellular alga indicate that RNA enriched in 8-OHG is accumulated in chloroplasts, although a similar localization in terrestrial multicellular plants would need confirmation. This modification occurs with the highest frequency in mRNA when compared to the other RNA types. In the process of seed dormancy breakage, 8-OHG formation is selective, limited to certain sets of transcripts. The selectivity of mRNA oxidation in response to stress factors would need experimental verification. Transcripts containing 8-OHG are translated with lower efficiency, which results in decreased levels of encoded proteins. Thus, 8-OHG formation constitutes one of the mechanisms of posttranscriptional regulation of gene expression.

3 Nitrative Modifications of RNA in Plants Nitric oxide (NO) has been described as a crucial gaseous signaling molecule with multiple functions and mechanisms of action in both plant and animal systems. Recent studies have implied that the phenomenon of protein, lipid, and nucleic acid nitration seems to represent a highly specific element of the NO-dependent signaling pathways in living organisms (Arasimowicz-Jelonek and Floryszak-Wieczorek 2019). One of the best-recognized nitrating agents in the cellular environment is peroxynitrite (ONOO¯), which is formed in the extremely rapid and diffusioncontrolled reaction of NO with superoxide (O2˙). Under physiological conditions, ONOO¯ is a highly reactive molecule, able to cross biological membranes and interact with biotargets in the surrounding cells within the radius of one or two cells (~5–20 μm) (Liaudet et al. 2009). It is well documented that ONOO¯ can covalently modify various biomolecules, significantly affecting their biochemistry (Jones 2012). In general, nitration is a chemical process leading to the introduction of a nitro group (-NO2) into a chemical compound. In the case of oligonucleotides, guanine moieties are preferentially nitrated, while adenine nitration is minor

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compared to its oxidation (Ihara et al. 2011). The reaction between ONOO¯ and guanine leads to the formation of several products, among which 8-OHG and 8-NG are the major ones. Therefore, the accumulation of 8-NG can function as a specific marker of nucleic acid nitration in the cellular environment (Ihara et al. 2011). Since nitration of guanine occurs mainly at the C8 position of the purine ring, the process can be considered as a selective one (Sodum and Fiala 2001). Guanine nitration in vivo in biological systems has been demonstrated mainly by immunoassays using anti-nitroguanine antibodies. In mammalian cells, the greater accumulation of 8-NG in DNA and RNA correlated with the formation of RNS was observed in cells under various pathological conditions. The formation of 8-NG was reported in livers of hamsters infected with a liver fluke Opisthorchis viverrini (Pinlaor et al. 2003) and in the human gastric mucosa infected with Helicobacter pylori (Ma et al. 2004). In turn, 8-NG formation in RNA was found in the pneumotropic virus infection in mice (Akaike et al. 2003) and in brain cells of mice exposed to arsenic (Piao et al. 2011). These studies have shown that 8-nitroguanine formation within the pool of nucleic acids could be a useful biomarker to evaluate the risk of infection- or inflammation-related carcinogenesis (Hiraku 2010). Although there are over a dozen studies describing the nitration of nucleic acids in animal and human systems (Murata et al. 2012), the information on the presence and functional role of nitrated nucleotides in other organisms is very limited. With regard to plants, nitrative modification of nucleic acids via ONOO¯ has been confirmed experimentally only in potato leaves inoculated with Phytophthora infestans (Izbiańska et al. 2018), in axes of embryos isolated from dormant apple seeds (Andryka-Dudek et al. 2019), and in tomato roots exposed to toxic non-proteinogenic amino acids (NPAAs) (Staszek and Gniazdowska 2020). The nitration of guanine has been detected in nucleotides embedded in total RNA (Izbiańska et al. 2018; Andryka-Dudek et al. 2019; Staszek and Gniazdowska 2020) as well as mRNA (Izbiańska et al. 2018). As noted above, total RNA and mRNA are even more susceptible to this modification than DNA, as in the case of oxidation via ROS (Liu et al. 2012). Additionally, it has been documented that compared with nitrated RNA, 8-NG in DNA is much less stable and can be spontaneously cut off from the DNA chain leaving the corresponding sites abasic instead (Ohshima et al. 2006). Looking for the functional role of ONOO¯ during the plant defense response to pathogen attack, Izbiańska et al. (2018) demonstrated for the first time that the nucleic acid phenomenon occurs in plant cells. The studies showed that only the resistant response was accompanied by a temporary limited increase of 8-NG within the total RNA and mRNA pools starting from the first hour postinoculation. This transient 8-NG accumulation was coincident with the first symptoms of programmed cell death (PCD) during hypersensitive responses (HR). Thus, the authors proposed that the nitration process in plants could be engaged in regulating the posttranscriptional gene expression and cell signaling towards active cell death. Additionally, in short-term experiments (24 h) on tomato seedlings exposed to toxic NPAA, Staszek and Gniazdowska (2020) documented that meta-tyrosine (m-Tyr) provoked an

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increasing level of 8-NG in comparison to nontreated plants; however, this effect was not observed in plants supplemented with another NPAA, canavanine (CAN). Interestingly, prolongation of seedling supplementation with both NPAAs resulted in a decreased level of nitrated guanine, which was particularly evident at the higher dose of these toxic compounds. The authors supposed that the lower level of RNA nitration at 72 h of seedling exposure to NPAAs could be associated with the conversion of 8-NG to 8-OHG in the further reaction with ONOO¯. The marker of nucleic acid nitration has also been detected in plant cells under normal conditions. For example, 8-NG was found within the pools of total RNA and mRNA in healthy leaves of various potato genotypes (Izbiańska et al. 2018). An increased level of nitrated RNA was also observed in axes of apple embryos during the transition from a dormant to a nondormant state (Andryka-Dudek et al. 2019). Based on the above, the modification of nucleotides via ONOO¯ is not only a marker of cellular dysfunction as in the case of animal and human systems, but it functions as an integral part of plant cell metabolism both in developmental and stress responses, similarly as it was proposed for oxidized mRNA.

4 Conclusions Many questions still need to be answered concerning plant oxidative and nitrative modifications, some of the most pending ones are presented in Table 2. So far, the participation of RNA oxidation/nitration in plant cell metabolism has been documented only in a few plant systems. Although oxidized/nitrated RNA may be dysfunctional and related to pathophysiological states, there is experimental evidence that the phenomenon is not a random process. To prove this assumption, the molecular mechanism(s) controlling the potential selectivity and fate of oxidized/ nitrated RNA should be essentially recognized. It can be anticipated that the nearby future will bring new exciting discoveries in the field of plant epitranscriptomics, including RNA oxidative and nitrative modifications. Table 2 Some important questions, related to RNA oxidative and nitrative modifications in plants, that require answers Is stress-dependent induction of 8-OHG and 8-NG formation a selective process? What is the mechanism of the selectivity of 8-OHG and 8-NG formation in particular transcripts? What is the destiny of 8-OHG- and 8-NG-enriched mRNAs? To what extent are oxidative/nitrative RNA modifications involved in plant growth regulation? What is the functional role of a rapid induction of 8-OHG and 8-NG formation in response to biotic and abiotic stresses? Is an increase in 8-OHG and/or 8-NG levels in transcripts a universal response to various stress factors? Could ROS/RNS-dependent modifications affect pre-mRNA splicing?

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Acknowledgements The studies concerning the 8-OHG role in plant response to stresses are financed by the National Science Centre, Poland, within the framework of project number 2019/33/ B/NZ9/00058. The studies concerning the role of 8-NG in plants are supported by the grant of the National Science Centre – project number 2017/25/B/NZ9/00905.

References Akaike T, Okamoto S, Sawa T, Yoshitake J, Tamura F, Ichimori K, Miyazaki K, Sasamoto K, Maeda H (2003) 8-nitroguanosine formation in viral pneumonia and its implication for pathogenesis. Proc Natl Acad Sci U S A 100:685–690 Andryka-Dudek P, Ciacka K, Wiśniewska A, Bogatek R, Gniazdowska A (2019) Nitric oxideinduced dormancy removal of apple embryos is linked to alterations in expression of genes encoding ABA and JA biosynthetic or transduction pathways and RNA nitration. Int J Mol Sci 20:1007 Arasimowicz-Jelonek M, Floryszak-Wieczorek J (2019) A physiological perspective on targets of nitration in NO-based signaling networks in plants. J Exp Bot 70:4379–4389 Bazin J, Langlade N, Vincourt P, Arribat S, Balzergue S, El-Maarouf-Bouteau H, Bailly C (2011) Targeted mRNA oxidation regulates sunflower seed dormancy alleviation during dry afterripening. Plant Cell 23:2196–2208 Castellani RJ, Nunomura A, Rolston RK, Moreira PI, Takeda A, Perry G, Smith MA (2008) Sublethal RNA oxidation as a mechanism for neurodegenerative disease. Int J Mol Sci 9:789–806 Chmielowska-Bąk J, Izbiańska K, Deckert J (2015) Products of lipid, protein and RNA oxidation as signals and regulators of gene expression in plants. Front Plant Sci 6:405 Chmielowska-Bąk J, Izbiańska K, Ekner-Grzyb A, Bayar M, Deckert J (2018) Cadmium stress leads to rapid increase in RNA oxidative modifications in soybean seedlings. Front Plant Sci 8:2219 Considine MJ, Sandalio LM, Foyer CH (2015) Unravelling how plants benefit from ROS and NO reactions, while resisting oxidative stress. Ann Bot 116:469–473 Das K, Roychoudhury A (2014) Reactive oxygen species (ROS) and response of antioxidants as ROS-scavengers during environmental stress in plants. Front Eviron Sci 2:53 El-Maarouf-Bouteau H, Meimoun P, Job C, Job D, Bailly C (2013) Role of protein and mRNA oxidation in seed dormancy and germination. Front Plant Sci 4 Evans M, Dizdaroglu M, Cooke MS (2004) Oxidative DNA damage and disease: induction, repair and significance. Mut Res 567:1–61 Gao F, Rampitsch C, Chitnis VR, Humphreys GD, Jordan MC, Ayele BT (2013) Integrated analysis of seed proteome and mRNA oxidation reveals distinct post-transcriptional features regulating dormancy in wheat (Triticum aestivum L.). Plant Biotech J 11:921–932 Hiraku Y (2010) Formation of 8-nitroguanine, a nitrative DNA lesion, in inflammation-related carcinogenesis and its significance. Environ Health Prev Med 15:63–72 Ihara H, Sawa T, Nakabeppu Y, Akaike T (2011) Nucleotides function as endogenous chemical sensors for oxidative stress signaling. J Clin Biochem Nutr 48:33–39 Izbiańska K, Floryszak-Wieczorek J, Gajewska J, Meller B, Kuźnicki D, Arasimowicz-Jelonek M (2018) RNA and mRNA nitration as a novel metabolic link in potato immune response to Phytophthora infestans. Front Plant Sci 9:672 Jones LH (2012) Chemistry and biology of biomolecule nitration. Chem Biol 19:1086–1092 Joudoi T, Shichiri Y, Kamizono N, Akaike T, Sawa T, Yoshitake J, Yamada N, Iwai S (2013) Nitrated cyclic GMP modulates guard cell signaling in Arabidopsis. Plant Cell 25:558–571 Kong Q, Lin CG (2010) Oxidative damage to RNA: mechanism, consequences, and diseases. Cell Moll Life Sci 67:1817–1829

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Labudda M, Różańska E, Czarnocka W, Sobczak M, Dzik JM (2018) Systemic changes in photosynthesis and reactive oxygen species homeostasis in shoots of Arabidopsis thaliana infected with the beet cys nematode Heterodera schachtii. Mol Plant Pathol 19:1690–1704 Li Z, Chen X, Liu Z, Ye W, Li L, Qian L, Ding H, Li P, Aung LHH (2020) Recent advances: molecular mechanism of RNA oxidation and its role in various diseases. Front Mol Biosci 7:184 Liaudet L, Vassalli G, Pacher P (2009) Role of peroxynitrite in the redox regulation of cell signal transduction pathways. Front Biosci 14:4809–4814 Liu M, Gong X, Alluri RK, Wu J, Sablo T, Li Z (2012) Characterization of RNA damage under oxidative stress in Escherichia coli. Biol Chem 393:123–132 Ma N, Adachi Y, Hiraku Y, Horiki N, Horiike S, Imoto I, Pinlaor S, Murata M, Semba R, Kawanishi S (2004) Accumulation of 8-nitroguanine in human gastric epithelium induced by Helicobacter pylori infection. Biochem Biophys Res Commun 319:506–510 Mittler R (2017) ROS are good. Trend Plant Sci 22:11–19 Murata M, Thanan R, Ma N, Kawanishi S (2012) Role of nitrative and oxidative DNA damage in inflammation-related carcinogenesis. J Biomed Biotechnol 2012:1–11 Ohshima H, Sawa T, Akaike T (2006) 8-nitroguanine, a product of nitrative DNA damage caused by reactive nitrogen species: formation, occurrence, and implications in inflammation and carcinogenesis. Antioxid Redox Signal 8:1033–1045 Piao F, Li S, Li Q, Ye J, Liu S (2011) Abnormal expression of 8-nitroguanine in the brain of mice exposed to arsenic subchronically. Ind Health 49:151–157 Pinlaor S, Yongvanit P, Hiraku Y, Ma N, Semba R, Oikawa S, Murata M, Sripa B, Sithithaworn P, Kawanishi S (2003) 8-nitroguanine formation in the liver of hamsters infected with Opisthorchis viverrini. Biochem Biophys Res Commun 309:567–571 Poulsen HE, Specht E, Broedbaek K, Henriksen T, Ellervik C, Mandrup-Poulsen T, Tonnesen M, Nielsen PE, Andersen HU, Weimann A (2012) RNA modifications by oxidation: a novel disease mechanism? Free Radic Biol Med 52:1353–1361 Schieber M, Chandel NS (2014). ROS function in redox signaling and review oxidative stress. Cur Biol 24:R453–R462 Simms CL, Hudson BH, Mosior JW, Rangwala AS, Zaher HS (2014) An active role for the ribosome in determining the fate of oxidized mRNA. Cell Rep 9:1256–1264 Sodum R, Fiala ES (2001) Analysis of peroxynitrite reactions with guanine, xanthine, and adenine nucleosides by high-pressure liquid chromatography with electrochemical detection: C8-nitration and -oxidation. Chem Res Toxicol 14:438–450 Staszek P, Gniazdowska A (2020) Peroxynitrite induced signaling pathways in plant response to non-proteinogenic amino acids. Planta 252:5 Zhan Y, Dhaliwal JS, Adjibade P, Uniacke J, Mazroui R, Zerges W (2015) Localized control of oxidized RNA. J Cell Sci 128:4210–4219

The Role of Ribonucleases in RNA Damage, Inactivation and Degradation Fabian Hia and Osamu Takeuchi

Abstract Nucleic acids are universally present in all forms of life on earth. Since their discovery more than 150 years ago, knowledge on the roles of nucleic acids, both DNA and RNA, has been gradually evolving. RNA, the less stable of the two, is a molecule capable of a wide range of functions, such as transmission of information, catalysis and regulation of gene expression among many others. The prevalence and diverse functions of RNA underlie the need for its regulation. As such, cells possess an arsenal of regulatory tools which modulate RNA. In this review, we focus on the role of ribonucleases acting specifically against damaged host RNA, pro-inflammatory messenger RNA and foreign RNA as part of defence mechanisms employed by mammalian cells against invading viruses. Keywords Ribonucleases · RNA damage · RNA degradation · Ribosome quality control · No-go-decay · Immune response · RNA-binding proteins · Antiviral response

1 Introduction Since the discovery of nucleic acids in 1869 by Johann Friedrich Miescher (Veigl et al. 2020), much progress has been made in unravelling the role of nucleic acids in biological systems. In particular, RNA since its discovery has been shown to act as more than just an informational template for the synthesis of proteins. Because of their multifaceted forms and roles, from ribosomal RNA (rRNA) to transfer RNA (tRNA) and non-coding RNA (ncRNA) among many others, RNA is present in all forms of life. As such, the regulation of RNA is a tightly controlled process with complex yet intricate mechanisms which process, edit, degrade and stabilize RNA, in order to maintain the physiological well-being of cells (Hia and Takeuchi 2020; F. Hia · O. Takeuchi (*) Department of Medical Chemistry, Graduate School of Medicine, Kyoto University, Kyoto, Japan e-mail: [email protected] © Springer Nature Switzerland AG 2021 I. Kotta-Loizou (ed.), RNA Damage and Repair, https://doi.org/10.1007/978-3-030-76571-2_5

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Teoh et al. 2020; Mino et al. 2019; Yang et al. 2020a). Importantly, the universality of RNA has also resulted in the evolution of biological defence mechanisms by cells that target and destroy non-host, exogenous RNA in the form of invading pathogens (Burroughs and Aravind 2016). While damage to host RNA has been traditionally exemplified through oxidative and alkylating reactions, RNA destruction also arises through targeted degradation via enzymatic effectors due to the need for physiological homeostasis, as well as from conflicts between biological systems. In these two cases, both exogenous and endogenous RNA are subjected to damage and subsequent degradation. In general, RNA damage can be defined as insults resulting in RNA being altered to non-functional or aberrant forms, inactivated or destroyed such that it (1) is unable to be fully translated, (2) loses its catalytic activity and (3) loses its ability to basepair or interact with other RNA or proteins. In this review, we would like to further discuss RNA damage and degradation through the lens of ribonucleases; ribonuclease activity results in cleaved RNA that is no longer functional. Ribonucleases are involved in the destruction of a wide variety of RNA molecules, from both endogenous sources, such as damaged RNA and pro-inflammatory messenger RNA (mRNA), as well as from exogenous sources in the form of viral RNA. Consequently, we have divided the review into two parts. The first discusses the role of ribonucleases in eliminating damaged RNA, mainly through ribosome quality control and its associated processes. In the second part, we discuss RNA damage with a focus on strategies employed by mammalian cells to target and inactivate host and viral RNA as part of the immune response.

2 Quality Control Mechanisms to Manage Damaged RNA Nucleic acids are constantly subjected to a variety of insults by the environment. While the insults upon DNA and their consequences have been extensively investigated extent, we have only begun to appreciate the mechanisms and effects of these insults on RNA. For a long time, the assumed transiency of RNA suggested that damage to RNA would not have significant impact on the viability of a cell. However, in the past two decades, RNA damage has been subjected to increasing scrutiny with many studies and reviews detailing the mechanisms by which RNA is damaged and repaired, as well as the consequences and diseases associated with RNA damage (Fimognari 2015; Nunomura et al. 2012a, b; Ding et al. 2012; Yan and Zaher 2019; Wurtmann and Wolin 2009; Simms and Zaher 2016). It is not surprising that a growing body of evidence suggests that quality control mechanisms are present in order to cope with damaged RNA (Simms and Zaher 2016; Hayakawa et al. 2002, 2010; Ishii et al. 2015, 2018, 2020; Simms et al. 2014). One of the common insults on RNA originates from endogenous reactive oxygen species (ROS). ROS are generated from a variety of sources, which include several steps in the electron transport chain (ETC) and phagocyte activity among others (Fimognari 2015; Yan and Zaher 2019; Wurtmann and Wolin 2009). The reaction of

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oxidative species such as •OH with ribonucleic acids results in the production of several distinct products such as 8-oxo-7,8-dihydroguanine (8-oxoG) and 5-hydroxycytidine (Weidner et al. 2011). In addition to oxidizing agents, alkylating compounds can react with oxygen and nitrogen atoms which comprise RNA, forming RNA adducts such as O6-alkylguanosine (O6-mG), O6-alkylthymidine (O6-mT), N1-methyladenosine (m1A) and N3-methylcytidine (m3C) (Yan and Zaher 2019; Wurtmann and Wolin 2009). Such modifications are inherently toxic to cells, inducing point mutations and slowing translation (Yan and Zaher 2019; Shan et al. 2007; Tanaka et al. 2007). To counter this, cells have evolved several mechanisms to resolve stalled ribosomes and dispose of damaged RNA, two of which have been gaining prominence recently—ribosome quality control (RQC) and no-go-decay (NGD). NGD in particular requires the nuclease activity of several ribonucleases which will be discussed in the following sections.

3 Ribosome Quality Control (RQC) and No-Go-Decay (NGD) RQC is a co-translational process aimed at resolving stalled ribosomes and destroying the nascent peptide, while NGD is a quality control system intended at destroying transcripts which cause ribosomal stalling (Fig. 1a). Both pathways are initiated following ribosome stalling and share common factors. Ribosomes have been shown to stall upon encountering RNA secondary structures, stretches of positively charged amino acid residues, poly(A) sequences, as well as nonoptimal or rare codons (Letzring et al. 2013; Hia et al. 2019; Han et al. 2020; Doma and Parker 2006; Juszkiewicz and Hegde 2017; Sundaramoorthy et al. 2017). As a single mRNA transcript can be translated concurrently by many ribosomes, a stalling event in the leading ribosome will result in a collision with the following ribosome: the two collided ribosomes are termed di-ribosomes or disomes (Han et al. 2020; Meydan and Guydosh 2020; Ikeuchi et al. 2019). In both yeast and humans, the disomes are recognized by special E3 ubiquitin ligases. Hel2, the yeast E3 ubiquitin ligase ubiquitinates ribosomal protein uS10, while the human ubiquitin ligase ZNF598 catalyzes ubiquitination of uS10 and eS10 (Juszkiewicz and Hegde 2017; Garshott et al. 2020; Matsuo et al. 2017). Concurrently, the NGD system is activated, recruiting endonucleases to cleave the problematic transcript. In contrast to regular mRNA degradation, endonucleolytic cuts are made proximal to the stalled ribosomes to produce two types of mRNA fragments—a 5’ NGD fragment and a 3’ NGD fragment (Doma and Parker 2006). The 5’ NGD fragment lacks a poly(A) tail and is subsequently degraded by the exosome complex in the cytoplasm, while the 3’ NGD fragment, lacking a cap structure, is degraded by the exonuclease Xrn1 (Doma and Parker 2006; Tsuboi et al. 2012). The ubiquitinated stalled ribosomes are then dissociated.

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Fig. 1 RQC and NGD are quality control mechanisms which deal with damaged RNA. (a) Stalled ribosomes are subject to collisions by trailing ribosomes resulting in the formation of disomes. The formation of disomes triggers RQC and NGD pathways which result in the dissociation of the ribosome, degradation of the nascent peptide and destruction of the defective transcript. The NGD pathway is illustrated in this figure. Domain architecture of (b) a potential NGD endonuclease, Cue2 and (c) a highly conserved 50 -30 exoribonuclease, Xrn1

While the exact details regarding ribosome dissociation are still unclear, studies have shown that Dom34 (Pelota in humans), in association with Hbs1 (Dom34: Hbs1), and the RQC trigger (RQT) complex have the ability to result in ribosome dissociation, respectively (Tsuboi et al. 2012; D’Orazio et al. 2019). Dom34:Hbs1

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has been previously shown in several studies to be involved in the non-stop decay mRNA surveillance pathway and can facilitate the dissociation of ribosomes (Graille and Seraphin 2012). Alternatively, the RQT complex, consisting of Shl1/Rqt2, Cue3/Rqt3 and Rqt4 (ASCC3, ASCC2 and ASCC1 in humans), also induces the dissociation of the ribosome subunits (Matsuo et al. 2017; Inada 2020; Hashimoto et al. 2020). It has been suggested that both pathways can act cooperatively, with the Dom34:Hbs1-mediated dissociation important for resolving ribosomes stalled at the 30 end of mRNAs or 5’ NGD fragments (Tsuboi et al. 2012; D’Orazio et al. 2019; Zinoviev et al. 2020). Listerin (Ltn1) then ubiquitinates the nascent chain targeting the protein arrest product for degradation by the proteasome (Brandman et al. 2012; Shao et al. 2013). For example, 8-oxoG has been demonstrated to have an effect on translation and stalls the translation machinery by reducing the rate of peptide formation due to an impediment in codon-anticodon interactions (Simms et al. 2014). Interestingly, in this study by Simms et al. (2014), 8-oxoG containing mRNA was stabilized in Dom34 mutant yeast, suggesting that 8-oxoG-modified mRNA is subjected to RQC and NGD. Indeed, Yan et al. demonstrated that the addition of oxidative and alkylating agents which damage RNA triggers RQC and NGD (Yan et al. 2019). In the following section, we will discuss Xrn1and the exosome complex in more detail.

4 Xrn1, a 50 –30 Exoribonuclease, and the Exosome, a 30 -50 Exoribonuclease Complex Regular mRNA turnover and surveillance is mediated via two main pathways; the 50 –30 and 30 – 50 decay pathways controlled by Xrn1 and the exosome complex, respectively (Langeberg et al. 2020; Hsu and Stevens 1993; Parker 2012). Xrn1 is a 175 kDa highly conserved cytosolic exoribonuclease responsible for the degradation of deadenylated-decapped substrates and intermediates from mRNA surveillance pathways, such as non-stop decay (NSD) and nonsense-mediated decay (NMD) (Coller and Parker 2004; Parker and Song 2004; Houseley and Tollervey 2009; Shoemaker and Green 2012). Structural studies of Xrn1 homologs in Kluyveromyces lactis and Drosophila melanogaster show that Xrn1 possesses a highly conserved core composed of two highly conserved regions at the N-terminus (Fig. 1c), which likely coordinate ion binding for catalysis (Jinek et al. 2011; Chang et al. 2011). In general, cytoplasmic mRNA degradation pathways are initiated by deadenylation, which involves the removal of the poly(A) tail by the Ccr4-Not and Pan2-Pan3 complexes (Coller and Parker 2004; Siwaszek et al. 2014). This is then followed by the removal of the 5’cap structure by the Dcp2 nudix hydrolase enzyme in a process termed as decapping (Coller and Parker 2004). The decapped monophosphorylated

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mRNA is then subjected to degradation by Xrn1 (Coller and Parker 2004). Several studies have shown that these factors act cooperatively, to couple deadenylation, decapping and degradation for efficient mRNA turnover (Chang et al. 2019; Nissan et al. 2010; Braun et al. 2012). In NGD, deadenylation and decapping is bypassed, since the 3’ NGD intermediate, as a result of endonucleolytic cleavage, possesses an exposed 50 end which can be directly targeted by Xrn1 (Tsuboi et al. 2012; Navickas et al. 2020). In contrast, the 50 fragment is truncated at the 30 end, does not possess a poly(A) tail and is subsequently degraded by the exosome complex (Tsuboi et al. 2012). The exosome is a highly conserved 30 -50 exoribonuclease complex localized in both the nucleus and the cytoplasm (Schmid and Jensen 2008; Weick and Lima 2020). Numerous subunits of the exosome are bound to its central core structure, forming a macromolecular complex responsible for the degradation and processing of myriad cellular RNAs (Schmid and Jensen 2008; Weick and Lima 2020). As such, the exosome is involved in a wide variety of cellular processes, from the maturation of functional mRNAs to the degradation of aberrant RNA molecules (Dziembowski et al. 2007; Zinder and Lima 2017; Allmang et al. 1999, 2000). In eukaryotes, while the core of the exosome complex appears to be inactive by itself, it functions in tandem with its subunits in the nucleus and cytoplasm to elicit the degradative abilities of the complexes (Dziembowski et al. 2007; Thoms et al. 2015; de la Cruz et al. 1998; Lykke-Andersen et al. 2009). In humans, the exosome core is a ring structure comprised of three different heterodimers, hRrp41-hRrp45, hRrp46hRrp43 and hMtr3-hRrp42, as well as three additional proteins, Rrp4, Csl4 and Rrp40, which stabilize the entire core complex (Shen and Kiledjian 2006; Liu et al. 2006). In particular, two exonucleases Dis3 (hDis3) and Rrp6 (EXOSC1) (Schmid and Jensen 2008; Zinder and Lima 2017; Robinson et al. 2015; Davidson et al. 2019) are associated with the core of the exosome, providing it with its degradative capability. Interestingly, the precise composition of the exosome complex is dependent on its cellular localization and function. Additional co-factors can readily associate with the exosome leading to a specific function; these co-factors are often spatially restricted to either the nucleus or the cytoplasm (Schmid and Jensen 2008; Schaeffer and van Hoof 2011). In the cytoplasm, the exosome associates with the Ski complex, a helicase complex which comprises proteins Ski2, Ski3 and Ski8, and together they mediate 30 –5’ mRNA decay (Schmidt et al. 2016). For instance, 5’ NGD intermediates without a poly(A) tail are degraded in this manner (Doma and Parker 2006; Shoemaker and Green 2012; Anderson and Parker 1998). Interestingly, Zinoviev et al. (2020) demonstrated that this cytoplasmic exosome is able to bind 80S ribosomes to release mRNA for degradation, suggesting that in addition to its established role in RNA clearance, the exosome can aid in resolving stalled ribosomes together with the Dom34:Hbs1 complex. In the following section, we will discuss the identity and roles of a potential NGD endonuclease in addition to Xrn1.

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5 Cue2 as a Potential NGD Endonuclease The activity of at least one endonuclease is required to produce NGD fragments in order for Xrn1 and the exosome complex to act on the exposed ends of their respective target mRNA fragments. The identity of the endoribonuclease was a mystery for some time since the discovery of the RQC and NGD pathways until recently. D’Orazio et al. (2019), using a reverse genetic screen in yeast, identified Cue2 as an endonuclease recruited to cleave the mRNA at the A-site of the colliding ribosome. Interestingly, while Xrn1 is the dominant factor in mRNA degradation, the inhibition of the Slh1 helicase activates the Cue2-mediated cleavage, suggesting that Cue2 is able to provide an alternative solution for the degradation of problematic transcripts (D’Orazio et al. 2019). One of the more compelling pieces of evidence implicating Cue2 as a NGD endonuclease lies in its domain structure (Fig. 1b). A 443 amino acid long protein, Cue2, possesses two ‘coupling of Ubiquitin conjugation to ER degradation’ (CUE) domains at its N-terminus. This is followed by two more putative ubiquitin-binding domains, an ‘ubiquitin-associated domain’ (UBA) and another putative CUE domain (D’Orazio et al. 2019). The presence of these ubiquitin-binding domains is crucial for RQC and NGD, suggesting that Cue2 binds to ubiquitinated collided ribosomes to execute its function. Additionally, Cue2 possesses a small MutSrelated (SM) hydrolase domain at the C-terminus, which has been shown to exhibit endonuclease activity in plants (Zhou et al. 2017). Glover et al. (2020), using Caenorhabditis elegans, illustrated that NONU-1, a homolog of Cue2, was required for the production of mRNA cleavage fragments similar to the NGD fragments observed in yeast. Similarly to Cue2, NONU-1 contains two CUE domains that allow binding of the protein to ubiquitin (and therefore ubiquitinated ribosomes), as well as an SMR domain which is required for NGD (Glover et al. 2020). Indeed, further investigations and biochemical analyses will shed light on the role of Cue2 as a NGD endonuclease. In the later sections, we explain how the degradative ability of the exosome is used by the immune system to degrade foreign RNA.

6 The Role of Ribonucleases in Targeting and Inactivating Host and Foreign RNA Thus far, we have described the role of ribonucleases in quality control of mRNA in a co-translational manner. Here, we now describe how ribonucleases specifically modulate the mammalian immune system in overcoming infections via a two-pronged strategy: (1) targeting host inflammatory mRNAs post-translationally and (2) directly attacking and damaging exogenous viral RNA. While ribonucleases involved in regulating the immune system are numerous, in this review, we concentrate on a select few and their roles in targeting either host transcripts, viral mRNAs or both.

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7 Regulatory RNase 1 (Regnase-1) Regulatory RNase 1 (Regnase-1), also known as Zc3h12a and Mcpip1, is an endoribonuclease encoded by the ZC3H12A gene and is essential for the regulation of the immune response, iron metabolism and degradation of viral RNA (Mino et al. 2019; Yoshinaga et al. 2017; Uehata et al. 2013; Cui et al. 2017; Nakatsuka et al. 2018, 2020; Matsushita et al. 2009). Consistent with its role as a modulator of the inflammatory response, we showed that mice which lack Regnase-1 (Regnase1/) possess increased cytokine and serum immunoglobulin levels, have heightened autoantibody production indicative of inflammatory autoimmune disease and die within 12 weeks after birth (Matsushita et al. 2009). Furthermore, these mice develop severe lymphadenopathy and splenomegaly and have increased levels of effector/memory T cells and plasma cell counts (Matsushita et al. 2009). Given the systemic severity of the diseases observed in murine models, the role of Regnase-1 in regulating pro-inflammatory cytokine levels is paramount. In this section, we discuss the role and mechanisms of Regnase-1 in degrading host cytokine mRNAs and its antiviral role in cleaving viral RNAs. Regnase-1 is part of the Cys-Cys-Cys-His (CCCH) zinc finger protein family (Liang et al. 2008). Notable CCCH-zinc finger proteins (e.g. Roquin and tristetraprolin) have been shown to be intimately involved in mRNA metabolism, specifically in the degradation of cytokine mRNAs (Maeda and Akira 2017). Furthermore, Regnase-1 sequence alignment revealed the presence of a ribonuclease domain known as N4BP1YacP nuclease (NYN) domain (part of the Pi1T N-terminus [PIN] domain superfamily) at the N-terminus (Fig. 2a), forming a negatively charged pocket eventually shown to be a functional Mg2+-dependent RNase (Matsushita et al. 2009). Using selective 20 -hydroxyl acylation and primer extension (SHAPE) analysis, together with computational prediction of Regnase-1 targets, we showed that Regnase-1 recognizes stem-loop structures which contain a pyrimidine-purinepyrimidine hairpin sequence and targets the mRNA for degradation (Mino et al. 2015). Interestingly, Regnase-1-mediated mRNA decay of immune-related transcripts requires the assistance of the helicase UPF-1 (Mino et al. 2015). Depletion of UPF1 in bone marrow-derived macrophages (BMDMs) resulted in an increase of Regnase-1 pro-inflammatory targets following lipopolysaccharide (LPS) stimulation (Mino et al. 2015). Further UPF1 knockdown and subsequent reconstitution experiments in HeLa cells revealed that the UPF1 helicase activity is essential for Regnase-1-mediated mRNA decay of inflammatory-related transcripts possessing specific stem-loop structures, in a manner similar to the degradation of nonsense mRNAs (Mino et al. 2015). We thus postulated that UPF1 allows Regnase-1 to degrade inflammatory mRNAs. In a further investigation to establish the sequential dynamics of Regnase-1mediated degradation, it was revealed that Regnase-1 initially binds to the stemloop structure on the 30 untranslated region (UTR) of inflammatory mRNAs (Mino et al. 2019). Without the unwinding by the UPF1 helicase, Regnase-1 is unable to

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CCCH-type zinc finger domain PIN

c PIN

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DSGxxS motif

KH domains forr binding ssRNA

PIN (NYN) domain

b Regnase-1 recognizes and binds stem-loop

Phosphorylated UPF1 binds Regnase-1 and unwinds stem-loop

Regnase-1 cleaves mRNA A at stem-loop

Fig. 2 NYN domain containing ribonucleases can target host and foreign RNA. (a) Regnase-1 contains a CCCH-type zinc finger domain, which can bind RNA, and a NYN domain (part of the PIN superfamily), which cleaves RNA endonucleolytically. (b) UPF1 enables Regnase-1 to cleave stem-loop structures of RNA. (c) Similarly to Regnase-1, N4BP1 contains a NYN domain which cleaves RNA. N4BP1 also possesses two KH domains which can bind single-stranded RNA

cleave the RNA (Mino et al. 2019). After phosphorylation at the T28 position by a PI3K-related protein kinase, SMG1, two points of interaction occur between UPF1 and Regnase-1 that promote the helicase unwinding activity of UPF1; the RNase domain of Regnase-1 binds to the SMG1-phosphorylated residue T28 in UPF1, while an intrinsically disordered segment in Regnase-1 binds to the RecA domain of UPF1 (Mino et al. 2019). Following unwinding of the stem-loop, Regnase-1 can then endonucleolytically cleave the RNA. Interestingly, stem-loop mutants, which possess short, two nucleotide stem-loops, could be cleaved by Regnase-1 without the aid of UPF1 (Mino et al. 2019). Regnase-1 without UPF1 was also capable of cleaving pre-unwound stem-loop RNAs (Mino et al. 2019). Indeed, Regnase-1 has been shown to be capable of cleaving unstructured single-stranded RNA (ssRNA) independent of nucleotide sequence in vitro (Wilamowski et al. 2018). These results suggest that stem-loop structures, while critical for Regnase-1 recognition can stop Regnase-1-mediated degradation or target RNAs, which can only occur after UPF1 unwinds the stem-loop (Fig. 2b). Regnase-1 has also been purported to possess antiviral activity, acting as a restriction factor in human immunodeficiency virus (HIV), hepatitis C virus (HCV) and coxsackievirus B3 (CV-B3) infections among others (Lin et al. 2013, 2014; Li et al. 2018; Qian et al. 2019). Liu et al. (2013) showed that through ectopic

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expression of Regnase-1 in cell lines, HIV-1 virus production is significantly decreased. Conversely, siRNA-mediated silencing of Regnase-1 results in higher viral titres (Liu et al. 2013). Subsequent infection experiments utilizing a human T-cell line, CEM-SS, also demonstrated that ectopic expression of Regnase-1 inhibits HIV-1 production (Liu et al. 2013). Northern blot analysis of HIV-1-infected human T-cell lines revealed a significant decrease in detectable HIV-1 RNA levels following expression of the monocyte chemotactic protein 1-induced protein 1 (MCPIP1) (Liu et al. 2013). Co-transfection studies showed that both human and monkey Regnase-1 could similarly restrict simian immunodeficiency virus (SIV) production (Li and Wang 2016). While the suppression of virus RNA levels and production are noteworthy in these two studies, whether Regnase-1 can directly bind and cleave HIV-1 remains unclear. It would be interesting to ascertain and test the efficacy of Regnase-1 on potential stem-loops within the HIV RNA sequences. In another study by Lin et al. (2014), Regnase-1 was shown to suppress replication of HCV. Similarly, the authors utilized an in vitro cleavage assay to demonstrate that Regnase-1 could target a previously known conserved stem-loop structure in the HCV 3’ UTR for degradation (Lin et al. 2014; Tanaka et al. 1996). The results concluded that in addition to modulating the pro-inflammatory response, Regnase-1 can directly target HCV RNA for degradation (Lin et al. 2014). Several other viral infection models have been used to illustrate the antiviral effects of Regnase-1, including infection by dengue virus (DEN), Japanese encephalomyocarditis virus (JEV), sindbis virus (SINV), influenza virus and adenovirus (Lin et al. 2013). Regnase-1 is able to reduce hepatitis B (HBV) viral load in hepatocytes by targeting HBV stem-loop structures (Li et al. 2020). Furthermore, in vitro cleavage assays demonstrated that Regnase-1 could cleave DEN and JEV RNA to prevent further replication of these viruses.

8 NEDD4-Binding Protein 1 (N4BP1) NEDD4-binding protein 1, encoded by the N4BP1 gene, was originally identified as a target of NEDD4, an E3 ubiquitin ligase (Sharma et al. 2010; Murillas et al. 2002). Initial studies revealed that following poly-ubiquitination, N4BP1 is targeted for degradation by the proteasome in promyelocytic leukaemia (PML) bodies (Sharma et al. 2010). Interestingly, N4BP1 is subjected to sumoylation and de-sumoylation, with the former competing with ubiquitination to confer stability to N4BP1 and the latter being a prerequisite step for its proteasome-mediated turnover (Sharma et al. 2010). The domain architecture of N4BP1 defines its degradative abilities (Fig. 2c). N4BP1 possesses two K homology (KH) domains, well-known for their singlestranded nucleic acid binding ability (Grishin 2001). In addition, similarly to Regnase-1, N4BP1 possess a highly conserved NYN nuclease domain (Sharma et al. 2010; Yamasoba et al. 2019; Anantharaman and Aravind 2006), previously reported to possess ribonuclease activity, implicating N4BP1 as a ribonuclease (Matsushita et al. 2009).

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Because of its localization in subnuclear compartments as well as the presence of nucleic acid binding and cleavage domains, N4BP1 was originally believed to have a functional role in nucleolar RNA processing (Sharma et al. 2010). However, emerging studies have revealed N4BP1 to additionally be capable of regulating tumour progression, modulating innate immune signalling and restricting viral replication (Yamasoba et al. 2019; Oberst et al. 2007; Gitlin et al. 2020; Spel et al. 2018; Nchioua et al. 2020). In this section of the review, we would like to focus on the role of N4BP1 as a ribonuclease in destroying viral RNA and containing excess pro-inflammatory transcripts. Gitlin et al. (2020) showed that N4BP1 is able to suppress a subset of cytokines and chemokines produced by the stimulation of TRIF-independent Toll-like receptors (TLRs). Conversely, N4BP1 is regulated by caspase-8 cleavage, which modulates excessive cytokine responses. Furthermore, it was demonstrated that tumour necrosis factor (TNF) production, as elicited by pathogens such as Streptococcus pneumoniae, induces caspase-8 cleavage of N4BP1, preventing N4BP1-mediated inhibition of immune responses to ensure a robust cytokine response by TRIFindependent TLRs (Gitlin et al. 2020). This study, in particular, exemplifies the role of N4BP1 as a ribonuclease checkpoint modulator in regulating immune responses by invading pathogens. N4BP1 also possesses a much more direct role in immune responses, having been implicated as another restriction factor in HIV infections. Our recent study (Yamasoba et al. 2019) demonstrated that N4BP1 which is strongly induced by type I interferons (IFNs), is a potent inhibitor of HIV-1 in both primary T cells and macrophages by restricting HIV-1 replication (Yamasoba et al. 2019). In particular, it was shown by RNA immunoprecipitation (RIP) assay that N4BP1 directly recognizes and binds to both spliced and unspliced HIV mRNA species (Yamasoba et al. 2019). Importantly, the role of N4BP1in suppressing HIV-1 is a potential contributing factor to the maintenance of HIV latency, the loss of N4BP1 resulting in reactivation of HIV (Yamasoba et al. 2019). Given the localization of N4BP1 in PML bodies (Sharma et al. 2010) together with the role of PML bodies in sequestering transcription activators (Lusic et al. 2013; Van Damme et al. 2010), it is possible that one potential role for N4BP1 is the degradation of HIV mRNA and the suppression of viral reactivation.

9 Zinc Finger Antiviral Protein (ZAP) and Co-Factor Nucleases The role of nucleases which can directly recognize and cleave target nucleic acids has been described and discussed above, and it is now apposite to discuss the role of several nucleases which can be recruited by accessory proteins to their targets. Initially, how these nucleases gain further specificity through the role of an accessory protein, zinc finger antiviral protein (ZAP), will be addressed.

96 Fig. 3 Certain nucleases require co-factor ZAP to direct them to their targets. (a) Schematic diagram of two isoforms of ZAP, ZAP-S and ZAP-L, both of which contain CCCH-type zinc finger domains. The longer isoform, ZAP-L, has a PARP-like domain. (b) KHNYN requires its KH-like and NYN domains to cleavage viral RNAs. However, this antiviral function is maximized when it is recruited by ZAP to viral RNAs

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a ZAP-S

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PARP PARP-like domain

KH-like

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KH-like domain

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While not traditionally known to be a functional nuclease, ZAP is capable of recruiting several nucleases to nucleic acid targets. As such, ZAP together with its associated nucleases is capable of restricting a broad range of viruses, from RNA viruses such as retroviruses to DNA viruses (Mao et al. 2013; Müller et al. 2007; Zhu et al. 2011; Li et al. 2015; Goodier et al. 2015). ZAP, also known as PARP13, belongs to the poly(ADP-ribose) polymerase (PARP) family, which is known to play roles in many cellular processes including transcription, replication and DNA repair among others (Morales et al. 2014; Meagher et al. 2019). ZAP is localized to the cytoplasm and possesses several isoforms (Li et al. 2019). Of these, two isoforms ZAP-Long (ZAP-L) and ZAP-Short (ZAP-S) have been well characterized (Fig. 3a). Both ZAP isoforms possess N-terminal RNA-binding domains consisting of 4 CCCH zinc finger domains (Meagher et al. 2019). Additionally, ZAP-L is a representative of the PARP family in that it possesses a C-terminal PARP-like domain. Interestingly, the ZAP-L PARP-like domain lacks the His-Tyr-Glu (H-YE) triad required for ADP-ribosylating activity (Kleine et al. 2008; Alemasova and Lavrik 2019). Conversely, ZAP-S lacks the PARP-like domain but is instead induced to a greater extent by IFNs (Li et al. 2019). Vertebrate genomes are known to exhibit CpG suppression, possessing fewer CpG-dinucleotides than expected (Karlin and Mrázek 1997). In a study aimed to identify how CpG suppression in HIV-1 affects its replication, Takata et al. (2017) showed that synonymous mutagenesis of HIV-1 to increase viral CpG-dinucleotides resulted in a significant inhibition of virus replication. Through a focused siRNA screen, the authors identified ZAP as a restriction factor that can bind to and restrict CpG-dinucleotide-enriched viral sequences (Takata et al. 2017). ZAP-L and ZAP-S possess distinct functions, with ZAP-L acting as a viral degradation factor, while ZAP-S acts as an immune function modulator (Schwerk et al. 2019). Schwerk et al.

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(2019) showed that ZAP-S, not ZAP-L, localizes to the cytoplasm, binding to the 3’UTR of host interferon mRNA to control expression. Conversely, ZAP-L is localized to the endolysosomes and plasma membranes, targeting viral replication during SINV infection (Schwerk et al. 2019). Not possessing any form of nuclease activity itself, ZAP exerts its antiviral function through the recruitment of nucleases. In the following section, the role of these nucleases, namely, KH and NYN domain containing (KHNYN) enzymes, as well as the previously mentioned exosome complex is reviewed.

10

ZAP and KHNYN

KHNYN was recently identified as a ZAP interacting factor through a yeast two-hybrid screen for both ZAP-L and ZAP-S (Ficarelli et al. 2019). Here Ficarelli et al. (2019) showed that KHNYN, together with a co-factor TRIM25, interacts with ZAP to inhibit a CpG-enriched version of HIV-1. Further studies revealed that both KH and NYN domains were required for KHNYN antiviral endonuclease activity (Ficarelli et al. 2019). KHNYN is evolutionary related to N4BP1 and has two isoforms, both possessing an N-terminal KH-like domain and a C-terminal endoribonuclease domain (Fig. 3b) (Castagnoli et al. 2019). The KH-like domain, also referred to as a CGIN1 domain, differs from the canonical KH domains since it possesses an additional small metalchelating module (Anantharaman and Aravind 2006; Ficarelli et al. 2019). Ficarelli et al. (2019) hypothesized that the insertion of this metal-chelating module in the KH-like domains may disrupt RNA binding and therefore suggests a different function. Deletion of the KH-like domain reduces antiviral activity against HIV-1 for both isoforms and results in the localization of the proteins in cytoplasmic foci (Ficarelli et al. 2019). In a subsequent investigation, it was shown that the extent of HIV-1 inhibition by KHNYN was correlated with ZAP sensitivity; ZAP antiviral activity was in turn subjected to location and sequence dependent limits (Ficarelli et al. 2020). These findings illustrate the intricacy and complexity of viral recognition by ZAP and subsequent viral mRNA degradation by KHNYN.

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ZAP and the Exosome Complex

Antiviral ZAP activity can also be attributed to its ability to recruit the RNA exosome to degrade its target mRNA (Zhu and Gao 2008; Chen et al. 2008; Guo et al. 2007). Guo et al. (2007) showed that ZAP, specifically its N-terminal portion, co-sediments with the exosome. Further investigation using immunoprecipitation revealed that the 224–254 amino acid region of ZAP was required for binding to exosome subunit hRrp46 (Guo et al. 2007). The association between the exosome and ZAP was further confirmed in a HIV-1 infection model; which Guo et al. (2007)

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showed that in addition to the exosome ZAP is also capable of recruiting a suite of mRNA degradation complexes, such as poly(A)-specific ribonuclease (PARN), and the decapping complex via a p72 helicase.

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The 20 ,50 -Oligoadenylate Synthetase (OAS)/RNAse L System

The 20 ,50 -oligoadenylate synthetase (OAS)/RNase L system is a potent interferoninducible pathway that senses foreign nucleic acids to initiate an antiviral response which cleaves single-stranded viral RNA (Chakrabarti et al. 2011; Sadler and Williams 2008). The OAS-RNase system was initially discovered in the late 1970s during studies on how interferon inhibits viral infections (Hovanessian et al. 1977; Kerr and Brown 1978; Slattery et al. 1979; Clemens and Williams 1978). OAS proteins act as pathogen recognition receptors (PRRs), sensing double-stranded RNA (dsRNA) (Kerr and Brown 1978). The OAS family of proteins consists of OAS1, OAS2, OAS3 and OAS-like protein (OASL); the first three proteins, while homologous to each other, contain one, two and three OAS domains, respectively, can sense dsRNA of different lengths (Sadler and Williams 2008; Schwartz and Conn 2019) and have been shown to synthesize 2-5A to activate RNase L (Sadler and Williams 2008; Ibsen et al. 2014; Ghosh et al. 1997; Rebouillat and Hovanessian 1999; Kwon et al. 2013). Following stimulation by dsRNA, OAS proteins synthesize adenosine oligomers (2-5A) through the polymerization of ATP (Fig. 4a) (Chakrabarti et al. 2011; Sadler and Williams 2008; Hovanessian and Justesen 2007). 2-5A molecules act as messengers to activate an endoribonuclease, RNase L which cleaves and inactivates RNA (Choi et al. 2015; Floyd-Smith et al. 1981; Wreschner et al. 1981). Additionally, fragmented RNA can be detected by other viral RNA sensing proteins such as retinoic acid-inducible gene I (RIG-I) and melanoma differentiation-associated gene 5 (MDA-5) (Sadler and Williams 2008; Choi et al. 2015; Malathi et al. 2010). Of interest in this review is the antiviral effector of this system RNase L, which was termed as a latent (thus ‘L’ in RNase L) endoribonuclease. RNase L is expressed in most mammalian cells and is composed of three domains (Fig. 4b)—an N-terminal regulatory ankyrin repeat domain (ARD), a protein kinase (PK)-like domain, and a C-terminal ribonuclease domain (RNASE) (Chakrabarti et al. 2011; Zhou et al. 2005). The PK-like domain has been shown to be non-functional, lacking key residues required for substrate binding (Dong et al. 1994, 2001). The ARD consists of ankyrin repeats which are typically known to mediate protein-protein interactions, controlling various cellular processes such as transcription, cell cycling and endocytosis among others (Mosavi et al. 2004; Barrick et al. 2008). These repeats are able to bind 2-5A, whose absence allows the ARD to inhibit the RNase domain of RNase L (Chakrabarti et al. 2011; Dong et al. 2001). As a result, RNase L remains inactive as a monomer, only dimerizing and activating after binding by

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a OAS proteins bind viral dsRNA to synthesize 2-5A

2-5A A activates RNase L

Activated RNase L cleaves viral RNA

Fragmented RNA A is detected by RIG-I and MDA5, inducing a type I interferon response

b

PK-like domain (Non-functional) PK-like

Ankyrin repeat domain

RNase RNase domain

Fig. 4 The OAS/RNase L system cleaves viral RNA and induces a type I interferon response in mammals. (a) The OAS/RNase L system cleaves and inactivates viral RNA, subsequently allowing the RNA fragments to be sensed by receptors of the innate immune system. (b) Domain architecture of RNase L

2-5A. Interestingly, ZAP deletion results in a higher constitutive expression of RNase L, while OAS3 deletion causes an upregulation of interferon-stimulated genes (ISGs), hinting at some form of cross talk between the two systems (Odon et al. 2019). The OAS/RNase L system has been intensely studied since its discovery. While there have been several excellent reviews written in the past with regard to its interaction with viruses (Chakrabarti et al. 2011; Li et al. 2016; Silverman 2007), we would like to describe several recent examples of the system and its efficiency in antagonizing viral replication. In a study to determine if the OAS/RNase L system enacts an antiviral response against HCV, Kwon et al. (2013) demonstrated that RNase L is able to exert its antiviral activity in human hepatoma cells. Additionally, only specific OAS proteins,

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OAS1 p46 and OAS p100, are able to mediate RNase L activity against the virus in the infection model (Kwon et al. 2013). In a separate study to determine RNase L cleavage sites in host cells and viruses, it was shown using deep sequencing that discrete regions in HCV RNA were notably susceptible to RNase L cleavage (Cooper et al. 2014). Varying effects of the OAS/RNase L system on other viruses have also been reported. Liao et al. (2020) showed that OAS2 is upregulated following Zika virus (ZIKV) infection and inhibits its replication. However, it was determined that the antiviral activity is potentially due to OAS2 upregulation through the activation of RIG-I, which initiates an antiviral interferon response independent of RNase L activity (Liao et al. 2020). In a separate study, Whelan et al. (2019) suggested that ZIKV can evade RNase L activity in the early stages of infection because it replicates at invaginations in the endoplasmic reticulum where it is protected from cleavage. Constantly subjected to targeting by the OAS/RNase L system, it was also demonstrated that interferon-resistant HCV genotypes possess mutations which reduce the typical numbers of UA and UU dinucleotides (cleavage sites for RNase L) in the HCV genome (Han and Barton 2002). Notably, HCV mRNA acquired silent mutations which eliminated UU and UA dinucleotides during interferon therapy, suggesting a strategy where HCV evades targeting by RNase L (Han and Barton 2002).

13

Conclusion

Beyond its initial perceived role in the central dogma of molecular biology, RNA has been demonstrated to fulfil many roles in the kingdoms of life. Its pervasiveness belies the many forms of regulation employed to control its role in crucial biological processes. While we have discussed how damaged RNA is disposed of via NGD, there also exist other possible mechanisms to avoid or repair the damage done to RNA (Wurtmann and Wolin 2009). Y-box-binding protein (YB-1), a stress granule localized protein, has been proposed to bind to 8-oxoG containing nucleotides and confer resistance to paraquat-induced oxidative damage when overexpressed in Escherichia coli, suggestive of a pathway to sequester damaged RNA (Hayakawa et al. 2002, 2010). Separately, E. coli and human AlkB demethylate methylated lesions in both DNA and RNA (Aas et al. 2003; Ougland et al. 2004). Given the physiological importance of RNA repair, further studies will help to clarify and identify possible new mechanisms that cells employ to deal with damaged RNA. In the second part of this review, we discussed how exogenous RNA is targeted by ribonucleases. It is important to note that the role of damaging and inactivating exogenous RNAs is not limited only to ribonucleases. While not discussed in this review, there exists a diversity of proteins which have a part in regulating the immune response and combating viral infections. For example, apolipoprotein B mRNA editing enzyme, catalytic polypeptide-like 3G (APOBEC3G) is a cytidine deaminase which has been demonstrated to restrict HIV-1 by genome editing,

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through deamination-dependent inhibition (Yang et al. 2020b; Salter et al. 2019; Delviks-Frankenberry et al. 2020). Moreover, in other kingdoms of life, weaponry besides nucleases has been employed to resolve biological conflicts in bacteria and archaea, such as CRISPR systems and restriction enzymes (Burroughs and Aravind 2016). In conclusion, the role of ribonucleases in processing RNA has an inevitable impact on the physiological well-being of cells. Critical processes like mRNA quality control have evolved over time to deal with compromised RNA. Accordingly, it is also of no surprise that mammalian systems have formulated ways to damage and degrade RNA of foreign origin as in the case of viral infections. Concurrently, the immune system is constantly modulated to ensure that foreign entities are efficiently cleared while preserving or minimizing harm to self. In the above-mentioned examples, ribonucleases take centre stage, fulfilling the roles of RNA inactivation and degradation. However, we would acknowledge that we have discussed but a fraction of an almost non-exhaustive list of ribonucleases, evolved to maintain physiological homeostasis in biological systems. Acknowledgements The authors thank all members of our laboratory for discussions. This work is supported by Japan Society for the Promotion of Science (JSPS) KAKENHI Grant Numbers JP18H05278 and 20F20115 and by AMED under Grant Number JP19gm4010002. Compliance with Ethical Standards Conflict of Interest The authors have no conflicts of interest to report.

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Cytoplasmic mRNA Recapping: An Unexpected Form of RNA Repair Daniel R. Schoenberg

Abstract The cap consists of a 7-methylguanosine residue joined to the first nucleotide of an mRNA through a 50 -50 triphosphate linkage. It is added co-transcriptionally to all mRNAs, and proteins binding to the cap modulate subsequent steps in pre-mRNA processing, export, quality control, localization, and translation. Eukaryotic cells have a number of enzymes that hydrolyze the cap structure, and loss of the cap was generally thought to be both irreversible and to commit the transcript to decay. However, this proved to be an oversimplification. Although capping was originally thought to be restricted to the nucleus, my lab identified cytoplasmic forms of all of the enzymes necessary to add a cap onto a decapped mRNA. These assemble on the common adapter protein, Nck1, which serves as a scaffold for a functional capping metabolon. This chapter describes the discovery of cytoplasmic capping and how the proteins that catalyze this process were identified. It tells how we discovered cap homeostasis, a process of decapping and recapping, and the relationship of this process to translational control, mRNA decay, and transcriptome complexity. Keywords mRNA cap · 7-methylguanosine cap · Cytoplasm · mRNA recapping · Translation · Transcriptome

1 The Nature of the Cap and Its Role in mRNA Metabolism The cap was first identified on viral RNAs, followed shortly thereafter on eukaryotic mRNAs (Furuichi 2015; Furuichi et al. 1975a; b; Moss 2017; Wei and Moss 1975; Wei et al. 1975). It consists of a 7-methylguanosine joined by a 50 ,50 -triphosphate linkage to the first transcribed nucleotide of pre-mRNAs, small nucleolar RNAs

D. R. Schoenberg (*) Department of Biological Chemistry and Pharmacology, The Ohio State University, Columbus, OH, USA e-mail: [email protected] © Springer Nature Switzerland AG 2021 I. Kotta-Loizou (ed.), RNA Damage and Repair, https://doi.org/10.1007/978-3-030-76571-2_6

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Fig. 1 The structure of the 50 cap is shown with the salient features plus the names of the key enzymes indicated.

(snoRNAs), primary microRNAs (pri-miRNAs), small nuclear RNAs (snRNAs), and long noncoding RNAs (lncRNAs, Fig. 1). This m7GpppN modification is characteristic of newly synthesized RNA polymerase II (Pol II) products but also some pre-tRNAs (Ohira and Suzuki 2016). For all but pre-tRNAs (for which the enzymology is not known), nuclear capping is catalyzed by capping enzyme (RNA guanylyltransferase and 50 -phosphatase, RNGTT) and cap methyltransferase (RNA guanine-7-methyltransferase, RNMT) bound to the C-terminal domain of the large subunit of RNA Pol II (Ho et al. 1998; Yue et al. 1997; Martinez-Rucobo et al. 2015). Addition of the cap begins with conversion of the 50 -triphosphate ends of newly synthesized RNA to a 50 -diphosphate by the N-terminal triphosphatase domain of RNGTT. RNGTT also forms a covalent complex with GMP at lysine 294 in the C-terminal triphosphatase domain. In the second step of capping, this GMP moiety is transferred onto the 50 -diphosphate end to generate a molecule with a GpppN terminus (Fig. 2a). The basic cap structure (termed Cap 0) is formed by RNMT-catalyzed methylation at the N7 position. In general, capped mRNAs undergo subsequent 2’-O methylation on the first (Belanger et al. 2010) and second (Werner et al. 2011) transcribed nucleotides. The resulting cap structures are termed Cap 1 and Cap 2, respectively (Fig. 1). These cap ribose modifications facilitate translation and protect mRNAs from being targeted by surveillance pathways that monitor for non-self-RNAs such as infecting virus (Hyde and Diamond 2015; Schuberth-Wagner et al. 2015). A number of additional cap and cap-like moieties have been identified by mass spectrometry, including GpppNm, m2,2,7GpppG, m7Gpppm6A, and m7Gpppm6Am and, less commonly, nicotinamide adenine dinucleotide (NAD), flavin adenine dinucleotide (FAD) (Sharma et al. 2020), UDP-glucose (UDP-Glc), and UDP-Nacetylglucosamine (UDP-GlcNac) caps (Wang et al. 2019; Galloway et al. 2020). Although it is conceivable that one or more of these moieties might be generated in

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Fig. 2 The nuclear (a) and cytoplasmic (b) capping reactions are shown with the enzymes that catalyze each step in this process

the cytoplasm, to date only the 7-methylguanosine cap has been shown to be added in the cytoplasm. By marking the mRNA 50 end, the cap serves as a unique binding site for proteins that modulate virtually every step in mRNA metabolism. These include nuclear processing (Gilmartin et al. 1988; Izaurralde et al. 1994; Schwer and Shuman 1996; Flaherty et al. 1997; Gonatopoulos-Pournatzis and Cowling 2014), export (Jarmolowski et al. 1994; Visa et al. 1996), translation (Lee et al. 2016; Tcherkezian et al. 2014; Lahr et al. 2017; Philippe et al. 2018; Topisirovic et al. 2011), microRNA silencing (Chapat et al. 2017), nonsense-mediated mRNA decay (Hosoda et al. 2005), and mRNA decay (Schoenberg and Maquat 2012). To date translation and mRNA decay are the major processes shown to be affected by cytoplasmic mRNA recapping.

2 Decapping and 50 Decay Decapping and endonuclease cleavage are the two major processes by which an mRNA can lose the cap (Schoenberg and Maquat 2012). In general, decapping is catalyzed by one or more Nudix (nucleoside diphosphate linked to another moiety X) proteins, the best known of which is Dcp2 (Grudzien-Nogalska and Kiledjian 2017). The human genome has 22 genes encoding Nudix family proteins, and these enzymes demonstrate varying degrees of substrate specify. For example, Dcp2 binds a subset of mRNAs that have a stem-loop structure located ten bases or less from the cap (Li et al. 2008), and Dcp2-mediated decapping is activated by its interaction with decay-activating proteins (Schoenberg and Maquat 2012). Nudt3, Nudt12, Nudt15, Nudt16, Nudt17, and Nudt19 have in vitro decapping activity, but in vivo evidence for decapping activity is limited to Dcp2, Nudt3, and Nudt16. Some of these

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enzymes can also cleave unmethylated caps (i.e., GpppN, Nudt2, Nudt3, Nudt12, Nudt15–17, Nudt19), NAD (Nudt12, Nudt16), and FAD (Nudt2, Nudt16), with Nudt16 appearing to be the most promiscuous decapping enzyme identified to date (Sharma et al. 2020). In addition, mammalian cells possess two scavenger decapping enzymes that hydrolyze the m7GpppN caps that remain after 30 decay of the mRNA body (Schoenberg and Maquat 2012), DcpS (Liu and Kiledjian 2005; Fuchs et al. 2020), and FHIT (Taverniti and Seraphin 2014). Separate from the Nudix enzymes is DXO, a multifunctional enzyme with decapping and 50 exonuclease activity (Chang et al. 2012). DXO can cleave NAD and FAD caps (Doamekpor et al. 2020) but not m7GpppN caps. It has the distinction of being the only enzyme to date shown to function in in vivo cap surveillance, targeting and degrading mRNAs with improperly methylated (i.e., GpppN) caps (Grudzien-Nogalska and Kiledjian 2017).

3 Early Evidence for Uncapped and Recapped Transcripts In general, decapping and endonucleolytic cleavage each generate decay products with a 50 -monophosphate end, making the resulting decapped transcript susceptible to degradation by the 50 -exonuclease Xrn1 (Schoenberg and Maquat 2012). Xrn1 was originally thought to be highly processive and to rapidly clear decapped mRNAs (Łabno et al. 2016). As such it seemed inconceivable that cells could have uncapped mRNAs, let alone decapped mRNAs that could undergo cytoplasmic recapping. Prior to our discovery of cytoplasmic capping, we undertook a study addressing whether AU-rich instability elements activated decay from either or both ends of the mRNA. This used the RNA Invader assay (Eis et al. 2001), which unfortunately is no longer available. RNA Invader was a highly sensitive FRET-based assay that used signal amplification to quantify RNA rather than target amplification used in PCR-based approaches. We developed Invader probes for each of the three exons of β-globin mRNA and used these to show that 3’-UTR instability elements activated decay from both ends of the mRNA (Murray and Schoenberg 2007). We then went a step further and looked at the interdependence of decapping and 30 decay by individually knocking down Dcp2 and the 30 -exonucleases Rrp41 and Rrp6 (EXOSC10, PM/Scl-100). As one might expect, after Dcp2 knockdown exon 3 decayed more rapidly than exons 1 and 2. In cells knocked down for Rrp41 or Rrp6, exons 1 and 2 were lost (most likely degraded by Xrn1 after decapping), but exon 3 remained remarkably stable. This meant (a) Xrn1 was not sufficiently processive to degrade through even this small mRNA, (b) cells had to maintain at least some population of uncapped mRNA, and/or (c) exon 3 mRNA may have acquired a 50 modification (cap?) that prevented its further degradation. At the time there were other indications for the existence of uncapped and recapped mRNAs. Perhaps the earliest came from work published in 1976 (Schibler and Perry 1976), which identified transcripts with 50 -monophosphate and 50 -diphosphate ends in RNA from cultured mouse cells. In 2008 two groups working in plants identified uncapped forms of mRNAs using an approach that involved

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ligating an adapter onto 50 -monophosphate ends (Jiao et al. 2008; Gregory et al. 2008). Pools of uncapped transcripts were subsequently identified in mammalian cells (Karginov et al. 2010; Mercer et al. 2010; Ni et al. 2010), and it was work in mammalian cells that provided the first evidence for recapped mRNAs. The earliest evidence appeared in a pair of 1992 papers studying the decay of β-globin mRNA in erythroid cells (Lim et al. 1992; Lim and Maquat 1992). Those authors created mice carrying a human β-globin transgene with a premature termination codon in the second exon. Expression of this transgene in erythroid cells was associated with the appearance of 50 -truncated degradation intermediates (Lim et al. 1992), which were unexpectedly stable. Based on recovery of these intermediates with anti-cap antibody and loss of recovery after treatment with tobacco acid pyrophosphatase they proposed, the 50 ends were protected by a cap or cap-like structure (Lim and Maquat 1992). Evidence for cytoplasmic capping next appeared in 2001 in work studying antisense inactivation of viral RNAs (Thoma et al. 2001). Base pairing of an antisense DNA activates cleavage by cellular RNase H, and in that study antisense-mediated cleavage products were stable and translated into N-terminally truncated protein products. Since cap-dependent initiation is the predominant form of translation, this finding raised the intriguing possibility that the downstream cleavage products were recapped.

4 The Discovery of Cytoplasmic Capping The discovery of cytoplasmic capping came about from work revisiting the nature of the 50 truncated decay intermediates of nonsense-containing β-globin mRNA. The initial observation in Lim et al. (1992) suggested these were generated in the cytoplasm. A collaboration between my lab, Audrey Stevens and Lynne Maquat showed this was indeed the case (Stevens et al. 2002), and my lab subsequently identified SMG6 as the responsible endonuclease (Mascarenhas et al. 2013). Using reagents that were not available in 1992, we also confirmed the findings in Lim and Maquat (1992) that decay intermediates of nonsense-containing β-globin mRNA indeed have a 7-methylguanosine cap (Otsuka et al. 2009). This presented a quandary; it was clear the decay products were generated in the cytoplasm, yet according to dogma capping only occurred in the nucleus. What processes were responsible for adding the cap in the cytoplasm? One possibility was the caps were taken from other mRNAs, similar to viral “cap snatching” (Dias et al. 2009; Gu et al. 2015; Clohisey et al. 2020). However, there was no evidence in the human genome for proteins with this activity. There was also no evidence for an additional capping enzyme beside RNGTT. This led us to ask whether there was a cytoplasmic pool of RNGTT. A salient feature of RNGTT is its ability to form a covalent intermediate with GMP bound to the ε-amino group of lysine 294 (K294) prior to transfer of this moiety onto RNA with a 50 -diphosphate end (Fig. 2). In the absence of an RNA substrate, this reaction stalls at the GMP bound state, making it possible to identify RNGTT in crude

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Fig. 3 Nuclear and cytoplasmic extracts of U2OS cells were incubated in vitro with α-32P-GTP to allow for covalent 32P labeling of RNGTT at the GMP-binding site. The input samples are shown in the left two lanes, with five times more cytoplasmic than nuclear protein. A western blot for U2AF65 shows limited contamination of the cytoplasmic fraction with nuclear protein. On the right side of the figure, each sample was incubated with streptavidin magnetic beads containing nonimmune IgG () or antibody to RNGTT (+), and recovered proteins were separated together with the input samples on SDS-PAGE. 32P-GMP labeled RNGTT was visualized by phosphorimager. The results identify a single protein of the anticipated size in both nuclear and cytoplasmic extracts

extracts by simply incubating with α-32P-GTP. After rigorously separating nuclei from cell lysates, this demonstrated the expected presence of RNGTT in the nucleus but also in the cytoplasm of every cell line studied (see Fig. 3 for an example). When incubated in a reaction containing ATP, immunoprecipitated cytoplasmic RNGTT was also able to add GMP onto RNA with a 50 -monophosphate end, but not onto RNA with a 50 -hydroxyl. This property was dependent on the C-terminal guanylyltransferase domain and was abolished by changing the GMP-binding lysine at position 294 to alanine (K294A). Importantly, the kinase and capping activities co-sedimented on a glycerol gradient, providing the first evidence for a cytoplasmic complex containing the enzymatic activities needed to restore the cap on decapped or endonuclease cleaved mRNAs.

5 The Proteins of the Cytoplasmic Capping Complex These findings raised the question of the proteins that constitute the cytoplasmic capping complex. Two fortuitous observations with transfected RNGTT-expressing transgenes led to the discovery of NCK1 as the scaffold for assembly of this

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Fig. 4 The left panel shows the organization of proteins within the cytoplasmic capping complex and how they assemble on adapter protein NCK1. The middle panel shows the relationship of each of these proteins to individual steps in cytoplasmic capping, and together with the right panel shows “cap homeostasis,” the cycling of mRNA between capped and uncapped states

complex. First was the finding that modifications to the C-terminus of RNGTT interfered with the ability of the immunoprecipitated complex to add α-32P-GMP onto 50 -monophosphate RNA (Mukherjee et al. 2014). Neither of these changes affected the covalent binding of GMP, so we reasoned they impacted the assembly of the cytoplasmic capping complex. Phylogenetic analysis of cap-end sequencing data suggested cytoplasmic capping was limited to higher metazoans. As it turns out, RNGTT of these organisms has a C-terminal proline-rich sequence whereas RNGTT of lower metazoans does not. Proteins with SH3 domains are the major binding partners of proline-rich sequence elements, and we quickly homed in on adapter protein NCK1 as a central component of the cytoplasmic capping complex. NCK1 has three consecutive SH3 domains and a terminal SH2 (phosphotyrosine-binding) domain and is best known for its role in transducing receptor tyrosine kinase signaling to the actin cytoskeleton (Bladt et al. 2003; Oser et al. 2010; Ger et al. 2011). Results in Mukherjee et al. (2014) showed that RNGTT binds to the third SH3 domain through its proline-rich C-terminus, and the second SH3 domain binds the kinase responsible for generating the 50 -diphosphate capping substrate from 50 -monophosphate RNA (Fig. 4). This physical juxtaposition of kinase and capping activities pointed to the cytoplasmic capping complex as a metabolon (Srere 1987; Wu and Minteer 2015), a physical grouping of enzymes able to catalyze sequential steps in a metabolic pathway. We next addressed N7 methylation of the nascent cytoplasmic cap. Although eukaryotic cells possess a number of RNA methyltransferases, RNMT is the only one that uniquely catalyzes cap N7 methylation. Much like RNGTT, it was generally thought that RNMT and cap methylation is restricted to the nucleus. In our work, immunofluorescence showed nuclear and cytoplasmic RNMT staining, and this was matched by the identification of cap methylation activity in cytoplasmic extracts and loss of this activity following RNMT knockdown. RNMT is activated by binding of a cofactor, termed RAM (or RAMAC) (Gonatopoulos-Pournatzis et al. 2011;

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Varshney et al. 2016). Cytoplasmic RNMT heterodimerizes with RAM, and RAM knockdown demonstrated that this interaction is required for optimal cap methylation activity. The presence of RNMT, RNGTT, and NCK1 in a single complex is exemplified by the gel filtration experiment in Fig. 5 (from Trotman et al. 2017), where all three proteins and cap methylation activity co-elute in a single complex. In nuclear capping RNMT and RNGTT are juxtaposed by their mutual binding to the phosphorylated C-terminal domain of the Pol II large subunit (Martinez-Rucobo et al. 2015). This raised the question how these two proteins could co-elute in a single cytoplasmic complex. There was early evidence that RNMT and RNGTT might directly interact (Pillutla et al. 1998), and this was confirmed in Trotman et al. (2017). The RNMT/RAM heterodimer is recruited to the cytoplasmic capping complex by the C-terminal domain of RNMT binding to RNGTT, and its presence provides the final evidence for this complex as a metabolon that catalyzes the stepwise conversion of RNA with a 50 -monophosphate end to one with a functional cap structure (Fig. 4). The identification of a functional cytoplasmic capping complex left a number of questions unanswered (see below), perhaps the most critical being the identity of the 50 -kinase that is required for generating 50 -diphosphate recapping substrates. It was conceivable that this might be one of the known nucleotide kinases, but analysis of the available enzymes showed none of these were the 50 -kinase. The search for the 50 -kinase used cross-linking and direct recovery and proximity-dependent biotinylation as a comprehensive approach to identifying all of the cytoplasmic proteins that interact with RNGTT. While this did not identify the sought-after 50 -kinase, it unexpectedly identified 66 proteins that interact with cytoplasmic RNGTT, 52 of which are RNA-binding proteins (Trotman et al. 2018). As an aside, nuclear RNGTT interacts with only four proteins (Youn et al. 2018). As another aside, our study also identified RNGTT as an HSP90 client protein and showed nuclear and cytoplasmic capping activity is reduced in cells treated with HSP90 inhibitors such as onalespib, which is used in cancer chemotherapy.

6 Cap Homeostasis as an RNA Repair Mechanism That Modulates Translation, mRNA Decay, and Translational Control Having identified how the cap gets restored on mRNAs in the cytoplasm, the obvious question is what purpose does this serve? The first inkling came in our initial study (Otsuka et al. 2009), where interference with cytoplasmic capping by the K294A form of RNGTT reduced the ability of cells to recover from a brief arsenite stress. Because translation and stress responses are intimately linked (Tahmasebi et al. 2018), we suspected this might be indicative of a role for cytoplasmic recapping in facilitating the transition between translating and non-translating states. Direct evidence for this came from the first study aimed at

Fig. 5 (a) Cytoplasmic extract from U2OS cells was separated on a calibrated Sephacryl S-200 gel filtration column with the indicated fractions analyzed by western blotting for RNMT, RNGTT, and NCK1. (b) In vitro cap methylation assay was performed on fractions 2–14 (representing the cytoplasmic capping complex) and fraction 32, which corresponds to the expected size of RNMT. In this assay RNA with a 32P-labeled cap is incubated with the indicated fractions plus S-adenosyl-methionine; the product is digested with P1 nuclease, separated on PEI cellulose thin layer and visualized by phosphorimager. The percent conversion is indicated under the autoradiogram and shown graphically in (c). These data are from Fig. 2 of Trotman et al. (2017) and are reproduced here under Creative Commons license CC BY

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identifying mRNA targets of cytoplasmic capping (Mukherjee et al. 2012). In that study cytoplasmic capping was again inhibited by induction of the K294A transgene. Cytoplasmic RNA recovered from uninduced and induced cells was depleted of rRNA, incubated in vitro with Xrn1, and analyzed on human exon microarrays. On these arrays each exon is represented by multiple probe sets, and bioinformatic tools were used to look at changes in signal intensity with 50 -30 polarity. Whereas capped mRNAs would have equivalent probe intensities across, each transcript uncapped mRNAs should display a polar pattern of low signal intensities at the 50 -most probe sets that increase toward the 30 -most probe sets. This resulted in identification of three classes of uncapped/recapped mRNAs. The first set was mRNAs that have an existing subpopulation of uncapped forms under normal conditions. We termed these transcripts “natively” uncapped. Another set of mRNAs had uncapped forms that appear only when cytoplasmic capping was inhibited by K294A induction. These were termed “capping inhibited” transcripts, and their identification laid the foundation for much of our subsequent work. The third population (“common”) consisted of mRNAs that already had a subpopulation of uncapped forms whose representation increased when cytoplasmic capping was inhibited with K294A. As one might expect, this study also identified transcripts whose steady-state levels decline when cytoplasmic capping is blocked. Their stabilization following Xrn1 knockdown linked cytoplasmic recapping to mRNA decay. Given the role of the cap in translation initiation, we were particularly interested in determining whether cytoplasmic recapping affects translation. We used sucrose gradients to monitor transcript distributions between polysomes and non-translating mRNPs in cells with normal cytoplasmic capping and in cells in which cytoplasmic capping was blocked with the K294A form of RNGTT. Absorbance traces of gradient fractions showed that inhibiting cytoplasmic capping resulted in increased absorbance of fractions at the top of the gradient (Fig. 6), indicative of an increase in the population of transcripts in non-translating mRNPs. RT-qPCR quantification of target and nontarget mRNAs across gradient fractions showed this shift corresponded to a change in the distribution of cytoplasmic capping targets (but not unaffected mRNAs) from polysomes to non-translating complexes (Mukherjee et al. 2012). Importantly, the transcripts that accumulated in the non-translating mRNP pool are uncapped, and they retain the same length poly(A) tails as their polysome-bound counterparts (Kiss et al. 2016). Because translation efficiency can be modulated by poly(A) tail length, this finding indicated that repair of the cap structure on these mRNAs is all that is needed to return them to the translating pool. We termed this cyclical process of cytoplasmic decapping and recapping “cap homeostasis” (see Fig. 4), and we hypothesized this represents a new nexus for regulating translation. Indirect support for this was subsequently shown by live cell imaging of individual mRNA translation events at the single-molecule level (Yan et al. 2016; Wu et al. 2016), where individual mRNAs cycled between translating and non-translating states as might be expected for transcripts undergoing cap homeostasis.

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Fig. 6 Cytoplasmic extracts from control (blue) and K294A-expressing cells (red) were fractionated on a 10–50% sucrose gradient, and the absorbance at 254 nm was monitored as fractions were collected. The locations of translating polysomes, 80S monosomes, and non-translating mRNP complexes are indicated, and text in the white boxes indicates changes in sedimentation of recapping targets and control mRNAs as a function of K294A expression

While inhibition of GMP addition with the K294A form of RNGTT provided a wealth of information, this approach has a number of drawbacks, all of which come down to its reliance on identifying uncapped transcripts. By definition uncapped RNAs are substrates for Xrn1, so one cannot be certain that the 50 ends identified using K294A to block recapping correspond to actual recapping sites (see below). Most uncapped RNAs are also unstable and can only be identified by knocking down Xrn1 (Mukherjee et al. 2012). Lastly, this approach depends on biochemical separation of capped versus uncapped RNA. These issues led us to develop a more straightforward approach, which also made it possible to directly map the position of recapped ends on target mRNAs.

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As noted above, cells possess surveillance enzymes that degrade mRNAs with improperly methylated caps (Grudzien-Nogalska and Kiledjian 2017), and cytoplasmic cap methylation is catalyzed by RNMT/RAM. RNMT has two domains. The N-terminal 120 amino acids contains nuclear targeting and regulatory sequences and is the target of a number of kinases (Aregger and Cowling 2013; Galloway and Cowling 2019). The remaining portion of RNMT which contains the catalytic domain is the binding site for the RNMT coactivator RAM (Varshney et al. 2016). We took advantage of this to create a form of RNMT that acts as a dominant negative inhibitor of cytoplasmic cap methylation (Trotman et al. 2017). This mutant RNMT consists of the methyltransferase domain (residues 121–476) with an inactivating mutation (D203A) in the S-adenosylmethionine binding site. Addition of the HIV Rev nuclear export signal (NES) resulted in a protein that is restricted to the cytoplasm that we termed “ΔN-RNMT.” ΔN-RNMT competes with endogenous RNMT for RAM and for binding to RNGTT in the cytoplasmic capping complex. As a result of ΔN-RNMT overexpression, we hypothesized that recapped mRNAs will have unmethylated caps and, as such, be degraded by cap surveillance enzymes. Indeed, this approach resulted in reduced steady-state levels of recapped transcripts, and this was readily detectable using RNA-Seq. Rather than sequence the entire transcriptome, we instead used a quantitative approach that focused on 30 end tags and limited analysis to transcripts whose levels changed more than 1.5-fold (Del Valle et al. 2020). This identified 5606 mRNAs half of which decreased as a function of ΔN-RNMT expression. A bioinformatics analysis of this cohort identified mRNAs with 50 -terminal oligopyrimidine tracts (5’-TOP) mRNAs as the single largest group of cytoplasmic capping targets. These mRNAs start with a run of 10–13 pyrimidine residues, always starting with C. They include transcripts for all of the ribosomal protein mRNAs and most translation factors (Meyuhas and Kahan 2015), and their translation is regulated by La-related protein (LARP1) binding to the cap and 5’-TOP sequence (Lahr et al. 2017; Fonseca et al. 2018; Philippe et al. 2018). As noted above, a major question is whether recapping is limited to the native 50 end or if it also occurs further downstream. Several 5’-TOP mRNAs were selected as examples to address this question using a technique that appends a double-stranded DNA adapter onto the 30 end of a first strand cDNA only when it is opposite the G of the cap (Del Valle and Schoenberg 2020). Gene-specific PCR primers were then used to examine the cap sites on three ribosomal protein mRNAs (RPS4X, RPS3, RPL8) and two initiation factor mRNAs (EIF3D, EIF3D), with the cap site identified by the sequence immediately adjacent to the ligated adapter. The three ribosomal protein mRNAs each generated a single PCR product and the cap sites mapped to their native 50 ends (Del Valle et al. 2020). PCR amplification of EIF3D and EIF3K generated two products, one at the anticipated size for capping at native 50 ends and one that was slightly smaller. Sequencing of these amplicons showed recapping occurred at the native 50 end but also within and downstream of the 5’-TOP sequence. This was the first definitive evidence for downstream recapping. The 5’-UTRs of 5’-TOP mRNAs are highly structured (Mizrahi et al. 2018), and structured regions within 5’-UTRs interfere

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with 50 -30 degradation by Xrn1 (Charley et al. 2018). Thus, structural elements may be responsible for the appearance of distinct recapping sites within the 5’-UTRs of EIF3D and EIF3K mRNAs. These findings also provide additional evidence of a regulatory role for cytoplasmic capping as 5’-TOP mRNAs that are recapped within or downstream of the polypyrimidine sequence should not bind LARP1 and thus evade translational regulation by LARP1. As noted above, inhibiting cytoplasmic capping reduced the ability of cells to recover from stress (Otsuka et al. 2009). LARP1 is involved in tethering 5’-TOP mRNAs to stress granules (Wilbertz et al. 2019), and since the cap is required for LARP1 binding, uncapped transcripts would likely not accumulate in stress granules and would either be degraded or prevented from returning to a translating state once the stress is removed.

7 Is There a Relationship Between Recapped Ends and CAGE Tags? Capped analysis of gene expression (CAGE) (Shiraki et al. 2003) was invented to identify transcription start sites. It was based on the understanding that capping only occurs at the site of transcription initiation; however, it also identified capped 50 ends downstream within the body of a large number of mRNAs (Fejes-Toth et al. 2009). The scope of CAGE tags has expended with the development of additional transcriptome-wide approaches for identifying capped ends (Afik et al. 2017; Batut et al. 2013; Djebali et al. 2012; Cartolano et al. 2016; Gu et al. 2012; Machida and Lin 2014). Djebali et al. (2012) concluded that approximately 70% of CAGE tags correspond to transcription start sites, with the remainder being downstream. This raised the question whether this large number of downstream capped ends is generated by cytoplasmic capping. Two reports provide support for a relationship between downstream CAGE tags and cytoplasmic recapping (Kiss et al. 2015; Berger et al. 2019). These studies used ligation-mediated PCR to capture uncapped 50 ends that appear following K294A inhibition of cytoplasmic capping, and in both cases newly formed uncapped ends either matched or were close to the locations of downstream CAGE tags. While these findings are consistent with cytoplasmic capping as a source of downstream CAGE tags, the reliance on detecting uncapped mRNAs makes any conclusions subject to the caveats noted above. Proof that cytoplasmic capping is responsible for downstream CAGE tags must await a transcriptome-wide evaluation of changes in downstream CAGE tags using an approach that does not depend on detection of uncapped transcripts, such as inhibiting cytoplasmic cap methylation (Del Valle et al. 2020).

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8 Does Cytoplasmic Capping Impact the Proteome? The existence of downstream CAGE tags raises the tantalizing possibility that mRNA recapping might expand the proteome by enabling translation initiation at locations downstream of canonical start sites (Trotman and Schoenberg 2019). There are examples of downstream initiation sites in ribosome profiling data (e.g., Lee et al. 2012; Calviello et al. 2020). However, it was not clear whether downstream initiation sites represent multiple initiation events on a single mRNA or single initiation events on 50 truncated isoforms of that mRNA. Proteomics data have also identified significant numbers of N-termini mapping downstream of canonical initiation sites (Kim et al. 2014; Na et al. 2018; Yeom et al. 2017). For example, Yeom et al. (2017) identified 13,095 N-termini from 5727 proteins in HEK293 cells and identified N-termini corresponding to 2789 known initiation sites, 203 signal peptide and/or propeptide cleavage sites, and 496 possible alternative translation initiation sites. They also identified 9608 N-termini of unknown origin. Therefore, it is conceivable that some of these novel N-termini might arise from recapping downstream of native or canonical start sites. As attractive as it is, to date there are no data to support this model. In collaboration with my colleague Vicki Wysocki, our lab has performed positional proteomics aimed specifically at identifying N-terminal peptides using two different approaches, PTAG (phospho tagging and TiO(2)-based depletion (Mommen et al. 2012) and Nrich (N-terminal peptides enrichment on the filter, Yeom et al. 2017). Each of these studies was performed multiple times using cells with normal cytoplasmic capping and cells in which recapping was blocked by induction of the inhibitory K294A form of RNGTT. Although we succeeded in identifying downstream N-termini, including many that were identified in those published studies, neither of these approaches provided convincing evidence that inhibition of cytoplasmic capping altered the representation of downstream N-termini. An alternative approach was to perform shotgun proteomics analysis comparing total protein and peptide representation in control cells with that of cells expressing the K294A inhibitor of cytoplasmic capping (Agana et al. 2020). Only 57 of the 3565 proteins identified in this analysis showed a significant change in abundance, with 29 proteins increasing and 28 proteins decreasing when cytoplasmic capping is blocked. There was no evidence of relatedness between these proteins by gene ontology analysis. Finally, if translation initiated downstream of canonical start codons, we expected to see an increase in the amount of some N-terminal peptides when cytoplasmic capping was blocked. However, there was no evidence for polar changes in amount for any of the 21,875 identified tryptic peptides. Based on these findings, it appears unlikely the mRNA recapping is a major driver of proteome complexity.

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9 Conclusion and Future Questions I end this chapter with several questions and hopefully some answers. Perhaps foremost is what purpose is served by repairing the cap on decapped mRNAs? Only a portion of the transcriptome undergoes mRNA recapping, and notably this includes many of the 5’-TOP mRNAs. Some 5’-TOP mRNAs (notably those encoding ribosomal proteins) are recapped only at their native 50 ends. This supports the model for cap homeostasis we put forward in Mukherjee et al. (2012), and as noted above, the cycling of caps on native 50 ends of specific mRNAs is consistent with in vivo on-off translational cycling of individual mRNAs observed in Yan et al. (2016), Wu et al. (2016). Therefore, maintenance of the cap on the mRNA 50 end is one, and perhaps the major, purpose of cytoplasmic capping and in that regard can be considered a form of RNA repair. The protein products of 5’-TOP mRNAs feature prominently in translation, and the translation of 5’-TOP mRNAs in turn is regulated by mTOR phosphorylation of LARP1 (Lahr et al. 2017). Both the cap and the 5’-TOP sequence are required for LARP1 binding (Philippe et al. 2018). Therefore, the observation in Del Valle et al. (2020) that EIF3D and EIF3K mRNAs undergo recapping within and downstream of the 5’-TOP sequence identifies a second function for cytoplasmic recapping: regulating translation by altering 5’-UTR sequences or structures. In this case, truncating the 5’-TOP sequence will interfere with binding and regulation by LARP1. In this regard EIF3D is particularly interesting as it is a cap-binding protein that functions with the eIF4G homolog DAP5 to mediate translation initiation of c-JUN (Lee et al. 2016) and mRNAs encoding proteins involved in DNA repair, translation, cell survival, and cell motility (de la Parra et al. 2018). As noted above, as attractive as it may be to think of cytoplasmic capping as the source of downstream CAGE tags and/or as a way of expanding proteome complexity, there is currently no proof for either of these. That does not rule out these possibilities, but the bar will be high. To be convincing one would have to demonstrate changes in representation of downstream CAGE tags in multiple ENCODE cell lines. A fascinating aspect of downstream CAGE tags is the large number of RNAs corresponding to capped 3’-UTRs (Mayr 2017). These can serve as sponges for microRNAs or RNA-binding proteins, or function in trans to alter gene expression (Ma and Mayr 2018), and it will be important to determine which if any of these are generated by cytoplasmic capping. mRNA recapping may also play a role in the cellular response to stress. We noted in Otsuka et al. (2009) that the ability of cells to recover from a brief arsenite stress was reduced by overexpression of the K294A inhibitory form of cytoplasmic capping enzyme. In the integrated stress response, 5’-TOP mRNAs are localized early to stress granules and P bodies (Wilbertz et al. 2019). The latter is enriched in Dcp1 and Dcp2, and in our 2009 review (Schoenberg and Maquat 2009), we hypothesized some mRNAs might be stored in these condensates in an uncapped state and that recapping might be involved in their return to the translating pool. This is supported by recent work using an in vitro reconstituted system where, in

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macromolecular condensates of Dcp1, Dcp2m and the activating protein Edc3, uncapped RNAs were retained after decapping (Tibble et al. 2021). The relationship between cap homeostasis, stress, macromolecular condensates (i.e., stress granules and P bodies), and translation will be an important area for future investigation. A lingering question is if (or how) cytoplasmic capping is regulated. We know from observations in Mukherjee et al. (2014) that the cytoplasmic capping complex assembles on the multidomain adapter protein NCK1. That study identified the third SH3 domain as the binding site for RNGTT (and with it RNMT/RAM and perhaps RNA-binding proteins), and the second SH3 domain as the binding site for the 50 -monophosphate kinase that is required for converting RNAs with 50 -monophosphate ends into a 50 -diphosphate capping substrate. Existence of this kinase has been known for years, but its identity remains unknown. This enzyme may prove to be a key mediator of a number of RNA repair processes, not just cytoplasmic recapping. We also don’t know the identity of proteins that are bound to the first SH3 domain of NCK1 or the phosphotyrosine-binding C-terminal SH2 domain. The existence of a protein mediator of signal transduction as the scaffold for the cytoplasmic capping complex suggests this process is regulated, likely in a cell and tissue specific manner. The final and perhaps largest question is the role mRNA recapping plays at the organismal level and in health. To date all of the work on cytoplasmic capping has been done using cultured cells. This reductionist focus was and remains important for defining the biochemical interactions needed for this process, the molecular basis of target selectivity, and the fate of recapped mRNAs. Currently, the only way to identify recapping events is to interfere with this process, and initial approaches that depended on identifying uncapped transcripts limited work to cultured cells. This changed with results in Del Valle et al. (2020) showing recapped mRNAs can be identified simply by RNA-Seq after inhibiting cytoplasmic cap methylation. This straightforward method of identifying recapped mRNAs makes it now possible to study the in vivo function of this cap repair process in specific cells and tissues and across the spectrum of animal models of development and disease. There is also the question whether cytoplasmic capping might play a role in the life cycle of RNA viruses. The timing of this chapter coincides with the global pandemic caused by SARS-CoV-2. Just like cellular mRNAs, SARS-CoV-2 genomic and subgenomic RNAs have an N7-methylguanosine cap (reviewed in Maranon et al. 2020). However, SARS-CoV-2 does not appear to hijack the cytoplasmic RNA recapping complex. Instead it has a guanylyltransferase in the nucleotidyltransferase domain (NiRAN) of the SARS-CoV-2 RNA-dependent RNA polymerase (nsp12, Yan et al. 2021), and the virus encodes its own cap N7 methyltransferase (nsp14) and a 2’-Omethyltransferase (nsp16). That does not rule the possibility that other RNA viruses might take advantage of cytoplasmic RNA recapping, and this remains a question for future study. Acknowledgments I wish to thank the people in my lab whose efforts over the years made the work described here possible. I particularly want to thank my long-time collaborator Ralf Bundschuh. I am a biochemist at heart, and Ralf oversaw and/or performed the bioinformatics

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that made this work possible. I also want to thank Vicki Wysocki and her former student Bernice Agana for their expertise, input, and help with all of the proteomics work, and Jackson Trotman and Mike Kearse for their helpful comments on this chapter. Research in the Schoenberg lab on cytoplasmic capping was funded by the US National Institutes of Health, grant GM084177 (to DRS).

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Part III

RNA Damage in Human Diseases

Adenosine-to-Inosine RNA Editing: A Key RNA Processing Step Rewriting Transcriptome in Normal Physiology and Diseases Priyankaa Pitcheshwar, Haoqing Shen, Jian Han, and Sze Jing Tang

Abstract Adenosine-to-inosine (A-to-I) RNA editing is the most prevalent type of RNA modification in mammals and is catalyzed by adenosine deaminase acting on the RNA (ADAR) family of enzymes that recognize double-stranded (ds) RNAs. Inosine mimics guanosine in base pairing with cytidine, thereby A-to-I RNA editing alters dsRNA secondary structure. Inosine is also recognized as guanosine by splicing and translation machineries, resulting to mRNA alternative splicing and protein recoding. Therefore, A-to-I RNA editing is an important mechanism that causing and regulating “RNA mutations” in both normal physiology and diseases, such as cancers. In this chapter, we reviewed the regulatory mechanisms of A-to-I RNA editing, from regulation of ADAR enzymes to the involvement of ADARinteracting secondary regulators. We also reviewed the roles of A-to-I RNA editing on miRNA-mediated gene silencing and RNA metabolism such as splicing, polyadenylation, and N6-Methyladenosine methylation, as well as the functions in apoptosis, immunity, and circadian rhythm. Keywords A-to-I RNA editing · ADAR · A-to-I RNA editing regulators · cross talk · miRNA biogenesis · splicing · cancer · immunity

Adenosine-to-inosine (A-to-I) RNA editing is the most prevalent type of RNA modification in mammals and is catalyzed by adenosine deaminase acting on the RNA (ADAR) family of enzymes that recognize double-stranded (ds) RNAs (Bass Priyankaa Pitcheshwar and Haoqing Shen contributed equally with all other contributors. P. Pitcheshwar · H. Shen Cancer Science Institute of Singapore, National University of Singapore, Singapore, Singapore Department of Anatomy, Yong Loo Lin School of Medicine, National University of Singapore, Singapore, Singapore J. Han (*) · S. J. Tang (*) Cancer Science Institute of Singapore, National University of Singapore, Singapore, Singapore e-mail: [email protected]; [email protected] © Springer Nature Switzerland AG 2021 I. Kotta-Loizou (ed.), RNA Damage and Repair, https://doi.org/10.1007/978-3-030-76571-2_7

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2002). ADAR proteins consist of variable numbers of dsRNA-binding motifs (dsRBMs) at their N-terminus and a C-terminal deaminase catalytic domain. The ADAR family consists of three structurally conserved members, ADAR1, ADAR2, and ADAR3; however, only ADAR1 and ADRA2 are catalytically active (Bass 2002; Chen et al. 2000). A-to-I editing can alter coding sequence, and these protein recoding events can eventually modify protein function or localization albeit rarely (Nishikura 2010; Zhang et al. 2019; Higuchi et al. 1993). Increased experimental evidence confirms the importance of ADAR1 and ADAR2 in cancer pathogenesis. Imbalances in the RNA editome induced by aberrant expression of ADAR proteins drive tumorigenesis. ADAR1 promotes or suppresses tumorigenesis in different cancers, while ADAR2 is mainly a tumor suppressor (Zhang et al. 2019). A handful of aberrant protein-recoding targets have been reported in cancer, and these editing events result in gain- or loss-of-function of oncogenes or tumor suppressors, respectively. For instance, the edited antizyme inhibitor 1 (AZIN1S367G) accelerates cell cycle progression and possesses stronger tumorigenic capabilities than the wild-type form (Chen et al. 2013). The edited zinc-finger protein GLI1 activates the hedgehog signaling pathway and promotes relapse of multiple myeloma (Shimokawa et al. 2013). In contrast, RNA editing renders a loss of metastasis suppressive function of the solute carrier family 22 member 3 (SLC22A3) (Fu et al. 2017). In addition, loss of the pro-apoptotic edited form of insulin-like growth factor-binding protein 7 (IGFBP7K95R) may promote tumorigenesis in esophageal squamous cell carcinoma (Chen et al. 2017). The edited podocalyxin-like protein 1 (PODXLH241R) confers a loss-of-function phenotype with reduced tumorigenesis in gastric cancer (Chan et al. 2016). An editing-mediated functional switch of recoding events was recently reported in the COPI coat complex subunit alpha (COPA). COPAI164V, switching from an oncogene to a tumor suppressor, exhibits a dominant negative effect and may repress the activation of the phosphatidylinositol 3-kinase/serine/ threonine protein kinase B/mammalian target of rapamycin (PI3K/Akt/mTOR) signaling pathway in hepatocellular carcinoma (Song et al. 2020).

1 Regulatory Mechanisms of A-to-I RNA Editing Although ADAR1 and ADAR2 are able to catalyze A-to-I RNA editing in vitro, the inconsistent correlation between the A-to-I editing frequencies and the endogenous ADAR expression levels (Brümmer et al. 2017) has emphasized the existence of spatiotemporal regulation of A-to-I RNA editing in vivo (Tan et al. 2017; QuinonesValdez et al. 2019). Multiple lines of evidence have demonstrated that this finetuning of A-to-I editing by secondary regulators occurs through several mechanisms (Hong et al. 2015).

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Editing Regulation by Altering ADAR Expression and Activity

The most straightforward regulatory strategy is via modulating the expression of ADAR proteins. The dynamics of the A-to-I RNA editome across different tissues during different developmental stages and in the context of different diseases is partially attributed to the ADAR mRNA and protein expression differences. Several mechanisms by which ADAR1 and ADAR2 expression is induced or repressed under certain circumstances have been reported. It is known that interferon (IFN) stimulation induces expression of the longer 150 kDa (p150) isoform of ADAR1 by alternative promoter usage (Patterson and Samuel 1995; George and Samuel 1999). The differential promoter usage of the ADAR1 p150 isoform allows transcription from an alternative first exon which contains a translational initiation signal, whereas the ADAR1 p110 isoform is translated from a downstream translational initiation signal in the second exon. The IFN-stimulated induction of the ADAR1 p150 isoform relying on the interferon-α/β receptor (IFNAR), Janus kinase 1 (JAK1), signal transducer and activator of transcription 2 (STAT2), and interferon regulatory factor 9 (IRF9)-dependent signaling (George et al. 2008; George and Samuel 2015), has demonstrated both anti- and proviral roles as it is able to edit viral RNA and triggers biological changes that affect virus-host interactions (Mannion et al. 2014). Cyclic adenosine monophosphate response element-binding protein (CREB) transactivates ADAR2 expression in neurons which edits the nuclear glutamate ionotropic receptor AMPA-type subunit 2 (GRIA2) pre-mRNA at a Q/R site and protects vulnerable cortical area 1 (CA1) pyramidal neurons from forebrain ischemia (Peng et al. 2006). Conversely, CREB represses ADAR1 expression in melanoma cells (Shoshan et al. 2015). Jun amino-terminal kinase 1 (JNK1) upregulates ADAR2 expression following glucose stimulation in pancreatic beta-cells (Yang et al. 2012). MicroRNAs, such as miR-17 and miR-432, have been shown to directly target ADAR1 mRNA for degradation in metastatic melanoma cells (Nemlich et al. 2013). In addition to the above, expression of ADARs can also be controlled by posttranscriptional processes such as splicing and editing. Two splicing isoforms of ADAR2, which differ with an Alu-J cassette insertion that causes a 40-amino acids difference in the deaminase domain, are expressed in human tissues (Gerber et al. 1997). Both isoforms show the same substrate specificity; however, the longer isoform shows decreased catalytic activity as compared with the shorter one. Interestingly, self-editing of ADAR2 transcript controls its own splicing, wherein the conversion of an intronic AA to AI dinucleotide at its proximal 30 acceptor site gives rise to an alternatively spliced transcript with a 47-nucleotide insertion in its coding region (Rueter et al. 1999). This negative autoregulatory mechanism of ADAR2 can modulate its own expression thus controlling A-to-I editing levels. ADAR’s expression can also be regulated at the posttranslational level. The WW domain-containing E3 ubiquitin protein ligase 2 (WWP2) binds to ADAR2 and catalyzes its ubiquitination and degradation, while peptidylprolyl cis/trans isomerase

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(PPIases), NIMA-interacting 1 (PIN1), stabilizes ADAR2 protein (Marcucci et al. 2011). In addition to having an effect on the expression of ADARs, posttranslational modifications can also affect their activity. Sumoylation of ADAR1 by small ubiquitin-like modifier 1 (SUMO1) alters the editing activity of ADAR1 (Desterro et al. 2005). ADAR1 p110 and ADAR2 can be phosphorylated by AKT kinases, namely, AKT-1, AKT-2, and AKT-3, at sites T738 and T553, respectively (Bavelloni et al. 2019). The catalytic activity of these phosphomimetic mutants of ADAR1 p110 (T738D) and ADAR2 (T553D) displayed a 50–100% reduction compared to their wild-type counterparts falling in line with previous evidence which highlighted the role of posttranslational modifications in mediating the editing activity of ADARs. Quite a number of fascinating regulatory mechanisms have also been proposed. From crystal structure studies, inositol hexakisphosphate (IP6) was discovered as a cofactor buried in the ADAR2 protein core, quintessential for protein folding, and in maintaining the catalytic activity of ADAR2 (Macbeth et al. 2005). The excitotoxic levels of glutamate induce cleavage of ADAR2 in neurons, and this leads to a loss of GRIA2 editing (Mahajan et al. 2011).

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Editing Regulation by Affecting Subcellular Localization of ADAR

Localization of ADARs determines their accessibility for their RNA substrates. Nascent RNA transcripts are synthesized in the nucleoplasm and edited before being exported to the cytoplasm; therefore changes in subcellular localization of ADARs affect the course of editing. The ADAR1 p110 isoform localizes predominantly in the nucleus, whereas the p150 isoform is largely cytoplasmic. Editing by ADAR1 p150 occurs efficiently in the cytoplasm, suggestive of its role as an IFN-stimulated gene in the antiviral response (Wong et al. 2003). The nuclear export signal (NES) located within the z-DNA-binding domain of the p150 isoform allows it to be recognized and exported by chromosomal region maintenance 1 (CRM1) (Poulsen et al. 2001). In addition, transportin-1 (TNPO1) and exportin-5 (XPO5) mediate the import and export of ADAR1 to the nucleus, respectively (Fritz et al. 2009). Stress, including UV irradiation and heat shock, induces cytoplasmic translocation of ADAR1, which is mediated by increased ADAR1 binding to XPO5 following ADAR1 phosphorylation by the MKK6-MSK-MAP pathway, including mitogen-activated protein kinase kinase 6 (MKK6), p38-mitogen- and stress- activated protein kinase (MSK), and mitogen-activated protein kinase (MAP) (Sakurai et al. 2017). Due to lack of N-terminal NES presented in ADAR1, ADAR2 localizes predominately in the nucleus and more specifically accumulates in the nucleolus (Desterro et al. 2003). Nucleolar sequestration of ADAR2, caused by its binding to ribosomal RNA, represses editing (Sansam et al. 2003). The karyopherin subunits α1 (KPNA1) and α3 (KPNA3) interact with ADAR2, and this may be responsible for its nuclear import or export (Maas and Gommans 2009). Furthermore, it has been

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demonstrated that nuclear import of ADAR2 is mediated by importin-α4 and is stabilized by interaction with PIN1 in the nucleus. This increased accessibility of ADAR2 to the editing substrates during neuronal maturation contributes to neuronal plasticity (Behm et al. 2017). Small nucleolar RNAs (snoRNAs) are a class of noncoding RNAs present in cells. A C/D box snoRNA MBII-52 (SNORD115) has been shown to regulate the serotonin 2C receptor (5-HT2CR) transcriptionally by decreasing the efficiency of ADAR2-mediated editing in the nucleolus (Vitali et al. 2005; Doe et al. 2009). All these studies support the notion that the accessibility of ADARs for their potential RNA substrates in specific cellular compartments is critical to A-to-I RNA editing.

1.3

Additional Layers of A-to-I RNA Editing Regulation

RNA-binding proteins (RBPs) are a multifaceted class of proteins involved in several aspects of RNA metabolism. Utilizing public RNA-Seq and eCLIP-Seq datasets from ENCODE, a number of RBPs were revealed as cell-type specific regulators with varying effects on A-to-I editing (Quinones-Valdez et al. 2019). Herein, DNA-binding protein 43, encoded by the TAR DNA-binding protein (TARDBP) gene, was identified as a negative regulator of A-to-I editing by affecting ADAR1 expression levels. Drosha, a core protein involved in the miRNA biogenesis pathway, behaved as an enhancer of A-to-I editing, presumably by interacting with ADAR1 in the nucleus and binding in close proximity to its substrates. Owing to their ability to form double-stranded structures, inverted Alu pairs provide the ideal topography for ADARs. Yet another repressor of A-to-I editing, the 60 kDa SS-A/ Ro ribonucleoprotein (Ro60; encoded by the TROVE2 gene), was also detected. It was speculated to affect editing by binding to substrate Alu elements. This repressive role for Ro60 is consistent in the context of systemic lupus erythematosus whereby Ro60 knockdown resulted in a global upregulation of the A-to-I editome. Recent studies have reported several ADAR interactors playing critical editing regulatory roles in human diseases. Widespread dysregulated RNA editing has been observed in autism spectrum disorder (ASD) brains with the Fragile X syndrome (Tran et al. 2019). The Fragile X proteins, Fragile X mental retardation protein (FMRP) and Fragile X-related protein 1 (FXR1P), bind to dysregulated RNA editing sites and knockdown of FMRP or FXR1P, either reduced or enhanced editing level of their respective sites. FMRP interacted with both ADAR1 and ADAR2, while FXR1P interacted with ADAR1 but not with ADAR2. Aberrant global RNA editing patterns are commonly found in human cancers, and a recent study of the ADAR interactor, death-associated protein 3 (DAP3), sheds light on how editing is dysregulated in cancer cells independent of dysregulated ADAR expression. DAP3 mainly interacts with the deaminase domain of ADAR2 and represses editing via disrupting association of ADAR2 with its target transcripts (Han et al. 2020). ADAR1 and ADAR2 form homodimers or heterodimers in the cell, and homodimerization is indispensable for their editing activity (Cho et al. 2003;

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Poulsen et al. 2006). While ADAR1 monomers homodimerize via their third dsRBD, ADAR2 homodimerizes via its first dsRBD (Poulsen et al. 2006). DAP3 interacts with ADAR1 through the dsRBD of ADAR1, affecting A-to-I RNA editing by interrupting ADAR1 homodimerization (Han et al. 2020). Similarly, serine- and arginine-rich splicing factor 9 (SRSF9) has been shown to selectively suppress brain-specific A-to-I RNA editing in primates by disrupting ADAR2 homodimerization (Shanmugam et al. 2018). Recently, an unbiased screening using BioID followed by mass spectrometry of ADAR1 and ADAR2 interactors revealed that the family of proteins containing domains associated with zinc fingers (DZF), including interleukin enhancer-binding factor 3 (ILF3), interleukin enhancerbinding factor 2 (ILF2), spermatid perinuclear RNA-binding protein (STRBP), and zinc-finger RNA-binding protein (ZFR), were strong trans-editing regulators that interacted with ADARs in an RNA-dependent fashion in both HeLa and M17 neuroblastoma cells (Freund et al. 2020). This observation was consistent with a previous study where ILF2 and ILF3 were shown to form a protein complex that interacts with ADAR1 (Nie et al. 2005) and function as negative regulators by binding RNA in the close proximity of the editing sites and globally suppressing A-to-I RNA editing levels (Quinones-Valdez et al. 2019). Preference and efficiency of dsRNA recognition by ADARs depends on both RNA sequences and their secondary structures. RNA folding and unwinding processes are important in rendering stability to dsRNA structures. Consistent with this it might be speculated that RBPs such as RNA helicases could possibly modulate RNA editing by affecting RNA-protein interactions or causing structure-specific changes in the RNA landscape. DEAH box protein 15 (DDX15) was the first RNA helicase reported to be involved in the regulation of A-to-I editing in Caenorhabditis elegans (Tariq et al. 2013). Recently, DEAH box helicase 9 (DHX9), an RNA helicase, was reported as the first bidirectional regulator of A-to-I editing in human cancer, with an intriguing substrate-specific bias that governed its opposite regulatory effect (Hong et al. 2018). Computational analysis of ENCODE RNA-Seq datasets from HepG2 and K562 cell lines has reported several helicases including DDX6, DDX19B, and DDX47 which have considerable effects on A-to-I editing, but how this operates remains to be investigated (Quinones-Valdez et al. 2019). These results suggest a potential role for RNA helicases as A-to-I editing regulators and require further investigation and definition. In addition to the above subset, several A-to-I editing regulators have been identified with unknown or vague regulatory mechanisms. For instance, a recent study showed that the depletion of zinc-finger protein at 72D (Zn72D) led to reduction of both ADAR proteins but had no effect on mRNA expression in Drosophila (Sapiro et al. 2020). Zn72D interacts with ADAR in an RNA-dependent manner, but the detailed mechanism behind its effect on ADARs reduction after its depletion remains unclear. From a high-throughput screen performed in yeast-based reporter systems, DSS1 (26S proteasome complex subunit deleted in split-hand/split-foot 1) was identified as a potential enhancer and was further speculated to be part of an RNP interaction network through which it could help stimulate editing by stabilizing or facilitating substrate folding and recognition

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by ADARs (Garncarz et al. 2013). Another study utilized publicly available Genotype-Tissue Expression (GTEx) project datasets and extensively profiled Ato-I editing in humans, mice, and primates. This led to the identification of the aminoacyl tRNA synthetase complex interacting multifunctional protein 2 (AIMP2) as a top negative regulator of A-to-I editing (Tan et al. 2017). It was shown to promote the degradation of the ADAR proteins and reduced editing specifically in the muscle tissue. With accumulating evidence multiple insights can be gleaned with regard to the complex regulatory network of A-to-I RNA editing. While several ADARinteracting proteins have been identified, it is important to note that not all are potent A-to-I editing regulators (Quinones-Valdez et al. 2019). The ADAR family of proteins play a critical role in a wide spectrum of RNA-related processes (Bahn et al. 2015), in addition to their roles in mediating innate immunity (Lamers et al. 2019; Yanai et al. 2020). Therefore, it is not surprising that they exhibit promiscuous binding ability. These ADAR-interacting partners could therefore affect other aspects of ADARs, if not A-to-I editing. This understanding emphasizes that interactions with ADARs are certainly not a prerequisite for A-to-I editing regulation. Regulators can certainly modulate the editing landscape in an indirect fashion by skipping the necessity to bind ADARs. Similarly, binding of a regulator in the close proximity of an editing site need not always induce changes in the A-to-I editing landscape (Han et al. 2020; Shanmugam et al. 2018). The poor correlation between Alu binding and the extent of editing also highlights that this is an aspect that is not a mandate for editing regulation (Quinones-Valdez et al. 2019). It is highly likely that regulators can exist in a complex and exert combinatorial control on A-to-I editing in a cell type-specific fashion. Investigations on a large number of A-to-I editing regulators have been reported, and progress is significant. However, an understanding of the mechanism of how these regulators function in vivo to modulate the A-to-I RNA editome is still lacking. Future work should concentrate on how these regulators operate, particularly identifying if the presence of specific domains or the existence of sequence and structural preferences contributes toward regulation of A-to-I editing. The interplay between ADARs and their secondary regulators is clear for maintenance of normal physiology. Alterations to this intricate regulatory network can have dramatic effects resulting in aberrant conditions such as cancer and neurodegenerative disorders. Thus, a thorough understanding of the A-to-I editing regulatory landscape will hopefully reveal therapeutic insight which can be harnessed to target specific diseases.

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2 Cross Talk of RNA Editing with Other RNA Processing and Cellular Pathways RNA editing generally occurs co-transcriptionally, and hence it cross-talks with or affects multiple downstream RNA processing pathways such as splicing, miRNA biogenesis, RNA modification, mRNA export, and stability (Fig. 1). In addition, ADAR1/2 and RNA editing are implicated in apoptosis, innate immunity, and circadian rhythm.

2.1

Roles of ADARs and RNA Editing in miRNA Targeting and Biogenesis

Early studies showed that ADAR2-mediated editing in miRNAs is enriched in the seed sequence in the brain and miRNA hypoediting is observed in glioblastoma which display low expression of ADAR2, suggesting a functional role of RNA editing in miRNA (Paul et al. 2017; Li et al. 2018). A-to-I editing at miRNA seed region which includes nucleotides 2–8 from the 50 -terminus can result in retargeting (Bartel 2009). ADAR2-mediated editing reverts miRNA function from promoting to inhibiting cell invasion and migration in glioblastoma. As examples, editing of miR-376a* enables its targeting to autocrine motility factor receptor (AMFR) and disables its targeting to tumor suppressor Ras-related protein Rap-2a (RAP2A) (Choudhury et al. 2012). In addition, ADAR2-mediated editing retargets miR-589–3p from the tumor suppressor protocadherin 9 (PCDH9) to ADAM metallopeptidase domain 12 (ADAM12) which promotes cell invasion (Cesarini et al. 2018). Alternatively, ADAR proteins regulate miRNA biogenesis through RNA editingdependent and editing-independent mechanisms. Editing at the regions near the Drosha cleavage sites of primary miRNAs (pri-miRNAs) such as pri-miR142, pri-let-7, and pri-miR-26a impairs processing of pri- to precursor miRNA (pre-miRNA) by the Drosha-DiGeorge critical region-8 protein (DGCR8) complex (Yang et al. 2006; Zipeto et al. 2016; Jiang et al. 2019). Similarly, editing of pri-miR151 at sites near the Dicer cleavage site inhibits the Dicer-transactivation response element RNA-binding protein (TRBP) complex, leading to an accumulation of edited intermediate pre-miRNA (Kawahara et al. 2007). Additionally, ADAR2mediated editing decreased maturation and expression of onco-miRNAs, miR-222/ 221, and miR-21 in glioblastoma (Tomaselli et al. 2015). RNA editing at these sites likely modifies the RNA secondary structure and hence affects the processing of pri-/ pre-miRNA. A recent study showed that mismatches or wobble base pairs present in the seed region of pri-miRNA inhibit productive Drosha cleavage, of a role of RNA editing in miRNA biogenesis (Li et al. 2020). Nevertheless, not all the editing sites located within the miRNA seed region are inhibitory for biogenesis since edited

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Fig. 1 Cross talk of ADAR proteins and their mediated RNA editing with other RNA processing pathways. (a) Involvement of RNA editing and ADARs in miRNA biogenesis. Upper panel: editing at regions (depicted as red front) near to Drosha or Dicer cleavage sites (depicted as red and purple arrows, respectively) inhibits miRNA processing. In addition, extensive editing at the 5p arm likely promotes the RISC incorporation of 3p miRNA. Lower panel:

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mature miRNAs with retargeting ability were detected. The last step of miRNA biogenesis is asymmetric strand selection of single-stranded guide RNAs from miRNA duplex, which will be incorporated into Argonaute proteins (Ago) to form the RNA silencing complex (RISC). Correlations of editing levels at the 5p and 3p arms of miRNAs and ratio of 5p/3p suggested that RNA editing in one strand may promote the selection of the other strand for RISC incorporation (Li et al. 2018). In addition, highly edited pri-miRNAs are likely degraded by Tudor-SN (TSN) or endonuclease V (Yang et al. 2006; Vik et al. 2013), suggesting a potential role of RNA editing in regulating miRNA expression level. Alternatively, ADAR1 can regulate miRNA biogenesis via editing-independent mechanisms. ADAR1 binding to the stem regions of pri-miRNAs competitively precludes the binding of DGCR8 (Quick-Cleveland et al. 2014) and inhibits pri-miRNA processing (Chen et al. 2015). In contrast, ADAR1 enhances the cleavage efficacy of Dicer and RISC loading of miRNAs by forming a complex with Dicer, which in turn increases miRNA expression regulating gene expression (Bahn et al. 2015; Qi et al. 2017; Ota et al. 2013) (Fig. 1a). Direct protein-protein interaction through the second dsRBD (dsRBD2) of ADAR1, and both the domain of unknown function 283 (DUF283) and the DEAD-box RNA helicase domain of Dicer induce conformational changes in Dicer, resulting in augmentation of Dicer turnover rate (Ota et al. 2013). When in complex with Dicer, ADAR1 is not able to catalyze RNA editing as is not in homodimer form (Ota et al. 2013). RNA editing in the 30 -untranslated region (UTR) may create or abolish miRNA targeting sites and modulate gene expression during development or cancers (Soundararajan et al. 2015; Zhang et al. 2016; Roberts et al. 2018). For example, editing in the phosphatase and actin regulator 4 (PHACTR4) and human aryl hydrocarbon receptor (AhR) 3’-UTRs creates new miRNA targeting sites recognized by miRNA-196a-3p and mir-378 in gastric and breast cancer, respectively (Cho et al. 2018; Nakano et al. 2016). In contrast, RNA editing in the mouse double minute 2 homolog (MDM2) and dihydrofolate reductase (DHFR) 3’-UTRs abolishes miRNA targeting sites, leading to an upregulation of their expression (Jiang et al. 2019; Zhang et al. 2016; Nakano et al. 2017).

Fig. 1 (continued) independent of RNA editing, binding of ADAR1 to the stem region of pri-miRNA precludes DGCR8-Drosha complex from binding and inhibits the subsequent cleavage reaction, while ADAR1 forms a complex with Dicer to promote Dicer cleavage efficiency. (b) Role of ADARs and editing in splicing. Upper panel: editing at the BPS or 3’ss results in intron retention or exonization, respectively. Editing that modulates splicing enhancer or silencer could either promote or repress exon inclusion. Lower panel: binding of ADAR2 to dsRNA involving 3’ss could block the recruitment of U2AF65 and inhibit exon inclusion. (c) ADAR1 likely regulates alternative polyadenylation by blocking binding of polyadenylation factor, CFIm68. (d) Competitive regulation between A-to-I editing and m6A modification. (e) ADAR1 regulates apoptosis through competitively antagonizing STAU binding to the 3’-UTR of anti-/pro-apoptotic mRNAs to prevent mRNA export or degradation

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ADARs and Splicing Regulation

RNA editing and splicing share the largest overlap in time and space during processing of pre-mRNA (Bentley 2014). Previous studies have reported effects of RNA on splicing using knockdown or overexpression of ADAR proteins, showing hundreds of splicing events promoted or repressed by ADAR proteins (Solomon et al. 2013; Hsiao et al. 2018; Tang et al. 2020). An exon is defined by the 50 splice site (5’ss; GU) and 30 splice site (3’ss; AG) which are recognized by U1 small nuclear ribonucleoproteins (snRNP) and U2 small nuclear RNA auxiliary factor 65 kDa subunit (U2AF65), respectively, while the branch point sequence (BPS) adenosine is essential to initiate splicing catalytic reaction which removes introns. Hence, modification of any of these essential cis-acting elements could alter splicing. A-to-I editing disrupts splicing by mutating the BPS adenosine to inosine, resulting in intron retention (Beghini et al. 2000). Alternatively, RNA editing-created novel 3’ss (AA!AG) can lead to usage of cryptic 3’ss or exonization of Alu sequences (Rueter et al. 1999; Hsiao et al. 2018; Lev-Maor et al. 2007). Alu exonization could be beneficial for expanding protein functional domains throughout evolution, yet it could be toxic to disrupt transcriptome integrity (Zarnack et al. 2013). Cryptic splicing (3’ss or intron retention) caused by RNA editing could cause frameshift and nonsense-mediated decay (NMD) which regulates gene expression. RNA editing also modulates exon inclusion through modifying auxiliary cis-acting elements (Goldberg et al. 2017; Chen et al. 2018) as well as affecting RNA secondary structure (Flomen et al. 2004; Mazloomian and Meyer 2015). A number of alternative splicing events regulated by ADARs, such as heterogeneous nuclear ribonucleoprotein L-like (HNRPLL), protein tyrosine phosphatase non-receptor type 6 (PTPN6), and signal transducer and activator of transcription 3 (STAT3) are implicated in tumorigenesis (Beghini et al. 2000; Goldberg et al. 2017; Chen et al. 2018), suggesting an additional role of ADARs in cancer development. In addition, ADAR proteins could promote or repress splicing using editing-independent mechanisms. As such, ADAR2 precludes binding of U2AF65 to the 3’ss and prevents recognition from the spliceosome resulting in exon skipping (Tang et al. 2020) (Fig. 1b). ADAR-mediated aberrant splicing, both editing-dependent and editingindependent, contributes to tumorigenesis (Tang et al. 2020; Beghini et al. 2000; Goldberg et al. 2017; Chen et al. 2018).

2.3

ADAR Directly Promotes Proximal Polyadenylation Site

ADAR1 is also involved in regulating alternative 3’-UTR usage as shown by 3’-UTR lengthening or shortening in ADAR1 knockdown cells. ADAR1 CLIP data showed binding enrichment in both core and extension regions of lengthened 3’-UTRs but not in shortened 3’-UTRs detected in ADAR1 knockdown sample, and the majority of binding sites are non-Alu. These results suggest that ADAR1 directly

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regulates lengthened 3’-UTRs and indirectly modulates shortened 3’-UTRs, probably through regulating polyadenylation factors. Cleavage and polyadenylation specificity factor 68 KDa subunit (CFIm68), which promotes usage of distal cleavage site (Zhu et al. 2018), displayed reduced binding in the ADAR1-regulated 3’-UTR as compared to control 3’-UTRs, suggesting that ADAR1 may preclude CFIm68 binding to favor the usage of proximal cleavage site (Bahn et al. 2015) (Fig. 1c).

2.4

ADAR and N6-Methyladenosine

Other than A-to-I editing, there are more than 100 different post-synthetic modifications occurring on RNA. Among them, A-to-I editing and N6-methyladenosine (m6A), which is the methylation on the position 6 aminate group of adenosines, are the most abundant modification in eukaryote cells. m6A is “written” by the m6A methyltransferase complex which consists of methyltransferase like 3 (METTL3), METTL14, and WT1-associated protein (WTAP) and is “erased” by alphaketoglutarate-dependent dioxygenase FTO and RNA demethylase ALKBH5. m6A is decoded by a specific group of RNA-binding proteins known as “readers” such as YTH domain-containing family proteins (YTHDF and YTHDC) to mediate downstream pathways. Thus far, m6A has been reported to play an important role in regulating RNA processing including translation efficiency, alternative splicing, RNA export and stability, and innate immune (Wang et al. 2014, 2015; Winkler et al. 2019; Roignant and Soller 2017), but the interaction between m6A and A-to-I editing remains little studied. Because both m6A and A-to-I editing act on adenosine and m6A requires the aminate group while A-to-I editing removes it, it is clear that these two modifications cannot occur at the same position, and they may compete with one another within the overlapping modification region caused by a spatial effect (Fig. 1d). However, due to technical limitation, this hypothesis was not confirmed until recently following m6A RNA immunoprecipitation (RIP) followed by deep sequencing, Xiang et al. (2018) compared m6A-positive and m6A-negative RNA transcripts and discovered that ADARs preferentially edit m6A-negative transcripts. Knockdown of m6A “writers” METTL3 and METTL14 as well as “eraser” results in globally massive A-to-I editing changes being either upregulated or downregulated (Xiang et al. 2018), implicating that other than through m6A, these proteins may regulate A-to-I editing levels from other aspects. It was claimed that knockdown of METTL3 or METTL14 does not affect ADARs expression level (Xiang et al. 2018). However, a recent study suggested that knockdown of METTL3 repressed A-to-I editing by decreasing the expression of ADAR1/2 and increasing the expression of ADAR3 (Visvanathan et al. 2019). Further investigations on these aspects are required.

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ADAR and Apoptosis

ADAR1 plays a role in regulating apoptosis through its binding to the 3’-UTRs of anti-apoptotic transcripts to regulate mRNA stability and localization. As mentioned earlier, following UV irradiation or heat shock induced stress, phosphorylation of ADAR1 by MKK6-p38-MSK MAP kinases promotes its binding to exportin-5 (XPO5) and nuclear export. Once translocated to the cytoplasm, ADAR1 competitively inhibits binding of Staufen1 (STAU1) to the inverted Alu repeats (IRAlus) in the 30 -UTRs of anti-apoptotic transcripts. This prevents activation of Staufen1mediated mRNA decay and suppresses apoptosis of stressed cells (Sakurai et al. 2017; Park and Maquat 2013; Lucas et al. 2018) (Fig. 1e). In addition to mRNA decay, STAU1 promotes the export and translation of 30 -UTR IRAlus containing mRNAs and prevents protein kinase R (PKR)-mediated innate immune response to cytoplasmic 30 -UTR IRAlus (Elbarbary et al. 2013). ADAR1 competes for transcript occupancy with STAU1 to facilitate nuclear retention of the X-linked inhibitor of apoptosis protein (XIAP) and MDM2 mRNAs, resulting in downregulation of protein levels. The majority of mutual targets of both ADAR1 and STAU1 are implicated in apoptosis. The antagonistic function of ADAR1 and STAU1 in regulating expression of these apoptotic genes diversifies the mechanistic functions of ADAR1 in development and tumorigenesis (Yang et al. 2017).

2.6

ADAR and Innate Immunity

ADAR1 is known to be a repressor of innate immune response by blocking the MDA5-MAVS type I IFN signaling pathway. Retinoic acid-inducible gene I (RIG-I) and melanoma differentiation-associated gene 5 (MDA5), which belong to the RIGI-like receptor (RLR) family, are pattern-recognition receptors (PRRs) for dsRNA. RIG-I and MDA5 both contain a DExD/H-box RNA helicase domain for dsRNA binding and two N-terminal caspase activation and recruitment domains (CARDs) which recruits adaptor mitochondrial antiviral signaling protein (MAVS) upon activation. Activation of MAVS results in phosphorylation and nuclear translocation of transcription factor IRF3, which subsequently drives IFN expression (Dias Junior et al. 2019). ADAR1 knockout mice die at embryonic day 11.5–14 with widespread IFN1 overexpression and impaired hematopoiesis (Mannion et al. 2014; Hartner et al. 2009). This embryonic lethality phenotype was rescued by deletion of either MAVS or MDA5. Nevertheless the rescued mice died within days after birth (Mannion et al. 2014; Liddicoat et al. 2015; Pestal et al. 2015). Furthermore, embryonic lethality could not be rescued by editing deficient ADAR1 (ADAR1E861A/E861A), suggesting editing is indispensable for the immune suppression function. Nevertheless, ADAR1E861A/E861A mice with loss of MDA5 can survive past weaning, implying the importance of a non-editing function of ADAR1

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(Liddicoat et al. 2015). Naturally occurring mutations in ADAR1 are associated with Aicardi-Goutieres syndrome (AGS) which is a severe and rare infant encephalopathy characterized by uncontrolled type I IFN expression in the brain (Rice et al. 2012; Crow and Manel 2015). Promiscuous ADAR1 editing is proposed to disrupt endogenous dsRNAs to prevent activation of the MDA5-MAVS pathway. Alternatively, ADAR1 hyperedited RNAs (I/U RNAs) interact with MDA5 or RIG-I and inhibit their activation (Vitali and Scadden 2010). PKR, which is one of the interferonstimulated genes (ISGs), is activated by dsRNAs, leading to dimerization and autophosphorylation of PKR followed by phosphorylation of eukaryotic initiation factor 2ɑ (eIF2α) (Garcia et al. 2007). ADAR1 blocks PKR activation and the subsequent translation repression through editing-dependent and editingindependent fashions (Vitali and Scadden 2010). Additionally, ADAR1 can inhibit the IFN-inducible oligoadenylate synthetase (OAS)-RNase L pathway which is activated by dsRNA (Li et al. 2017). Following dsRNA binding, OAS proteins produce 20 ,50 -oligoadenylates (2-5A) which then activates RNase L. RNase L cleaves both endogenous and viral RNAs, leading to translation arrest, apoptosis, and IFN production preventing viral replication (Chakrabarti et al. 2011).

2.7

ADAR2 and Circadian Rhythm

The circadian clock is sustained by the transcriptional-translational feedback loops (TTFLs), in which heterodimers of circadian locomotor output cycles protein kaput (CLOCK) and brain and muscle ARNT-like 1 (BMAL1) proteins act via enhancer box (E-box) regulatory sequences to initiate daytime expression of period (PER) and cryptochrome proteins (CRY). In turn, PER and CRY suppress CLOCK-BMAL1 activity at their E-boxes to reduce their own gene transcription and subsequent degradation of PER and CRY in the circadian night allowing reinitiation of the circadian cycle (Takahashi 2017). ADAR2 and ADAR2-mediated RNA editing are important in circadian rhythm as ADAR2-knockout mice exhibited short-period rhythms in locomotor activity and gene expression as well as abnormal accumulation of CRY2 (von Schantz and Archer 2003; Terajima et al. 2017). ADAR2 suppresses translation of CRY2 through promoting miRNA biogenesis including let-7. In addition, circadian ADAR2-mediated A-to-I editing in a variety of transcripts is regulated by the expression of ADAR2 whose expression is controlled by CLOCKBMAL1 (Terajima et al. 2017). The diverse roles of RNA editing make it a potent target for cancer, viral infection, and immune diseases. Nevertheless, there are many questions left to be answered such as the editing regulatory mechanism, substrate specificity, and detailed mechanistic function of ADARs in other cellular pathways. Continued study of these topics will be beneficial for therapeutic development.

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RNA-Mediated Metabolic Defects in Microsatellite Expansion Diseases Nan Zhang

Abstract Many incurable neurological and neuromuscular diseases are caused by microsatellite expansions consisted of 3–6 nucleotide repeat units. The expanded RNA and/or the translated proteins cause metabolic defects and toxicity at multiple cellular levels, and such dyshomeostasis has dramatic impacts on neuronal health. This review sets to provide new insights into RNA-mediated toxicity, repeatassociated non-ATG translation, microRNA-mediated pathology, nucleocytoplasmic transport alterations, and liquid-liquid phase separation in microsatellite expansion diseases. Keywords Microsatellite expansion diseases · Polyglutamine diseases · Neurodegeneration · CAG repeat · Huntington’s diseases · RAN translation · Liquid-liquid phase separation · RNA foci · Stress granules

1 Overview of Microsatellite Expansion Diseases Repetitive DNA comprises over half of the human genome and can be broadly categorized into two classes: tandem repeats and interspersed repeats (Hannan 2018; Treangen and Salzberg 2011). A tandem repeat is a series of consecutive blocks of nucleotides confined to a specific locus (Dumbovic et al. 2017), within which there are three subclasses: satellites (can occupy 100 kb to >1 Mb, commonly found near centrosomes and telomeres), minisatellites [repeat units range from 9 to 80 nucleotides (nt)], and microsatellites [repeat units are typically 200) expansion in the 50 -UTR of the FMR1 gene that encodes FMRP (fragile X mental retardation protein) (Kremer et al. 1991; Oberle et al. 1991). The CGG expansion induces silencing of FMR1 by DNA hypermethylation and histone hypoacetylation and hypermethylation (Oberle et al. 1991; Coffee et al. 1999; Kumari et al. 2012; Sutcliffe et al. 1992), leading to an absence of FMRP in patients. FMRP is essential for mRNA transport, synaptic plasticity, stress granule formation, DNA damage response, and RNA processing (Alpatov et al. 2014; Anderson and Kedersha 2006; Dury et al. 2013; Hagerman et al. 2017; He and Portera-Cailliau 2013). 3. Toxic RNA GOF, as exemplified by myotonic dystrophy type 1 (DM1) and C9orf72-amyotrophic lateral sclerosis and frontotemporal dementia (ALS/FTD). The expanded repeats (CTG repeats in the 30 -UTR of DMPK in DM1 and GGGGCC repeats in intron 1 of C9orf72 in C9orf72-ALS/FTD) form nuclear RNA aggregates (foci) that sequester various RNA-binding proteins (RBPs) from performing their normal functions (DeJesus-Hernandez et al. 2011; Mankodi et al. 2001; Miller et al. 2000; Renton et al. 2011). With the recent discovery of the uncanonical repeat-associated non-AUG (RAN) translation from many bidirectionally transcribed repeat expansions (Cleary et al. 2018; Cleary and Ranum 2017), the distinction between the three pathological categories are blurred, and their contribution to neurodegeneration may be combinatorial in some cases.

2 RNA Structure and RNA-Mediated Toxicity The majority of microsatellite expansions are CG-rich repeats that can readily form mismatched hairpin loops and contribute to RNA foci formation in the nucleus (Conlon et al. 2016; de Mezer et al. 2011; Kiliszek and Rypniewski 2014). These

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nuclear RNA foci may drive pathogenesis through sequestration and nuclear retention of specific RBPs. RBP sequestration may occur in a stoichiometric fashion where repeat expansion creates an excess of RBP binding sites, or RBPs may have a higher affinity for the secondary or tertiary structure of the expanded RNA. The sequestration model is well illustrated in DM1. DM1 is a multisystemic disease that targets tissues including the skeletal, cardiac, and smooth muscles, central nervous system (CNS), and eyes. The CUG-expanded RNA of the DMPK gene forms hairpins and foci that sequester a class of splicing regulatory RBPs—muscleblind 1–3 (MBNL1–3)—at the periphery of the nuclear splicing speckles and block their splicing activity (Goodwin et al. 2015; Holt et al. 2007; Mooers et al. 2005). MBNL1 actively participates in RNA foci formation by binding to distorted GC bases or unpaired UU bases (Ho et al. 2005; Konieczny et al. 2014; Lambert et al. 2014; Yildirim et al. 2015). Depletion of MBNL1 has been linked to aberrant splicing as well as mRNA mislocalization and microRNA misprocessing (Rau et al. 2011; Taliaferro et al. 2016). Functional disruption of MBNL2 has been associated with neurological features such as learning difficulties and daytime sleepiness (Charizanis et al. 2012), while depletion of MBNL3 causes defects in muscle regeneration and a spectrum of age-related pathologies (such as abnormal glucose metabolism) (Choi et al. 2016; Poulos et al. 2013). CUG foci formation also leads to protein kinase C (PKC)-mediated hyperphosphorylation and elevated expression of CUG-binding protein 1 (CUGBP1 or CELF1) (Kuyumcu-Martinez et al. 2007). Collectively, the inhibition of MBNL1 and upregulation of CUGBP1 in DM1 drive splicing of a variety of transcripts from adult to fetal isoforms. Many of the misspliced genes are components of the sodium/calcium current regulation, intra-/intercellular transport, and sarcomere/cytoskeleton structure and function (Chau and Kalsotra 2015; Dixon et al. 2015), thus unable to fulfill their normal cellular functions. Similar to CUGBP1, a recent study shows that overexpression of heterogeneous nuclear ribonucleoprotein A1 (HNRNP A1)—not previously linked to DM1—also triggers DM1 muscle pathology and a shift of RNA targets to an earlier developmental pattern in mice (Li et al. 2020). Antisense CAG foci have also been reported in adult and congenital DM1 patients and mice (Huguet et al. 2012; Michel et al. 2015). These foci do not appear to colocalize with their sense counterparts nor sequester any MBNL. Whether they are regulated by the same ribonucleoprotein complex as the sense foci and contribute to disease pathology is largely unknown (Paul et al. 2011). Repeat expansions with a high guanine content can form G-quadruplexes in which four guanines form a planar tetrad, and two or more tetrads stack to form a G-quadruplex (Fig. 2) (Burge et al. 2006). The most common mutation responsible for inherited ALS and FTD consists of a GGGGCC expansion in intron 1 of the C9orf72 gene (DeJesus-Hernandez et al. 2011; Renton et al. 2011), and the expanded GGGGCC repeats form G-quadruplexes (Fratta et al. 2012). An RNA toxic role has been implicated in C9orf72-ALS/FTD, because (1) both sense and antisense nuclear RNA foci have been identified in patient tissues, induced pluripotent stem cells (iPSCs), iPSC-derived neurons, Drosophila, and mice (CooperKnock et al. 2015; Donnelly et al. 2013; Jiang et al. 2016; Lagier-Tourenne et al. 2013; Liu et al. 2016; Peters et al. 2015; Zu et al. 2011); (2) the expanded RNA

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sequesters a variety of RBPs, including Purine-rich element-binding protein-alpha (Purα), adenosine deaminase B2 (ADARB2), AIY/REF export factor (ALYREF), heterogeneous nuclear ribonucleoprotein F (HNRNP F), HNRNP A1, HNRNP H, serine and arginine-rich splicing factor 2 (SRSF2), among others (Cooper-Knock et al. 2015, 2014; Donnelly et al. 2013; Sareen et al. 2013; Zhang et al. 2015); and (3) the expanded sense RNA alone is sufficient to recapitulate neurodegeneration in Drosophila and mice (Liu et al. 2016; Xu et al. 2013). Interestingly, antisense CCCCGG RNA foci have been shown to preferentially accumulate in vulnerable cell types (Liu et al. 2016), and correlate with a nuclear loss of TAR DNA binding protein 43 (TDP-43) (Cooper-Knock et al. 2015). Given that the C9ORF72 protein performs important functions in autophagy and its inactivation provokes or accelerates early death in mice (Nassif et al. 2017; Zhu et al. 2020), a protein LOF may synergize with RNA GOF toxicity in C9orf72-ALS/FTD. The GGGGCC repeat expansions also form R loops at the site of transcription where the nascent RNA hybridizes to the complementary DNA strand (Fig. 2) (Lin et al. 2010b; Reddy et al. 2011). R loops have also been observed in FXS (CGG repeats) and FRDA (GAA repeat) (Groh et al. 2014) and may contribute to disease phenotype by impeding transcription (Haeusler et al. 2014), blocking translation (Huertas and Aguilera 2003), disrupting chromatin remodeling (Castellano-Pozo et al. 2013), promoting repeat instability (Lin et al. 2010b), and causing nucleolar stress (Haeusler et al. 2014).

3 Repeat Associated Non-AUG Translation In addition to causing RNA/protein-related toxicity, microsatellite expansions also support an uncanonical mode of translation initiation—RAN translation—where repetitive peptides are synthesized from the expanded RNA in the absence of an AUG start codon (Zu et al. 2011). This paradigm-shifting discovery was first made in SCA8 where the antisense transcript from ATXN8 (ATXN8OS) contains a short AUG-driven open reading frame encompassing a CAG expansion. The removal of the AUG or placement of stop codons upstream of the CAG repeat did not abolish polyQ peptide synthesis. Instead, translation occurred in all three reading frames of the repeat, additionally generating non-AUG initiated polyserine (polyS from the AGC frame) and polyalanine (polyA from the GCA frame) (Zu et al. 2011). Because many microsatellite expansions can be bidirectionally transcribed (Banez-Coronel and Ranum 2019; Batra et al. 2010; Gendron et al. 2013; Zu et al. 2013; Mori et al. 2013a), both sense and antisense repeats could potentially give rise to a cocktail of six RAN peptides, thus adding another layer of complexity to disease mechanisms. Currently, twelve microsatellite expansion diseases undergo RAN translation in cell or animal models, including DM1, DM2, C9orf72-ALF/FTD, fragile X tremor ataxia syndrome (FXTAS), fragile X premature ovarian insufficiency (FXPOI), HD, Huntington disease-like 2 (HDL2), SCA2, SCA3, SCA8, SCA31, and Fuchs endothelial corneal dystrophy (FECD) (Zu et al. 2011; Banez-Coronel and Ranum

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2019; Ayhan et al. 2018; Ishiguro et al. 2017; Scoles et al. 2015; Soragni et al. 2018; Todd et al. 2013; Zhang and Ashizawa 2017). Very little is known about the underlying mechanism of RAN translation initiation. It is possible that it shares common features with the m7G cap-dependent scanning mechanism and/or a cap-independent internal ribosome initiation site (IRES)-like mechanism for initiation. The scanning mechanism begins when a 40S ribosomal subunit binds to the ternary complex (comprised of the methionine initiator tRNA and GTP-bound eIF2αβγ), forming the 43S preinitiation complex (PIC). The 43S PIC is recruited to the 50 -m7G cap of the transcript by the eIF4F complex (composed of the cap-binding protein eIF4E, the helicase eIF4A, and the scaffold protein eIF4G) and scans from 50 to 30 in an ATP-dependent fashion to locate the first AUG start codon (Kearse and Wilusz 2017). While originally described for viral elements, the IRES initiation occurs on highly structured RNA and directly recruits the 43S PIC to the transcript independent of the m7G cap (Kearse and Wilusz 2017). For FXTAS, RAN translation from CGG repeats in the 50 leader of FMR1 requires a functional 50 -m7G cap and is eIF4E- and eIF4Adependent, suggesting that it resembles the canonical scanning mechanism except that a near-cognate start codon (ACG or GUG) is used for initiation (Kearse et al. 2016). RAN translation from GGGGCC repeats in C9orf72-ALS/FTD is more complex. Initiation across expanded GGGGCC repeats is cap-, eIF4A-, and eIF4E-dependent and utilizes an upstream near-cognate CUG codon (Green et al. 2017; Tabet et al. 2018). However, an IRES-driven translation has also been shown to operate on uncapped, spliced GGGGCC repeats following its export to the cytoplasm (Cheng et al. 2018). Thus, RAN translation initiation may be different across repeat types or even in different reading frames of the same repeat. RAN translation may contribute to overall disease pathology in FXTAS, HD, and C9orf72-ALS/FTD. FXTAS is caused by a 55–200 CGG repeat in the 50 -UTR of FMR1 on the X chromosome and primarily affects older men who develop progressive tremor, gait ataxia, parkinsonism, and dementia (Hagerman and Hagerman 2016; Hagerman et al. 2001; Jacquemont et al. 2007). A pathological hallmark of FXTAS is the accumulation of ubiquitin-positive nuclear inclusions in both neurons and astrocytes (Greco et al. 2006, 2002). The expanded CGG repeats undergo RAN translation, producing polyglycine (polyG in the GGC frame) and polyalanine (polyA in the GCG frame) but not polyarginine (polyR in the CGG frame) in cell culture (Banez-Coronel and Ranum 2019). The polyG peptide colocalizes with p62and ubiquitin-positive inclusions in patient brains and cause neuronal cell death possibly by interfering with proteasome function and nuclear lamina integrity (Todd et al. 2013; Buijsen et al. 2016; Sellier et al. 2017). In the antisense direction, polyproline (polyP from the CCG frame), polyR (from the CGC frame), and polyA (from the GCC frame) are expressed from the expanded CCG RNA. Similar to polyG, both polyP and polyA accumulate as ubiquitin-positive inclusions in patient brains, suggesting their contribution to disease (Krans et al. 2016). Clear evidence supporting the role of RAN translation in neurotoxicity and pathology in FXTAS comes from mouse studies. Mice expressing both expanded CGG RNA and polyG showed inclusion formation, motor dysfunction, Purkinje cell loss, and

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decreased survival (Sellier et al. 2017). In contrast, no significant pathology was observed in mice expressing the same CGG expansion but unable to produce polyG due to the absence of near-cognate initiation sites (Sellier et al. 2017). HD is an autosomal dominant disorder caused by a CAG repeat expansion in exon 1 of Huntingtin (HTT). Patients generally exhibit involuntary movements and cognitive and psychiatric alterations with adult onset (Dickey and La Spada 2018). Although a mutant HTT GOF primarily defines the disease pathology, four RAN translation products have been detected in patient brains: polyA (from the GCA frame) and polyS (from the AGC from) from the sense transcript and polyleucine (polyL from the CUG frame) and polycysteine (polyC from the UGC frame) from the antisense transcript (Banez-Coronel et al. 2015). These RAN products abundantly accumulate in most affected HD brain regions, including the caudate/putamen and, in juvenile cases, the cerebellum, as well as in brain regions that show atrophy, astrogliosis, and Caspase 3 and microglial activation (Banez-Coronel et al. 2015). Moreover, prominent RAN-positive staining was observed in white matter regions where polyQ aggregates are absent or minimal, suggesting a role of RAN products in white matter abnormalities observed in HD patients (Banez-Coronel et al. 2015; Bohanna et al. 2011; Fennema-Notestine et al. 2004; Paulsen et al. 2010; Reading et al. 2005). In C9orf72-ALS/FTD, the GGGGCC expanded RNA is transcribed bidirectionally to produce five potential RAN products, polyglycine/alanine (polyGA), polyglycine/arginine (polyGR), polyproline/alanine (polyPA), polyproline/arginine (polyPR), and polyglycine/proline (polyGP), with the most efficient expression from the polyGA reading frame and lesser expression in the polyGR and polyGP reading frames (Green et al. 2017; Tabet et al. 2018; Sonobe et al. 2018). The C9orf72-RAN products colocalize with p62 but are TDP-43 negative (Gendron et al. 2013; Ash et al. 2013; Mori et al. 2013b). PolyGR and polyPR are extremely toxic to multiple model systems including cultured cells, zebrafish, Drosophila, and mice (BanezCoronel and Ranum 2019). It has been demonstrated that polyGR and polyPR mimic pre-mRNA splicing RBPs that contain serine:arginine (SR) repeats and bind to HNRNP A2 hydrogel. When applied to cells, these two RAN peptides translocate to the nucleus, bind to the nucleolus, and impair both pre-mRNA splicing and ribosomal RNA biogenesis, leading to cell death (Fig. 2) (Kwon et al. 2014). However, whether RAN translation is the primary driver of the disease is still under debate. For instance, RAN aggregates are relatively rare in ALS vulnerable cells yet abundant in non-affected brain regions (Banez-Coronel and Ranum 2019). Conflicting results were also observed in BAC transgenic mice when C9orf72-RAN products were expressed at endogenous levels. Phenotypes due to RAN accumulation in different mouse models range from no apparent manifestations to neurodegeneration with earlier disease onset (Jiang et al. 2016; Liu et al. 2016; Peters et al. 2015; O’Rourke et al. 2015). RAN translation may exacerbate disease progression through a feed-forward loop (Green et al. 2017; Cheng et al. 2018): both FXTAS- and C9orf72-RAN products induce endoplasmic reticulum (ER) and oxidative stresses that lead to increased eIF2α phosphorylation and global translation repression, while their own expression increases upon ER stress and eIF2α

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phosphorylation. Additionally, increased R-loop formation and DNA breaks as well as defective DNA damage repair associated with the expanded GGGGCC repeats may feed into this loop by further stressing the cell (Cleary et al. 2018).

4 Emerging Pathological Roles of microRNA microRNAs (or miRNAs, 21 nt) are small non-coding RNAs that act as posttranslational regulators of genes. They are transcribed in the nucleus by polymerase II or III as capped and polyadenylated primary-miRNA (pri-miRNA). The pri-miRNA is then processed in the nucleus by the Drosha/DGCR8 microprocessor complex into a shorter hairpin-containing precursor-miRNA (pre-miRNA). Interestingly, Drosha but not DGCR8 is mislocalized with polyGA and polyGP to form cytoplasmic inclusions in the hippocampus, frontal cortex, and cerebellum in C9orf72-ALS/ FTD patients, suggesting that disrupted pre-miRNA processing may contribute to disease pathology (Porta et al. 2015). After exported to the cytoplasm by Exportin 5, the pre-miRNA is cleaved by Dicer that removes the hairpin. The guide strand of the miRNA duplex is then loaded onto the RNA-induced silencing complex (RISC), and depending on the level of sequence complementarity, the target mRNA is silenced via either translational repression or mRNA degradation through unknown mechanisms (Aleman et al. 2007; Eulalio et al. 2008; Filipowicz et al. 2008). The brain expresses more miRNAs than any other organ with great regional and spatiotemporal specificity (Rajgor 2018). Given the key regulatory roles of miRNA in neuronal development and plasticity (McNeill and Van Vactor 2012; Rajgor and Hanley 2016; Saraiva et al. 2017), it is not surprising that their malfunction contributes to various age-dependent neurodegenerative diseases (Maciotta et al. 2013; Shah et al. 2018). Aberrant expression or dysregulation of miRNA has been linked to the pathology of multiple polyQ diseases. One of the molecular phenotypes in HD patients is transcriptional misregulation in the striatum and distinct cortical regions (Hodges et al. 2006). Hoss et al. identified five significantly upregulated miRNAs (miRNA-10b, miRNA-196a, miRNA-615, miRNA-196b, and miRNA-483) in the prefrontal cortex from 28 HD patients (Hoss et al. 2015). Marti et al. found that miRNA-100, miRNA-151-3p, miRNA-16, miRNA-219-2-3p, miRNA-27b, miRNA-451, and miRNA-92a were upregulated in the striatum and frontal cortex from 5 HD patients, while in the same regions miRNA-128, miRNA-139-3p, miRNA-222, miRNA-382, miRNA-433, and miRNA-485-3p were significantly downregulated (Marti et al. 2010). By analyzing Brodmann’s area four from 19 HD cortices, Packer et al. found that miRNA-132 was upregulated, while miRNA-9/9*, miRNA-29b, and miRNA-124a were downregulated (Packer et al. 2008). The transcription repressor—RE1-silencing transcription factor (REST)—binds to neuron-restrictive silencer elements (NRSEs) in more than 1800 human promoters, many of which drive gene expression in neuronal function, survival, and differentiation (Abrajano et al. 2009; Bruce et al. 2004; Zuccato et al. 2003). One of

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the REST targets is the brain-derived neurotrophic factor (BDNF)—an important survival factor for the striatal neurons that die in HD. In normal individuals, REST is bound by wildtype HTT and sequestered in the cytoplasm, making it unavailable for NRSE occupancy and gene repression in the nucleus (Fig. 2) (Landles and Bates 2004). In HD patients, the mutant HTT aggregates and fails to engage REST, leading to an increased nuclear presence of REST and shutdown of BDNF production. A similar regulatory mechanism may also apply to many abnormally expressed miRNAs in HD brains—such as miRNA-132, miRNA-124, miRNA-9/9*, miRNA-29a, miRNA-29b, and miRNA-330—because they are all targets of REST. For instance, miRNA-132 is vital for neuronal outgrowth and sprouting and is significantly elevated in HD brains (Packer et al. 2008). Its overexpression, whether due to REST activation by mutant HTT or by downregulation of methyl CpG-binding protein 2 (MeCP2), has been linked to BDNF reduction (Su et al. 2015; Yao et al. 2017). miRNA-124 is a well-known regulator of the REST complex that represses neuronal-specific genes in nonneuronal cells, including miRNA-124 itself (Ballas et al. 2005; Conaco et al. 2006). miRNA-124, along with miRNA-9/9*, promotes neurogenesis by synergizing with neuronal-specific transcription factors (Yoo et al. 2011). Force expression of miRNA-124 drives trans-differentiation of C2C12 myoblasts to neuronal-like cells (Watanabe et al. 2004) and murine fibroblasts to neurons (Xue et al. 2013). This conversion process in murine is facilitated by the REST-miRNA-124-PTBP1 (polypyrimidine tract binding protein 1) loop where expression of miRNA-124 knocks down PTBP1 protein levels and dismantles the REST complex, thereby initiating neuronal-specific splicing events (Hu et al. 2018). For human cells to become fully committed to neuronal fate, the maturation process requires a second regulatory loop via BRN2-PTBP2-miRNA-9 (Xue et al. 2016). BRN2/POU3F2 (POU class 3 homeobox 2) transcriptionally activates miRNA-9, which in turn knocks down PTBP2 protein levels. Given that both miRNA-124 and miRNA-9 are significantly reduced in HD brains (Packer et al. 2008), it is possible that neuronal-specific conversion and maturation are inhibited in HD patients, thus contributing to neurodegeneration and brain atrophy. Interestingly, the miRNA-124-PTBP1 regulatory loop has been successfully targeted for phenotypic improvement in mice. HD mice injected with miRNA-124 showed increased neurogenesis in the striatum and cortex and slowed down disease progression (Liu et al. 2015). In a different study, downregulation of Ptbp1 converted glia to dopaminergic neurons and alleviated motor dysfunctions in a Parkinson’s mouse model (Zhou et al. 2020). miRNAs also play important roles in SCA3. SCA3 is the most common dominantly inherited ataxia in the world and is caused by a CAG repeat expansion in exon 10 of ATXN3 (Paulson 2009, 2012). By analyzing SCA3 patient serums, Shi et al. found that miRNA-25, miRNA-29a, and miRNA-125b were significantly reduced, while miRNA-34b expression was elevated (Shi et al. 2014). miRNA-25 binds to the 3’-UTR of ATXN3 mRNA and reduces both wild type and mutant ATXN3 protein levels. Expression of miRNA-25 increases SCA3 cell viability and suppresses early apoptosis (Huang et al. 2014). miRNA-9, miRNA-181a, and miRNA-494 also bind

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to the 30 -UTR of ATXN3, and their expression is downregulated in SCA3 cell and mouse models (Carmona et al. 2017). Overexpression of the three miRNAs through viral delivery leads to mutant ATXN3 reduction and neuronal rescue. Taken together, miRNAs not only contribute to microsatellite disease pathology but also have great potential as diagnostic and progression biomarkers due to their stability and abundance in biofluids (Dong and Cong 2019). However, caution should be taken using transgenic animals for miRNA profiling, because miRNA expression varies significantly from case to case and throughout the development of the animal [as observed in HD YAC128 and R6/2 mice (Lee et al. 2011)]; thus, a single miRNA could have differential impacts on disease progression.

5 Defects in Nucleocytoplasmic Transport The diverse functions of mRNA are in part determined by their subcellular localization. The nucleocytoplasmic transport system that shuttles RNA through the nuclear pore complex (NPC) is particularly important in highly compartmentalized and morphologically complex cells such as neurons. The bulk mRNA export from the nucleus uses the nuclear export factor 1 (NXF1)-mediated pathway. This process begins by deposition of the translation export complex (TREX) at the 50 -cap of the nascent mRNA (Masuda et al. 2005). TREX recruits NXF1 through its subunit ALYREF—a protein that associates with the 50 -end of mRNA to prevent degradation by nuclear exosomes and is sequestered by the expanded GGGGCC repeats in C9orf72-ALS/FTD (Cooper-Knock et al. 2014; Freibaum et al. 2015). NXF1 then facilitates NPC docking and RNA translocation by interacting with phenylalanineglycine repeat-containing nups (FG-nups) (Guo et al. 2019; Kim and Taylor 2017). The FG-nups reversibly crosslink with one another and form a hydrogel that functions as a sieve for nuclear transport receptors and their cargos. Importin and exportin receptors regulate cargo transport with the help of the RanGTP-RanGDP system (Hoelz et al. 2011). Ran (Ras-related nuclear protein) is a small GTPase that can switch between a GTP- (RanGTP) or GDP-bound (RanGDP) state. The nucleotide state of Ran is determined by GEF (guanine exchange factor) and GAP (guanine-activating protein). RanGEF stimulates the exchange of GDP for GTP and along with RanGTP is enriched in the nucleus. RanGAP accelerates GTP hydrolysis and along with RanGDP is enriched in the cytoplasm. Segregation of RanGEF and RanGAP sets up a gradient of RanGTP to dictate the directionality of nucleocytoplasmic transport. Several studies have revealed a novel disease mechanism stemmed from nucleocytoplasmic transport dysregulation on many levels in C9orf72-ALS/FTD model systems. (1) The nuclear export adaptor SR-rich splicing factor 1 (SRSF1) mediates mRNA nuclear export through binding to NXF1. Sequestration of SRSF1 by the expanded C9orf72 RNA inappropriately licensed repeat RNA export for RAN translation in the cytoplasm. Thus, depleting SRSF1 or preventing its RNA

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sequestration and interaction with NXF1 alleviates RAN-mediated neurotoxicity in Drosophila and patient-derived motor neurons (Hautbergue et al. 2017). (2) Screening of disease modifiers in Drosophila expressing (GGGGCC)30 identified RanGAP as a potent suppressor of neurodegeneration (Zhang et al. 2015). Direct binding of RanGAP to expanded C9orf72 repeats causes RanGAP mislocalization, disruption in Ran gradient, and NPC pathology. (3) RanGAP1 (the human orthologue of fly RanGAP) and the nuclear pore protein POM121 mislocalize and aggregate with polyGA peptides in a mouse model (Fig. 2) (Zhang et al. 2016). (4) Screening in Drosophila expressing (GGGGCC)58 identified 18 disease modifiers that are components of the NPC and nuclear exosomes (Freibaum et al. 2015). Two of these modifiers are Nup50 (human orthologue NUP50, which promotes protein nuclear import) and Ref1 (human orthologue ALYREF, which promotes mRNA nuclear export). Deletion of Nup50 enhanced the degenerative rough eye phenotype, while deletion of Ref1 suppressed (GGGGCC)58-induced toxicity. However, the important question remains: Do the expanded GGGGCC RNA or RAN products cause the nuclear pore defects? (5) An attempt to answer this question was carried out in yeast expressing codon-optimized polyPA without using the expanded GGGGCC repeats (Jovicic et al. 2015). Many identified genes that are RAN toxicity modifiers are also components of the nucleocytoplasmic transport as well as ribosome biogenesis, ubiquitination, proteasome, mitochondria, transcription, among others. (6) A similar study in Drosophila that specifically focused on polyPR identified multiple RAN toxicity modifiers encoding the NPC, importins/exportins, RanGTP regulators, and arginine methyltransferases (Boeynaems et al. 2016). In contrast to the previous fly study (Freibaum et al. 2015), Nup50 was identified as a suppressor rather than an enhancer in this work. (7) RAN peptides (PolyPRs) bind to the FG repeats in Nup54 and Nup98 and block the central channel of the NPC in Xenopus laevis (Fig. 2) (Shi et al. 2017). Defects in nucleocytoplasmic transport have also been observed in HD. Grima et al. demonstrated that NUP62—a FG-nup essential for protein nuclear import— and RanGAP1 aggregated with mutant HTT in R6/2 and zQ175 mice (Grima et al. 2017). Both proteins mislocalized in HD brain tissue and patient iPSC-derived neurons. Overexpression of RanGAP1 conferred neuroprotection, consistent with its toxicity suppressor function as observed in flies (Zhang et al. 2015). In this study, HD RAN peptides also contribute to defects in nuclear import. Mutant HTT can partially sequester nucleocytoplasmic transport factors—Gle1 and RanGAP1—in a dosage-dependent manner as shown in HTTQ7/Q7, HTTQ7/Q175, and HTTQ175/Q175 mice (Gasset-Rosa et al. 2017). In aged mice, RanGAP1-positive aggregates shifted to perinuclear and cytoplasmic areas. Mutant HTT markedly accelerated defects in nuclear envelop integrity, nucleocytoplasmic transport, and mRNA accumulation in the striatum and cortex of mice. Taken together, nucleocytoplasmic transport dysfunction may contribute to age-related neurodegeneration and aging phenotypes across multiple microsatellite expansion diseases.

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6 Liquid-Liquid Phase Separation (LLPS) and Stress Granules Cells use phase-separated RNA-RBP membraneless organelles to carry out important functions such as stress control (stress granules, SG), RNA degradation (P bodies), RNA transport (transport granules), spliceosome maturation (Cajal bodies), and ribosomal RNA biogenesis (nucleoli) (Wolozin and Ivanov 2019). The formation of liquid droplets of membraneless organelles that partition from the surrounding medium is mediated by intrinsically disordered regions (IDRs) in RBPs. IDRs typically have low sequence complexity and are enriched in Ala, Arg, Gly, Gln, Ser, Glu, Lys, and Pro (Uversky and Dunker 2010). The binding of an IDR-containing RBP to its partners through weak multivalent interactions drives up the local concentration of these proteins and favors phase separation. In addition, some IDR-containing proteins can self-assemble—this is particularly relevant to polyQ proteins. When the local concentration of IDR-containing proteins is too high or the local interactions are too strong, the liquid droplets may shift from a highly dynamic state to less dynamic hydrogels and ultimately to rigid irreversible aggregates (Wolozin and Ivanov 2019; Rodriguez and Todd 2019). This transition process may occur over time and contribute to neurodegeneration. Like RBPs, RNA can form foci through phase separation. The expanded CUG, CAG, and GGGGCC RNA can achieve gelation through multivalent base paring without requiring protein components in vitro or in cells (Jain and Vale 2017). RNA gelation only occurs at a boundary condition of increasing valency, which could explain why many microsatellite disease phenotypes are only triggered above a certain repeat threshold. In addition, the RNA recognition motifs (RRMs) within RBPs also regulate phase separation because they are the sites of protein and RNA binding and posttranslational modifications. For instance, arginine methylation of the RRM in FUS (fused in sarcoma) promotes chaperone binding, which prevents phase separation (Wolozin and Ivanov 2019). RNA also promotes phase separation of the RRM domains, possibly by engaging RRMs in a multivalent fashion via several RNA motifs. The above observations suggest that both protein-protein and RNA-protein interactions synergistically contribute to phase separation of membraneless organelles (Li et al. 2012; Molliex et al. 2015). Stress granules are specialized membraneless organelles that prevent the translation of unnecessary transcripts during stress and triage transcripts for degradation or translation when the stressor has passed (Anderson and Kedersha 2006). During SG formation, many nuclear RBPs—such as T cell intracellular antigen 1 (TIA1), FUS, TDP-43, ataxin 2 (ATXN2)—translocate to the cytoplasm and spread diffusely in neurons without phase separation. Core SGs—consisting of stalled PICs, mRNA, and specific RBPs such as FMRP, TIA1-related protein (TIAR), Ras GTP-activating protein-binding protein 1 (G3BP1) and TIA—begin to form (Wolozin and Ivanov 2019). Since the translationally stalled RNA is not decorated with ribosomes, it can nucleate a “shell” of secondary RBPs. After the stressor has passed, SG resolution begins with disaggregates (such as valosin-containing protein (VCP) or nuclear

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import receptors (NIRs)) redistributing the RBPs (Wolozin and Ivanov 2019; Guo et al. 2018; Yoshizawa et al. 2018). With persistent stress, the phase-separated RBPs may potentially remain aggregated as inert pathological remnants. Other proteins, such as p62 (SQSTM1, Sequestosome 1), bind to SG and many RBPs become posttranslationally modified by adducts such as phosphate or ubiquitin (Wolozin and Ivanov 2019). Thus, SGs could function as a genesis for pathological inclusions in microsatellite expansion diseases. Indeed, expression of C9orf72-RAN poly(GR)100 peptides in mice has led to impaired SG dynamics and neurotoxicity independent of TDP-43 pathology, nucleocytoplasmic transport defects, and nucleolar stress (Zhang et al. 2018). In addition, the arginine-rich C9orf72-RAN peptides undergo phase separation, induce a spontaneous assembly of SG, impair the dynamic exchange between SG and the cytoplasm, and alter the assembly of Cajal bodies and nuclear speckles (Boeynaems et al. 2017; Lee et al. 2016).

7 Conclusions and Future Perspectives Microsatellite expansion diseases vary widely in clinical manifestations, neuropathology, and genetic background. However, it has become increasingly clear that disruption of one or more mechanisms that tightly regulate RNA homeostasis and stability occurs early in the disease cascade and contributes to downstream pathology. Many of the discussed toxicity and metabolic defects may be low grade, and cells could cope with these subtle changes through compensatory mechanisms. Over time these mechanisms may slowly erode and lead to degeneration and ultimately cell death, especially in long-lived nondividing neurons. Many therapeutic attempts have focused on excising the repeat DNA either allele-specifically or allele nonspecifically with CRISPR-Cas9 (Dastidar et al. 2018; Monteys et al. 2017; Provenzano et al. 2017; van Agtmaal et al. 2017; Wang et al. 2018). Unintended consequences of permanent gene editing—such as repeat instability and inversion at target sites (Monteys et al. 2017; Wang et al. 2018), off-target cleavage, p53-dependent response to double-stranded breaks (Ihry et al. 2018), immunity against Cas proteins and adeno-associated viruses (AAV) (Chew 2018; Wagner et al. 2019), and AAV packaging limitation (Bengtsson et al. 2017; Yang et al. 2016)—warrant further analysis before safely applied to humans, especially to the CNS. Other “safer” strategies that eliminate the repeat-containing RNA by using antisense nucleotide (ASO) gapmers (Kordasiewicz et al. 2012; McLoughlin et al. 2018; Moore et al. 2017, 2019; Friedrich et al. 2018; Niu et al. 2018; Scoles et al. 2017; Tabrizi et al. 2019), bleomycin-cleavage modules (Rzuczek et al. 2017), or deactivated Cas9 (dCas9) conjugated to a PIN (PilT N-terminus) domain (Batra et al. 2017) are also under investigation. Beyond RNA-destruction therapies, aberrant interactions between RBPs and expanded RNAs can be modulated by small molecules (e.g., DM1) (Childs-Disney et al. 2013; Gareiss et al. 2008; Nguyen et al. 2015; Rzuczek et al. 2015; Warf et al. 2009); gene silencing can be reversed by dCas9-Tet1 recruitment (e.g., FXS) (Liu et al. 2018); repeat transcription could be

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blocked by dCas9 and translation by RNase H-incompatible ASOs (e.g., DM1 and polyQ diseases) (Liu et al. 2018; Gagnon et al. 2011; Hu et al. 2009; Kourkouta et al. 2019; Pinto et al. 2017; Toonen et al. 2017); and mutant polyQ proteins could be allele-specifically lowered by autophagosome-tethering compounds (e.g., polyQ diseases) (Li et al. 2019). An in-depth understanding of disease mechanisms and the native function of each protein involved in disease pathways will help sharpen the therapeutic tools and minimize off-target effects.

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