RNA Modifications: Methods and Protocols (Methods in Molecular Biology, 2298) 1071613731, 9781071613733

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Methods in Molecular Biology 2298

Mary McMahon Editor

RNA Modifications Methods and Protocols

METHODS

IN

MOLECULAR BIOLOGY

Series Editor John M. Walker School of Life and Medical Sciences University of Hertfordshire Hatfield, Hertfordshire, UK

For further volumes: http://www.springer.com/series/7651

For over 35 years, biological scientists have come to rely on the research protocols and methodologies in the critically acclaimed Methods in Molecular Biology series. The series was the first to introduce the step-by-step protocols approach that has become the standard in all biomedical protocol publishing. Each protocol is provided in readily-reproducible step-bystep fashion, opening with an introductory overview, a list of the materials and reagents needed to complete the experiment, and followed by a detailed procedure that is supported with a helpful notes section offering tips and tricks of the trade as well as troubleshooting advice. These hallmark features were introduced by series editor Dr. John Walker and constitute the key ingredient in each and every volume of the Methods in Molecular Biology series. Tested and trusted, comprehensive and reliable, all protocols from the series are indexed in PubMed.

RNA Modifications Methods and Protocols

Edited by

Mary McMahon Department of Urology, University of California, San Francisco, San Francisco, CA, USA

Editor Mary McMahon Department of Urology University of California, San Francisco San Francisco, CA, USA

ISSN 1064-3745 ISSN 1940-6029 (electronic) Methods in Molecular Biology ISBN 978-1-0716-1373-3 ISBN 978-1-0716-1374-0 (eBook) https://doi.org/10.1007/978-1-0716-1374-0 © Springer Science+Business Media, LLC, part of Springer Nature 2021 The chapter 20 is licensed under the terms of the Creative Commons Attribution 4.0 International License (http:// creativecommons.org/licenses/by/4.0/). For further details see license information in the chapter. 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 Humana imprint is published by the registered company Springer Science+Business Media, LLC, part of Springer Nature. The registered company address is: 1 New York Plaza, New York, NY 10004, U.S.A.

Preface Post-transcriptional RNA modifications offer a wealth of chemical and functional diversity to RNA and are emerging as key regulators of gene expression and function, as well as governors of important physiological processes. Collectively referred to as the epitranscriptome, over 150 distinct types of modifications have been identified on diverse classes of both coding and non-coding RNA. Advances in methodologies to map RNA modifications have significantly propelled the discovery of new types of modifications especially within messenger RNA, as well as the identification of new substrates of seemingly well-characterized RNA-modifying enzymes. Moreover, a new wave of research has supported the discovery of novel biological functions of modified residues and of “writers,” “readers,” and “erasers” of various modifications, alterations of which are becoming increasingly linked to disease pathologies. As the need to better understand RNA modifications is increasing, so too is the need to accurately identify, measure, and study modified residues. This book describes some of the most recent advances and up-to-date methodologies to detect, quantify, analyze, and elucidate the biological function of different types of RNA modifications. Importantly, the methodologies and tools described herein can be applied to a wide variety of organisms and can be used to address biological and clinical questions. We hope these methods and protocols will serve those working directly in the fields of epitranscriptomics and post-transcriptional gene regulation, as well as scientists and clinicians interested in bioinformatic tools to study RNA modifications and techniques to dissect their roles in physiology and disease. San Francisco, CA, USA

Mary McMahon

v

Contents Preface . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Contributors. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

PART I

BIOINFORMATIC TOOLS TO STUDY RNA MODIFICATIONS

1 RNA Post-Transcriptional Modification Mapping Data Analysis Using RNA Framework. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Ilaria Manfredonia and Danny Incarnato 2 An Informatics Pipeline for Profiling and Annotating RNA Modifications. . . . . . Qi Liu, Xiaoqiang Lang, and Richard I. Gregory

PART II

3 15

DETECTING RNA MODIFICATIONS USING NANOPORE DIRECT RNA SEQUENCING

3 EpiNano: Detection of m6A RNA Modifications Using Oxford Nanopore Direct RNA Sequencing. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Huanle Liu, Oguzhan Begik, and Eva Maria Novoa 4 Adaptation of Human Ribosomal RNA for Nanopore Sequencing of Canonical and Modified Nucleotides. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Miten Jain, Hugh E. Olsen, Mark Akeson, and Robin Abu-Shumays

PART III

v ix

31

53

NEXT-GENERATION SEQUENCING APPROACHES TO DETECT AND CAPTURE MODIFIED RNAS

5 AlkAniline-Seq: A Highly Sensitive and Specific Method for Simultaneous Mapping of 7-Methyl-guanosine (m7G) and 3-Methyl-cytosine (m3C) in RNAs by High-Throughput Sequencing . . . . . . . . . . . . . . . . . . . . . . . . . . 77 Virginie Marchand, Lilia Ayadi, Vale´rie Bourguignon-Igel, Mark Helm, and Yuri Motorin 6 Transcriptome-Wide Detection of Internal N7-Methylguanosine . . . . . . . . . . . . . 97 Li-Sheng Zhang, Chang Liu, and Chuan He 7 miCLIP-MaPseq Identifies Substrates of Radical SAM RNA-Methylating Enzyme Using Mechanistic Cross-Linking and Mismatch Profiling . . . . . . . . . . . 105 Vanja Stojkovic´, David E. Weinberg, and Danica Galonic´ Fujimori 8 Mapping RNA Modifications Using Photo-Crosslinking-Assisted Modification Sequencing . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 123 Bryan R. Cullen and Kevin Tsai 9 Quantitative and Single-Nucleotide Resolution Profiling of RNA 5-Methylcytosine . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 135 Jun Li, Xingyu Wu, Trung Do, Vy Nguyen, Jing Zhao, Pei Qin Ng, Alice Burgess, Rakesh David, and Iain Searle

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10

Contents

A Small RNA-Seq Protocol with Less Bias and Improved Capture of 20 -O-Methyl RNAs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 153 Erwin L. van Dijk and Claude Thermes

PART IV ASSESSING RNA MODIFICATIONS USING QPCR- AND MOLECULAR BIOLOGY-BASED METHODS 11 12

13

14

15

Assessing 20 -O-Methylation of mRNA Using Quantitative PCR . . . . . . . . . . . . . . Brittany A. Elliott and Christopher L. Holley Relative Quantification of Residue-Specific m6A RNA Methylation Using m6A-RT-QPCR . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Ane Olazagoitia-Garmendia and Ainara Castellanos-Rubio Monitoring the 5-Methoxycarbonylmethyl-2-Thiouridine (mcm5s2U) Modification Utilizing the Gamma-Toxin Endonuclease . . . . . . . . . . . . . . . . . . . . . Jenna M. Lentini and Dragony Fu Analysis of Queuosine tRNA Modification Using APB Northern Blot Assay. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Cansu Cirzi and Francesca Tuorto Detecting ADP-Ribosylation in RNA . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Deeksha Munnur and Ivan Ahel

PART V 16

17

18 19

20

185

197

217 231

MASS SPECTROMETRY- AND NMR-BASED METHODS FOR RNA MODIFICATIONS ANALYSIS

Detecting Internal N7-Methylguanosine mRNA Modifications by Differential Enzymatic Digestion Coupled with Mass Spectrometry Analysis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Xue-Jiao You and Bi-Feng Yuan A General LC-MS-Based Method for Direct and De Novo Sequencing of RNA Mixtures Containing both Canonical and Modified Nucleotides . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Ning Zhang, Shundi Shi, Xiaohong Yuan, Wenhao Ni, Xuanting Wang, Barney Yoo, Tony Z. Jia, Wenjia Li, and Shenglong Zhang Quantification of Modified Nucleosides in the Context of NAIL-MS. . . . . . . . . . Matthias Heiss, Kayla Borland, Yasemin Yoluc¸, and Stefanie Kellner A Method to Monitor the Introduction of Posttranscriptional Modifications in tRNAs with NMR Spectroscopy . . . . . . . . . . . . . . . . . . . . . . . . . . . Alexandre Gato, Marjorie Catala, Carine Tisne´, and Pierre Barraud

PART VI

171

247

261

279

307

APPROACHES TO ASSESS KINETICS, DETERMINANTS, AND FUNCTIONS OF RNA MODIFICATIONS

Effects of mRNA Modifications on Translation: An Overview . . . . . . . . . . . . . . . . 327 Bijoyita Roy

Contents

21

22

23

ix

Assaying the Molecular Determinants and Kinetics of RNA Pseudouridylation by H/ACA snoRNPs and Stand-Alone Pseudouridine Synthases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 357 Dominic P. Czekay, Sarah K. Schultz, and Ute Kothe Investigating Pseudouridylation Mechanisms by High-Throughput in Vitro RNA Pseudouridylation and Sequencing . . . . . . . . . . . . . . . . . . . . . . . . . . . 379 Nicole M. Martinez and Wendy V. Gilbert Targeted RNA m6A Editing Using Engineered CRISPR-Cas9 Conjugates. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 399 Xiao-Min Liu and Shu-Bing Qian

Index . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

415

Contributors ROBIN ABU-SHUMAYS • Biomolecular Engineering Department and Genomics Institute, University of California, Santa Cruz, CA, USA IVAN AHEL • Sir William Dunn School of Pathology, University of Oxford, Oxford, UK MARK AKESON • Biomolecular Engineering Department and Genomics Institute, University of California, Santa Cruz, CA, USA LILIA AYADI • Universite´ de Lorraine, CNRS, INSERM, EpiRNASeq Core Facility, UMS2008/US40 IBSLor, Nancy, France; Universite´ de Lorraine, CNRS, UMR7365 IMoPA, Nancy, France PIERRE BARRAUD • Expression ge´ne´tique microbienne, UMR 8261, CNRS, Universite´ de Paris, Institut de biologie physico-chimique (IBPC), Paris, France OGUZHAN BEGIK • Centre for Genomic Regulation (CRG), The Barcelona Institute of Science and Technology, Barcelona, Spain; Department of Neuroscience, Garvan Institute of Medical Research, Darlinghurst, NSW, Australia; St. Vincent’s Clinical School, UNSW Sydney, Darlinghurst, NSW, Australia KAYLA BORLAND • Department of Chemistry, Ludwig Maximilians University Munich, Munich, Germany VALE´RIE BOURGUIGNON-IGEL • Universite´ de Lorraine, CNRS, INSERM, EpiRNASeq Core Facility, UMS2008/US40 IBSLor, Nancy, France; Universite´ de Lorraine, CNRS, UMR7365 IMoPA, Nancy, France ALICE BURGESS • Department of Molecular and Biomedical Sciences, School of Biological Sciences, The University of Adelaide, Adelaide, SA, Australia AINARA CASTELLANOS-RUBIO • Department of Genetics, Physical Anthropology and Animal Physiology, University of the Basque Country (UPV-EHU), Leioa, Spain; Biocruces Bizkaia Health Research Institute, Barakaldo, Spain; Ikerbasque, Basque Foundation for Science, Bilbao, Spain; CIBER de Diabetes y Enfermedades Metabolicas Asociadas (CIBERDEM), Instituto de Salud Carlos III, Madrid, Spain MARJORIE CATALA • Expression ge´ne´tique microbienne, UMR 8261, CNRS, Universite´ de Paris, Institut de biologie physico-chimique (IBPC), Paris, France CANSU CIRZI • Division of Epigenetics, German Cancer Research Center (DKFZ), Heidelberg, Germany; Faculty of Biosciences, University of Heidelberg, Heidelberg, Germany BRYAN R. CULLEN • Department of Molecular Genetics and Microbiology, Duke University Medical Center, Durham, NC, USA DOMINIC P. CZEKAY • Department of Chemistry and Biochemistry, Alberta RNA Research and Training Institute, University of Lethbridge, Lethbridge, AB, Canada RAKESH DAVID • Department of Molecular and Biomedical Sciences, School of Biological Sciences, The University of Adelaide, Adelaide, SA, Australia TRUNG DO • Department of Molecular and Biomedical Sciences, School of Biological Sciences, The University of Adelaide, Adelaide, SA, Australia BRITTANY A. ELLIOTT • Department of Medicine, Duke University Medical Center, Durham, NC, USA DRAGONY FU • Department of Biology, Center for RNA Biology, University of Rochester, Rochester, NY, USA

xi

xii

Contributors

DANICA GALONIC´ FUJIMORI • Department of Cellular and Molecular Pharmacology, University of California San Francisco, San Francisco, CA, USA; Department of Pharmaceutical Chemistry, University of California San Francisco, San Francisco, CA, USA ALEXANDRE GATO • Expression ge´ne´tique microbienne, UMR 8261, CNRS, Universite´ de Paris, Institut de biologie physico-chimique (IBPC), Paris, France WENDY V. GILBERT • Department of Molecular Biophysics and Biochemistry, Yale University, New Haven, CT, USA RICHARD I. GREGORY • Stem Cell Program, Division of Hematology/Oncology, Boston Children’s Hospital, Boston, MA, USA; Department of Biological Chemistry and Molecular Pharmacology, Harvard Medical School, Boston, MA, USA; Department of Pediatrics, Harvard Medical School, Boston, MA, USA; Harvard Initiative for RNA Medicine, Boston, MA, USA; Harvard Stem Cell Institute, Cambridge, MA, USA CHUAN HE • Department of Chemistry, The University of Chicago, Chicago, IL, USA; Howard Hughes Medical Institute, The University of Chicago, Chicago, IL, USA; Department of Biochemistry and Molecular Biology, Institute for Biophysical Dynamics, The University of Chicago, Chicago, IL, USA MATTHIAS HEISS • Department of Chemistry, Ludwig Maximilians University Munich, Munich, Germany MARK HELM • Institute of Pharmacy and Biochemistry, Johannes Gutenberg University Mainz, Mainz, Germany CHRISTOPHER L. HOLLEY • Department of Medicine, Duke University Medical Center, Durham, NC, USA DANNY INCARNATO • Department of Molecular Genetics, Groningen Biomolecular Sciences and Biotechnology Institute (GBB), University of Groningen, Groningen, Netherlands MITEN JAIN • Biomolecular Engineering Department and Genomics Institute, University of California, Santa Cruz, CA, USA TONY Z. JIA • Earth-Life Science Institute, Tokyo Institute of Technology, Meguro-ku, Tokyo, Japan; Blue Marble Space Institute of Science, Seattle, WA, USA STEFANIE KELLNER • Department of Chemistry, Ludwig Maximilians University Munich, Munich, Germany UTE KOTHE • Department of Chemistry and Biochemistry, Alberta RNA Research and Training Institute, University of Lethbridge, Lethbridge, AB, Canada XIAOQIANG LANG • Precision Medicine Research Center, West China Hospital, Sichuan University, Chengdu, Sichuan, China JENNA M. LENTINI • Department of Biology, Center for RNA Biology, University of Rochester, Rochester, NY, USA JUN LI • Department of Molecular and Biomedical Sciences, School of Biological Sciences, The University of Adelaide, Adelaide, SA, Australia WENJIA LI • Department of Computer Science, New York Institute of Technology, New York, NY, USA CHANG LIU • Department of Chemistry, The University of Chicago, Chicago, IL, USA; Howard Hughes Medical Institute, The University of Chicago, Chicago, IL, USA HUANLE LIU • Centre for Genomic Regulation (CRG), The Barcelona Institute of Science and Technology, Barcelona, Spain QI LIU • Stem Cell Program, Division of Hematology/Oncology, Boston Children’s Hospital, Boston, MA, USA; Department of Biological Chemistry and Molecular Pharmacology, Harvard Medical School, Boston, MA, USA

Contributors

xiii

XIAO-MIN LIU • School of Life Science and Technology, China Pharmaceutical University, Nanjing, China ILARIA MANFREDONIA • Department of Molecular Genetics, Groningen Biomolecular Sciences and Biotechnology Institute (GBB), University of Groningen, Groningen, The Netherlands VIRGINIE MARCHAND • Universite´ de Lorraine, CNRS, INSERM, EpiRNASeq Core Facility, UMS2008/US40 IBSLor, Nancy, France NICOLE M. MARTINEZ • Department of Molecular Biophysics and Biochemistry, Yale University, New Haven, CT, USA YURI MOTORIN • Universite´ de Lorraine, CNRS, INSERM, EpiRNASeq Core Facility, UMS2008/US40 IBSLor, Nancy, France; Universite´ de Lorraine, CNRS, UMR7365 IMoPA, Nancy, France DEEKSHA MUNNUR • Sir William Dunn School of Pathology, University of Oxford, Oxford, UK PEI QIN NG • Department of Molecular and Biomedical Sciences, School of Biological Sciences, The University of Adelaide, Adelaide, SA, Australia VY NGUYEN • Department of Molecular and Biomedical Sciences, School of Biological Sciences, The University of Adelaide, Adelaide, SA, Australia WENHAO NI • Department of Biological and Chemical Sciences, New York Institute of Technology, New York, NY, USA EVA MARIA NOVOA • Centre for Genomic Regulation (CRG), The Barcelona Institute of Science and Technology, Barcelona, Spain; Department of Neuroscience, Garvan Institute of Medical Research, Darlinghurst, NSW, Australia; St. Vincent’s Clinical School, UNSW Sydney, Darlinghurst, NSW, Australia; Universitat Pompeu Fabra (UPF), Barcelona, Spain ANE OLAZAGOITIA-GARMENDIA • Department of Genetics, Physical Anthropology and Animal Physiology, University of the Basque Country (UPV-EHU), Leioa, Spain; BioCruces Bizkaia Health Research Institute, Barakaldo, Spain HUGH E. OLSEN • Biomolecular Engineering Department and Genomics Institute, University of California, Santa Cruz, CA, USA SHU-BING QIAN • Division of Nutritional Sciences, Cornell University, Ithaca, NY, USA BIJOYITA ROY • RNA and Genome Editing, New England Biolabs Inc., Ipswich, MA, USA SARAH K. SCHULTZ • Department of Chemistry and Biochemistry, Alberta RNA Research and Training Institute, University of Lethbridge, Lethbridge, AB, Canada IAIN SEARLE • Department of Molecular and Biomedical Sciences, School of Biological Sciences, The University of Adelaide, Adelaide, SA, Australia SHUNDI SHI • Department of Chemical Engineering, Columbia University, New York, NY, USA VANJA STOJKOVIC´ • Department of Cellular and Molecular Pharmacology, University of California San Francisco, San Francisco, CA, USA CLAUDE THERMES • Institute for Integrative Biology of the Cell, UMR9198, CNRS CEA Univ Paris-Sud, Universite´ Paris-Saclay, Gif sur Yvette Cedex, France CARINE TISNE´ • Expression ge´ne´tique microbienne, UMR 8261, CNRS, Universite´ de Paris, Institut de biologie physico-chimique (IBPC), Paris, France KEVIN TSAI • Department of Molecular Genetics and Microbiology, Duke University Medical Center, Durham, NC, USA; Institute of Biomedical Sciences, Academia Sinica, Taipei, Taiwan FRANCESCA TUORTO • Division of Biochemistry, Mannheim Institute for Innate Immunoscience (MI3), Medical Faculty Mannheim, Heidelberg University,

xiv

Contributors

Mannheim, Germany; Center for Molecular Biology of Heidelberg University (ZMBH), Mannheim, Germany ERWIN L. VAN DIJK • Institute for Integrative Biology of the Cell, UMR9198, CNRS CEA Univ Paris-Sud, Universite´ Paris-Saclay, Gif sur Yvette Cedex, France XUANTING WANG • Department of Chemical Engineering, Columbia University, New York, NY, USA DAVID E. WEINBERG • Department of Cellular and Molecular Pharmacology, University of California San Francisco, San Francisco, CA, USA XINGYU WU • Department of Molecular and Biomedical Sciences, School of Biological Sciences, The University of Adelaide, Adelaide, SA, Australia YASEMIN YOLUC¸ • Department of Chemistry, Ludwig Maximilians University Munich, Munich, Germany BARNEY YOO • Department of Chemistry, Hunter College, City University of New York, New York, NY, USA XUE-JIAO YOU • Sauvage Center for Molecular Sciences, Department of Chemistry, Wuhan University, Wuhan, China BI-FENG YUAN • Sauvage Center for Molecular Sciences, Department of Chemistry, Wuhan University, Wuhan, China XIAOHONG YUAN • Department of Biological and Chemical Sciences, New York Institute of Technology, New York, NY, USA LI-SHENG ZHANG • Department of Chemistry, The University of Chicago, Chicago, IL, USA; Howard Hughes Medical Institute, The University of Chicago, Chicago, IL, USA NING ZHANG • Department of Biological and Chemical Sciences, New York Institute of Technology, New York, NY, USA; Department of Chemical Engineering, Columbia University, New York, NY, USA SHENGLONG ZHANG • Department of Biological and Chemical Sciences, New York Institute of Technology, New York, NY, USA JING ZHAO • Department of Molecular and Biomedical Sciences, School of Biological Sciences, The University of Adelaide, Adelaide, SA, Australia

Part I Bioinformatic Tools to Study RNA Modifications

Chapter 1 RNA Post-Transcriptional Modification Mapping Data Analysis Using RNA Framework Ilaria Manfredonia and Danny Incarnato Abstract RNA post-transcriptional modifications (PTMs) are progressively gaining relevance in the study of codingindependent functions of RNA. RNA PTMs act as dynamic regulators of several aspects of the RNA physiology, from translation to half-life. Rising interest is supported by the advance of high-throughput techniques enabling the detection of these modifications on a transcriptome-wide scale. To this end, here we illustrate the usefulness of RNA Framework, a comprehensive toolkit for the analysis of RNA PTM mapping experiments, by reanalyzing two published transcriptome-scale datasets of N1-methyladenosine (m1A) and pseudouridine (Ψ) mapping, based on two different experimental strategies. Key words RNA post-transcriptional modifications, RNA immunoprecipitation, High-throughput sequencing, m1A, N1-methyladenosine, Pseudouridine

1

Introduction Although the discovery of the very first RNA post-transcriptional modification (PTM) dates back to 1951 [1] and, since then, over 150 PTMs have been identified [2], the study of RNA PTMs remained elusive for over 50 years. This was largely due to a lack of suitable methods for detection of RNA PTMs, especially on low-abundant transcripts. Thanks to the advent of high-throughput sequencing methods, it is now possible to identify, localize, and contextualize RNA PTMs. Over the past decade, RNA PTMs have been implicated in modulating RNA structure by affecting the formation of both intra- and intermolecular interactions, half-life, translation, protein binding, and many other aspects of RNA physiology [3]. Currently available sequencing-based techniques for mapping RNA PTMs can be broadly divided into two major categories. The first category involves techniques that rely on the use of a specific antibody to enrich for PTM-containing RNA fragments, dubbed RNA immunoprecipitation (RIP) approaches. The second

Mary McMahon (ed.), RNA Modifications: Methods and Protocols, Methods in Molecular Biology, vol. 2298, https://doi.org/10.1007/978-1-0716-1374-0_1, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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Ilaria Manfredonia and Danny Incarnato

category, instead, includes techniques based on the exploitation of individual chemical properties of the modified nucleobases, with the aim of achieving single-base resolution mapping. A major bottleneck of these experiments is represented by data analysis. To this end, we have recently introduced RNA Framework as a generalized toolkit for the analysis of NGS-based RNA PTM mapping experiments [4]. Herein we guide the reader through the steps necessary for the successful analysis of data derived from both IP-based and single-base resolution experiments, by exploiting two previously published datasets.

2 2.1

Materials RNA Framework

RNA Framework (obtainable from our Git repository: https:// github.com/dincarnato/RNAFramework) is implemented in Perl. It requires Perl v5.12 (or greater), with ithreads support and a 64-bit architecture system running Linux or any other UNIXbased OS. The following software and packages are also required: 1. Bowtie v1.1.2 or greater (http://bowtie-bio.sourceforge.net/ index.shtml), and/or Bowtie v2.2.7 or greater (http://bowtiebio.sourceforge.net/bowtie2/index.shtml). 2. SAMTools v1.2 or greater (http://www.htslib.org/). 3. BEDTools v2.0 or greater (https://github.com/arq5x/ bedtools2/). 4. Cutadapt v2.1 or greater (http://cutadapt.readthedocs.io/en/ stable/index.html). 5. ViennaRNA Package v2.2.0 or greater (http://www.tbi.univie. ac.at/RNA/). 6. Perl non-CORE modules (https://metacpan.org/): – DBD::mysql. – RNA (installed by the ViennaRNA package). – XML::LibXML. – Config::Simple. To start using RNA Framework, first clone it from our GitHub repository by typing: $ git clone https://github.com/dincarnato/RNAFramework

This will create the “RNAFramework” folder. RNA Framework executables can then be simply added to user’s PATH by typing: $ export PATH=$PATH:$(pwd)/RNAFramework

High-Throughput Mapping of RNA Modifications

5

Table 1 List of SRA accession IDs for the datasets used in this chapter Accession ID

Description 1

Reference

SRR2086044

m A-seq (H. sapiens, polyA+)

[5]

SRR2086045

Input (H. sapiens, polyA+)

[5]

SRR1327248

CMC (S. cerevisiae, total RNA)

[6]

SRR1327249

CMC+ (S. cerevisiae, total RNA)

[6]

2.2

Data Retrieval

To illustrate a typical data analysis workflow with RNA Framework, we chose two published datasets. The first one is m1A-seq [5], a RIP approach applied to map N1-methyladenosine (m1A) in H. sapiens polyadenylated transcripts. The second dataset is Pseudo-Seq [6], a single-base resolution method based on the ability of N-cyclohexyl-N0 -(2-morpholinoethyl)carbodiimide metho-p-toluenesulfonate (CMC) to generate an alkali-resistant adduct with the N3 of pseudouridine (Ψ) residues, resulting in reverse transcription (RT) drop-off and which can be applied to map Ψ in S. cerevisiae ribosomal RNAs (rRNAs). 1. Data can be obtained from the NCBI Sequence Read Archive (https://www.ncbi.nlm.nih.gov/sra). Using the NCBI SRA Toolkit (https://www.ncbi.nlm.nih.gov/sra/docs/ toolkitsoft/) it is possible to obtain the raw FASTQ files by typing: $ fastq-dump -A

2. SRA accession IDs used in this chapter are reported in Table 1.

3

Methods For a detailed list of all program-specific parameters, all tools can be invoked with the “-h” (or “--help” flag). Also, a detailed documentation of RNA Framework and its components can be found online at https://rnaframework.readthedocs.io. Bowtie (either version 1 or 2) is the default read aligner used by RNA Framework [7, 8]. Alternatively, the user can employ any other read alignment tool, as long as it is able to report alignments in SAM/BAM format. In this case, the next two paragraphs can be skipped and it is possible to proceed directly to Subheading 3.3.

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Ilaria Manfredonia and Danny Incarnato

3.1 Generation of the Reference Index

1. Data derived from any of the aforementioned PTM mapping techniques (IP-based and single-base resolution) can be mapped to the reference transcriptome using Bowtie v1, a fast ungapped read aligner. Bowtie reference transcriptome index generation is automatized through the rf-index module. Besides including a set of pre-built reference indexes (displayable with the “-lp” flag and downloadable through the “-pb ” parameter, where is the code identifying the desired index), rf-index further allows the creation of tailored reference indexes. For this purpose, it relies on querying the UCSC genome database (https://genome.ucsc.edu) for a userspecified genome assembly and gene annotation. A complete list of available genome assemblies is available at https:// genome.ucsc.edu/FAQ/FAQreleases.html. 2. For the purpose of this chapter, we are going to need two reference indexes: one on H. sapiens RefSeq protein-coding transcripts and one on S. cerevisiae rRNAs. To obtain the pre-built indexes, type: $ rf-index -pb 6 # H. sapiens transcriptome $ rf-index -pb 3 # S. cerevisiae rRNAs

3. Alternatively, the user can generate a new reference transcriptome index using the desired genome assembly and gene annotation, for example: $ rf-index -g hg38 -a refGene -n -co # H. sapiens transcriptome

In this example, the reference index will be created using the hg38 assembly and the refGene (RefSeqGene) annotation. The “-n” flag instructs rf-index to use gene symbols instead of gene IDs (where possible). If multiple transcript isoforms are available, only the longest will be picked as representative of the gene. The “-co” flag instead allows selecting only protein-coding transcripts. It is worth noting that, more recently, approaches for singlebase resolution mapping of PTMs based on reverse transcription (RT) conditions favoring the read-through on sites of PTM-adduct formation have been proposed [9, 10]. Analogously to mutational profiling strategies for RNA structure mapping, these methods result in the recording of read-through sites as mutations or insertions and deletions (indels) in the resulting cDNA molecules. As Bowtie v1 is not able to handle indels, it is advisable to use Bowtie v2 for the analysis of these experiments. This can be easily done by invoking both rf-index and rf-map with the “-b2” (or “--bowtie2”) flag. However, we are not going to focus on the analysis of these types of experiments in this chapter.

High-Throughput Mapping of RNA Modifications

7

3.2 Mapping of Reads

Following index generation, it is possible to proceed to read mapping using the rf-map module. All the necessary read preprocessing steps, including adaptor clipping and low-quality base trimming, are automatized as part of this tool. As a result, a sorted BAM file will be generated for each sample being analyzed. Before proceeding with read mapping, it is also advisable to check base qualities using the FastQC tool (https://www.bioinformatics. babraham.ac.uk/projects/fastqc/).

3.2.1 Mapping of m1A-Seq Reads

To map m1A-seq data, simply type: $ rf-map -ca3 GATCGGAAGAGCACACGTCTGA -cq5 20 -bm 7 -ba -bi Hsapiens_refGene_longest_bt2/reference

-o

rf_map_m1Aseq/

SRR208604*.fastq

Through the “-ca3” parameter it is possible to specify the sequence of the 30 adaptor to be clipped (in this case corresponding to the Illumina TruSeq adaptor). Quality trimming (Phred Q30 quality, will be found in the “clean_outdir” directory created using the “mkdir” command. “--phred33” indicates the encode type of sequence quality, which can be identified in the FastQC report (see Note 4). 6. Short read mapping: Map the cleaned sequence generated in the previous step to the reference genome sequence using a splicing-aware short read aligner tool, such as STAR [21] and TopHat [22], which can generate mapping results in SAM/BAM format. Here, STAR is used to align the cleaned sequences (INPUT and m6A IP samples) to human reference genome sequences (hg19): $ STAR --runThreadN 8 --runMode genomeGenerate --genomeDir hg19 --genomeFastaFiles hg19.fa --sjdbGTFfile gencode.v19.annotation.gtf $ for sraID in SRR6686554 SRR6686555 SRR6686557 SRR6686558 SRR6686560 SRR6686561 SRR6686563 SRR6686564; do STAR --genomeDir hg19 --sjdbGTFfile gencode.v19.annotation.gtf --outSAMtype BAM SortedByCoordinate --outFilterMultimapNmax 1 -outFilterMismatchNmax 2 --runThreadN 8 --readFilesCommand zcat --readFilesIn clean_outdir/${sraID}_trimmed.fq.gz --outFileNamePrefix $sraID; done

During the mapping process, a maximum of two mismatches (--outFilterMismatchNmax 2) is allowed and unique mapped reads are reported (--outFilterMultimapNmax 1). “hg19” is the name of the genome sequence index which is built by genomeGenerate model of STAR. “gencode.v19.

A Pipeline for Profiling and Annotating RNA Modifications

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annotation.gtf” is the gene annotation for hg19 reference genome in GENCODE [23]. The BAM mapping files with file extension “Aligned.sortedByCoord.out.bam” will be generated in the current working directory. 7. Index the BAM files: Index the BAM files using Samtools: $ for sraID in SRR6686554 SRR6686555 SRR6686557 SRR6686558 SRR6686560 SRR6686561 SRR6686563 SRR6686564; do samtools index ${sraID}Aligned.sortedByCoord.out.bam; done

Here, the BAM files are further indexed, which can be used to visually check the modification peaks in the Integrative Genomics Viewer (IGV) [24]. 8. Modification site/peak calling: Call the m6A RNA modification peaks based on INPUT and m6A IP sample using MACS2 [18] or exomePeak [19] (see Note 5). Here, MACS2 is used to call the modifications: $ inputs=(SRR6686554 SRR6686555 SRR6686560 SRR6686561) $ ips=(SRR6686557 SRR6686558 SRR6686563 SRR6686564) $ for(( i=0;i 1.5): prediction = ’mod’ else: prediction = ’unm’

Acknowledgments We thank all members of the Novoa lab for their valuable insights and discussion. We thank Rebeca Medina for obtaining the TapeStation image used for Fig. 1. OB is supported by an international PhD fellowship (UIPA) from the University of New South Wales. This work was supported by the Australian Research Council (DP180103571 to EMN) and the Spanish Ministry of Economy, Industry and Competitiveness (MEIC) (PGC2018-098152-A-100 to EMN). We acknowledge the support of the MEIC to the EMBL partnership, Centro de Excelencia Severo Ochoa, and CERCA Program/Generalitat de Catalunya.

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References 1. Cohn WE, Volkin E (1951) Nucleoside-50 -phosphates from ribonucleic acid. Nature 167:483–484 2. Adams JM, Cory S (1975) Modified nucleosides and bizarre 50 -termini in mouse myeloma mRNA. Nature 255:28–33 3. Desrosiers R, Friderici K, Rottman F (1974) Identification of methylated nucleosides in messenger RNA from Novikoff hepatoma cells. Proc Natl Acad Sci U S A 71:3971–3975 4. Dubin DT, Taylor RH (1975) The methylation state of poly A-containing messenger RNA from cultured hamster cells. Nucleic Acids Res 2:1653–1668 5. Perry RP, Kelley DE, Friderici K, Rottman F (1975) The methylated constituents of L cell messenger RNA: evidence for an unusual cluster at the 50 terminus. Cell 4:387–394 6. Jia G, Fu Y, Zhao X, Dai Q, Zheng G, Yang Y, Yi C, Lindahl T, Pan T, Yang Y-G, He C (2011) N6-methyladenosine in nuclear RNA is a major substrate of the obesity-associated FTO. Nat Chem Biol 7:885–887 7. Zheng G, Dahl JA, Niu Y, Fedorcsak P, Huang C-M, Li CJ, Va˚gbø CB, Shi Y, Wang W-L, Song S-H, Lu Z, Bosmans RPG, Dai Q, Hao Y-J, Yang X, Zhao W-M, Tong W-M, Wang X-J, Bogdan F, Furu K, Fu Y, Jia G, Zhao X, Liu J, Krokan HE, Klungland A, Yang Y-G, He C (2013) ALKBH5 is a mammalian RNA demethylase that impacts RNA metabolism and mouse fertility. Mol Cell 49:18–29 8. Dominissini D, Moshitch-Moshkovitz S, Schwartz S, Salmon-Divon M, Ungar L, Osenberg S, Cesarkas K, Jacob-Hirsch J, Amariglio N, Kupiec M, Sorek R, Rechavi G (2012) Topology of the human and mouse m6A RNA methylomes revealed by m6A-seq. Nature 485:201–206 9. Meyer KD, Saletore Y, Zumbo P, Elemento O, Mason CE, Jaffrey SR (2012) Comprehensive analysis of mRNA methylation reveals enrichment in 30 UTRs and near stop codons. Cell 149:1635–1646 10. Hu Y, Ouyang Z, Sui X, Qi M, Li M, He Y, Cao Y, Cao Q, Lu Q, Zhou S, Liu L, Liu L, Shen B, Shu W, Huo R (2020) Oocyte competence is maintained by m6A methyltransferase KIAA1429-mediated RNA metabolism during mouse follicular development. Cell Death Differ. https://doi.org/10.1038/s41418-0200516-1 11. Lee H, Bao S, Qian Y, Geula S, Leslie J, Zhang C, Hanna JH, Ding L (2019) Stagespecific requirement for Mettl3-dependent

m6A mRNA methylation during haematopoietic stem cell differentiation. Nat Cell Biol 21:700–709 12. Zhao BS, He C (2015) Fate by RNA methylation: m6A steers stem cell pluripotency. Genome Biol 16:43 13. Geula S, Moshitch-Moshkovitz S, Dominissini D, Mansour AA, Kol N, SalmonDivon M, Hershkovitz V, Peer E, Mor N, Manor YS, Ben-Haim MS, Eyal E, Yunger S, Pinto Y, Jaitin DA, Viukov S, Rais Y, Krupalnik V, Chomsky E, Zerbib M, Maza I, Rechavi Y, Massarwa R, Hanna S, Amit I, Levanon EY, Amariglio N, Stern-Ginossar N, Novershtern N, Rechavi G, Hanna JH (2015) Stem cells. m6A mRNA methylation facilitates resolution of naı¨ve pluripotency toward differentiation. Science 347:1002–1006 14. Lence T, Akhtar J, Bayer M, Schmid K, Spindler L, Ho CH, Kreim N, AndradeNavarro MA, Poeck B, Helm M, Roignant J-Y (2016) m6A modulates neuronal functions and sex determination in Drosophila. Nature 540:242–247 15. Haussmann IU, Bodi Z, Sanchez-Moran E, Mongan NP, Archer N, Fray RG, Soller M (2016) m6A potentiates Sxl alternative pre-mRNA splicing for robust Drosophila sex determination. Nature 540:301–304 16. Helm M, Motorin Y (2017) Detecting RNA modifications in the epitranscriptome: predict and validate. Nat Rev Genet 18:275–291 17. Li X, Xiong X, Yi C (2016) Epitranscriptome sequencing technologies: decoding RNA modifications. Nat Methods 14:23–31 18. Linder B, Grozhik AV, Olarerin-George AO, Meydan C, Mason CE, Jaffrey SR (2015) Single-nucleotide-resolution mapping of m6A and m6Am throughout the transcriptome. Nat Methods 12:767–772 19. Novoa EM, Mason CE, Mattick JS (2017) Charting the unknown epitranscriptome. Nat Rev Mol Cell Biol 18:339–340 20. Jonkhout N, Tran J, Smith MA, Schonrock N, Mattick JS, Novoa EM (2017) The RNA modification landscape in human disease. RNA 23:1754–1769 21. Liu H, Begik O, Lucas MC, Ramirez JM, Mason CE, Wiener D, Schwartz S, Mattick JS, Smith MA, Novoa EM (2019) Accurate detection of m6A RNA modifications in native RNA sequences. Nat Commun 10:4079 22. Lorenz DA, Sathe S, Einstein JM, Yeo GW (2020) Direct RNA sequencing enables m6A

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detection in endogenous transcript isoforms at base-specific resolution. RNA 26:19–28 23. Parker MT, Knop K, Sherwood AV, Schurch NJ, Mackinnon K, Gould PD, Hall AJ, Barton GJ, Simpson GG (2020) Nanopore direct RNA sequencing maps the complexity of Arabidopsis mRNA processing and m6A modification. elife 9. https://doi.org/10.7554/eLife.49658 24. Price AM, Hayer KE, McIntyre ABR, Gokhale NS, Della Fera AN, Mason CE, Horner SM, Wilson AC, Depledge DP, Weitzman MD

(2019) Direct RNA sequencing reveals m6A modifications on adenovirus RNA are necessary for efficient splicing. bioRxiv 865485 25. Li H (2018) Minimap2: pairwise alignment for nucleotide sequences. Bioinformatics 34:3094–3100 26. Loman NJ, Quick J, Simpson JT (2015) A complete bacterial genome assembled de novo using only nanopore sequencing data. Nat Methods 12:733–735

Chapter 4 Adaptation of Human Ribosomal RNA for Nanopore Sequencing of Canonical and Modified Nucleotides Miten Jain, Hugh E. Olsen, Mark Akeson, and Robin Abu-Shumays Abstract Historically, RNA has been sequenced as cDNA copies derived from reverse transcription of cellular RNA followed by PCR amplification. Recently, RNA sequencing using nanopores has emerged as an alternative. Using this technology, individual cellular RNA strands are read directly as they are driven through nanoscale pores by an applied voltage. The speed of translocation is regulated by a helicase that is loaded onto each RNA strand by an adapter that also facilitates capture by the nanopore electric field. Here we describe a technique for adapting human ribosomal RNA subunits for nanopore sequencing. Using this strategy, a single Oxford Nanopore MinION run delivered 470,907 sequence reads of which 396,048 aligned to ribosomal RNA, with 28S, 18S, 5.8S, and 5S coverage of 6053, 369,472, 16,058, and 4465 reads, respectively. Example alignments that reveal putative nucleotide modifications are provided. Key words Ribosome, RNA, Nanopore, Sequencing, Single molecule

1

Introduction Nanopore sequencing was invented in 1989 and first implemented as a commercial device in 2014 [1]. Briefly, arrays of hundreds to thousands of independently addressable nanopores are formed in thin films on an application-specific integrated circuit (ASIC). An applied voltage produces an ionic current through each pore. When an RNA or DNA strand is captured and translocated single file through the pore, the current changes in discrete steps on the millisecond timescale. These steps correspond to the sequence of nucleotides passing through the pore. Neural networks trained on known DNA or RNA sequences are used to convert the ionic current segment series into nucleotide sequences. Typical median single-strand read accuracies are 94% for DNA and 87% for RNA. The nanopore sensor reads RNA directly, permitting detection of base modifications in the context of surrounding canonical bases [2–6]. One of the main challenges to developing computational methods for base modification detection is the requirement for

Mary McMahon (ed.), RNA Modifications: Methods and Protocols, Methods in Molecular Biology, vol. 2298, https://doi.org/10.1007/978-1-0716-1374-0_4, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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high-confidence training data. Additionally, the heterogeneity and abundance of RNA modifications further increase the complexity. Software tools like nanopolish [7] and Guppy [8] have recently enabled detection of 5-methylcytosine in nanopore genomic DNA data. Many groups are working on improved computational methods for detecting base modifications, both for RNA and DNA. Modified ribonucleotides regulate ribosome function through tuning of RNA folding, and through interactions with ribosomal proteins and tRNAs [9–12]. Therefore, substantial current research involves using nanopore sequencing as a tool for identification of base modifications in rRNA. To date a majority of the modified bases in human 18S rRNA have been associated with differences between canonical and modified base ionic current signals (personal communication). Here, we provide a detailed description of our nanopore-based human nuclear-encoded rRNA sequencing protocol, with the key steps diagrammed in Fig. 1.

Fig. 1 Schematic of the human rRNA adaption and nanopore sequencing protocol. 18S rRNA is given as an example. (a) (i) Total RNA extraction using TRIzol and chloroform. (ii) Annealing and ligation of a nanoporespecific adapter to the 30 end of human 18S rRNA (red). Thirteen nucleotides of the adapter are reverse complements to 13 nucleotides of the target strand. (iii) Ligation of the nanopore sequencing adapter that has the motor protein preloaded. (iv) Addition of the adapted library to the nanopore for sequencing. (v) Basecalling of individual nanopore RNA strands using Guppy. This process yields individual RNA strand sequence reads. (b) A representative ionic current trace for a human 18S rRNA strand processed on the nanopore. Ionic current components: (i) strand capture; (ii) ONT and 18S rRNA-splint adapter translocation; (iii) human 18S rRNA translocation; and (iv) exit of the strand into the transcompartment. (c) Expanded view of the region where the ionic current transitions from the nanopore adapter to the 30 end of the adapted 18S rRNA

Nanopore Sequencing of Human Nuclear rRNA

2

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Materials

2.1 Total RNA Isolation from Flash-Frozen Cell Pellets

1. TRI Reagent. 2. 1-Bromo-3-chloro-propane or chloroform. 3. Isopropanol. 4. 100% Ethanol. 5. Nuclease-free water. 6. 10 TE (Tris-EDTA): 100 mM Tris–HCl pH 7.6, 10 mM EDTA. Sterile filter the stock solutions through 0.2μm filters and dilute to 1 as needed. 7. Qubit™ HS RNA kit. 8. Flash-frozen cell pellet from cell line of interest.

2.2 Nanopore Sequencing of Biological Human rRNA

1. NEB Quick Ligase Buffer and T4 DNA Ligase (2000 U/μL). 2. ONT SQK-RNA002 kit (or newer). 3. Tris-NaCl-EDTA buffer: 10 mM Tris–HCl, pH 8, 1 mM EDTA, 50 mM NaCl. 4. 100% Ethanol. 5. Beckman Coulter Agencourt RNAClean XP Beads. 6. Magnet for bead-based purifications. 7. Nuclease-free water. 8. Oligomers for preparing biological human rRNA splints. The sequence for individual oligomers is below: Top strand: This oligomer is common across all the splints. Note that this oligomer needs to have a 50 -phosphate group for ligation in sequencing library preparation: 50 -/5PHOS/GGCTTCTTCTTGCTCTTAGGTAGTA GGTTC-30 Human 5S rRNA bottom strand: 50 -CCTAAGAGCAAGAAGAAGCCAAAGCCTACAGCA-30 Human 5.8S rRNA bottom strand: 50 -CCTAAGAGCAAGAAGAAGCCAAGCGACGCTCAG-30 Human 18S rRNA bottom strand: 50 -CCTAAGAGCAAGAAGAAGCCTAATGATCCTTCC-30 Human 28S rRNA bottom strand: 50 -CCTAAGAGCAAGAAGAAGCCGACAAACCCTTGT-30

2.3 Generating and Sequencing Canonical rRNAs (Optional, Only Needed for IVT rRNAs)

1. 10 Tris-NaCl-EDTA buffer: 100 mM Tris–HCl pH 8, 10 mM EDTA, 500 mM NaCl. Filter sterilize the stock solutions through 0.2μm filters and dilute to 1 as needed. 2. Nuclease-free water. 3. Oligonucleotides for IVT templates: Prepare oligonucleotides as 100μM stocks in nuclease-free water or TE. The

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oligonucleotides produce IVT products for which all but ~12 50 -most nucleotides can be sequenced. For suggestions to address this see Note 1. For 5S and 5.8S rRNA, 5 and 5.8S top strand (the T7 promoter site is underlined): 50 -CATCATCATTTAATACGACTCACTATAG-30 5S bottom strand: 50 -AAAGCCTACAGCACCCGGTATTCCCAGGCGG TCTCCCATCCAAGTACTAACCAGGCCCGACCCTGC TTAGCTTCCGAGATCAGACGAGATCGGGCGCGTTC AGGGTGGTATGGCCGTAGACTATAGTGAGTCGTATT AAATGATGATG-30 5.8S bottom strand: 50 -AAGCGACGCTCAGACAGGCGTAGCCCCGGGA GGAACCCGGGGCCGCAAGTGCGTTCGAAGTGTCGA TGATCAATGTGTCCTGCAATTCACATTAATTCTCGCA GCTAGCTGCGTTCTTCATCGACGCACGAGCCGAGTG ATCCACCGCTAAGAGTCGCTATAGTGAGTCGTATT AAATGATGATG-30 4. HiScribe™ T7 Quick High Yield RNA Synthesis kit. 5. RNasin (40 units/μL) or equivalent RNase inhibitor. 6. DNase I (RNase-free) (2000 units/mL) and 10 DNase I buffer. 7. 100% Ethanol. 8. Beckman Coulter Agencourt RNAClean XP Beads. 9. Qubit™ dsDNA BR Assay Kit. 10. Qubit™ RNA BR Assay Kit. 11. Q5 DNA polymerase 2 master mix. 12. Genomic DNA from a human cell line (1–2μg). 13. 18S PCR primers: Prepare primers as 100μM stocks in nuclease-free water or TE. The primers produce an IVT template for an RNA for which all but ~12 50 -most nucleotides can be sequenced. For suggestions to address this see Note 1. 18S forward primer (the T7 promoter site is underlined): 50 -CATCATCATTTAATACGACTCACTATAGTACCTG GTTGATCCTGCCAGTAGC-30 . 18S reverse primer: 50 -TAATGATCCTTCCGCAGGTTCACCTACGGAAACC-30 14. 10 TBE (Tris-borate-EDTA): 890 mM Tris base, pH 7.6, 890 mM boric acid, 20 mM EDTA. Sterile filter the stock solutions through 0.2μm filters and dilute to 1 as needed. 15. 50 TAE (Tris-acetate-EDTA): 2 M Tris base, 1 M glacial acetic acid, 50 mM EDTA. Sterile filter the stock solutions through 0.2μm filters and dilute to 1 as needed.

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16. Agarose gel standards DNA and RNA ladders. 17. SYBR™ Gold. 18. 6 gel electrophoresis loading dye with 1:1000 SYBR™ Gold. 19. Molecular biology-grade agarose. 20. RNase AWAY™ or other product for eliminating surface RNases. 21. pcDNA 3.1(+) plasmid containing a 28S rRNA gene made by Taoka et al. [13]. Requests for this plasmid can be directed to the authors of this manuscript. 22. XhoI (20,000 units/mL) and 10 buffer. 23. Razor blades for gel excision. 24. Magnet for bead-based purifications. 25. D-tube dialyzer MWCO 3.5 kD MIDI columns (Novagen) or other product for gel purification. 26. E. coli poly(A) polymerase (5000 units/mL) with 10 buffer. 27. 10 mM Adenosine 50 -triphosphate (ATP). 28. Molecular biology-grade bis-acrylamide 29:1 (40% solution): for 5 and 5.8S rRNA gel purification. 29. 3 M NaOAC pH 5.2, nuclease free. 30. Glycogen, RNA grade (20 mg/mL). 31. NEB Quick Ligase Buffer and T4 DNA Ligase (2000 U/μL). 32. ONT SQK-RNA002 (or newer). 2.4 Nanopore Sequencing Hardware and Software, and Reagents

3

Please refer to the Oxford Nanopore Technologies website for details: https://nanoporetech.com/getting-started-with-minion. For new users, the MinION Starter Pack is a good option.

Methods For general RNA work practices see Note 2.

3.1

RNA Isolation

1. Add 4 mL of TRI Reagent per frozen pellet of 5  107 cells, and vortex immediately. 2. Incubate this sample at room temperature for 5 min. 3. Add 400μL BCP (1-bromo-3-chloro-propane) or 200μL CHCl3 (chloroform) per mL of sample, followed by vigorous mixing by inversion. 4. Incubate this mixture at room temperature for 5 min and then mix vigorously again. 5. Spin the mixed sample for 10 min at 12,000  g (4  C).

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6. Pool the aqueous phase in a LoBind Eppendorf tube and add equal volume of isopropanol. 7. Mix the tube followed by incubation at room temperature for 15 min. 8. Spin for 15 min at 12,000  g (4  C). 9. Remove the supernatant. 10. Wash the RNA pellet by adding 750μL 80% ethanol. 11. Spin for 5 min at 12,000  g (4  C). 12. Remove the supernatant. 13. Air-dry the pellet for 10 min. 14. Resuspend the pellet in nuclease-free water (100μL final volume) or TE buffer and quantify RNA using Qubit or similar. 15. The RNA can be stored at 80  C. 3.2 Oligomer Splint Preparation for RNA Adaptation

The sequence for individual oligomers is shown in Subheading. 2.2. 1. To make an oligomer splint adapter, the top and the bottom strands (four annealing reactions total) are hybridized at 10μM each in TNE buffer. 2. Heat the mixture at 75  C for 1 min before slowly cooling to room temperature in a thermocycler.

3.3 Splint Annealing, Ligation, and Cleanup

The following steps are taken from the Oxford Nanopore Direct RNA Sequencing protocol using the SQK-RNA002 kit (see Note 3). 1. Prepare the RNA in nuclease-free water by transferring 1000 ng RNA to a 1.5 mL Eppendorf DNA LoBind tube. Adjust the volume to 9μL with nuclease-free water. Mix thoroughly by flicking the tube to avoid unwanted shearing. Spin down briefly in a microfuge. 2. In a 0.2 mL thin-walled PCR tube, mix the reagents in the order outlined below. The custom rRNA splint adapter mix is made by combining 1μL volume from each of the 10μM oligomer splints (5S, 5.8S, 18S, 28S). Reagent

Volume

NEBNext quick ligation reaction buffer (5)

3.0μL

RNA CS (RCS), 110 nM (optional)

0.5μL

RNA

6.5μL (or 6.0μL if adding RCS) (continued)

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Reagent

Volume

Custom rRNA splint adapter mix

4.0μL

T4 DNA Ligase (2000 U/μL)

1.5μL

Total

15.0μL

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3. Mix by pipetting and spin down. 4. Incubate the reaction for 10 min at room temperature. 5. Add 25μL of nuclease-free H2O to bring up the volume to a total of 40μL. 6. Resuspend the stock of Agencourt RNAClean XP beads by vortexing. 7. Add 72μL (1.8) of resuspended RNAClean XP beads to the reaction and mix by pipetting. 8. Incubate on a Hula mixer (rotator mixer) for 5 min at room temperature. 9. Prepare 200μL of fresh 70% ethanol with nuclease-free water. 10. Spin down the sample and place on the magnetic stand. Keep the tube on the magnet, and pipette off the supernatant. 11. Keep the tube on magnet and wash the beads with 150μL of freshly prepared 70% ethanol without disturbing the pellet as described below. 12. Keeping the magnetic rack on the benchtop, rotate the beadcontaining tube by 180 . Wait for the beads to migrate toward the magnet and form a pellet. 13. Rotate the tube 180 again (back to the starting position) and wait for the beads to pellet. 14. Remove the 70% ethanol using a pipette, and discard. 15. Spin down and place the tube back on the magnet. Pipette off any residual 70% ethanol. 16. Remove the tube from the magnetic rack and resuspend pellet in 20μL nuclease-free water. 17. Incubate for 5 min at room temperature. 18. Pellet the beads on a magnet until the eluate is clear and colorless. 19. Pipette 20μL of eluate into a clean 1.5 mL Eppendorf DNA LoBind tube. 3.4 Nanopore Sequencing Adapter Ligation and Cleanup

The following steps are taken from the Direct RNA Sequencing protocol using the SQK-RNA002 kit, both of which are supplied by Oxford Nanopore Technologies.

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1. In a clean 1.5 mL Eppendorf DNA LoBind tube, mix the reagents in the following order: Reagent

Volume

Splint-annealed RNA from Subheading 3.3 (step 19)

20.0μL

NEBNext quick ligation reaction buffer (5)

8.0μL

RNA adapter (RMX)

6.0μL

Nuclease-free water

3.0μL

T4 DNA Ligase (2000 U/μL)

3.0μL

Total

40.0μL

2. Mix by pipetting. 3. Incubate the reaction for 10 min at room temperature. 4. Resuspend the stock of Agencourt RNAClean XP beads by vortexing. 5. Add 60μL (1.5) of resuspended RNAClean XP beads to the adapter ligation reaction and mix by pipetting. 6. Incubate on a Hula mixer (rotator mixer) for 5 min at room temperature. 7. Spin down the sample and pellet on a magnet. Keep the tube on the magnet, and pipette off the supernatant. 8. Add 150μL of the wash buffer (WSB) from the SQK-RNA002 kit to the beads. Close the tube lid and resuspend the beads by flicking the tube. 9. Return the tube to the magnetic rack, allow beads to pellet, and pipette off the supernatant. 10. Repeat steps 8 and 9. 11. Remove the tube from the magnetic rack and resuspend pellet in 21μL elution buffer from the kit by gently flicking the tube. 12. Incubate for 10 min at room temperature. 13. Pellet the beads on a magnet until the eluate is clear and colorless. 14. Remove and retain 21μL of eluate into a clean 1.5 mL Eppendorf DNA LoBind tube. 15. [Optional] Quantify 1μL of nanopore-adapted rRNA using the Qubit fluorometer RNA HS assay (recovery aim ~200 ng). 3.5 Nanopore Sequencing

For this part we recommend following the instructions provided by Oxford Nanopore Technologies. Below are the steps to follow for the SQK-RNA002 kit. The steps involved here include:

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1. Priming the flow cell. 2. Loading the nanopore-adapted rRNA. 3. Starting the sequencing run. 3.6

Basecalling

Nanopore data are basecalled using Guppy software that is provided by ONT. Guppy has a model to be used for Direct RNA Sequencing runs. Please follow instructions from ONT for using the newest version of Guppy.

3.7 Data Analysis and Visualization

Nanopore basecalling yields sequence reads in FASTQ format. These can be aligned to the reference human rRNA sequences (in FASTA format) using minimap2 (with -ax map-ont setting) [14]. This process will yield a SAM file (human-readable alignment format) that can then be converted to a sorted BAM file (machinereadable alignment file) using SAMtools [15]. Once this is done, the sorted BAM file and the reference sequence FASTA file can be uploaded into Integrative Genomics Viewer (IGV) [16] for visual inspection or used for downstream analyses. IGV can be downloaded from the Broad Institute at no cost (Fig. 2) (see Notes 4 and 5).

3.8 Anticipated Throughput

A conventional nanopore sequencing experiment in our laboratory using GM12878 cell line RNA yielded 470,907 reads, of which 396,048 aligned to one of the four rRNA reference sequences (5S, 5.8S, 18S, and 28S). The breakdown of aligned sequences by specific rRNA is shown in Table 1.

3.9 Optional: Nanopore Sequencing of Human rRNA Copies Composed of Canonical Nucleotides

IVT-derived rRNA strands can be analyzed using nanopore devices and used to train algorithms that identify modified rRNA nucleotides. In summary, RNA synthesis of the canonical controls is performed by in vitro transcription (IVT) using the HiScribe™ T7 Quick High Yield RNA Synthesis kit per the manufacturer’s protocol. For the 5S and 5.8S rRNAs, synthetic DNA oligonucleotides are used for the templates. For the 18S rRNA, a PCR template is amplified from human genomic DNA. For the 28S rRNA, a plasmid template constructed by Taoka et al. [13] is used. The IVT products are purified prior to sequencing. Gel purification is recommended for all IVTs and is required for making 28S rRNA. The library preparation for the purified 5S, 5.8S, and 18S IVT products follows the same procedures described for biological rRNA described starting in Subheading 3.3. The gel-purified 28S IVT product is polyadenylated with E. coli poly(A) polymerase. Following cleanup of the polyadenylated 28S IVT product, it is sequenced using the standard SQK-RNA002 kit protocol.

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Fig. 2 IGV alignment of human 18S rRNA reads. Reads were aligned to the 18S rDNA reference genome sequence [13] using minimap2. The top axis is the nucleotide position in the 1858 nt long 18S rRNA reference sequence [13]. Gray color represents alignment coverage where read and reference sequences agree. In the body of the alignment, purple colors represent insertions, white spaces with black lines represent deletions, and the red, blue, green, and orange colors represent base-specific mismatches in the alignment. Each horizontal line is an individual nanopore strand read. The density plot below the reference coordinates represents alignment coverage. The expanded view shows examples of features within a 40 nt region. These include alignment coverage, insertions, deletions, mismatches, and alignment coverage density, as described above. The consistent C miscalls at position 406 reveal a likely uridine-to-pseudouridine modification

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Table 1 Number of aligned nanopore strand reads per rRNA class

3.9.1 Synthesis of 5S and 5.8S rRNA

rRNA

Number of reads

5S

4465

5.8S

16,058

18S

369,472

28S

6053

1. In two PCR tubes, one for the 5S and the other for the 5.8S rRNA, combine 0.7μL 10 Tris-NaCl-EDTA, 165 pmol (1.65μL of 100μM) of the top-strand oligonucleotide, 33 pmol (0.66μL of 50μM) of bottom-strand oligonucleotide, and nuclease-free H20 (4.69μL) to a total volume of 7μL. 2. Heat the mixture to 75  C for 15 s and slowly cool to 25  C in a thermocycler to anneal the top and bottom DNA strands. 3. Assemble the IVT reactions in PCR tubes using the HiScribe™ T7 Quick High Yield RNA Synthesis kit reagents as follows: Reagent

Volume

Annealed top and bottom DNA strands from step 2

7.0μL

NTP buffer

10.0μL

T7 RNA polymerase mix

2.0μL

RNasin (40 units/μL) or equivalent RNase inhibitor

1μL

Total

20.0μL

4. Allow reaction to proceed for 16 h at 37  C in a thermocycler. 5. Add 3μL of 10 DNase I buffer, 1μL DNase I (RNase-free) (2000 units/mL), and 6.0μL of nuclease-free water to bring the reaction volume to 30μL. Incubate at 37  C for 15 min. 6. Add 10μL of nuclease-free H2O to bring the volume to 40μL. Transfer solution to a 1.5 mL Eppendorf DNA LoBind tube. 7. Follow the procedure for Agencourt RNAClean XP bead purification as described in Subheading 3.9.2 below. Use 1.8 beads (72μL) and use 70% ethanol for washes. 8. Determine the concentration of the eluate using the Qubit™ fluorometer RNA BR assay. See Note 6 regarding concentration determination method and see Note 7 regarding monitoring reactions with gels. 9. Purify the RNA as described in Subheading 3.9.7 below.

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3.9.2 RNA Ampure XP Bead Purifications (for Use in Subheadings 3.9.1, 3.9.2, 3.9.3, 3.9.4, 3.9.5, 3.9.6, 3.9.7, and 3.9.8)

1. Resuspend the stock of Agencourt RNAClean XP beads by vortexing. 2. Based on the size of the rRNA species, different ratios of beads to solution volume are used: (a) 1.8 for 5S and 5.8S rRNA, for example 72μL of beads for a 40μL solution (b) 0.8 for 18S rRNA (c) 0.6 for 28S rRNA Add the appropriate volume of beads and mix by pipetting up and down. 3. Incubate on a Hula mixer (rotator mixer) for 5 min at room temperature. 4. Make 500μL fresh ethanol solution for washes to be used in step 6: (a) For 5 and 5.8S make 80% ethanol: 400μL 100% ethanol +100μL nuclease-free water. (b) For 18S and 28S make 70% ethanol: 350μL 100% ethanol +150μL nuclease-free water. 5. Spin down the sample and pellet on a magnet. Keep the tube on the magnet, and pipette off the supernatant. 6. Keep the tube on the magnet and wash the beads with 150μL of freshly prepared 70% (or 80%) ethanol without disturbing the pellet as described below. 7. Keeping the magnetic rack on the benchtop, rotate the beadcontaining tube by 180 . Wait for the beads to migrate toward the magnet and form a pellet. 8. Rotate the tube 180 again (back to the starting position) and wait for the beads to pellet. 9. Remove the ethanol solution using a pipette, and discard. 10. Spin down and place the tube back on the magnet. Pipette off any residual ethanol solution. 11. Repeat steps 6–10. 12. Remove the tube from the magnetic rack and resuspend pellet in 15μL nuclease-free water by pipetting up and down. 13. Incubate on a Hula mixer (rotator mixer) for 5 min at room temperature. 14. Return to the magnet and wait for the pellet to form and the solution to clear. 15. Remove and retain ~15μL of eluate to a clean 1.5 mL Eppendorf DNA LoBind tube.

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Generate the DNA template with PCR using Q5 master mix (2). 1. Input the Following PCR Protocol into Thermocycler: Initial denaturation: 30 s at 98  C 33 cycles of: 5 s denaturation at 98  C 10 s annealing of 72  C 100 s of extension at 72  C Final extension: 120 s at 72  C 2. Assemble the Reaction in a PCR Tube on Ice: Reagent

Volume

Q5 master mix (2)

25μL

Genomic DNA from human cell line

X μL (2.4μg)

18S forward primer (10μM stock)

2.5μL

18S reverse primer (10μM stock)

2.5μL

Nuclease-free H20

50  (X + 30) μL

Total

50.0μL

3. Run PCR using the protocol described in step 1. 4. Verify the presence of a single amplified band of correct size (~1.8 kb) by running 1μL of the PCR reaction with a standard ladder on a 1% agarose gel using TAE or TBE buffer. Given that the amplification product is in PCR buffer, it runs more slowly than the standard ladder and may appear as high as 3 kb compared with the ladder. 5. Assuming that a robust single band is seen on the gel for the PCR reaction, transfer the solution to a clean 1.5 mL Eppendorf DNA LoBind tube. 6. Follow the procedure for Agencourt RNAClean XP bead purification described in Subheading 3.9.2. Use 0.8 beads (40μL) and use 70% ethanol for washes. 7. Determine the concentration of bead-purified PCR template using the Qubit™ dsDNA BR assay kit. 3.9.4 Synthesis of Canonical 18S rRNA from the PCR-Derived DNA Template

1. Assemble the IVT reaction using the HiScribe™ T7 Quick High Yield RNA Synthesis kit reagents as follows:

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Reagent

Volume

Purified DNA from Subheading 3.9.3

X μL (0.5–1μg)

NTP buffer

10.0μL

T7 RNA polymerase mix

2.0μL

RNasin (40 units/μL) or equivalent RNase inhibitor

1μL

Nuclease-free water

20  (X + 13) μL

Total

20.0μL

2. Run reaction for 2–3 h at RT. 3. Add 3μL of 10 DNase I buffer, 1μL DNase I (2000 units/ mL) (RNase-free), and 6.0μL of nuclease-free water to bring the reaction volume to 30μL. Incubate at 37  C for 15 min. 4. Add 10μL of nuclease-free H2O to bring the volume to 40μL. Transfer solution to a 1.5 mL DNA LoBind tube. 5. Follow the procedure for Agencourt RNAClean XP bead purification as described in Subheading 3.9.2. Use 0.8 beads (32μL) and use 70% ethanol for washes. 6. Determine the concentration using the Qubit™ fluorometer RNA BR assay kit. 7. Purify the RNA as described in Subheading 3.9.7 below. 3.9.5 Preparing a Linearized DNA Plasmid Bearing a 28S Gene

A pcDNA3.1(+) plasmid containing the human 28S gene, described by Taoka et al. [13], is used for synthesizing the canonical 28S rRNA. The plasmid is transformed into competent DH5-alpha E. coli and plasmid DNA isolated using a standard miniprep protocol. Preparation of linearized template: 1. Set up a XhoI restriction digest using 4μg of the plasmid as follows (see Note 8): Reagent

Volume

28S containing plasmid (4μg)

X μL

10 buffer for XhoI

10.0μL

Xho I (20,000 units/mL)

10.0μL

Nuclease-free H20

100  (X + 20)μL

Total

100.0μL

2. Digest for 2 h at 37  C.

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3. Clean up and concentrate the reaction product using Agencourt RNAClean XP bead purification as described in Subheading 3.9.2. Use 0.6 beads (60μL) and use 70% ethanol for washes. 4. Prepare samples for electrophoresis and gel purification (see Note 9). The samples include (A) DNA ladder with 6 DNA loading dye containing 1:1000 SYBR™ Gold; (B) DNA digest with 6 DNA loading dye containing 1:1000 SYBR™ Gold: this is done by using 15μL of bead-purified digested DNA (step 3) + 3μL 6 loading dye containing 1:1000 SYBR™ Gold; and C) ~200 ng uncut plasmid in 6 loading dye containing 1:1000 SYBR™ Gold. 5. Prepare a 0.8% TAE/agarose gel in 1 TAE using a comb that can accommodate 20μL of sample per well. If 50 mL is appropriate for your gel rig, measure out 0.4 g agarose and add it to 50 mL 1 TAE. Microwave the solution to dissolve the agarose. Allow to cool so that it may be handled. Pour into gel rig using appropriate comb. Wait for gel to solidify. Put gel in gel box and cover with 1 TAE to indicated fill line so that there is about 1 cm of buffer covering the gel. Carefully load samples A, B, and C in wells. With the negative electrode toward the base of the gel, run gel at approximately 90 min at 120 V or until bands are well separated. 6. Place the gel on a piece of Saran Wrap in a darkroom. Wear face shield, glasses, lab coat, and gloves. Using a handheld UV source, note positions of the uncut and cut plasmid samples in their respective lanes. Having identified the linearized DNA (~11 kB) band for sample B, excise it using a clean razor blade. Shave off excess gel material from the excised band with the razor blade. Work efficiently to avoid damaging the DNA from prolonged UV exposure. 7. Purify the DNA from the gel. Several commercially available products and do-it-yourself protocols are available for gel purification. We use the D-tube Dialyzer Midi columns according to the manufacturer’s protocol (see Note 10). 8. Regardless of the specific method used for gel purification, the final volume of the purified DNA should be ~15μL in nucleasefree water or TE. 9. Quantitate the DNA concentration using Qubit™ dsDNA BR Assay kit. 3.9.6 Synthesis of 28S rRNA from the Plasmid Template

1. Assemble the IVT reaction using the HiScribe™ T7 Quick High Yield RNA Synthesis kit reagents in a PCR tube as follows:

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Reagent

Volume

Purified linearized DNA from Subheading 3.9, step 5

X μL (1.5–2.0μg)

NTP buffer

20.0μL

T7 RNA polymerase mix

4.0μL

RNasin (40 units/ul) or equivalent RNase inhibitor, optional

2.0μL

Nuclease-free water

40  (X + 26)μ L

Total

40.0μL

2. Run the reaction for 3 h at 37  C. 3. Add 5μL of 10 DNase I buffer, 2μL DNase I (2000 units/ mL), and 3.0μL of nuclease-free water to bring the reaction volume to 50μL. Incubate at 37  C for 15 min. 4. Transfer solution to a 1.5 mL DNA LoBind tube. 5. Follow the procedure for Agencourt RNAClean XP bead purification as described in Subheading 3.9.2. Use 0.6 beads (30μL) and use 70% ethanol for washes. 6. Determine the concentration using the Qubit fluorometer RNA BR assay kit. 7. Gel purify the RNA as described in Subheading 3.9.7 below. 3.9.7 Purification of Canonical Transcripts

1. The IVT products must be purified prior to sequencing. In our experience, gel purification is the method that results in the best read coverage and throughput. This is especially true for the 28S IVT product. Other options for purification include phenol/chloroform extraction, followed by ethanol precipitation, and use of spin columns (see Note 11). 2. The general scheme for gel purification involves running the IVT sample along with a sizing ladder on a denaturing PAGE or a non-denaturing agarose gel. The gel is pre- or post-stained with SYBR™ Gold and the appropriately sized RNA (121 nt for 5S, 155 nt for 5.8S, 1.8 kb for 18S, and 5 kb for 28S) is identified and excised. 3. Following gel excision, we have had success using the D-tube Dialyzers MWCO 3.5 kD according to the manufacturer’s protocol to purify the RNA. This involves electroelution and ethanol precipitation. Other gel purification kits require different steps and may need to be optimized. 4. Following purification determine the concentration of the sample using Qubit.

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5. Once purified, the 5S, 5.8S, and 18S RNA are ready for sequencing (Subheading 3.9.9). 6. The purified 28S RNA is next polyadenylated as described below in Subheading 3.9.8. 3.9.8 Polyadenylation of the 28S rRNA IVT Product

1. Set up a 40μL reaction containing 3–4μg of gel-purified canonical 28S rRNA. Reagent

Volume

Gel-purified canonical 28S rRNA from Subheading 3.9.6

X μL (3.0–4.0μg)

10 buffer

4.0μL

10 mM ATP

4.0μL

E. coli poly(A) polymerase

2.0μL

Nuclease-free H20

40  (X + 10) μL

Total

40.0μL

2. Run for 1 h at 37  C. 3. Follow the procedure for Agencourt RNAClean XP bead purification as described in Subheading 3.9.2. Use 0.6 beads (24μL) and use 70% ethanol for washes. 4. Measure the concentration using a Qubit™ BR RNA Assay kit. One μg of this material will be used for the library (see Subheading 3.9.10). 3.9.9 Library Preparation Using Splint Adapters for Canonical 5S, 5.8S, and 18S rRNAs

Splint annealing follows the procedures described in Subheading 3.2. Please note that the 28S splint is not needed. The custom rRNA splint adapter mix listed below is made by combining 1μL volume from each of the 10μM oligomer splints (5S, 5.8S, 18S). The following amounts of purified IVT rRNA are recommended: 500 ng of 5S, 500 ng of 5.8S, and 350 ng of 18S. 1. Set up the first ligation as follows: Reagent

Volume

NEBNext quick ligation reaction buffer (5)

3.0μL

RNA CS (RCS), 110 nM (optional)

0.5μL

IVT rRNAs for 5S, 5.8S, 18S

7.5μL (or 7.0μL if adding RCS)

Custom rRNA splint adapter mix

3.0μL

T4 DNA Ligase (2000 U/μL)

1.5μL

Total

15.0μL

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2. Follow the procedures beginning in Subheading 3.3, step 3, through Subheading 3.7 for the sequencing, basecalling, data analysis, and visualization of the data. They are the same as described for the biological rRNAs. For information on expected throughput see Note 12. 3.9.10 Library Preparation for Polyadenylated Canonical 28S rRNA

Because the sample is polyadenylated, the standard ONT mRNA library protocol (SQK-RNA002) is used. We suggest starting with 1μg of polyadenylated 28S IVT RNA for the library. 1. Set up the first ligation as follows in a 1.5 mL DNA LoBind tube. Reagent

Volume

NEBNext quick ligation reaction buffer (5)

3.0μL

RNA CS (RCS), 110 nM (optional)

0.5μL

RNA-1μg polyadenylated 28S IVT rRNA X μL (or X–0.5μL if adding RCS) RTA adapter

1.0μL

Nuclease-free H20

Bring to 13.5μL

T4 DNA Ligase (2000 U/μL)

1.5μL

Total

15.0μL

2. Follow the procedures beginning in Subheading 3.3, step 3, through Subheading 3.7 to complete the library preparation, sequencing, basecalling, data analysis, and visualization of the data. For information on expected throughput see Note 13.

4

Notes 1. Concerning 50 coverage of canonical (IVT) rRNAs, in our current design, the 50 -most ~12 nucleotides do not get sequenced. One improvement to address this would be to design the IVT template to produce a transcript with an additional ~15 nt 50 of the rRNA start. For the 5S and 5.8S rRNAs, the bottom oligonucleotides could be redesigned. For example, for 5S the top strand would remain the same: 50 -CATCATCATTTAATACGACTCACTATAG-30 But the bottom strand could be: 50 -AAAGCCTACAGCACCCGGTATTCCCAGGCGG TCTCCCATCCAAGTACTAACCAGGCCCGACCCTGC TTAGCTTCCGAGATCAGACGAGATCGGGCGCGTTC AGGGTGGTATGGCCGTAGANNNNNNNNNNNNNNN CTATAGTGAGTCGTATTAAATGATGATG-30

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where N is a randomly chosen sequence. This would produce sequence 50 of the start of the rRNA. For 18S, one could design the forward PCR primer to generate an amplicon with sequence 50 to the start of the rRNA. This is shown below where N indicates nucleotides between the T7 promoter and 18S start. T7 promoter 18S start 50 -CATCATCATTTAATACGACTCACTATAGNNNNNNNNNNNNTACCTGGTTGATCCTGCCAGTAGC-30

To pursue this, PCR optimization will be required. Likely the best results would come from using the sequences just 50 to the 18S start found in the genomic DNA sequence for the positions indicated by “N” above. 2. When working with RNA, wear gloves, use pipette tips with barrier filters, use a designated RNA bench if possible, use designated pipetman, clean surfaces with RNase AWAY™ or other RNase surface decontaminant, and use nuclease-free water. 3. There are some alterations (highlighted in bold) in this protocol to ensure recovery of the smaller rRNA molecules (5S and 5.8S). 4. It is important to be mindful that there are 17 gene copies for 5S rRNA, 6 gene copies for 5.8S rRNA, 5 gene copies for 18S rRNA, and 5 gene copies for 28S rRNA in the human nuclear genome. Some of the gene copies have identical sequences (e.g., 5S and 5.8S), and others vary in sequence composition (e.g., 18S and 28S). For the purpose of alignments, we use one gene copy for each rRNA. Our recommendation is to use the reference sequences curated by Taoka et al. [13]. 5. Conventional nanopore sequencing cannot read ~12 nucleotides at the 50 end of each strand due to the architecture of the helicase/pore interface. 6. Concerning concentration determination: Nanodrop vs. Qubit. We have noticed that using of nanodrop to estimate the concentration of IVT-generated rRNA overestimates the concentration of the transcript. We think this is because of unincorporated ribonucleotides remaining after cleanup contributing to the measurement. This issue is avoided by determining the concentration using Qubit. This also is the case following polyadenylation where ATP carried over from the reaction contributes to the concentration read by nanodrop overestimating the actual amount of transcript present.

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7. The production of IVT rRNAs for sequencing is a multistep process. Therefore, particularly when running the procedure for the first time, or when troubleshooting, it is worthwhile to monitor reactions by running small amounts of reactions on gels to assess the quality of the RNAs produced. The downside of this is that it takes time and can decrease the amount of final material available for sequencing. If this is an issue, multiple reactions can be set up and pooled to insure adequate amounts of product. 8. Alternatively, SpeI may be used to linearize the plasmid for IVT [13]. 9. Regarding the gel purification of linearized 28S plasmid DNA following XhoI digest, purification of the linearized DNA is necessary because uncut plasmid present can dominate the IVT reaction producing unwanted concatenated sequences. 10. We have used the D-tube Dialyzers MWCO 3.5 kD (Novagen) Midi columns for gel purifications of DNA and RNA according to the manufacturer’s protocol. We are able to get good recovery; however the procedure is relatively lengthy compared to some other gel purification kits we have not evaluated. It is best to run the electroelution steps in TAE as opposed to TBE. After electroelution the sample is ethanol precipitated with 3 volume of ethanol overnight at 20  C. The ethanol precipitation includes 0.3 M NaOAc pH 5.2 and 1μL of glycogen is added to help follow the position of the pellet. Following precipitation and 70% ethanol washes, the pellet is dried, brought up in 15μL nuclease-free water, and quantitated by Qubit. 11. In our hands, particularly for long transcripts, such as the 28S rRNA, gel excision and electroelution of the appropriate-sized transcript from a gel help to ensure complete sequence coverage. Smaller incomplete transcripts are more efficiently captured during nanopore sequencing and can prevent the detection of full-length product in a mixture. We recommend this for the purification of all IVT rRNAs. In the case of 5S and 5.8S, the unpurified IVT reactions can be run on denaturing PAGE gels (8%) prior to excision. For the 18S and 28S IVT reactions, the samples are run on 0.8% TAE agarose gels. For these gels care is taken to clean the gel box and combs with RNase away before casting the gel. Given that the IVT reactions contain incomplete products and due to losses in purification, run a minimum of 4μg of the IVT reaction for purification. For best success, for each rRNA IVT run several lanes of 4μg IVT, excise the appropriate band slices, purify, and pool. We recommend running a gel to verify that a transcript of the correct size is produced and that the purification method

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chosen is effective in enriching for the transcript size of interest. It is always better to have too much material than too little. When possible, set up multiple IVT reactions to safeguard against losses and to ensure success. 12. Regarding throughput for IVT runs for 5S, 5.8S, and 18S rRNA, in the past, the number of reads per flow cell was especially low for 5S (3589 reads) and 5.8S rRNA (5471 reads) compared to 18S rRNA (90,081 reads). The current protocol as described herein uses 500 ng per flow cell of gel-purified 5S and 5.8S canonical rRNA which should improve those yields. 13. 28S canonical rRNA expected throughput and explanation of why a plasmid-based template was used: The expected throughput using the current protocol is 291,929 reads. We found that using the linearized 28S-containing plasmid for the template, followed by polyadenylation of the resultant IVT product, gave more reads and better quality alignments than when we used a PCR-based template for IVT. PCR amplification of the 28S was challenging. We were not able to generate an amplicon using the Q5 polymerase (NEB) but were successful using the Primestar GXL polymerase (Takara). Using of RNA synthesized from the 28S PCR template followed by the splint ligation resulted in 4230 reads compared with 291,929 for the method described starting in Subheading 3.9.5. References 1. Deamer D, Akeson M, Branton D (2016) Three decades of nanopore sequencing. Nat Biotechnol 34:518–524 2. Garalde DR, Snell EA, Jachimowicz D et al (2018) Highly parallel direct RNA sequencing on an array of nanopores. Nat Methods. https://doi.org/10.1038/nmeth.4577 3. Workman RE, Tang AD, Tang PS et al (2019) Nanopore native RNA sequencing of a human poly(A) transcriptome. Nat Methods 16:1297–1305 4. Smith AM, Jain M, Mulroney L et al (2019) Reading canonical and modified nucleobases in 16S ribosomal RNA using nanopore native RNA sequencing. PLoS One 14:e0216709 5. Kim D, Lee J-Y, Yang J-S et al (2020) The architecture of SARS-CoV-2 transcriptome. Cell 181 e10:914–921 6. Viehweger A, Krautwurst S, Lamkiewicz K et al Direct RNA nanopore sequencing of fulllength coronavirus genomes provides novel

insights into structural variants and enables modification analysis. https://doi.org/10. 1101/483693 7. Simpson JT, Workman RE, Zuzarte PC et al (2017) Detecting DNA cytosine methylation using nanopore sequencing. Nat Methods. https://doi.org/10.1038/nmeth.4184 8. Methylation calling — Medaka 1.0.3 documentation. https://nanoporetech.github.io/ medaka/methylation.html. Accessed 10 Jul 2020 9. Liang X-H, Liu Q, Fournier MJ (2009) Loss of rRNA modifications in the decoding center of the ribosome impairs translation and strongly delays pre-rRNA processing. RNA 15:1716–1728 10. King TH, Liu B, McCully RR, Fournier MJ (2003) Ribosome structure and activity are altered in cells lacking snoRNPs that form pseudouridines in the peptidyl transferase center. Mol Cell 11:425–435

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11. Narla A, Ebert BL (2010) Ribosomopathies: human disorders of ribosome dysfunction. Blood 115:3196–3205 12. Lafontaine DLJ (2015) Noncoding RNAs in eukaryotic ribosome biogenesis and function. Nat Struct Mol Biol 22:11–19 13. Taoka M, Nobe Y, Yamaki Y et al (2018) Landscape of the complete RNA chemical modifications in the human 80S ribosome. Nucleic Acids Res 46:9289–9298

14. Li H (2018) Minimap2: pairwise alignment for nucleotide sequences. Bioinformatics. https:// doi.org/10.1093/bioinformatics/bty191 15. Li H, Handsaker B, Wysoker A et al (2009) The sequence alignment/map format and SAMtools. Bioinformatics 25:2078–2079 16. Robinson JT, Thorvaldsdo´ttir H, Winckler W et al (2011) Integrative genomics viewer. Nat Biotechnol 29:24–26

Part III Next-Generation Sequencing Approaches to Detect and Capture Modified RNAs

Chapter 5 AlkAniline-Seq: A Highly Sensitive and Specific Method for Simultaneous Mapping of 7-Methyl-guanosine (m7G) and 3-Methyl-cytosine (m3C) in RNAs by High-Throughput Sequencing Virginie Marchand, Lilia Ayadi, Vale´rie Bourguignon-Igel, Mark Helm, and Yuri Motorin Abstract Epitranscriptomics is an emerging field where the development of high-throughput analytical technologies is essential to profile the dynamics of RNA modifications under different conditions. Despite important advances during the last 10 years, the number of RNA modifications detectable by next-generation sequencing is restricted to a very limited subset. Here, we describe a highly efficient and fast method called AlkAniline-Seq to map simultaneously two different RNA modifications: 7-methyl-guanosine (m7G) and 3-methyl-cytosine (m3C) in RNA. Our protocol is based on three subsequent chemical/enzymatic steps allowing the enrichment of RNA fragments ending at position n + 1 to the modified nucleotide, without any prior RNA selection. Therefore, AlkAniline-Seq demonstrates an outstanding sensitivity and specificity for these two RNA modifications. We have validated AlkAniline-Seq using bacterial, yeast, and human total RNA, and here we present, as an example, a synthetic view of the complete profiling of these RNA modifications in S. cerevisiae tRNAs. Key words 7-Methyl-guanosine, 3-Methyl-cytosine, High-throughput sequencing, RNA modification mapping, Bacteria

1

Introduction RNA modifications are extremely diverse throughout evolution and present both in noncoding and in coding RNAs. Mapping of these RNA modifications at single-nucleotide resolution represents a big step forward since only 7 (m6A, m6Am, m1A, m5C, hm5C, Nm, and ψ) [1–9] out of >150 RNA modifications can be detected by high-throughput sequencing. Therefore, there is an urgent need to develop new methods for analyzing many still insufficiently studied RNA modifications. One attractive candidate is 7-methylguanosine (m7G), a modified nucleotide which does not disrupt

Mary McMahon (ed.), RNA Modifications: Methods and Protocols, Methods in Molecular Biology, vol. 2298, https://doi.org/10.1007/978-1-0716-1374-0_5, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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Virginie Marchand et al. A. CULTURE, RNA EXTRACTION AND QUALITY CONTROL Culture human HEK cells

bacteria E. coli

TRIzol reagent

RNA extraction

yeast S. cerevisiae

Hot acid phenol extraction RNA precipitation

RNA precipitation

RNA quantity and quality control 10 9 8 7 6 5 4 3 2 1 0

Total RNA from yeast S. cerevisiae

absorbance

RNA quality control (Capillary electrophoresis)

18S 25S rRNA

tRNA

tRNA

wavelength

B. ALKANILINE-SEQ AlkAniline Treatment

25S 18S

260 270 280 290 300 310 320 330

220 230 240 250

RNA quantity (Spectrophotomer)

Ladder (nt)

5’ P

OH3’

Total RNA from yeast S. cerevisiae

Mild Alkaline Hydrolysis 5’ P

P3’ 5’HO

OH 3’

RNA precipitation

OH3’ 5’HO

OH 3’

RNA precipitation

OH 3’

RNA precipitation

Extensive 5’ and 3’ dephosphorylation 5’HO

Aniline cleavage 5’HO

OH3’ 5’HO

X 3’ 5’ P

Library preparation and clean-up PCR

3’ Adaptor ligation P5

RT primer hybridization P5 P5

BC P7

5’ Adaptor ligation P5 P5

BC P7 BC P7

First strand cDNA synthesis

Clean up

Library quantification and quality control Library quality control (Capillary electrophoresis)

Library quantity (Fluorometer)

Ladder (bp)

C. BIOINFORMATIC PIPELINE

P5

N₊₁

Scoring

BC P7

Normalized cleavage = 6*1000/14 = 428 units Stop-rao = 6/(6+2) = 0.75 Starng reads = 6 6 Passing reads = 2

N₊₁

5’-end counts

Multiplexing and Sequencing

Total 5’-ends = 14 posions

Bioinformatic workflow

parameters

Trimming (Trimmomac) TruSeq3-SE.fa:2:30:7 LEADING:30 TRAILING:30 SLIDINGWINDOW:4:15 MINLEN:8 AVGQUAL:30

Alignment (Bowe 2.0) -D 15 -R 2 -N 0 -L 10 -i S,1,1.15

Keep only mapped reads

Conversion to BED (bedtools)

Count 5’-ends and coverage (awk)

Score calculaon and graphs (R)

samtools view -h -F 4 -b

Fig. 1 Overview of experimental and analysis steps of AlkAniline-Seq protocol. (a) Human total RNA is extracted using the standard TRIzol protocol; extraction of total RNAs from bacterial and yeast cells is

Mapping of m7G and m3C in RNA

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base-pairing and thus may be potentially present even in proteincoding regions of mRNAs. A specific detection of m7G in RNA was described in the 1970’s and consists of a two-step chemical approach, combining sodium borohydride (NaBH4) reduction followed by aniline cleavage [10, 11]. However, we found that coupling these reactions to a next-generation sequencing technique leads to a high background signal and numerous false discovery hits [12]. Therefore, we designed a completely novel strategy (named AlkAniline-Seq) to map m7G (and as revealed later, also m3C) residues in RNA by combining a three-step protocol: a mild alkaline hydrolysis, an extensive 30 - and 50 -dephosphorylation, and finally aniline cleavage of the RNA chain. A key feature in our protocol is the direct 50 -adapter ligation to the 50 -phosphate resulting from phosphodiester bond scission induced by aniline cleavage at the RNA abasic site (Fig. 1). In contrast to other NGS approaches, this provides the basis for both high sensitivity and specificity of the AlkAniline-Seq technology [12]. Here, we describe in detail the procedure for AlkAniline-Seq from the RNA extraction to the bioinformatic analysis, and as an example, we show the complete profiling of m7G/m3C tRNA modifications in yeast S. cerevisiae (Fig. 2).

2

Materials Prepare all solutions using RNase-free water. Wear gloves to prevent degradation of RNA samples by RNases.

ä Fig. 1 (continued) performed using the “hot acid phenol” protocol. Quantification and quality of RNA are assessed by spectrophotometer and RNA integrity is evaluated using capillary electrophoresis. (b) RNAs are subjected to mild alkaline hydrolysis, generating fragments of about 200–300 nt in length. Fragments are extensively dephosphorylated to remove all possible 50 - and 30 -phosphate residues from RNA. Afterwards, RNAs are subjected to aniline cleavage at the abasic sites generated by decomposition of m7G/m3C residues upon alkaline hydrolysis. Library preparation is performed by direct ligation of pre-adenylated 30 -adapter, followed by RT primer annealing and ligation of 50 -adapter and RT primer extension. The resulting cDNA is converted to sequencing library by second-strand DNA synthesis and limited PCR step for barcoding and inclusion of Illumina P5 and P7 sequences. Quantification of the library is performed using a fluorometer and the quality is assessed by capillary electrophoresis. (c) Scoring of AlkAniline-Seq signals is done by calculation of both normalized cleavage (ratio of reads starting at a given RNA position to total number of reads mapped to this RNA) and stop-ratio, corresponding to the ratio of reads starting at a given position to reads overlapping it. Normalized cleavage provides exceptional selectivity, while stop-ratio is very sensitive, but captures numerous false-positive hits. Bioinformatic analysis consists of trimming step to keep adapter-free reads, bowtie 2.0 end-to-end alignment followed by counting of reads mapped to different positions in RNA and calculation of AlkAniline-Seq scores

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Fig. 2 AlkAniline-Seq signals for m7G and m3C residues in yeast S. cerevisiae tRNAs. (a) m7G residues detected in 10 m7G46-modified yeast tRNAs. Initiator tRNAMeti also has m7G46 residue but is not illustrated here since it has a low coverage in sequencing. tRNA positions are numbered sequentially from 50 - to 30 -end; however specific tRNA numbering scheme (with 17a, 20ab, and extra nucleotides in variable loop) is not

Mapping of m7G and m3C in RNA

2.1 Total RNA Extraction 2.1.1 Yeast and Bacteria Total RNA Extraction by Hot Acid Phenol

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1. Yeast or bacteria cell culture (10 mL of culture grown to an OD600 of 0.7–2). 2. RNase-free 1.5 mL microcentrifuge tubes. 3. RNase-free water. 4. AE buffer: 50 mM NaOAc in water, pH 5.2, 10 mM EDTA. 5. 10% (w/v) SDS. 6. Acid phenol, pH 4.5. 7. Phenol:chloroform:isoamyl alcohol mix (25:24:1, v/v). 8. Chloroform. 9. 3 M NaOAc, pH 5.2. 10. 96% Ethanol. 11. 80% Ethanol. 12. Dry ice. 13. Refrigerated tabletop centrifuge. 14. Water bath or heating block set to 65  C.

2.1.2 Human Total RNA Extraction by TRIzol™

1. Human HEK 293 cells (8–10  106 cells grown to 90% confluence in a cell culture dish). 2. 1 PBS (8.1 mM Na2HPO4, 1.47 mM KH2PO4, 137 mM NaCl, and 2.7 mM KCl). 3. RNase-free 1.5 mL microcentrifuge tubes. 4. RNase-free water. 5. TRIzol™ reagent. 6. Chloroform. 7. Refrigerated tabletop centrifuge. 8. Isopropanol. 9. Glycoblue™ coprecipitant: 15 mg/mL. 10. 75% Ethanol. 11. Cell scraper.

ä Fig. 2 (continued) respected. Bar plot represents normalized cleavage score for a given RNA species in four yeast strains (WT—wild-type BY 4741 strain, ΔTRM140—strain deleted for m3C32:tRNA-methyltransferase, ΔTRM8/ΔTRM82—two strains deleted for genes encoding subunits of heterodimeric m7G46:tRNA-methyltransferase). Position of m7G46 is indicated by a gray dot. Some of the yeast tRNA species also contain adjacent dihydrouridine (D47), purple dot. This residue is also partially cleaved during mild alkaline hydrolysis (see small peaks at position 47) but becomes visible only in the absence of a major signal corresponding to m7G. Some weak cleavage signals are also visible in the region 16–20, and correspond to D residues in the D-loop of tRNA. (b) Two yeast tRNAs containing m3C32 residues (shown as pink dot). Closely related isoforms of yeast tRNASer (AGA, CGA, and TGA) are collapsed into unique tRNASer(NGA) sequence. tRNA positions are numbered as in (a)

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2.2 RNA Quantification and Quality Assessment

1. UV-visible spectrophotometer for small volumes: Any kind of UV-visible spectrophotometer allowing measurements of 1 μL samples. We use NanoDrop™ 2000. 2. RNase-free 1.5 mL microcentrifuge tubes. 3. RNase-free water. 4. Agilent 2100 Bioanalyzer or 2200 TapeStation (Agilent Technologies) or Experion (BioRad) or LabChip GX (Caliper): We use an Agilent 2100 Bioanalyzer. 5. Agilent RNA 6000 Pico kit (quantitative range 50–5000 pg/μL). 6. Chip priming station. 7. Tabletop centrifuge.

2.3

AlkAniline-Seq

2.3.1 Alkaline Hydrolysis

1. Sodium bicarbonate buffer: 100 mM Sodium bicarbonate, pH 9.2. 2. RNase-free water. 3. Individual RNase-free 0.2 mL PCR tubes. 4. PCR thermal cycler (we use Agilent SureCycler 8000). 5. RNase-free 1.5 mL microcentrifuge tubes. 6. 96% Ethanol. 7. 15 mg/mL Glycoblue™ coprecipitant. 8. 3 M NaOAc, pH 5.2. 9. 80% ethanol. 10. Dewar containing liquid nitrogen. 11. Refrigerated tabletop centrifuge.

2.3.2 Extensive RNA Dephosphorylation and RNA Precipitation

1. RNase-free 0.2 mL PCR tubes, strips of 8. 2. Flat PCR caps, strips of 8. 3. PCR thermal cycler. 4. 5 U/μL Antarctic phosphatase. 5. 40 U/μL RiboLock RNase inhibitor. 6. RNase-free 1.5 mL microcentrifuge tubes. 7. Phenol:chloroform:isoamyl alcohol mix (25:24:1, v/v/v). 8. Chloroform. 9. 15 mg/mL Glycoblue™ coprecipitant. 10. 96% Ethanol. 11. 3 M NaOAc, pH 5.2. 12. 80% Ethanol. 13. Refrigerated tabletop centrifuge.

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1. Aniline: 1 M in acetic acid, pH 4.5. 2. RNase-free 1.5 mL microcentrifuge tubes. 3. RNase-free water. 4. Agitating heating block (we use Eppendorf Thermomixer®). 5. 15 mg/mL Glycoblue™ coprecipitant. 6. 96% Ethanol. 7. 3 M NaOAc, pH 5.2. 8. 80% Ethanol. 9. Refrigerated tabletop centrifuge.

2.4 Library Preparation

1. NEBNext® Multiplex Small RNA Library Prep Set for Illumina® (set 1 or 2) (see Note 1). 2. RNase-free 0.2 mL PCR tubes, strips of 8. 3. Flat PCR caps, strips of 8. 4. Thermal cycler.

2.5 Library Purification

1. GeneJET PCR Purification Kit or equivalent. 2. RNase-free 1.5 mL microcentrifuge tubes. 3. RNase-free 1.5 mL DNA low-binding tubes. 4. Tabletop centrifuge.

2.6 Library Quantification and Quality Assessment

1. Any kind of fluorometer able to quantify DNA library with high sensitivity (e.g., Qubit® 2.0 fluorometer). 2. Qubit® dsDNA HS Assay kit (0.2–100 ng). 3. Thin-walled polypropylene tubes of 500 μL compatible with the fluorometer (e.g., Qubit® Assay Tube or Axygen® PCR-05-C tubes). 4. Agilent 2100 Bioanalyzer (Agilent Technologies). 5. Agilent HS DNA kit (quantitative range 5–500 pg/μL). 6. Chip priming station. 7. RNase-free 1.5 mL microcentrifuge tubes. 8. Tabletop centrifuge.

2.7 Library Sequencing

1. Any kind of Illumina sequencers (starting from MiSeq to NovaSeq). 2. Any appropriate sequencing kit for a single read length of 35–50 nt.

2.8 Bioinformatic Analysis

1. Unix (Linux) server (we use Illumina Compute Dell R815 server).

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2. Adapter trimming software Trimmomatic (current version 0.36 http://www.usadellab.org/cms/?page¼trimmomatic). 3. Alignment software Bowtie 2.0 (current version 2.2.9 http:// bowtie-bio.sourceforge.net/bowtie2/index.shtml). 4. Samtools (current version 1.9, http://www.htslib.org/doc/ samtools.html). 5. Bedtools v2.25.0 est/index.html).

(https://bedtools.readthedocs.io/en/lat

6. R environment ver. 3.3.3 for calculations of normalized cleavage and stop-ratio scores and data analysis.

3

Methods

3.1 Total RNA Extraction 3.1.1 Yeast and Bacteria Total RNA Extraction by Hot Acid Phenol

The following protocol details total RNA isolation from yeast/ bacteria using hot acid phenol and is adapted from [13]. 1. Transfer yeast/bacteria cell culture in 1.5 mL microcentrifuge tubes and pellet cells by centrifugation at 1200  g for 5 min at room temperature. Discard the supernatant. 2. Resuspend cells in 1 mL of RNase-free water. Centrifuge for 1 min at full speed at room temperature. Discard the supernatant. 3. Resuspend the cell pellet in 400 μL of AE buffer. 4. Add 40 μL of 10% SDS and vortex until the pellet is completely resuspended. 5. Add 440 μL of acid phenol. Vortex. 6. Incubate for 4 min at 65  C and then cool the mixture rapidly on dry ice for 2–3 min. 7. Centrifuge the samples for 10 min at full speed at room temperature. Carefully transfer the aqueous (upper) phase to a new 1.5 mL microcentrifuge tube. 8. Add 420 μL of phenol:chloroform:IAA, vortex, and centrifuge for 10 min at full speed at room temperature. 9. Transfer the aqueous phase to a new 1.5 mL centrifuge tube. Add 400 μL of chloroform. Vortex and centrifuge at full speed at room temperature for 10 min. 10. Transfer the aqueous phase to a new 1.5 mL centrifuge tube. Add 40 μL of 3 M NaOAc and 1 mL of 96% ethanol. Place at 80  C for at least 30 min. 11. Centrifuge for 30 min at full speed at 4  C. 12. Discard the supernatant and wash pellet with 500 μL of 80% ethanol. 13. Centrifuge for 5 min at full speed at 4  C.

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14. Discard the supernatant; centrifuge again for a short spin. 15. Remove any remaining liquid with a pipette. 16. Incubate samples with open lid for 2 min at 37  C or 5 min at room temperature. 17. Resuspend the pellet with 10 μL of RNase-free water and pool your samples. 18. Quantify yeast or bacteria total RNA samples by measuring A260nm using a UV spectrophotometer (see Note 2) (see Subheading 3.2). Check the quality of the corresponding samples by using the Agilent 2100 Bioanalyzer (see Subheading 3.2). 3.1.2 Human Total RNA Extraction by TRIzol™

Isolate total RNA using TRIzol™ following the manufacturer’s instructions. 1. Wash HEK 293 cells grown in a cell culture dish with 1.5 mL 1 PBS. 2. After PBS removal, add 1 mL of TRIzol™ directly to the cell culture dish to lyse the cells, scrap the cells, and pipet the lysate up and down several times to homogenize. 3. Incubate for 5 min at room temperature to get complete RNP dissociation. 4. Add 200 μL chloroform, vortex, and incubate for 5 min at room temperature. 5. Centrifuge for 15 min at 12,000  g at room temperature. 6. Transfer the aqueous phase containing RNA to a new 1.5 mL microcentrifuge tube, add 500 μL of isopropanol and 1 μL of Glycoblue™, and mix by inverting the tube up and down several times. 7. Incubate for 10 min at room temperature. 8. Centrifuge for 10 min at 12,000  g at 4  C. 9. Discard the supernatant and wash pellet with 1 mL of 75% ethanol. 10. Centrifuge for 5 min at 12,000  g at 4  C. 11. Discard the supernatant, and centrifuge again for a short spin. 12. Remove any remaining liquid. 13. Incubate with open lid for 2 min at 37  C or 5 min at room temperature. 14. Resuspend the pellet with 50 μL of RNase-free water. 15. Quantify human total RNA samples by measuring A260nm using a UV spectrophotometer (see Subheading 3.2). Check

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the quality of your samples by using the Agilent 2100 Bioanalyzer (see Subheading 3.2). 3.2 RNA Quantification and Quality Control 3.2.1 RNA Quantification

Carry out all procedures at room temperature.

1. On a Nanodrop 2000 start screen, select the “Nucleic Acid” application. 2. After the wavelength verification test, select the type of sample to measure, in this case “RNA.” 3. Prepare the blank/buffer solution used for sample resuspension but without any trace of RNA (e.g., RNase-free water). 4. Load 1 μL of the blank solution to the bottom pedestal, lower the arm, and click on the “Blank” button. 5. Wipe the upper and lower pedestal using a dry wipe, load 1 μL of one of your samples of interest to the bottom pedestal, lower the arm, and click “Measure.” 6. Analyze the data obtained for your different RNA samples. For “pure” RNA, the ratio of A260/A280 should be 2; the ratio of A260/A230 should be in the range of 1.8–2.2 (see Notes 3 and 4).

3.2.2 RNA Quality Assessment

1. Before starting, equilibrate all solutions of the kit at room temperature for at least 30 min in the dark. Vortex and spin down before use. 2. Transfer 550 μL of gel matrix (red-cap vial) into a spin filter provided in the kit. 3. Centrifuge for 10 min at 1500  g at room temperature. 4. Prepare 65 μL aliquots of the gel and store them at 4  C for a maximum of 1 month. 5. Prepare the gel-dye mix by mixing 1 μL of RNA dye concentrate to a gel aliquot. 6. Centrifuge for 10 min at 13,000  g at room temperature. 7. Dilute your RNA samples for quantification on the Nanodrop to 3–5 ng/μL with RNase-free water to be within the optimal range concentration of the assay. 8. Add 1 μL of your diluted RNA samples to 11 different 1.5 mL tubes already containing 5 μL of RNA marker (green-cap vial) (see Note 5). Mix by pipetting up and down. 9. Mix 1 μL of the ladder (see Note 6) with 5 μL of RNA marker (green-cap vial). Mix by pipetting up and down.

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10. Prepare the chip priming station. Adjust the syringe clip to the highest top position. 11. Load 9 μL of the gel-dye mix in the well marked with a “G” surrounded by a black circle. 12. Close the chip priming station properly and press the plunger of the syringe until it is held by the clip. 13. Wait for 30 s and then release the clip. 14. Wait for 5 s until the plunger stops and pull it slowly back to the 1 mL position of the syringe. 15. Open the chip priming station and load 9 μL of the gel-dye mix in the two other wells marked “G.” 16. Load 9 μL of the conditioning solution (white-cap vial) in the well marked “CS.” 17. Load 6 μL of the diluted ladder in the well marked with a ladder. 18. Load 6 μL of the diluted RNA samples in the wells marked 1–11. 19. Inspect the chip and make sure that no liquid spills are present on the edges of the wells. 20. Insert the chip in the Agilent 2100 Bioanalyzer and close the lid (see Note 7). 21. Select the following assay “Eukaryote Total RNA Pico series II” in the 2100 Expert Software screen. 22. Press “Start” to begin the chip to run (see Note 8). 23. After the run, immediately remove the chip and clean the electrodes with the electrode cleaner filled with 350 μL of RNase-free water. 24. Analyze the results of the chip (see Fig. 1). 3.3

AlkAniline-Seq

3.3.1 Alkaline Hydrolysis

1. Prepare one 1.5 mL tube per sample to be analyzed (“precipitation tube”) containing 10 μL of NaOAc, 1 μL of Glycoblue™, and 1 mL of 96% ethanol for subsequent precipitation of the sample (store at 20  C until further use). 2. Dilute your RNA samples to a concentration of 10 ng/μL with RNase-free water. 3. To individual PCR tubes, add 10 μL of each of your diluted RNA samples, and keep on ice until further use. 4. Add 10 μL of bicarbonate buffer and mix by pipetting up and down. 5. Incubate in a thermal cycler preheated at 95  C. Start a timer and incubate for 5 min (see Note 9). 6. Proceed with the next sample every 30 s.

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7. Stop each reaction after the required time at 95  C by spinning down the PCR microtube and add the whole sample into the corresponding 1.5 mL precipitation tube from Step 1. 8. Mix by inverting the tube several times and snap freeze in liquid nitrogen. 9. Recover tubes from liquid nitrogen and centrifuge your samples for 30 min at 4  C at full speed in a microcentrifuge. 10. Remove supernatant and make sure not to lose the pellet. 11. Wash with 600 μL of 80% ethanol. 12. Centrifuge your samples for 10 min at 4  C at full speed. 13. Remove supernatant. 14. Centrifuge your samples for a short spin. 15. Remove any remaining liquid. 16. Incubate your samples with open lid for 2 min at 37  C or 5 min at room temperature. 17. Resuspend the pellet with 16 μL of RNase-free water. 3.3.2 Extensive Dephosphorylation and RNA Precipitation

1. Combine 16 μL of your treated RNA samples in a PCR tube with 2 μL of phosphatase buffer, 1 μL of RiboLock RNase Inhibitor, and 1 μL of Antarctic phosphatase. 2. Mix by pipetting up and down. 3. Incubate the PCR tubes for 1 h at 37  C and then for 5 min at 70  C (to inactivate the phosphatase) and store for indefinite hold at 4  C in a thermal cycler. 4. Add 180 μL of RNase-free water and 200 μL of phenol:chloroform:IAA mix, and vortex. 5. Centrifuge for 10 min at full speed at room temperature. 6. Transfer the supernatant in a new 1.5 mL tube, add 200 μL of chloroform, and vortex. 7. Centrifuge for 10 min at full speed at room temperature. 8. Transfer the supernatant in a new 1.5 mL tube and precipitate the sample by adding 20 μL of 3 M NaOAc, 1 μL of Glycoblue, and 1 mL 96% ethanol. 9. Incubate at 80  C for 30 min and centrifuge your samples for 30 min at 4  C at full speed. 10. Take out the supernatant and wash the pellet with 500 μL of 80% ethanol. 11. Centrifuge for 10 min at full speed at 4  C. 12. Remove supernatant and dry the pellet for 2 min at 37  C.

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1. Resuspend the pellet in 20 μL of aniline. 2. Incubate for 15 min at 60  C in the dark. 3. Stop the reaction by adding 180 μL of RNase-free water, 20 μL of NaOAc, 1 μL of Glycoblue, and 600 μL of 96% ethanol and mix by inverting up and down. 4. Incubate at 80  C for at least 1 h. 5. Centrifuge for 30 min at 4  C at full speed. 6. Remove supernatant and wash the pellet with 500 μL of 80% ethanol. 7. Centrifuge for 10 min at full speed at 4  C. 8. Take out the supernatant and air-dry the pellet for 2 min at 37  C. 9. Resuspend the pellet in 6 μL of RNase-free water.

3.4 Library Preparation

Upon opening the NEBNext® Multiplex Small RNA Library Prep Set for Illumina®, resuspend 50 SR adapter (yellow-cap vial) in 120 μL of RNase-free water and store at 80  C. 1. Mix 6 μL of RNA sample with 1 μL of 30 SR adapter (green-cap vial) (previously diluted ½ in RNase-free water) in a PCR tube. 2. Incubate for 2 min at 70  C in a preheated thermal cycler. Transfer immediately to ice. 3. Add 10 μL of 30 ligation buffer (green-cap vial) and 3 μL of 30 ligation enzyme (green-cap vial). 4. Incubate for 1 h at 25  C in a thermal cycler. 5. Add 4.5 μL of RNase-free water and 1 μL of SR RT primer (pink-cap vial) (previously diluted ½ in RNase-free water). 6. Incubate for 5 min at 75  C, 15 min at 37  C, and 15 min at 25  C. 7. Within the last 15 min of incubation, add 1.1*n (n ¼ number of samples) μL of the 50 SR adapter (yellow-cap vial) (previously diluted ½ in RNase-free water) in an individual PCR tube. 8. Denature the 50 SR adapter in a thermal cycler for 2 min at 70  C and immediately place the tube on ice (see Note 10). 9. Add 1 μL of 50 SR adapter (previously denatured step 8), 1 μL of 50 ligation reaction buffer (yellow-cap vial), and 2.5 μL of ligase enzyme mix (yellow-cap vial). 10. Incubate for 1 h at 25  C in a thermal cycler. 11. Add the following components to the adapter-ligated RNA mix from the previous step, 8 μL of first-strand synthesis reaction buffer (red-cap vial), 1 μL of murine RNase inhibitor (red-cap vial), and 1 μL of ProtoScript II reverse transcriptase (red-cap vial), and mix well by pipetting up and down.

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12. Incubate for 1 h at 50  C. 13. Immediately proceed to PCR amplification (see Note 11). Add the following components to the RT reaction mix from the previous step: 50 μL of LongAmp Taq Master Mix (blue-cap vial), 2.5 μL of SR primer (blue-cap vial), 2.5 μL of index primer (see Note 12), and 5 μL of RNase-free water. Mix well. 14. Perform the following PCR cycling conditions: 1 cycle of initial denaturation for 30 s at 94  C, 12–15 cycles of denaturation for 15 s at 94  C, annealing for 30 s at 62  C, extension for 15 s at 70  C, and 1 cycle of final extension for 5 min at 70  C and store at 4  C for indefinite hold. 3.5 Purification of the Library

Using the GeneJET PCR Purification Kit, carry out all procedures at room temperature. 1. Transfer the PCR mix to a 1.5 mL tube and add 100 μL of binding buffer. Mix thoroughly. 2. Transfer the solution to the purification column. Centrifuge at full speed for 30 s. Discard the flow-through. 3. Add 700 μL of wash buffer to the column and centrifuge at full speed for 30 s. Discard the flow-through. 4. Centrifuge the empty column for an additional 1 min. 5. Transfer the column to a clean 1.5 mL DNA low-binding tube. Add 30 μL of elution buffer to the center of the column membrane and centrifuge at full speed for 1 min. 6. Store the purified library at 20  C until further use.

3.6 Library Quantification

1. Before starting, incubate all solutions of the Qubit dsDNA HS assay kit at room temperature for at least 30 min. The kit provides the concentrated assay reagent, dilution buffer, and pre-diluted standards. 2. Prepare the dye working solution by diluting the concentrated assay reagent 1:200 in dilution buffer. Prepare 200 μL of working solution for each sample and two additional standards. 3. Prepare the two standards annotated “C” and “D” by mixing 10 μL of standard with 190 μL of working solution. 4. Add working solution to 1 μL of library sample to obtain 200 μL in total. 5. Vortex the tubes for 2 s and incubate for 2 min at room temperature. 6. Insert the tubes into the Qubit® 2.0 Fluorometer and proceed with measurements: on the home screen of the Qubit® 2.0 Fluorometer, choose the type of assay (e.g., “HS DNA”) for which you want to perform a new calibration. 7. Press “Yes” to read new standards.

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8. When indicated, insert the standard tube and press “Read.” Standards #1 and #2 correspond to standards “C” and “D,” respectively. 9. Once the calibration is done, insert each sample and press “Read” to make the measurements. Check that the value of your samples is within the assay’s range, and press “Calculate Stock Conc” (see Note 13). 3.7 Library Quality Assessment

1. Before starting the experiments, incubate all solutions of the Agilent High Sensitivity DNA kit at room temperature for at least 30 min in the dark. Vortex them and spin them down before use. 2. Add 15 μL of high-sensitivity DNA dye concentrate (blue-cap vial) into a high-sensitivity DNA gel matrix vial (red-cap vial) (see Note 14). 3. Vortex for 10 s and transfer the gel-dye mix to the center of the spin filter. 4. Centrifuge for 10 min at 2240  g. 5. Add 1 μL of each of your library to 11 different tubes of 1.5 mL already containing 5 μL of RNA marker (green-cap vial). Mix by pipetting up and down. 6. Mix 1 μL of the ladder (yellow-cap vial) with 5 μL of highsensitivity DNA marker (green-cap vial). Mix by pipetting up and down. 7. Prepare the chip priming station. Adjust the syringe clip to the lowest top position. 8. Load 9 μL of the gel-dye mix in the well marked with a “G” surrounded by a black circle. 9. Close the chip priming station properly and press the plunger of the syringe until it is held by the clip. 10. Wait for 1 min and then release the clip. 11. Wait for 5 s until the plunger stops and pull it slowly back to the 1 mL position of the syringe. 12. Open the chip priming station and load 9 μL of the gel-dye mix in the three other wells marked “G.” 13. Load 6 μL of the diluted ladder in the well marked with a ladder. 14. Load 6 μL of the diluted library samples in the wells labeled 1–11. 15. Insert the chip in the Agilent 2100 Bioanalyzer, close the lid, and select the following assay “High Sensitivity DNA” in the 2100 Expert Software screen.

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16. Press “Start” to begin the chip to run. 17. After the run, immediately remove the chip and clean the electrodes with the electrode cleaner filled with 350 μL of RNase-free water. 18. Analyze the results of the chip. 3.8 Library Sequencing

1. For sequencing, libraries are multiplexed and diluted to 6–8 pM final concentration. 2. Recommended sequencing depth or coverage for RNAs is ~5–10 million reads/sample. 3. Sequencing length may vary from 35 to 50 nt in a singleread mode.

3.9 Bioinformatic Analysis

1. Trim adapter sequences of raw reads (FastQ files) using Trimmomatic with the following parameters: java -jar trimmomatic-0.35.jar SE -phred33 input.fq.gz output.fq.gz ILLUMINACLIP :TruSeq3-SE:2:30:10 LEADING:30 TRAILING:30 SLIDINGWINDOW:4:15 AVGQUAL:30 MINLEN:17

(see Note 15).

2. Align the trimmed reads to the appropriate reference sequence (E. coli or yeast rRNA/tRNA dataset, described in [14]) using bowtie2 with the following parameters: bowtie2 -D 15 -R 2 -N 0 -L 10 -i S,1,1.15 -x -U --S.

The use

of soft trimming is not recommended. 3. Mapped reads are extracted from the *.sam file by RNA ID and converted to *.bed format using bedtools v2.25.0. 4. Count the 50 -ends in the produced *.bed file using Unix awk command: awk ’{print $2}’ | sort | uniq -c | awk ’{print $3,$2,$1,$4}’ | sort --n. 5. Calculate normalized cleavage and stop-ratio scores for each position. Normalized cleavage is calculated as a proportion of reads starting at a given position divided by the total number of reads mapped to a given RNA species (1000) and stop-ratio corresponds to the number of reads starting at a given position divided by the number of reads covering this position in RNA (Fig. 1). Normalized cleavage varies from 10–25 for background values to 1000 if all reads in RNA start at one single position. Stop-ratio varies from 0 to 1; values >0.75 generally correspond to m7G/m3C signals in stoichiometrically modified RNA (Fig. 2).

Mapping of m7G and m3C in RNA

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Notes 1. The kit NEBNext® Multiplex Small RNA Library Prep Set for Illumina® (set 1) includes a set of 12 barcoding primers (numbered 1–12) that will be used for multiplexing reactions during PCR amplification. There is also a version set 2 with primers (numbers 13–24). If you do not need these barcoding primers, you may order a similar kit without the primers and use any other source of barcoding primers (Illumina or NEB). 2. The typical amount obtained with 1 mL of a haploid wild-type yeast culture (BY4741 or BY4742) or bacteria culture (DH5α) grown to an OD600 of 1 is about 15–30 μg. 3. If your RNA sample is diluted with RNase-free water instead of 10 mM Tris-EDTA (TE) pH 8.0, the ratio of A260/A280 may be below 2.0 due to the lower pH of water [15]. A ratio of A260/A280 of 1.8 for samples diluted in RNase-free water is considered “pure” for RNA. 4. If your RNA sample is contaminated by phenol or chaotropic salts (e.g., guanidinium thiocyanate used in TRIzol™ extraction or other protocols), this will result in a ratio of A260/ A230 below 1.8. Another round of phenol:choroform:isoamyl alcohol (PCA) extraction and two successive steps of chloroform extraction followed by ethanol precipitation are recommended in this case before alkaline digestion. 5. In case you are working with less than 11 samples, add 1 μL of RNase-free water to the empty wells. 6. The ladder loaded in the Pico RNA chip is provided in a separate package and should be prepared before the beginning of the experiment: spin down the tube and transfer 10 μL to a RNase-free tube. Heat for 2 min at 70  C. Cool down on ice and add 90 μL of RNase-free water. Prepare 5 μL aliquots using the Safe-Lock PCR tubes provided in the kit and store at 70  C. Before use, thaw one tube and keep it on ice. The ladder is quite stable at 70  C and may be used for at least 4 months. 7. RNase contamination problems of the Bioanalyzer electrodes are very frequent and will affect the RNA integrity number of your samples. Therefore, if the Agilent 2100 Bioanalyzer is also frequently used to run DNA chips, it is strongly recommended to use a dedicated electrode cartridge only for RNA assays. In addition, we recommend for each chip to load an internal RNA control (total RNA preparation with a known RIN >9). If you encounter contamination problems, soak the electrode cartridge into an RNaseZap® decontamination solution for at

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least 10 min, then rinse the electrodes with RNase-free water, and let them dry out overnight. 8. The Agilent 2100 Bioanalyzer is very sensitive to vibrations and this may affect your results. Therefore, make sure that no vibrations occur during the run. 9. Fragmentation time should be adjusted for each RNA preparation depending on the species and quality of the RNA used. We recommend testing 3–4 different times of fragmentation to define the appropriate conditions for mild hydrolysis. 10. Do not leave the heated adapter on ice for more than 5–10 min before proceeding to the next step; this may impact your library preparation. 11. We recommend proceeding immediately with PCR amplification. However, if it is not possible, inactivate the RT by heating for 15 min at 70  C and cool down the reaction at 4  C for 1–3 h or safely store the reactions at 20  C overnight. 12. Make sure to use only combinations of compatible primers for barcoding. Most Illumina sequencers use a green laser (or LED) to read G and T nucleotides and a red laser (or LED) to read A and C nucleotides. Within each sequencing cycle, at least one nucleotide for each color channel must be read in the index to ensure proper reading of the barcode sequence. Use as a reference the following guide (ScriptSeq™ Index PCR primers, Illumina) for verification of barcode compatibility or check compatibility with Illumina Experimental Manager software. 13. This quantification step is crucial. Make sure to quantify all your libraries properly since an under- or overestimated quantification will interfere with subsequent sequencing read proportion and quality. 14. The high-sensitivity DNA gel-dye mix is stable for 1 month at 4  C protected from light. 15. MINLEN parameter can vary, but we use the minimal length of 17 nt during trimming to avoid ambiguously mapped reads.

Acknowledgments This work was supported by a joint ANR-DFG grant HTRNAMod (ANR-13-ISV8-0001/HE 3397/8-1) to MH and YM. References 1. Hussain S, Aleksic J, Blanco S, Dietmann S, Frye M (2013) Characterizing 5-methylcytosine in the mammalian epitranscriptome. Genome Biol 14:215

2. Novoa EM, Mason CE, Mattick JS (2017) Charting the unknown epitranscriptome. Nat Rev Mol Cell Biol 18:339–340

Mapping of m7G and m3C in RNA 3. Schwartz S (2016) Cracking the epitranscriptome. RNA N Y N 22:169–174 4. Helm M, Motorin Y (2017) Detecting RNA modifications in the epitranscriptome: predict and validate. Nat Rev Genet 18:275–291 5. Molinie B, Wang J, Lim KS, Hillebrand R, Lu Z-X, Van Wittenberghe N, Howard BD, Daneshvar K, Mullen AC, Dedon P et al (2016) m(6)A-LAIC-seq reveals the census and complexity of the m(6)A epitranscriptome. Nat Methods 13:692–698 6. Schwartz S, Motorin Y (2017) Nextgeneration sequencing technologies for detection of modified nucleotides in RNAs. RNA Biol 14:1124–1137 7. Meyer KD, Jaffrey SR (2014) The dynamic epitranscriptome: N6-methyladenosine and gene expression control. Nat Rev Mol Cell Biol 15:313–326 8. Li X, Peng J, Yi C (2017) Transcriptome-wide mapping of N1-methyladenosine methylome. Methods Mol Biol Clifton NJ 1562:245–255 9. Schwartz S, Bernstein DA, Mumbach MR, Jovanovic M, Herbst RH, Leo´n-Ricardo BX, Engreitz JM, Guttman M, Satija R, Lander ES et al (2014) Transcriptome-wide mapping reveals widespread dynamic-regulated pseudouridylation of ncRNA and mRNA. Cell 159:148–162

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10. Zueva VS, Mankin AS, Bogdanov AA, Baratova LA (1985) Specific fragmentation of tRNA and rRNA at a 7-methylguanine residue in the presence of methylated carrier RNA. Eur J Biochem 146:679–687 11. Wintermeyer W, Zachau HG (1975) Tertiary structure interactions of 7-methylguanosine in yeast tRNA Phe as studied by borohydride reduction. FEBS Lett 58:306–309 12. Marchand V, Ayadi L, Ernst FGM, Hertler J, Bourguignon-Igel V, Galvanin A, Kotter A, Helm M, Lafontaine DLJ, Motorin Y (2018) AlkAniline-Seq: profiling of m7G and m3C RNA modifications at single nucleotide resolution. Angew Chem Int Ed Engl 57 (51):16785–16790 13. Schmitt ME, Brown TA, Trumpower BL (1990) A rapid and simple method for preparation of RNA from Saccharomyces cerevisiae. Nucleic Acids Res 18:3091–3092 14. Marchand V, Pichot F, Thu¨ring K, Ayadi L, Freund I, Dalpke A, Helm M, Motorin Y (2017) Next-generation sequencing-based ribomethseq protocol for analysis of tRNA 2’-O-methylation. Biomol Ther 7(1):13 15. Wilfinger WW, Mackey K, Chomczynski P (1997) Effect of pH and ionic strength on the spectrophotometric assessment of nucleic acid purity. BioTechniques 22:474–476. 478–481

Chapter 6 Transcriptome-Wide Detection of Internal N7-Methylguanosine Li-Sheng Zhang, Chang Liu, and Chuan He Abstract m7G-seq detects internal 7-methylguanosine (m7G) sites within mRNAs and noncoding RNAs by misincorporation signatures. A chemical-assisted sequencing approach selectively converts internal m7G sites into abasic sites, triggering misincorporation at these sites in the presence of a specific reverse transcriptase. The further enrichment of m7G-induced abasic sites by biotin pull-down reveals hundreds of internal m7G sites in human mRNA. The misincorporation ratio before pull-down enrichment can be used for estimating the methylation fraction of some highly methylated m7G sites. Key words 7-Methylguanosine, Misincorporation

1

m7G-seq,

RNA

epitranscriptomics,

mRNA

methylation,

Introduction N7-methylguanosine (m7G) is a well-known RNA modification at the mRNA cap region [1, 2] that stabilizes transcripts against exonucleolytic degradation [3] and affects translation [4]. In addition, m7G can exist internally at position 46 of human cytoplasmic tRNAs [5] and position 1639 of human 18S rRNA [6], installed by METTL1-WDR4 complex and WBSCR22, respectively [7, 8]. These internal m7G methylations display functional roles in RNA processing and have been linked to human diseases [9, 10]. To investigate the existence and distribution of internal m7G within human mRNAs, we developed a chemical-assisted method termed “m7G-seq” to sequence internal m7G at base resolution in the forms of misincorporation signatures. To map internal m7G at base precision, m7G-seq targets the unique chemical reactivity of m7G (Fig. 1). Due to the positive charge on the five-membered ring, NaBH4-mediated reduction converts m7G into reduced m7G selectively [11], without affecting unmodified G. Subsequent heating (55
C) at acidic condition (pH 4.5) induces depurination of the reduced m7G, generating

Mary McMahon (ed.), RNA Modifications: Methods and Protocols, Methods in Molecular Biology, vol. 2298, https://doi.org/10.1007/978-1-0716-1374-0_6, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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Fig. 1 Schematic diagram displaying the chemical reactivity of positively charged m7G under NaBH4 reduction, depurination, and biotin labeling reactions in m7G-seq. Only the reduced m7G can further generate abasic sites and then biotinylated AP sites in the presence of biotin hydrazide under acidic conditions. Biotinylated AP sites, before or after pull-down, induce misincorporation when performing reverse transcription using HIV RT

an RNA abasic site which can be further captured by biotin-ligated hydrazide to produce a biotinylated RNA. After reverse transcription using HIV reverse transcriptase (RT), the abasic sites (or biotinylated abasic sites) are read as predominantly T as well as C [12]. Thus, we can identify the internal m7G sites based on these misincorporation signatures at single-base resolution [13].

2

Materials Prepare all solutions using RNase-free water. Purchase, prepare, and store all buffers at room temperature or 20
C (following the manufacturer’s instructions). Properly follow all waste disposal regulations when disposing of waste materials.

2.1 Preparation of Fragmented mRNA

1. Biological samples: Cells or tissue of interest. 2. DPBS buffer: DPBS, no calcium, no magnesium (Gibco™, 14,190,144). 3. TRIzol™ Reagent. 4. Chloroform. 5. Isopropanol. 6. 70% Ethanol.

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7. Dynabeads mRNA DIRECT kit. 8. Qubit™ RNA HS Assay Kit. 9. RNase-free water (DEPC-treated, DNase, RNase free/Mol. Biol.). 10. Ambion 10 Fragmentation reagent. 11. Zymo Research Oligo Clean & Concentrator. 2.2

mRNA Decapping

1. 10 Decapping Reaction Buffer and Tobacco Decapping Plus 2 enzyme. 2. 20 U/μL SUPERaselIn™ RNase Inhibitor.

2.3 End Repair and 30 -Adapter Ligation

1. 10 T4 Polynucleotide Kinase Reaction Buffer. 2. 10 U/μL T4 Polynucleotide Kinase. 3. 10 CutSmart Buffer and Shrimp Alkaline Phosphatase (rSAP) enzyme. 4. RNA 30 -adapter: 50 rApp- AGATCGGAAGAGCGTCGTG 3SpC3 (synthesized by IDT). 5. 10 T4 RNA Ligase Reaction Buffer and T4 RNA ligase 2-truncated KQ. 6. PEG8000. 7. 50 -Deadenylase. 8. RecJf (NEB, M0264L). 9. Zymo Research RNA Clean and Concentrator.

2.4 Conversion of m7G Site into Abasic Site

1. 1.0 M Sodium borohydride (NaBH4) (Sigma-Aldrich, 213,462-25G) solution: Freshly prepared in water. 2. 50 mM EZ-Link Hydrazide-Biotin (in DMSO). 3. 1 M MES buffer, pH 4.5: Dissolve 1 pack BupH™ MES Buffered Saline Packs (Thermo Scientific™, 28,390) in 50 mL H2O and adjust the pH to 4.5 using AcOH. 4. Dynabeads™ MyOne™ Streptavidin C1 beads and 2 B&W buffer. 5. 1 IP wash buffer: 50 mM Tris–HCl, pH 7.4, 300 mM NaCl, 0.0 5% (v/v) NP-40. 6. 1 proteinase K digestion buffer: 50 mM Tris–HCl, pH 7.4, 75 mM NaCl, 5 mM EDTA, 1% (w/v) SDS. 7. Proteinase K, recombinant, PCR grade.

2.5 Reverse Transcription

1. RT primer: 50 - ACACGACGCTCTTCCGATCT -30 (synthesized by IDT). 2. 10 AMV Reverse Transcriptase Reaction Buffer and AMV Reverse Transcriptase.

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3. 10 mM dNTP solution mix. 4. RNaseOUT™ Recombinant Ribonuclease Inhibitor. 5. Recombinant LS05003).

HIV

reverse

transcriptase

(Worthington,

6. RNase H. 2.6 cDNA 30 -Ligation and PCR Amplification

1. cDNA 30 -linker: 50 Phos- NNNNNNAGATCGGAAGAGCA CACGTCTG-3SpC3 (synthesized by IDT). 2. T4 RNA Ligase 1 (ssRNA Ligase), High Concentration (NEB, M0437M). 3. NEBNext Multiplex Oligos.

3

Methods

3.1 Preparation of Fragmented mRNA

1. Starting from one 15 cm plate for cells of interest, wash cells once with 5 mL ice-cold DPBS buffer. Isolate cellular total RNA using TRIzol reagent following the manufacturer’s protocol and using isopropanol precipitation. 2. With the purified cellular total RNA, extract mRNA by two rounds of polyA+ purification with Dynabeads mRNA DIRECT kit following the manufacturer’s protocol. mRNA concentration is measured using Qubit™ RNA HS Assay Kit with a Qubit 2.0 fluorometer. 3. Starting with 10μg of human mRNA (Step 2 above), dissolve the mRNA in 18μL RNase-free water followed by adding 2μL 10 Fragmentation Buffer. Heat the mixture at 70
C for 15 min. mRNA will fragmented into 50–100 nt (see Note 1). 4. Fragmented mRNA is purified with Oligo Clean & Concentrator and eluted with RNase-free water.

3.2

mRNA Decapping

1. Using a maximum of 6μg fragmented mRNA dissolved in 34μL RNase-free water, add 5μL 10 Decapping Reaction Buffer, 1μL 50 mM MnCl2, and 2μL SUPERaselIn™ RNase Inhibitor. Mix well and then add 8μL decapping enzyme. Mix well and incubate the reaction at 37
C for 2 h. 2. Decapped mRNA is purified using the Oligo Clean & Concentrator and eluted with 40μL RNase-free water.

3.3 End Repair and 30 -Adapter Ligation

1. After decapping the fragmented mRNA, 40μL RNA is mixed with 5μL 10 T4 Polynucleotide Kinase Reaction Buffer. Mix well and add 5μL T4 PNK. Mix well and incubate the mixture at 37
C for 1 h (see Note 2).

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2. 30 -End-repaired RNA is extracted from the solution using the Oligo Clean & Concentrator and eluted with 40μL RNase-free water. 3. 40μL RNA is then mixed with 5μL 10 CutSmart Buffer. Mix well and add 5μL rSAP enzyme. Mix well and the 50 -30 -dephosphorylation step is conducted at 37
C for 1.5 h (see Note 3). 4. Dephosphorylated RNA is extracted from the solution with RNA Clean and Concentrator and eluted in 22μL RNase-free water. 5. To start the RNA 30 -adapter ligation [14], incubate the repaired and dephosphorylated RNA fragments (22μL) with 1.6μL 100μM of the RNA 30 -adapter at 70
C for 2 min and transfer immediately to ice. 6. Add 5μL 10 T4 RNA Ligase Reaction Buffer, 15μL 50% PEG8000, and 2μL SUPERaselIn™ RNase Inhibitor to the RNA-adapter mixture. Mix well and add 4μL T4 RNA ligase 2. Mix well and incubate at 25
C for 2 h followed by 16
C for 12 h (see Note 4). 7. The reaction is then diluted to 94μL with RNase-free water, and the 50 -ends of excess adapters are digested by adding 4μL 50 -deadenylase and incubating at 30
C for 1 h, followed by the addition of 2μL RecJf for ssDNA digestion at 37
C for another hour. 8. 30 -End-ligated RNA is then extracted using RNA Clean and Concentrator and eluted with 20μL RNase-free water. Save 20 ng as “input.” 3.4 Conversion of m7G Sites into Abasic Sites

1. The 30 -end-ligated RNA (in a volume of 20μL) is subject to reduction by adding 20μL of 1.0 M NaBH4 solution. Incubate the reaction at 25
C for 1 h, with occasional low-speed shaking (see Note 5). 2. Quench the reaction using 300μL RNA-binding buffer provided in the RNA Clean and Concentrator and then extract the RNA according to the manufacturer’s protocol. Elute RNA into 35μL RNase-free water. 3. Add 5μL MES buffer to the eluted RNA (35μL last step). Mix immediately with 10μL EZ-Link Hydrazide-Biotin. Incubate the mixture at 55
C for 1 h (see Note 6). 4. Extract RNA using the Oligo Clean & Concentrator and elute with 20μL RNase-free water. Save 20 ng as “before pulldown.” 5. Using the streptavidin C1 beads, wash 10μL of beads twice with 1 B&W buffer (according to the manufacturer’s protocol, see Note 7), and resuspend in 20μL 2 B&W buffer. Mix

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the bead suspension with 20μL RNA (Step 4) and then incubate at 4
C for 20 min. 6. After the biotin pull-down, wash beads five times with 1 IP wash buffer and proceed to proteinase K digestion by adding 45μL 1 proteinase K digestion buffer and 5μL proteinase K to the beads. Mix well and incubate at 55
C for 30 min under high-speed shaking. 7. The flow-through is saved and RNA recovered with the RNA Clean and Concentrator. Elute RNA in 12μL RNase-free water. Save as “pull-down.” 3.5 Reverse Transcription

1. The 30 -ligated RNA (as “input”), RNA before pull-down assay (as “before pull-down”), and eluted RNA after pull-down (as “pull-down”) are subjected to reverse transcription. RNAs are dissolved in 12μL RNase-free water and incubated with 1μL of 2.0μM RT primer at 65
C for 2 min, followed by moving immediately onto ice. 2. Add 2μL 10 mM dNTPs, 2μL 10 AMV Reverse Transcriptase Reaction Buffer, and 1μL RNaseOUT recombinant RNase inhibitor to the 13μL RNA-primer mixture. Mix well and add 2μL recombinant HIV reverse transcriptase. Mix well and incubate the reaction at 37
C for 1.5 h (see Note 8). 3. Add 1μL RNase H to the mixture and incubate at 37
C for 20 min. 4. cDNAs is purified with the Oligo Clean & Concentrator and eluted with 7μL RNase-free water.

3.6 cDNA 30 -Ligation and PCR Amplification

1. The purified cDNA is then subject to cDNA 30 -adaptor ligation [14]. The cDNA is first denatured with 1μL of 50μM cDNA 30 -linker at 75
C for 2 min, followed by transferring immediately to ice. 2. Add 3μL 10 T4 RNA Ligase Reaction Buffer, 15μL 50% PEG8000, and 3μL 10 mM ATP to the 8μL cDNA-adapter mixture. Mix well and add 1μL of T4 RNA ligase 1. Mix well and incubate the reaction at 25
C for 12 h (see Note 9). 3. 30 -Ligated cDNA is purified with the Oligo Clean & Concentrator and eluted with 20μL RNase-free water. 4. The library is then PCR amplified with the universal primer and indexed primers using the NEBNext Multiplex Oligos for Illumina. All libraries are sequenced on an Illumina NextSeq 500 with single-end 80 bp read length.

3.7 Data Processing and Analysis

Proceed with data processing and analysis to identify m7G-seqinduced mutations as previously described [13].

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Notes 1. 10 Fragmentation Buffer, which is a buffer based on Zn2+, is used for generating RNA fragments of 50–100 nt. Traditional fragmentation buffer (based on Mg2+) might give RNA fragments with lengths of 100–150 nt. Fragmented RNA longer than 100 nt may bring difficulties to the subsequent RNA 30 -ligation and cDNA 30 -ligation steps, where shorter size is optimal for efficient ligation. 2. The chemical-assisted RNA fragmentation leaves damaged structures at the 30 -ends, which need to be further repaired by T4 PNK enzyme. PNK repair generates OH group at the 30 -ends. 3. The decapping reaction produces a monophosphate group at the 50 -end of cap-containing RNA fragments. These 50 -monophosphate groups are removed by subsequent alkaline phosphatase (rSAP). This procedure ensures that the 50 -monophosphate will not react with hydrazide in the step of converting internal m7G into an abasic site. Only RNA fragments with internal m7G modification will be enriched by the biotin pull-down. 4. For RNA 30 -ligation, we incubate the reaction at 16
C for 12 h to ensure ligation efficiency. Due to the long incubation time, we include SUPERaselIn™ RNase Inhibitor to protect RNA degradation. However, SUPERaselIn™ RNase Inhibitor contains Na+ which is harmful for most ligation reactions. Here, we used T4 RNA ligase 2 (truncated KQ) as a robust enzyme that tolerates a low concentration of Na+ in the reaction mixture. 5. 20μL of 1.0 M NaBH4 (freshly prepared in water) solution is used as a 2 buffer for the reduction reaction at internal m7G sites. Adding 20 μL of 1.0 M KBH4 (freshly prepared in water) instead of NaBH4 could further enhance the reduction efficiency [15]. 6. When mixing the RNA and EZ-Link Hydrazide-Biotin in MES buffer, heating may trigger the generation of abasic sites. However, a heating temperature above 55
C might lead to unexpected cleavage at abasic sites. 7. When washing the streptavidin C1 beads, it is advised to add SUPERaselIn™ RNase Inhibitor (used as 20) into 1 B&W buffer. This will ensure that the washed beads are RNase free. 8. In the reverse transcription reaction, the wild-type HIV reverse transcriptase displays an excellent behavior in generating misincorporation at abasic sites. Other RT enzymes such as ProtoScript II RT, SuperScript II RT, SuperScript III RT, and AMV RT gave much lower misincorporation rates.

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9. For the cDNA 30 -linker ligation, T4 RNA ligase 1 is employed, which is commonly used as a ssRNA ligase. In this case, a higher PEG8000 concentration (a final 25% v/v) is applied to ensure an efficient ligation. The PEG8000 concentration in RNA 30 -ligation in the presence of T4 RNA ligase 2 (truncated KQ) should be kept at around 15% v/v.

Acknowledgments This work was supported by NIH HG008935 (C.H.). The Mass Spectrometry Facility of the University of Chicago is funded by National Science Foundation (CHE-1048528). We thank Dr. Pieter W. Faber and the Genomics Facility of the University of Chicago for their generous help with high-throughput sequencing. References 1. Cowling VH (2009) Regulation of mRNA cap methylation. Biochem J 425:295–302 2. Furuichi Y (2015) Discovery of m7G-cap in eukaryotic mRNAs. Proc Jpn Acad Ser B Phys Biol Sci 91:394–409 3. Murthy KG, Park P, Manley JL (1991) A nuclear micrococcal-sensitive, ATP-dependent exoribonuclease degrades uncapped but not capped RNA substrates. Nucleic Acids Res 19:2685–2692 4. Muthukrishnan S, Both GW, Furuichi Y, Shatkin AJ (1975) 50 -terminal 7-methylguanosine in eukaryotic mRNA is required for translation. Nature 255:33–37 5. Guy MP, Phizicky EM (2014) Two-subunit enzymes involved in eukaryotic posttranscriptional tRNA modification. RNA Biol 11:1608–1618 6. Sloan KE, Warda AS, Sharma S, Entian KD, Lafontaine DLJ, Bohnsack MT (2017) Tuning the ribosome: the influence of rRNA modification on eukaryotic ribosome biogenesis and function. RNA Biol 14:1138–1152 7. Leulliot N, Chaillet M, Durand D, Ulryck N, Blondeau K, van Tilbeurgh H (2008) Structure of the yeast tRNA m7G methylation complex. Structure 16:52–61 8. Haag S, Kretschmer J, Bohnsack MT (2015) WBSCR22/Merm1 is required for late nuclear pre-ribosomal RNA processing and mediates N7-methylation of G1639 in human 18S rRNA. RNA 21:180–187 ˜ unap K, Kasper L, Kurg A, Kurg R (2013) 9. O The human WBSCR22 protein is involved in

the biogenesis of the 40S ribosomal subunits in mammalian cells. PLoS One 8:e75686 10. Shaheen R, Abdel-Salam GMH, Guy MP, Alomar R, Abdel-Hamid MS, Afifi HH, Ismail SI, Emam BA, Phizicky EM, Alkuraya FS (2015) Mutation in WDR4 impairs tRNA m7G46 methylation and causes a distinct form of microcephalic primordial dwarfism. Genome Biol 16:210 11. Wintermeyer W, Zachau HG (1970) A specific chemical chain scission of tRNA at 7-methylguanosine. FEBS Lett 11:160–164 12. Kupfer PA, Leumann CJ (2005) RNA abasic sites: preparation and trans-lesion synthesis by HIV-1 reverse transcriptase. Chembiochem 6:1970–1973 13. Zhang LS, Liu C, Ma H, Dai Q, Sun HL, Luo G, Zhang Z, Zhang L, Hu L, Dong X, He C (2019) Transcriptome-wide mapping of internal N7-methylguanosine methylome in mammalian mRNA. Mol Cell 74:1304–1316 14. Li X, Xiong X, Zhang M, Wang K, Chen Y, Zhou J, Mao Y, Lv J, Yi D, Chen XW, Yi C (2017) Base-resolution mapping reveals distinct m1A methylome in nuclear- and mitochondrial-encoded transcripts. Mol Cell 68:993–1005 15. Lin S, Liu Q, Lelyveld VS, Choe J, Szostak JW, Gregory RI (2018) Mettl1/Wdr4-mediated m7G tRNA methylome is required for normal mRNA translation and embryonic stem cell self-renewal and differentiation. Mol Cell 71:244–255

Chapter 7 miCLIP-MaPseq Identifies Substrates of Radical SAM RNA-Methylating Enzyme Using Mechanistic Cross-Linking and Mismatch Profiling Vanja Stojkovic´, David E. Weinberg, and Danica Galonic´ Fujimori Abstract The family of radical SAM RNA-methylating enzymes comprises a large group of proteins that contains only a few functionally characterized members. Several enzymes in this family have been implicated in the regulation of translation and antibiotic susceptibility, emphasizing their significance in bacterial physiology and their relevance to human health. While few characterized enzymes have been shown to modify diverse RNA substrates, highlighting potentially broad substrate scope within the family, many enzymes in this class have no known substrates. The precise knowledge of RNA substrates and modification sites for uncharacterized family members is important for unraveling their biological function. Here, we describe a strategy for substrate identification that takes advantage of mechanism-based cross-linking between the enzyme and its RNA substrates, which we named individual-nucleotide-resolution cross-linking and immunoprecipitation combined with mutational profiling with sequencing (miCLIP-MaPseq). Identification of the position of the modification site is achieved using thermostable group II intron reverse transcriptase (TGIRT), which introduces a mismatch at the site of the cross-link. Key words RNA methylation, Radical SAM, Substrate identification, Methyl adenosine, RlmN, Cfr, TGIRT

1

Introduction There are more than 100 chemically distinct RNA modifications, out of which methylation is the most common. RNA methylation is ubiquitous across all domains of life; however, the exact location, biological function, and corresponding RNA-methylating enzymes are poorly understood. Recent strategies that combine immunoprecipitation of modified RNA or chemical treatment of RNA with next-generation sequencing have allowed mapping of the location and abundance of a subset of RNA modifications, such as N6-methyladenosine (m6A), 5-methylcytosine (m5C), and pseudouridine (Ψ) [1–8]. These approaches take advantage of either the unique chemical reactivity of the methyl group (e.g., detection of

Mary McMahon (ed.), RNA Modifications: Methods and Protocols, Methods in Molecular Biology, vol. 2298, https://doi.org/10.1007/978-1-0716-1374-0_7, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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m5C via RNA bisulfite sequencing) [3] or the availability of modification-specific antibodies (e.g., transcriptome-wide identification of m6A and Ψ) [1, 4, 9]. Additionally, strategies based on UV cross-linking and immunoprecipitation, known as CLIP-based methods, have been used to identify enzyme-substrate pairs for RNA-modifying enzymes [10–14]. These methods employ UV irradiation to generate covalent protein-RNA adducts that can be subsequently isolated and enriched to identify RNA-interacting partners. Despite many advances that resulted from CLIP-based methods, the main disadvantage of UV cross-linking is its low cross-linking efficiency. An alternative approach developed for substrate identification for some RNA methyltransferases exploits the formation of a covalent catalytic intermediate between the enzyme and its RNA substrate [3, 15]. While inherently limited to enzyme families that form a covalent intermediate in their catalytic mechanisms, this approach allows for highly efficient cross-linking of enzyme-substrate pairs. This strategy has earlier been applied to NSun RNA methyltransferase family members, which methylate cytosines in RNA to yield m5C. The covalent enzyme-substrate intermediate is trapped either by using a 5-azacitidine (Aza) analog, as in Aza-IP [3], or by mutation of the key cysteine residue in these enzymes that is necessary for the resolution of the covalent intermediate, as in methylation-iCLIP (miCLIP) approach [15]. Radical SAM RNA-methylating enzymes employ a radicalbased mechanism to generate 2-methyladenosine (m2A, RlmN enzymes) and 8-methyladenosine (m8A, Cfr enzymes). Mechanistic studies by our group [16] and others [17–20] have revealed that substrate methylation by radical SAM RNA-methylating enzymes proceeds through an enzyme-substrate covalent intermediate distinct from those formed by RNA m5C methyltransferases [21]. The hallmark of the methylation is formation of a methylene-bridged covalent intermediate between a Cys residue in the enzyme (C355 in E. coli RlmN) and amidine carbon of the adenosine substrate (Fig. 1) [16–20, 22, 23]. Subsequently, the enzyme-RNA covalent adduct is resolved by a second conserved cysteine (C118 in E. coli RlmN) [16]. Mutation of C118 (C118A) stabilizes the proteinRNA intermediate, enabling isolation of the enzyme-RNA covalent pairs by immunoprecipitation (Fig. 1) [16]. By combining this key mechanistic feature with next-generation sequencing, we have developed a novel strategy where individual-nucleotide-resolution cross-linking and immunoprecipitation are combined with mutational profiling with sequencing (miCLIP-MaPseq) [24]. This method can allow for the identification of substrates and modification sites for any member of the radical SAM RNA-methylating enzyme family. The method was developed and validated using the most well-characterized member of the family, RlmN from E. coli that is known to modify 23S rRNA, as well as a subset of tRNA substrates [25, 26].

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Fig. 1 Mechanistic scheme for RlmN-mediated methylation of RNA showing key steps. The stable covalent intermediate trapped by C118A mutation is shown

miCLIP-MaPseq relies on immunoprecipitation of a stable covalent complex between the mutant enzyme and RNA, followed by high-throughput RNA sequencing (Fig. 2). Following isolation, the protein-RNA species are digested using Proteinase K, which leaves a peptide scar on the RNA at the site of the protein-RNA cross-link formation. RNA is then size-selected on a denaturing TBE-urea gel (Fig. 3). RNA species larger than 300 nucleotides are fragmented prior to dephosphorylation [27], while smaller RNA fragments are dephosphorylated without prior fragmentation. After size selection, RNAs are converted to cDNA using the TGIRT reverse transcriptase, and the resulting library is subjected to PCR amplification and high-throughput sequencing. One main advantage of miCLIP-MaPseq is that it uses TGIRT to generate cDNA. This reverse transcriptase is highly processive and introduces a mismatch when it encounters the protein scar on RNA and thus allows identification of methylation sites using mutational profiling. Here, we provide a detailed miCLIP-MaPseq protocol that can be easily modified and implemented to identify substrates of any member of the radical SAM RNA-methylating family.

2

Materials

2.1 Cell Lysis and Target Protein Immunoprecipitation

1. Lysis buffer: 50 mM Tris–HCl pH 7.5, 150 mM NaCl, 1 mM EDTA, 1% (v/v) Triton X-100. 2. 100 mM PMSF. 3. TBS buffer: 50 mM Tris–HCl pH 7.5, 150 mM NaCl.

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Fig. 2 Schematic representation of library preparation strategy for identification of substrates and methylation sites of RlmN. Red bars represent the fraction of mismatches at a specific nucleotide on substrate RNA

4. Glycine buffer: 100 mM glycine–HCl pH 3.5. 5. Stringent-TBS wash: 50 mM Tris–HCl pH 7.5, 500 mM NaCl. 6. Resin recycle solution: 50% Glycerol, 50% TBS, 0.02% sodium azide. 7. 10 mM Tris pH 7.5. 8. RQ1 RNase-free DNase. 9. Anti-FLAG M2 resin. 10. IP dilution solution: 12 μL of 5 mg/mL 1
FLAG peptide and 388 μL of 10 mM Tris pH 7.5. 11. LB medium. 12. 1.5 mL Eppendorf tubes. 13. E. coli strain encoding a FLAG-tagged wild-type RNA-modifying enzyme of interest and the corresponding FLAG-tagged mutant RNA-modifying enzyme. For the example described herein, use E. coli BW25113 strain encoding FLAG-tagged wild-type RlmN and E. coli BW25113 FLAG-tagged C118A RlmN. 2.2

RNA Isolation

1. Proteinase K. 2. GlycoBlue. 3. 3 M Sodium acetate pH 5.5. 4. Isopropanol.

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Fig. 3 Gel analysis of isolated RNA after immunoprecipitation and Proteinase K treatment of FLAG-tagged C118A RlmN. RNA was size selected into four regions (A-D) as indicated on the gel, and each region was individually sequenced. Lanes 1–3: Isolated RNA after immunoprecipitation and Proteinase K treatment of FLAG-tagged C118A RlmN, where the amount of sample loaded in lane 1 is half of the amount loaded in each of the lanes 2 and 3; lane 4: low-range singlestranded RNA markers

5. 80% Ethanol. 6. Novex 10% TBE-urea precast gel. 7. 1
TBE running buffer: Dilute 10
TBE running buffer to 1
using DEPC-treated water. 8. 2
RNA loading dye. 9. Low-range ssRNA ladder. 10. SYBR Gold Nucleic Acid Gel Stain. 11. 18G
1 ½ syringe needle. 12. Costar SpinX column. 13. Nuclease-free water. 14. RNase-free non-stick 0.5 mL tubes and RNase-free 1.5 mL tubes. 15. 100% Ethanol.

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2.3 RNA Fragmentation

1. 10
Fragmentation Reagent mix. 2. Nuclease-free water. 3. RNase-free PCR tubes.

2.4 Library Preparation

1. T4 polynucleotide kinase. 2. SUPERase-In. 3. TGIRT-III template-switching kit. 4. 10
PNK buffer: 70 mM Tris–HCl pH 7.5, 10 mM MgCl2, 5 mM DTT. 5. 5
TGIRT reaction buffer: 100 mM Tris–HCl pH 7.5, 2.25 M NaCl, 25 mM MgCl2. 6. Novex 8% TBE precast gel. 7. 5
GelPilot DNA Loading Dye. 8. 50 DNA Adenylation Kit. 9. Zymo Oligo Clean & Concentrator kit. 10. Thermostable 50 AppDNA/RNA Ligase. 11. 10
NEBuffer 1. 12. 50 mM MnCl2. 13. MiniElute PCR Purification Kit. 14. Phusion High-Fidelity DNA Polymerase. 15. Deoxynucleotide (dNTP) Solution Mix. 16. 10 mM Tris, pH 8. 17. Oligos used for library preparation: R2 RNA (provided in a kit by InGex; 3SpC3 is a C3 Spacer phosphoramidite): 50 -rArGrA rUrCrG rGrArA rGrArG rCrArC rArCrG rUrCrU rGrArArCrUrCrCrArG rUrCrA rC/3SpC3/-30 . R2R DNA (provided in a kit by InGex): 50 -GTG ACT GGA GTT CAG ACG TGT GCT CTT CCG ATC TN (N ¼ equimolar A, T, G, C). R1R DNA (IDT; 3SpC3 is a C3 Spacer phosphoramidite): 50 -/5Phos/GAT CGT CGG ACT GTA GAA CTC TGA ACG TGT AG/3SpC3/-30 . Illumina multiplex primer (IDT): 50 -AAT GAT ACG GCG ACC ACC GAG ATC TAC ACG TTC AGA GTT CTA CAG TCC GAC GAT C-30 . Illumina barcode primer (IDT): 50 -CAA GCA GAA GAC GGC ATA CGA GAT [barcode] GTG ACT GGA GTT CAG ACG TGT GCT CTT CCG ATC T-30 .

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1. KAPA library quantification kit. 2. 96-Well PCR plate. 3. BioRad CFX Connect real-time PCR. 4. Agilent 2100 Bioanalyzer.

2.6

3

Sequencing

1. Illumina HiSeq4000.

Methods

3.1 Expression of the FLAG-Tagged Enzyme

A C-terminal DYKDDDDK octapeptide (FLAG) sequence is fused to the genomic version of rlmN or rlmN containing C118A mutation as described previously [16]. 1. Inoculate 1 L of LB medium with 1:100 from an overnight culture of E. coli BW25113 encoding either the FLAG-tagged WT RlmN or the FLAG-tagged C118A RlmN. 2. Grow the cells at 37  C, 200 rpm, for 90 min for WT RlmN, or 150 min for C118A RlmN (see Note 1). 3. Harvest the cells by centrifugation at 1610
g (rotor F9S 4
1000y) for 15 min at 4  C. Flash freeze the pellets in liquid nitrogen, and either store them at 80  C or immediately proceed with the next step.

3.2 Lysis and DNase Treatment

1. Thaw ~1.5 g of cells and resuspend them in 4 mL of cold lysis buffer. Add 40 μL of 100 mM PMSF. 2. While keeping the cells on ice, sonicate the cells using the microtip on a power setting 3 and duty cycle 50%, for three 40-s pulses with 1-min breaks between pulses. The probe should be positioned approximately 0.5 cm from the bottom of the tube and should not be touching the tube sides in order to avoid foaming. 3. Divide 4 mL sample among five 1.5 mL Eppendorf tubes. Add 18 μL RQ1 RNase-free DNase to each tube and incubate at 37  C for 15 min. Spin down at 19,722
g (Sorvall Legend Micro 21R) for 10 min at 4  C. Without disturbing the pellet remove the supernatant to a new 1.5 mL tube. If not immediately proceeding with the next step, store the samples at 20  C.

3.3 Immunoprecipitation 3.3.1 Resin Preparation

1. Thoroughly suspend the anti-FLAG M2 affinity resin and immediately transfer 375 μL to a new Eppendorf tube. Centrifuge the resin at 8000
g, for 1 min at 4  C. Let the resin settle for 1 min. Remove the supernatant making sure not to transfer any resin. 2. Wash the resin twice with 1 mL of TBS buffer. For each wash, add 1 mL of TBS buffer, resuspend the resin by gently pipetting, centrifuge the resin at 8000
g for 1 min at 4  C, and then let it settle for 1 min.

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3. Wash the resin once with 1 mL glycine buffer (see Note 2). 4. Wash the resin three times with TBS buffer. For each wash, the process is the same as in Step 2. 3.3.2 Binding of FLAG-Tagged Protein

1. Add approximately 75 μL of resuspended resin into each tube containing ~800 μL sample. Let the samples incubate with the resin on a rotator for at least 2 h at 4  C.

3.3.3 Elution of FLAG-Tagged Protein

1. Centrifuge the samples at 8000
g, for 1 min at 4  C. Let the resin settle for 1 min. Remove the supernatant. 2. Wash the resin three times with 500 μL of stringent TBS wash buffer. 3. Add 75 μL of IP dilution solution into each sample. Gently resuspend and rotate samples for at least 1 h at 4  C (see Note 3). 4. Centrifuge the samples at 8000
g, for 1 min at 4  C. Let the resin settle for 2 min. 5. Carefully transfer the supernatant to a new Eppendorf tube. Put both resin and supernatant on ice. Store resin without glycerol at 4  C. Store supernatant at 20  C.

3.3.4 Recycling of the Resin

1. Resuspend used resin in 500 μL of TBS buffer. Centrifuge the resin at 8000
g, for 1 min at 4  C. Remove the supernatant. 2. Wash the resin three times with 1 mL of glycine buffer. Centrifuge the resin at 8000
g, for 1 min at 4  C. Let the resin settle for 1 min. 3. Wash the resin five times with 1 mL of resin recycle solution. Centrifuge the resin at 8000
g, for 1 min at 4  C. Store the resin at 20  C.

3.4 Proteinase K Treatment

1. Thaw the sample obtained after immunoprecipitation on ice. Remove 10 μL for subsequent Western blot analysis. Add 9.6 U of Proteinase K into the rest of the sample and incubate the reaction for 2 h at 37  C. 2. Divide each sample into two Eppendorf tubes (~230–250 μL). Precipitate RNA by adding 1000 μL of isopropanol, 25 μL of sodium acetate pH 5.5, and 2 μL of GlycoBlue co-precipitant into each tube. Leave the tubes overnight at 20  C. 3. The next day precipitate RNA by centrifugation at 20,000
g for 30–40 min at 4  C. 4. Carefully remove the supernatant and wash the pellet with 750 μL of 80% cold ethanol. Precipitate RNA by centrifugation at 20,000
g for 40 min at 4  C. Carefully remove the supernatant and air-dry the pellets (see Note 4). 5. Resuspend pellets with 40 μL nuclease-free water (see Note 5).

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1. Set up the gel apparatus and pre-run the 10% TBE-urea gel at 180 V for at least 20 min in 1
TBE running buffer. 2. Add 25 μL of 2
RNA loading dye to 25 μL of sample. Prepare molecular weight marker solution by mixing 1 μL of ssRNA low-range ladder to 9 μL of nuclease-free water, followed by 10 μL of 2
RNA loading dye. Heat the samples at 92  C for 4 min. In two wells load 20 μL of sample and in one well 10 μL. Load the marker in the last lane (see Note 6). 3. Run the gel at 180 V until the lower (dark blue) dye is close to the bottom, approximately 70 min. Incubate the gel with 50 mL of TBE buffer containing 5 μL SYBR-Gold dye for 5 min. Wash the gel twice with nuclease-free water. 4. Visualize and record the gel under UV light. 5. Prepare 0.5 mL RNase-free non-stick tubes by piercing a hole in the bottom using an 18G syringe needle. Cut the gel on transilluminator as indicated in Fig. 3. Put gel pieces into 0.5 mL tubes and then place the tubes into a 1.5 mL collection tube. Centrifuge at 20,000
g, for 3 min at 4  C. Remove 0.5 mL tube and add 300 μL nuclease-free water. 6. Shake samples on the thermomixer at 157
g for 10 min at 68  C, and then freeze them on dry ice for 10 min. Thaw samples at room temperature for 10 min and then incubate on thermomixer at 157
g for 10 min at 68  C. 7. Cut the tips of the P1000 barrier tips, transfer the sample (with gel pieces) onto a Costar SpinX column, and spin at 20,000
g for 3 min at 4  C. 8. Add to each tube 2 μL of GlycoBlue, 33 μL of 3 M sodium acetate pH 5.5, and 900 μL of 100% ethanol. Vortex to mix. Put at 20  C overnight. Next morning precipitate RNA as described in Subheading 3.4.

3.6 RNA Fragmentation

Fragment RNAs longer than 300 nt to 50–200 nt long fragments using the following protocol: 1. Resuspend RNA obtained in Subheading 3.5 in 11 μL of nuclease-free water. Transfer to PCR tubes and place in the thermocycler. 2. Heat the samples for 2 min at 95  C to denature RNA. 3. Add 1 μL of 10
Fragmentation Reagent mix. If you are dealing with larger number of tubes, keep the tubes on ice. This will ensure that most of the RNA stays denatured. Keeping samples at the room temperature will allow for the slow refolding of RNA.

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4. Return the samples to thermomixer and incubate them for 2 min at 95  C. Add 1 μL of stop solution and place the samples on ice. 5. Purify the RNA on 10% TBE-urea gel. Extract 50–200 nt long fragments and precipitate RNA as described in Subheading 3.5 (see Note 7). 3.7 Library Preparation 3.7.1 RNA 30 End Dephosphorylation

3.7.2 TemplateSwitching Reaction Using TGIRT-III

1. Determine RNA concentration by NanoDrop. Low RNA concentration should be measured by Qubit or Bioanalyzer. 2. Prepare 11 μL reaction mixture by mixing 7 μL of RNA sample with 1.1 μL of 10
PNK buffer, 1 μL of Superase-In, and 2 μL of T4 polynucleotide kinase. Incubate reaction mixture at 37  C for 1 h, followed by 3 min at 90  C to inactivate the enzyme. 1. In a PCR tube prepare 17.5 μL reaction mixture by mixing nuclease-free water with 4 μL of 5
TGIRT reaction buffer, 2 μL of 50 mM DTT, ~50 ng of dephosphorylated RNA sample, and 2 μL 10
TGIRT-III RT/template-primer substrate mix (see Note 8). 2. Preincubate reaction mixture at room temperature for 30 min, and then add 2.5 μL of 10 mM dNTPs. 3. Incubate the reaction at 60  C for 60 min (see Note 9). 4. Add 1 μL of 5 M NaOH and incubate the sample at 65  C for 15 min. 5. Cool to room temperature and neutralize sample with 1 μL of 5 M HCl. 6. To each sample add 100 μL of 10 mM Tris pH 8.0, 13 μL of 3 M sodium acetate pH 5.5, 3 μL of GlycoBlue, and 600 μL of 100% ethanol. Incubate overnight at 20  C. Next morning precipitate cDNA as described in Subheading 3.4. 7. Pre-run 8% TBE gel at 155 V for at least 20 min. 8. Resuspend precipitated cDNA in 5 μL of 10 mM Tris pH 8.0 and add 1.25 μL of 5
DNA loading dye. Run gel at 155 V for 40–45 min. 9. Size-select cDNA using 10 bp ladder as a guide. Add 300 μL of nuclease-free water to cut gel pieces, and extract cDNA from gel pieces by following the general protocol presented in Subheading 3.5. Precipitate cDNA overnight, at 20  C, by adding 3 μL of GlycoBlue, 33 μL of sodium acetate pH 5.5, and 900 μL of 100% ethanol. Next morning precipitate cDNA as described in Subheading 3.4 (see Note 10).

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1. In a PCR tube prepare 20 μL reaction mixture by combining 2 μL of 10
50 DNA Adenylation Reaction Buffer, 2 μL of 1 mM ATP, 1 μL 100 μM R1R DNA, and 2 μL of Mth RNA Ligase. We usually set up 4–8 parallel reactions. 2. Incubate reactions in a thermocycler at 65  C for 1 h. 3. Incubate samples at 85  C for 5 min to inactivate the enzyme. 4. Clean up the adenylated R1R DNA with an Oligo Clean & Concentrator kit and elute in 10 μL of nuclease-free water to obtain 10 μM adenylated R1R DNA. If doing several adenylation reactions in separate PCR tubes, combine them for a cleanup since higher elution volume helps with consistent and efficient recovery of adenylated oligos. 5. Check the extent of adenylation by running the sample on a 20% TBE-7 M-urea gel.

3.7.4 Thermostable Ligation

1. In a PCR tube prepare 20 μL reaction mixture by combining 2 μL of 10
NEBuffer 1, 2 μL of 50 mM MnCl2, 4 μL of 10 μM adenylated R1R DNA, 10 μL of cDNA from templateswitching, and 2 μL of Thermostable 50 AppDNA/RNA Ligase. 2. Incubate reactions in thermocycler at 65  C for 2 h. 3. Incubate samples at 90  C for 3 min to inactivate the enzyme. 4. Clean up the ligated cDNA with a MiniElute PCR Purification Kit and elute in 23 μL of nuclease-free water.

3.7.5 PCR Amplification

1. In an Eppendorf tube prepare a 53 μL reaction mixture by combining 29.5 μL of nuclease-free water, 10 μL of 5
Phusion HF buffer, 1 μL of 10 μM Illumina multiplex primer, 1 μL of 10 μM Illumina barcode primer, 10 μL of cDNA, 1 μL of 10 mM dNTPs, and 0.5 μL of Phusion High-Fidelity DNA Polymerase. 2. Divide reaction mixture among three PCR tubes. Heat cDNA at 98  C for 5 s, then amplify it for 15, 18, or 21 cycles of 98  C for 5 s, 60  C for 10 s, and 72  C for 12 s. 3. Mix 17 μL of PCR product with 4.25 μL of 5
DNA loading dye, load on an 8% TBE gel, and run the gel at 155 V for 45 min. Stain the gel with SYBR gold. 4. Size-select amplified DNA using 10 bp ladder as a guide. Add 300 μL of nuclease-free water to cut gel pieces, and extract DNA from gel pieces by following the general protocol presented in Subheading 3.5. Precipitate DNA overnight, at 20  C, by adding 3 μL of GlycoBlue, 33 μL of sodium acetate pH 5.5, and 900 μL of 100% ethanol. Next morning precipitate as described in Subheading 3.4. Resuspend each library in 10 μL of 10 mM Tris pH 8.

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3.7.6 qPCR Quantification

1. For the quantification of the libraries we use KAPA library quantification kit. If the kit is being used for the first time, add 1 mL of 10
Primer Premix to the bottle of 2
KAPA SYBR FAST qPCR Master Mix (5 mL) and mix by vortexing. Aliquot this solution and store at 20  C. 2. Determine the total number of reactions that will be performed. Usually we run six DNA standards in triplicate and each library dilution in duplicate. Using a NanoDrop, estimate the concentration of each library and determine which dilutions to prepare to stay within the dynamic range of the assay. A 1:5000 and 1:10,000 dilution usually fall around the midpoint of the assay standards. 3. Prepare 1:5, 1:50, 1:5000, and 1:10,000 library dilutions in 10 mM Tris pH 8 buffer. 4. For each reaction, prepare the following in a 96-well PCR plate: 6 μL of Master Mix containing primers, and 4 μL of either DNA standard or specific library dilution. Seal the plate with optical adhesive film. 5. Run the plate with the following program in the qPCR machine: 95  C for 5 min, and 35 cycles of 95  C for 30 s, and 60  C for 45 s. 6. Use the KAPA analysis template to calculate slope and intercept of the standard curve, to convert the average Cq score for each library dilution to pM, to calculate the average size-adjusted concentration (in pM) for each dilution, and to calculate the size-adjusted concentration for the original undiluted library. 7. Prepare 15 μL of 10 nM library solution, containing up to 20 libraries. Store individual libraries in RNase-free low-retention Eppendorf tubes at 20  C (see Note 11). 8. Check the quality of the library on Bioanalyzer prior to submitting sample for sequencing on an Illumina HiSeq4000 or similar (see Note 12).

3.8 Sequencing Read Mapping and Analysis 3.8.1 Sequence Processing and Alignment

1. Prior to bioinformatic analysis it is important to de-multiplex sequences if multiple samples were run within one sequencing lane. De-multiplexing for our samples was performed by the Center for Advanced Technology at UCSF. 2. Upload sequencing data to the Galaxy web platform and use the public server at usegalaxy.org to analyze the data [28] (see Note 13). 3. Process reads with FASTQ Groomer [29] and then remove adapters using Clip tool also available through Galaxy web platform.

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4. Go to Ensembl Bacteria Genome Database (EMBL-EBI), https://bacteria.ensembl.org/index.html, and download E. coli BW25113 FASTA file and gtf file. 5. Align sequences greater than 15 bp to the genome using Bowtie 2 with default options [30, 31]. Default settings for “sensitive-local” are the default option in “local-mode” (details are -D 15 -R 2 -N 0 -L 20 -i S,1,0.75) (see Note 14). 3.8.2 Enrichment Analysis of Reads

1. Determine the raw counts per gene by using HTSeq-count script, which is available through Galaxy web platform [32]. Select intersection-nonempty mode to handle reads overlapping more than one feature. Summarize counts from regions A-D per replicate. Use this file to perform the enrichment analysis. 2. For enrichment analysis of reads mapped to any set of genes use DESeq2 module. In DESeq2 specify the factor levels that will be analyzed (e.g., sample vs. control), and select all replicates belonging to a specific factor level. As an input data use summarized HTSeq-count data as described above. Use parametric fitting and leave on the following options: outliers replacement, outliers filtering, and independent filtering. For the control samples, we generated a library from the rRNAdepleted total RNA isolated from E. coli BW25113 strain (see Note 15).

3.8.3 Analysis of Stop Sites and Mismatches

1. Download Integrated Genomic Viewer (IGV) [33]. Open E. coli BW25113 genome file. Open all BAM files and their corresponding BAM_index files. 2. Select the gene of interest and determine the percent of mismatches for a specific nucleotide by cumulative analysis of all biological replicates (Fig. 4). 3. To determine the 50 end of the reads (stop sites) use script “make_wiggle” to convert sorted and indexed BAM files to wiggle files. This script was developed by the Weissman lab at UCSF and is readily available through Plastid [36]. The results can be readily visualized with IGV.

4

Notes 1. Optimal expression time for an enzyme should be determined empirically prior to proceeding to the next step. 2. Resin cannot stay in glycine buffer for longer than 20 min. 3. When dealing with a very small sample volume, combine all the samples from a single experiment into a single tube prior to leaving the sample on a rotator.

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Fig. 4 Examples of read profiles for specific RNAs displayed in Integrative Genomic Viewer. (a) Read profiles for tRNAGlnUUG displayed in Integrative Genomic Viewer (IGV) [33–35]. (b) Read profiles for tRNAHisGUG displayed in IGV. tRNAs were isolated after immunoprecipitation of FLAG-tagged C118A RlmN. The depth

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4. After centrifugation remove the ethanol using P1000 pipette. Recap the tube, and pulse centrifuge to bring down the remaining ethanol. Remove the remaining liquid using P200 pipette. Leave the tube open at room temperature before proceeding to another sample. By the time all the samples are finished, the pellets should be sufficiently dry. Do not overdry the pellet since it can be hard to re-solubilize RNA. 5. Perform Western blot analysis to ensure that the enzyme was successfully digested. We use monoclonal anti-FLAG M2-peroxidase (HRP) antibody. 6. Prior to loading the samples flush the remaining urea out of the wells using P1000 pipette. This will decrease smearing and abnormal band shapes. Additionally, it is advisable to load a smaller amount of sample in one of the lanes to better see discrete bands, since loading a large amount of sample can lead to increased smearing in the gel. 7. Under these conditions, the extent of fragmentation will depend on the initial amount of RNA. If substantial amount of RNA is not successfully fragmented, extract the RNA longer than 300 nt, and repeat the fragmentation step. Make sure to decrease the time for the re-fragmentation step (e.g., from 2 min to 1 min, or less). 8. Add RNA sample and enzyme/template-primer mix last. 9. For long or heavily modified RNAs, such as tRNAs, it is necessary to run the reaction for 60 min. For short RNAs 5–15 min is usually sufficient, but the exact time should be determined empirically. 10. In case no pellet is observed after the first centrifugation step, add 1 μL of GlycoBlue, place sample on dry ice for at least 30 min, thaw sample at room temperature, and then repeat centrifugation step. Only then remove the supernatant and perform the wash step with cold 80% ethanol. 11. When submitting multiple samples within one sequencing lane, approximately equal amounts of each library should be added.

ä Fig. 4 (continued) of the reads (counts) displayed at a specific locus is represented as a gray bar chart (top panel). Alignment of individual reads is represented in the bottom panel. Known modifications are represented using abbreviations and were taken from the MODOMICS database [35]. Abbreviations: 4-thiouridine (s4U), dihydrouridine (D), queuosine (Q), 7-methylguanosine (m7G), 20 -O-methylguanosine (Gm), 20 -O-methyluridine (Um), 5-carboxymethylaminomethyl-2-selenouridine (cmnm5se2U), 5-methyluridine (m5U), 2 2-methyladenosine (m A), and pseudouridine (Ψ)

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12. For our application and for cost-effectiveness, 50-nucleotide single-end runs are sufficient. 13. When uploading large sequencing files, FileZilla, an opensource software, can be used. 14. If applying this method to a different system, we suggest aligning sequences to the genome of interest using both Bowtie 1 and Bowtie 2 under various settings and then comparing results. 15. DESeq2 considers the variability between the replicates and normalizes read counts to account for differences in sequencing depth between samples, reporting fold change values between the sample and the control. In our analysis, we use a fourfold increase in abundance and adjusted P value of C mutation at the crosslinked site. As with PAR-CLIP, computational screening for this T > C conversion in the sequencing reads allows the identification

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Fig. 1 Overview of PA-mod-seq. (a) Flowchart of the steps involved in the method, with PA-m6A-seq as an example. (b) Example of PA-m6A-seq results visualized in Integrated Genome Viewer (IGV), m6A sites mapped on the first 500 nts of Simian virus 40 (SV40) VP1 mRNA (GEO database accession #GSE106698) [14]. Red/ blue bars in the upper coverage pileup track denote sites of cross-link-induced T > C conversions, with the height of red/blue bars proportional to the occurrence of T and C residues. Blue bars in the bottom individual read track denote the location of T > C conversions in each read. Note that a diverse variety (3+) of T > C conversion sites are expected in a good m6A peak

of reads that truly derive from antibody-bound RNA fragments, thus allowing the elimination of almost all background reads. Furthermore, an RNase footprinting step is included during immunoprecipitation, so that any RNA that is not bound and protected by the antibody will be degraded. This step not only further decreases the observed background but also increases the modification mapping resolution down to the size of the antibody, which protects ~32 nts of bound RNA [11]. During early testing of this

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method with nonspecific IgG control antibodies, we were unable to build any sequencing libraries from IgG immunoprecipitated samples, demonstrating the ability of RNase to effectively degrade background RNA fragments. Overall, PA-m6A-seq results in modification maps of RNA-seq read peaks ~32 nts wide that are confirmed as antibody bound due to the presence of UV cross-linkinginduced T > C conversions, with very low numbers of background reads between peaks. Through the use of different antibodies, we have successfully adapted this method to map multiple different RNA modifications including not only N6-methyladenosine (m6A) but also 5-methylcytidine (m5C) and N4-acetylcytidine (ac4C), and envision further adaptation to map other types of modifications as antibodies become available [13, 16, 17]. To accommodate the variety of modifications that can be mapped with this method, below we refer to this method as PA-mod-seq.

2

Materials

2.1 Tissue Culture and RNA Preparation

1. 0.2 M 4-Thiouridine (4SU) (Sigma T4509 or Carbosynth NT06186): Prepare a 0.2 M stock solution by dissolving 250 mg of 4SU in 4.8 mL DMSO. Aliquot in small volumes, and keep solution at 80  C (see Note 1). 2. Trizol reagent. 3. Chloroform. 4. Isopropanol. 5. 70% Ethanol. 6. GlycoBlue Coprecipitant. 7. RNase-free H2O. 8. Poly(A)Purist MAG Kit.

2.2 Immunoprecipitation and Cross-Linking

1. IPP buffer: 10 mM Tris–HCl pH 7.4, 150 mM NaCl, 0.1% NP-40 in RNase-free H2O. 2. 40 U/μL RNaseIn. 3. Modification-specific antibody: We have successfully used the following antibodies: anti-m6A (SySy #202003), anti-m5C (Diagenode #C15200081), anti-ac4C (Abcam #ab252215). 4. 12-Well tissue culture plate. 5. Stratagene UV Stratalinker 2400 with 365 nm light source. 6. 1000 U/μL RNase T1. 7. Protein G magnetic beads. 8. DynaMag-2 Magnet. 9. Phosphate-buffered saline (PBS).

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10. PAR-CLIP IP wash buffer: 50 mM HEPES-KOH, pH 7.5, 300 mM KCl, 0.05% NP40 in RNase-free H2O. 11. PAR-CLIP high-salt wash buffer: 50 mM HEPES-KOH, pH 7.5, 500 mM KCl, 0.05% NP40 in RNase-free H2O. 2.3

RNA End Repair

1. 10 U/μL Calf intestinal phosphatase (see Note 2). 2. 10 NEB CutSmart Buffer. 3. Phosphatase wash buffer: 50 mM Tris–HCl pH 7.5, 20 mM EGTA-NaOH pH 7.5, 0.5% NP40 in RNase-free H2O. 4. PNK buffer without DTT: 50 mM Tris–HCl pH 7.5, 50 mM NaCl, 10 mM MgCl2 in RNase-free H2O. 5. 1 M DTT. 6. ATP. 7. T4 Polynucleotide Kinase (T4-PNK).

2.4

RNA Elution

1. 20 mg/mL Proteinase K. 2. 4x Proteinase K buffer: 200 mM Tris–HCl pH 7.5, 300 mM NaCl, 25 mM EDTA-NaOH pH 8, 4% SDS in RNase-free H2O. 3. Trizol LS reagent.

2.5 Sequencing Library Preparation

3 3.1

1. NEB Next Small RNA Library Prep Set for Illumina. 2. Novex 10-well 6% TBE gel.

Methods Tissue Culture

Start with an amount of cells that produce ~10 μg of RNA for the immunoprecipitation step (see Notes 3 and 4). Passage cells so they are ~70% confluent and actively growing the day before harvest. Actively growing cells are essential to ensure efficient 4SU uptake and incorporation into RNA. 1. Add 4SU directly to the cell culture media to a final concentration of 100 μM (see Note 1). 2. 16–24 h later, collect cells as appropriate for the cell type: scrape off attached cells or collect suspension cells. Spin cells down at 500  g for 10 min to pellet (see Note 5). 3. Wash cell pellet once with ice-cold PBS, spin down at 500  g for 10 min, and remove PBS. 4. Lyse cell pellet directly in Trizol using 1 mL for every 107 cells. Cells in Trizol can be stored at 80  C.

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3.2 Total RNA Extraction Using Trizol

1. Aliquot every 1 mL of Trizol-cell lysate into 1.5 mL tubes. 2. Add 200 μL chloroform to every 1 mL of Trizol, shake vigorously for 15 s, and incubate at room temperature for 3 min. 3. Centrifuge at 12,000  g for 15 min at 4  C. 4. Collect the upper aqueous phase into a new tube with 500 μL isopropanol (avoid collecting the white interphase), and incubate at room temperature for 10 min. 5. Pellet RNA by centrifugation at 12,000  g for 15 min at 4  C. 6. Precipitate RNA with dH2O:NaOAc:EtOH ¼ 1:0.1:2.2 volume (350:35:770 μL each), and then add 1 μL of GlycoBlue. Mix well and incubate at 80  C for 30 min (or on dry ice for 15 min) (see Note 6). 7. Pellet RNA again at 12,000  g for 20 min at 4  C. 8. Wash pellet with 1 mL 70% ethanol, and centrifuge at 12,000  g for 10 min at 4  C. 9. Remove supernatant. Do a quick spin and carefully remove residual ethanol with a clean pipette. 10. Resuspend RNA pellet in RNase-free H2O, using 25 μL for every 1 mL of starting Trizol.

3.3 Poly(A) Purification

Depending on the cell type, roughly 1–2.5% of total RNA is poly (A)+ mRNA. An IP reaction requires 8–12 μg of RNA; thus start by using ~600 μg of total RNA for poly(A) purification, aiming for a yield of 10 μg poly(A) + mRNA. 1. Follow the Poly(A)Purist MAG Kit instructions to isolate poly (A)+ RNA. Resuspend the resulting RNA pellet in 30 μL dH2O. 2. Measure the concentration of RNA with a Nanodrop spectrophotometer prior to starting the IP.

3.4 Immunoprecipitation and Cross-Linking

1. Prepare IP mix by combining in a 1.5 mL tube, on ice: 10 μg RNA (we use poly(A)-purified RNA from Subheading 3.3 above), 20 μL RNaseIn (a total of 800 units), 7.5 μg of modification-specific antibody, and 800 μL of IPP buffer (see Note 7). 2. Seal the tube lid with parafilm, and rotate at 4  C for 2 h to overnight to allow the antibodies bind to the modified RNA. 3. Transfer the IP mix to a 12-well tissue culture plate well on ice. Irradiate twice with 365 nm UV 2500  100 μJ/cm2 in a UV Stratalinker with the lid off, and then transfer IP mix back into a 1.5 mL tube (the original sample tube can be reused) (see Note 8). 4. Pre-warm a water bath in a cold room to 22  C.

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5. Dilute 2 μL RNase T1 in 58 μL IPP buffer to 0.1 U/μL (1/30 dilution). Add 3 μL of the 1/30 diluted RNase T1 to each IP tube, and digest for 15 min in the 22  C water bath. Invert the tubes every 5 min to mix, and then cool tubes for 5 min on ice (leave the 22  C water bath on for later use). 6. While waiting for the RNase digestion, transfer 90 μL of protein G magnetic beads per IP to a new 1.5 mL tube, place on a magnetic rack for 2 min, and then remove buffer. Wash the beads by resuspending beads in 1 mL PBS, incubating on the magnetic rack for 2 min, and removing PBS. Wash a total of two times. Then resuspend beads in the original bead volume (90 μL per IP) of IPP buffer. 7. After the RNase/IP mix has cooled, add 90 μL of pre-washed protein G beads to each IP tube, and then rotate at 4  C for 1 h. 8. While waiting for the beads to bind antibody-RNA complexes, aliquot 10 mL each of PAR-CLIP IP wash buffer and high-salt wash buffer. Add 5 μL of 1 M DTT to each aliquot (add DTT shortly before use). 9. After the bead capture incubation (step 7 above), isolate the bead-antibody-RNA complexes by placing the IP tubes on a magnetic rack for 2 min, and remove the supernatant. (This supernatant contains the population of RNA not captured by the antibody, so collect and save for analysis if needed.) 10. Wash the bead complexes with IP wash buffer: Resuspend beads in 1 mL IP wash buffer, incubate tubes on the magnetic rack for 2 min, and then carefully remove buffer without touching beads. Repeat this wash a total of three times. 11. Resuspend beads in 100 μL of IP wash buffer, and add 1.5 μL of RNase T1 to a final concentration of 15 U/μL. Digest for 15 min in the 22  C water bath. Invert the tubes every few minutes, and then cool the tubes on ice for 5 min (the dephosphorylation reaction mix can be prepared while waiting for this step). 12. Wash the beads three times with the high-salt wash buffer as before. 3.5

RNA End Repair

1. Prepare dephosphorylation reaction mix by adding 5 μL CIP (final concentration 0.5 U/μL), 10 μL of 10x NEB CutSmart Buffer, and 85 μL of H2O. Resuspend beads from each IP in 100 μL of this reaction mix. 2. Incubate the dephosphorylation reaction on a tube shaker at 800 rpm in a 37  C incubator for 10 min. 3. Wash beads twice with 500 μL of phosphatase wash buffer as before. 4. Wash beads twice with 500 μL PNK buffer without DTT.

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5. Prepare a PNK reaction mix, for each IP: 10 μL of T4-PNK (final 1 U/μL), 88.5 μL of PNK buffer without DTT, 0.5 μL of 1 M DTT (final 5 mM), and 1 μL of 100 μM ATP (final 10 mM). 6. Resuspend beads in PNK reaction mix, 100 μL per IP. Incubate the PNK reaction on a tube shaker at 800 rpm in a 37  C incubator for 30 min. 7. Wash beads three times with 500 μL of PNK buffer without DTT. 3.6

RNA Elution

1. Prepare the Proteinase K elution mix, for each IP: 75 μL of 4x Proteinase K buffer, 225 μL of H2O, and 4.2 μL of Proteinase K (~85 μg). 2. Resuspend the washed beads in Proteinase K elution mix (300 μL per IP), and then incubate at 50  C for 90 min, tapping the tubes to mix every 15 min (alternatively, use an Eppendorf thermoshaker set at 900 rpm). 3. After proteinase digestion, all RNA originally bound to antibodies should be in the supernatant (~300 μL). Transfer supernatant to a fresh tube. 4. Remove Proteinase K using Trizol LS: Add 900 μL Trizol LS to the 300 μL eluate, and mix well. 5. Add 240 μL of chloroform, vortex for 15 s, wait for 2 min, and then centrifuge at 12,000  g for 15 min at 4  C to separate the organic phase from the aqueous phase. 6. Collect the upper aqueous phase into a new tube, and mix with 600 μL of isopropanol and 1 μL of GlycoBlue Coprecipitant. Wait for 10 min, and then centrifuge at 12,000  g for 20 min at 4  C to pellet RNA (see Note 9). 7. (Optional cleanup) Resuspend RNA pellet in 300 μL of H2O, and then add 30 μL of sodium acetate, 1 μL of GlycoBlue, and 660 μL of 100% ethanol. Precipitate at 20  C for 2 h or overnight. Then centrifuge at 12,000  g for 30 min at 4  C (see Note 10). 8. Wash pellet with 70% ethanol, vortex briefly, and centrifuge at 12,000  g for 10 min at 4  C. 9. Resuspend each pellet in 15 μL of H2O.

3.7 Sequencing Library Preparation

1. Prepare sequencing libraries with the NEB Next Small RNA Library Prep Set for Illumina following the kit instructions. The expected RNA size is the RNase footprint of the antibody, which is typically ~32–50 nt. With the ligated 50 and 30 adapters totaling 120 nt, it is necessary to isolate bands ~150–170 nt on a TBE polyacrylamide gel. We have found the Invitrogen Novex 6% TBE gel to give the best separation between our desired product and adapter dimers (120 nt).

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2. With expected pulled-down RNA sizing of 32–50 nt, we have found Illumina sequencing at the 50 nt single-read mode to be sufficient. While paired-end sequencing may increase the data quality, the difference is not noticeable when sufficient read depth is obtained. 3.8 Sequencing Data Analysis

Sequencing result analysis typically involves trimming off adaptor sequences, discarding any reads shorter than 15 nts, aligning sequencing reads to the genome sequence of interest, screening for T > C conversions, and reformatting the alignment information for visualization in the Integrated Genome Viewer (IGV) (see example in Fig. 1b). 1. We use the FASTX toolkit [18] for sequencing read preprocessing; this includes screening for reads with a FASTQ quality score >Q33, removing adapter sequences, and selecting for a minimum read length of 15 nts. Alternatively, we have used Cutadapt [19] with good results. 2. We then use Bowtie [20, 21] for alignment of the sequencing reads to the genome of interest. If you are interested in human cellular transcripts, for example, you can align your reads to a human genome build such as hg19. If you are interested in viral transcripts, you should pre-align the reads to the host genome, then take the host non-aligning reads, and align them to the viral genome. If your model system of interest has a heavily spliced transcriptome, it might be informative to align to the transcriptome with a splice-aware aligner, such as TopHat [22, 23] or STAR [24]. 3. To screen for T > C conversions, freely available analysis packages such as PARalyzer [25] are available. If your model system of interest has a simple genome like a virus, then a simple script can be used to screen for T > C conversions. Note that if you only use a simple T > C conversion screen with no statistics, we would recommend manually looking at each peak with the following criteria for reliable peaks: Each peak needs to consist of more than three distinct reads (varying in length or alignment location so that they are unlikely to be PCR duplicate reads), with at least three different locations of T > C conversions. 4. The Integrated Genome Viewer (IGV) from the Broad Institute [26] can be used to visualize sequencing results. Running Bowtie with the “--sam” argument will give results output in the SAM format. IGV reads alignments in the BAM format more efficiently, with a requirement that BAM files be presorted and indexed. The SAMtools suite [27] can be used to convert SAM to BAM, and sort and index the files for loading into IGV.

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Notes 1. If the model system has an unusually low abundance of uridines in the genes of interest, 6-thioguanosine (6SG, Sigma #858412 or Carbosynth #NT04480) can be used as an alternative to 4SU. While we have not tested 6SG to date, it was previously noted to have lower cross-linking efficiency, and will result in G > A conversions [15]. 2. We have used NEB CIP that has recently been discontinued. One potential alternative is alkaline phosphatase from Roche (#11097075001, 20 U/μL). The unit definitions from both NEB and Roche are the same: the amount of enzyme that hydrolyzes 1 μmol of p-nitrophenylphosphate (pNPP) in a 1 mL reaction at 37  C. Thus half the volume (2.5 μL) of the Roche CIP would be needed per reaction. 3. We have obtained from 1.1  108 HIV-1-infected CEM T cells 700 μg of total RNA, which yields 10 μg of poly(A)+ RNA (at 1.4% poly(A)+). For 293 T cells, we typically start with 10  15 cm plates at 50% confluency, whereas a good starting point for lymphoblastoid cell lines (Epstein-Barr virus immortalized B cells) would be 1.8  108 cells grown at a concentration of 6  105 cells/mL. 4. For HIV-1 studies, we typically use CEM T cells infected with HIV-1, remove the input virus, resuspend cells in fresh media 1 day postinfection (dpi), supplement with 4SU 2 dpi, and harvest at 3 dpi. 5. If mapping of RNA modifications on the genomic RNA of viruses is desired, as can be done with HIV-1, the supernatant of infected cells can also be collected for isolation of viral particles. Viral particles can be pelleted through a 20% sucrose cushion, and the virion pellet can then be lysed in Trizol to extract virion RNA, as described by Eckwahl et al. [28]. Virion RNA does not need to be poly(A) purified and can be directly used for immunoprecipitation. 6. The RNA re-precipitations in Subheading 3.2, steps 6 and 7, are optional if you typically do not see high salt contamination after Trizol extraction, or if the downstream poly (A) purification is not needed, as with virion-extracted RNA. 7. We recommend saving a 0.5 μg + aliquot of the input RNA used for the IP reaction. This input sample can be analyzed by RNA-seq run alongside the PA-mod-seq IP sample to measure the relative expression level of each transcript. 8. As UV does not penetrate plastic plate lids well, it is essential to cross-link with the lid off.

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9. The RNA pellet at this step will be small yet visible. If the pellet is not visible, add an additional 100 μL of isopropanol and 1 μL of GlycoBlue, mix well, and repeat the 20-min centrifugation step. A preincubation of the tube at 80  C or on dry ice prior to centrifugation may also enhance precipitation. 10. This optional re-precipitation is to ensure minimum salt contamination going into the library preparation. If the RNA pellet from the previous step is very small and hard to spot, we omit this step to avoid loss of the pellet.

Acknowledgments This research was funded in part by NIH grants R01-DA046111 and U54-GM103297 to B.R.C., along with a Duke University Center for AIDS Research (CFAR, P30-AI064518) pilot award to K.T. References 1. Li S, Mason CE (2014) The pivotal regulatory landscape of RNA modifications. Annu Rev Genomics Hum Genet 15:127–150. https:// doi.org/10.1146/annurev-genom-090413025405 2. Wang GG, Allis CD, Chi P (2007) Chromatin remodeling and cancer, part II: ATP-dependent chromatin remodeling. Trends Mol Med 13(9):373–380. https:// doi.org/10.1016/j.molmed.2007.07.004 3. Kennedy EM, Courtney DG, Tsai K, Cullen BR (2017) Viral epitranscriptomics. J Virol 91 (9). https://doi.org/10.1128/JVI.02263-16 4. Desrosiers R, Friderici K, Rottman F (1974) Identification of methylated nucleosides in messenger RNA from Novikoff hepatoma cells. Proc Natl Acad Sci U S A 71 (10):3971–3975. https://doi.org/10.1073/ pnas.71.10.3971 5. Lavi S, Shatkin AJ (1975) Methylated simian virus 40-specific RNA from nuclei and cytoplasm of infected BSC-1 cells. Proc Natl Acad Sci U S A 72(6):2012–2016. https://doi.org/ 10.1073/pnas.72.6.2012 6. Meyer KD, Saletore Y, Zumbo P, Elemento O, Mason CE, Jaffrey SR (2012) Comprehensive analysis of mRNA methylation reveals enrichment in 3’ UTRs and near stop codons. Cell 149(7):1635–1646. https://doi.org/10. 1016/j.cell.2012.05.003 7. Dominissini D, Moshitch-Moshkovitz S, Salmon-Divon M, Amariglio N, Rechavi G (2013) Transcriptome-wide mapping of N(6)-

methyladenosine by m(6)A-seq based on immunocapturing and massively parallel sequencing. Nat Protoc 8(1):176–189. https://doi.org/10.1038/nprot.2012.148 8. Dominissini D, Moshitch-Moshkovitz S, Schwartz S, Salmon-Divon M, Ungar L, Osenberg S, Cesarkas K, Jacob-Hirsch J, Amariglio N, Kupiec M, Sorek R, Rechavi G (2012) Topology of the human and mouse m6A RNA methylomes revealed by m6A-seq. Nature 485(7397):201–206. https://doi.org/ 10.1038/nature11112 9. Arango D, Sturgill D, Alhusaini N, Dillman AA, Sweet TJ, Hanson G, Hosogane M, Sinclair WR, Nanan KK, Mandler MD, Fox SD, Zengeya TT, Andresson T, Meier JL, Coller J, Oberdoerffer S (2018) Acetylation of cytidine in mRNA promotes translation efficiency. Cell 175(7):1872–1886. e1824. https://doi.org/ 10.1016/j.cell.2018.10.030 10. McIntyre ABR, Gokhale NS, Cerchietti L, Jaffrey SR, Horner SM, Mason CE (2020) Limits in the detection of m6A changes using MeRIP/m6A-seq. Sci Rep 10(1):6590. https://doi.org/10.1038/s41598-02063355-3 11. Chen K, Lu Z, Wang X, Fu Y, Luo GZ, Liu N, Han D, Dominissini D, Dai Q, Pan T, He C (2015) High-resolution N(6)-methyladenosine (m(6) A) map using photo-crosslinkingassisted m(6) A sequencing. Angew Chem Int Ed Engl 54(5):1587–1590. https://doi.org/ 10.1002/anie.201410647

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12. Courtney DG, Kennedy EM, Dumm RE, Bogerd HP, Tsai K, Heaton NS, Cullen BR (2017) Epitranscriptomic enhancement of influenza a virus gene expression and replication. Cell Host Microbe 22(3):377–386. e375. https://doi.org/10.1016/j.chom.2017.08. 004 13. Kennedy EM, Bogerd HP, Kornepati AV, Kang D, Ghoshal D, Marshall JB, Poling BC, Tsai K, Gokhale NS, Horner SM, Cullen BR (2016) Posttranscriptional m(6)a editing of HIV-1 mRNAs enhances viral gene expression. Cell Host Microbe 19(5):675–685. https:// doi.org/10.1016/j.chom.2016.04.002 14. Tsai K, Courtney DG, Cullen BR (2018) Addition of m6A to SV40 late mRNAs enhances viral structural gene expression and replication. PLoS Pathog 14(2):e1006919. https://doi. org/10.1371/journal.ppat.1006919 15. Hafner M, Landthaler M, Burger L, Khorshid M, Hausser J, Berninger P, Rothballer A, Ascano M Jr, Jungkamp AC, Munschauer M, Ulrich A, Wardle GS, Dewell S, Zavolan M, Tuschl T (2010) Transcriptome-wide identification of RNA-binding protein and microRNA target sites by PAR-CLIP. Cell 141(1):129–141. https://doi.org/10.1016/j.cell.2010.03.009 16. Courtney DG, Tsai K, Bogerd HP, Kennedy EM, Law BA, Emery A, Swanstrom R, Holley CL, Cullen BR (2019) Epitranscriptomic addition of m5C to HIV-1 transcripts regulates viral gene expression. Cell Host Microbe 26 (2):217–227.e216. https://doi.org/10. 1016/j.chom.2019.07.005 17. Tsai K, Jaguva Vasudevan AA, Martinez Campos C, Emery A, Swanstrom R, Cullen BR (2020) Acetylation of cytidine residues boosts HIV-1 gene expression by increasing viral RNA stability. Cell Host Microbe 28 (2):306–312.e6. https://doi.org/10.1016/j. chom.2020.05.011 18. Gordon A, Hannon G (2010) FastX toolkit. http://hannonlab.cshl.edu/fastx_toolkit/ 19. Martin M (2011) Cutadapt removes adapter sequences from high-throughput sequencing reads 2011. EMBnet J 17(1):3. https://doi. org/10.14806/ej.17.1.200 20. Langmead B (2010) Aligning short sequencing reads with Bowtie. Curr Protoc Bioinformatics

Chapter 11:Unit 11, 17. doi:https://doi.org/ 10.1002/0471250953.bi1107s32 21. Langmead B, Trapnell C, Pop M, Salzberg SL (2009) Ultrafast and memory-efficient alignment of short DNA sequences to the human genome. Genome Biol 10(3):R25. https:// doi.org/10.1186/gb-2009-10-3-r25 22. Trapnell C, Pachter L, Salzberg SL (2009) TopHat: discovering splice junctions with RNA-Seq. Bioinformatics 25(9):1105–1111. https://doi.org/10.1093/bioinformatics/ btp120 23. Trapnell C, Roberts A, Goff L, Pertea G, Kim D, Kelley DR, Pimentel H, Salzberg SL, Rinn JL, Pachter L (2012) Differential gene and transcript expression analysis of RNA-seq experiments with TopHat and Cufflinks. Nat Protoc 7(3):562–578. https://doi.org/10. 1038/nprot.2012.016 24. Dobin A, Davis CA, Schlesinger F, Drenkow J, Zaleski C, Jha S, Batut P, Chaisson M, Gingeras TR (2013) STAR: ultrafast universal RNA-seq aligner. Bioinformatics 29(1):15–21. https:// doi.org/10.1093/bioinformatics/bts635 25. Corcoran DL, Georgiev S, Mukherjee N, Gottwein E, Skalsky RL, Keene JD, Ohler U (2011) PARalyzer: definition of RNA binding sites from PAR-CLIP short-read sequence data. Genome Biol 12(8):R79. https://doi. org/10.1186/gb-2011-12-8-r79 26. Robinson JT, Thorvaldsdottir H, Winckler W, Guttman M, Lander ES, Getz G, Mesirov JP (2011) Integrative genomics viewer. Nat Biotechnol 29(1):24–26. https://doi.org/10. 1038/nbt.1754 27. Li H, Handsaker B, Wysoker A, Fennell T, Ruan J, Homer N, Marth G, Abecasis G, Durbin R (2009) Genome project data processing S: the sequence alignment/map format and SAMtools. Bioinformatics 25 (16):2078–2079. https://doi.org/10.1093/ bioinformatics/btp352 28. Eckwahl MJ, Arnion H, Kharytonchyk S, Zang T, Bieniasz PD, Telesnitsky A, Wolin SL (2016) Analysis of the human immunodeficiency virus-1 RNA packageome. RNA 22 (8):1228–1238. https://doi.org/10.1261/ rna.057299.116

Chapter 9 Quantitative and Single-Nucleotide Resolution Profiling of RNA 5-Methylcytosine Jun Li, Xingyu Wu, Trung Do, Vy Nguyen, Jing Zhao, Pei Qin Ng, Alice Burgess, Rakesh David, and Iain Searle Abstract RNA has coevolved with numerous posttranscriptional modifications to sculpt interactions with proteins and other molecules. One of these modifications is 5-methylcytosine (m5C) and mapping the position and quantifying the level in different types of cellular RNAs and tissues is an important objective in the field of epitranscriptomics. Both in plants and animals bisulfite conversion has long been the gold standard for detection of m5C in DNA but it can also be applied to RNA. Here, we detail methods for highly reproducible bisulfite treatment of RNA, efficient locus-specific PCR amplification, detection of candidate sites by sequencing on the Illumina MiSeq platform, and bioinformatic calling of non-converted sites. Key words Bisulfite conversion, Epitranscriptome, Fluidigm Access Array, Illumina, Next-generation sequencing, 5-Methylcytosine

1

Introduction Cellular RNAs can be modified, or decorated, with more than 120 chemically and structurally distinct nucleoside modifications [1]. The emerging field of epitranscriptomics [2] has been enabled by the development of high-throughput mapping methods for RNA modifications, typically based on second-generation sequencing. Transcriptome-wide positions of N1-methyladenosine (m1A, [3–5]), N6-methyladenosine (m6A, [6, 7]), 5-methylcytosine (m5C, [8]), and pseudouridine [9] have each been reported in this way. To detect m5C in RNA, a range of methods have been developed, including the indirect (aza-IP [10], miCLIP [11]) immunoprecipitation of methylated RNA or direct methods (meRIP, [7]). Of particular interest here, the bisulfite conversion approach previously used for DNA has been adapted to RNA [12, 13]. Bisulfite conversion of nucleic acids takes advantage of the differential chemical reactivity of m5C compared to

Mary McMahon (ed.), RNA Modifications: Methods and Protocols , Methods in Molecular Biology, vol. 2298, https://doi.org/10.1007/978-1-0716-1374-0_9, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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Fig. 1 Protocol overview showing the workflow for either parallel or single amplicon amplification for effective detection of m5C. (a) Parallel amplification and sequencing of up to 2304 amplicons across 48 tissues and 48 primer pairs. Forty-eight different tissues can be selected, total RNA isolated and purified, spiked with MGFP in vitro-transcribed control RNA and bisulfite converted. Bisulfite-converted RNA is reverse transcribed

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unmethylated cytosines; unmethylated cytosines are deaminated to uracil while m5C remains as a cytosine. The RNA bisulfite conversion method has been applied to animals and plants [8, 14] using second-generation sequencing, for example Illumina, based transcriptome-wide readout and has mapped thousands of novel candidate m5C sites in a diverse array of RNAs, including mRNAs and long noncoding RNAs (lncRNAs). Here, we detail protocols for RNA bisulfite conversion, locusspecific PCR amplification of up to 2304 amplicons, and bioinformatics calling of converted or non-converted sites. Sequencing of PCR amplicons is conveniently done on the Illumina MiSeq, as this affords multiplexing of multiple distinct amplicons while still achieving ample read depth for estimating the proportion of m5C at targeted positions. For instance, each of the 96 Fluidigm indexed adapters could be assigned to a separate RNA derived from different tissues, and 96 multiple PCR amplicons per sample could be included in the sequencing pool, potentially generating thousands of independent quantitative measurements of the m5C levels in a single MiSeq run (Fig. 1).

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Materials Prepare all solutions using RNase-free and DNase-free H2O and analytical grade reagents. Store and prepare all reagents at room temperature unless indicated otherwise. Prepare and perform bisulfite conversion, cDNA synthesis, and PCR amplification experiments in an RNase-free area. Follow all state or national safety and waste disposal regulations when performing experiments.

2.1 Total RNA Extraction and In Vitro Transcription

1. TRIzol™ reagent. 2. Chloroform. 3. Isopropanol. 4. 75% Ethanol. 5. Ultrapure™ H2O. 6. Monster Green® Fluorescent protein phMGFP vector.

ä Fig. 1 (continued) (RT) to cDNA using gene-specific RT primers that include the positive control MAG5 (AT5G47480) and negative control MGFP. Target regions are PCR amplified using a Fluidigm Access Array Integrated Fluidic Circuit (IFC); up to 2304 amplicons are harvested and eluted pools are quantified. Equal concentrations of the pools are combined into a final pool, purified using AMPure beads, accurately quantified, PhiX control library spiked-in, and subjected to sequencing on the Illumina MiSeq platform. (b) Single amplicon amplification and sequencing. A single tissue is selected, RNA isolated and purified in triplicate, spiked with MGFP in vitro-transcribed control RNA and bisulfite converted. Bisulfite conversion and cDNA synthesis are the same as outlined above except a specific target RT primer is used. The target amplicon is PCR amplified, triplicate amplicons are pooled, size and concentration are assessed on a Shimadzu MultiNA, and amplicons are pooled at equal concentration. Pooled amplicons are purified, PhiX control library spiked-in, and subjected to sequencing on the Illumina MiSeq platform

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7. XbaI restriction enzyme. 8. HiScribe™ T7 In Vitro Transcription Kit. 9. TURBO™ DNase. 10. Phase Lock Gel ™ QuantBio (2.0 mL). 11. UltraPure ™ Phenol:Water (3.75:1 v/v). 12. 100% Ethanol. 13. 3 M sodium acetate, pH 5.2. 14. 5 mg/mL Glycogen. 15. Agilent RNA 6000 Nano Kit. 16. Biological tissue samples (animal or plant). 2.2 Sodium Bisulfite Conversion

1. Sodium bisulfite solution: 40% (w/v) Sodium metabisulfite, 0.6 mM hydroquinone, final pH 5.1. To prepare the sodium bisulfite solution, prepare the following: 0.6 M Hydroquinone: Weigh 66 mg hydroquinone and place into a 1.5 mL tube. Add H2O to 1 mL and cover in foil to protect from light. Place in an orbital shaker to dissolve. 40% (w/v) Sodium bisulfite: Dissolve 4 g sodium metabisulfite in 10 mL H2O in a 50 mL falcon tube and vortex until it completely dissolves. Add 10μL 0.6 M hydroquinone to the 40% sodium bisulfite solution, vortex, and adjust pH to 5.1 with 10 M NaOH. Filter the solution through a 0.2μm filter. Cover in foil to protect from light (see Note 1). 2. 1 M Tris–HCl, pH 9.0. 3. Micro Bio-Spin ™ P-6 Gel Columns. 4. Mineral oil. 5. 75% Ethanol. 6. 100% Ethanol. 7. 3 M sodium acetate, pH 5.2. 8. 5 mg/mL Glycogen.

2.3

cDNA Synthesis

1. SuperScript ™ III Reverse Transcriptase. 2. 10 mM Mixed dNTPs. 3. Single-target priming: 20μM Gene-specific oligo for each amplicon. Here is an example of a positive control (MAG5) and a negative control (mGFP) (all C should be converted to U) primer sequence: MAG5: CACACACACCCATACATCCAC. mGFP: AACAAAAAAATTAACCCCATC 4. Pool target priming: Up to 48 primers at 20μM each. Design for target genes of interest.

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1. KAPA HiFi DNA polymerase. 2. First PCR primers: Examples of positive control (MAG5) and negative control (mGFP) first round primer primers are shown (underlined sequence ¼ tag, BC ¼ 8 nt barcode): MAG5 F: ACACTGACGACATGGTTCTACAGGTAAAGGT AAAATTGGGTAATGAG. MAG5 R: TACGGTAGCAGAGACTTGGTCT-[BC]-AGACCAAGTCT CTGCTACCGTA. mGFP F: ACACTGACGACATGGTTCTACAGAGGGTGAT GGGAAAGGTAAG. mGFP R: TACGGTAGCAGAGACTTGGTCTCAATCATCC ACACCCTTCATC 3. Second PCR primers (underlined sequence ¼ tag, BC ¼ 8 nt barcode): P5_CS1_F: AATGATACGGCGACCACCGAGATCTACACT GACGACATGGTTCTACA P7_BC_CS2_R: CAAGCAGAAGACGGCATACGAGAT -[BC]-TACGGTAGCAGAGACTTGGTCT 4. 10 mM Mixed dNTPs. 5. Fluidigm Access Array Integrated Fluidic Circuit (IFC) 48.48. 6. FastStart ™ High Fidelity PCR System, dNTPack. 7. 20 Access Array Loading Reagent. 8. 1 Access Array Harvest Solution. 9. 1 Access Array Hydration Reagent v2. 10. Access Array Barcode primers for Illumina Sequencers-384: Single Direction.

2.5 MultiNA Microelectrophoresis System

1. DNA-500 Kit (Shimadzu Corporation).

2.6 PCR Amplicon Purification and Quantification

1. Agencourt AMPure XP beads.

2.7 Library Sequencing Components

1. 0.2 M NaOH.

3

2. Qubit ™ dsDNA Broad Range Assay Kit. 3. KAPA Library Quantification Kit (Universal).

2. Illumina MiSeq Reagent Kit v3 (150 or 600 cycles) (see Note 2).

Methods Carry out all procedures described below at room temperature unless otherwise stated.

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3.1 RNA Extraction, Purification, and DNase Treatment

Total RNA is extracted and purified directly from tissue with 1 mL of TRIzol™ as per the manufacturer’s protocol. RNA is then treated with TURBO™ DNase as per the manufacturer’s protocol. Assess the integrity of the RNA by using a RNA 6000 Nano Chip on the Agilent 2100 Bioanalyzer according to the manufacturer’s protocol.

3.2 Generation of the MGFP In Vitro Transcript Spike-in Control

1. Linearize the phMGFP vector by using the restriction enzyme XbaI and purify the linearized DNA vector according to the HiScribe T7 In Vitro Transcription Kit protocol. 2. Perform in vitro transcription according to the HiScribe T7 In Vitro Transcription Kit protocol by using 1μg of linearized DNA. An incubation period of 4 h at 37  C with the kit components is sufficient. 3. Add 2 U TURBO™ DNase and incubate at 37  C for 30 min. 4. Transfer the reaction to a Phase Lock Gel™ tube and make the volume of the reaction up to 100μL with ultrapure H2O. 5. Add an equal volume of phenol:water and chloroform, shake vigorously for 15 s, and centrifuge at 15,000  g for 5 min. 6. Add the same volume of chloroform as in step 5 to the tube, shake vigorously for 15 s, and centrifuge at 15,000  g for 5 min again. 7. Transfer the aqueous phase to a clean 1.5 mL tube. Add 1/10 volume 3 M sodium acetate, 3 volumes of 100% ethanol, and 1μL glycogen; vortex; and precipitate the RNA overnight at 80  C. 8. Centrifuge RNA at 17,000  g at 4  C for 60 min and carefully remove the supernatant. 9. Add 1 mL 75% ethanol to the RNA, invert five times, and centrifuge at 7500  g at 4  C for 10 min (see Note 3). 10. Carefully remove the supernatant and let the pellet air-dry for approximately 15 min (see Note 4). 11. Resuspend the RNA in 25μL of ultrapure H2O. 12. Optional step: Treat 5μg of in vitro-transcribed MGFP transcript with 2 U TURBO™ DNase according to the manufacturer’s protocol at 37  C for 30 min. 13. Assess the integrity and size of the MGFP in vitro transcripts by using an RNA 6000 Nano Chip on the Agilent 2100 Bioanalyzer according to the manufacturer’s protocol (see Note 5).

3.3 Bisulfite Conversion of RNA

1. Add 1/2000 of the MGFP RNA transcript to 2μg DNasetreated purified total RNA. Increase the volume of the RNA sample to 20μL with ultrapure H2O. 2. Denature RNA by heating to 75  C for 5 min in a heat block.

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3. Preheat the sodium bisulfite solution to 75  C, add 100μL to the RNA, vortex thoroughly, and briefly spin in a microcentrifuge (13,000  g for 1 min). 4. Overlay the reaction mixture with 100μL of mineral oil. Cover the tube in aluminum foil to protect the reaction mixture from light (see Note 6). 5. Incubate at 75  C for 4 h in a heat block. 6. About 15 min before the bisulfite conversion reaction is complete, prepare two Micro Bio-Spin Columns for each conversion reaction by allowing the Tris solution in the column to drain into a collection tube. Discard the Tris flow-through, place the column back into the collection tube, and centrifuge at 1000  g for 2 min. Transfer each column to a clean 1.5 mL tube (see Note 7). 7. Remove the bisulfite reaction mixture from the heat block and gently transfer the aqueous layer (that is under the mineral oil) containing the sodium bisulfite/RNA mixture to the Micro Bio-Spin column (see Note 8). 8. Centrifuge at 1000  g for 4 min. 9. Carefully transfer the eluate into the second Micro Bio-Spin column placed in a 1.5 mL tube and repeat step 8. 10. Preheat the temperature of the heat block to 75  C in preparation for step 12. 11. Add an equal volume of 1 M Tris–HCl pH 9.0 to the second eluate, vortex, spin briefly, and then overlay with 175μL of mineral oil. Cover the tube in aluminum foil to protect the reaction mixture from light. 12. Incubate at 75  C for 1 h in the heat block. 13. Transfer the bottom aqueous layer containing the RNA to a clean 1.5 mL tube. 14. Precipitate the bisulfite-treated RNA by following steps 7–11 in Subheading 3.2 and resuspend the bisulfite-converted RNA in H2O (see Note 9). 3.4 Bisulfite Oligonucleotide Primer Design for cDNA Synthesis and PCR

1. For efficient parallel amplification of 48 target amplicons on the Fluidigm Access Array, use targeted cDNA synthesis to reduce the amplification of spurious amplicons. Targeted cDNA synthesis is achieved by designing reverse transcriptase (RT) primers 30–40 nt 30 of the cytosine(s) to be assayed. N. B.: Design the RT primers such that they avoid areas of bisulfite-converted cytosines as inefficient BS conversion may result in unconverted cytosines and biasing of later amplification. See Fig. 2.

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Fig. 2 Overview of bisulfite conversion of RNA, reverse transcription to cDNA, and PCR amplification. (a) In the in vitro-transcribed MGFP sequence, unmodified cytosines (underlined) are converted to uracil, reverse transcribed (RT) by reverse transcriptase to cDNA, and then PCR amplified. RT and PCR primers are designed to avoid stretches of converted cytosines to prevent preferential amplification of converted sequences which may incorrectly indicate efficient bisulfite conversion. (b) In MAG5 control and other candidate sequences, primers are designed to span areas containing converted cytosines to preferentially amplify converted sequences. C3349 is methylated in Arabidopsis thaliana and serves as an over-conversion control. Flanking cytosines are not methylated and should be completely converted. Primers are designed with a Tm of 59–61  C, preferably with a 30 G nucleotide and to amplify PCR products of 170–200 bp

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Fig. 3 Overview of first and second PCR amplification of target regions. (a) For the first PCR, the forward PCR primer is designed with the gene-specific sequence (GS) and universal forward tag called Common Sequence, CS1 (50 - TACGGTAGCAGAGACTTGGTCT -30 ), and reverse PCR primer is designed with the gene-specific sequence (GSS) and universal reverse tag called Common Sequence CS2 (5’-ACACTGACGACATGGTTCTACA -30 ). (b) For the second PCR, the forward primer is designed with the CS1 and Illumina P5 sequences and the reverse primer contains the CS2, barcoding, and Illumina P7 sequences. The Fluidigm barcodes or indexes are 10 nt in length

2. Design primers for the first round of PCR amplification so that small amplicons are 170–200 bp, to allow efficient amplification (see Notes 10 and 11). As the G/C content in the template is low, design long primers to ensure that a Tm is in the rage of 59–61  C. Add the CS1 sequence (50 -TACGGTAGCA GAGACTTGGTCT -30 ) to the forward primer gene-specific sequence (GSS) and CS2 (50 - ACACTGACGACATGGTTC TACA -30 ) to the reverse primer GSS. For the second PCR amplification, use the forward primer containing the complementary sequences to the P5 Illumina flow cell combined with CS1 (P5_CS1) and the reverse primer containing the barcode, and complementary sequences to the P7 Illumina flow cell combined with CS2 (P7_BC_CS2) primer (see Note 12). See Fig. 3. 3.5

cDNA Synthesis

1. Mix 500 ng of bisulfite-converted RNA, 1μL of 1 mM dNTP mix, and 2μL of 10 pooled primer mix and add ultrapure H2O to a final volume of 13μL. Incubate the mix at 65  C for 5 min to denature the RNA. 2. Reverse transcribe the bisulfite-converted RNA using SuperScript™ III Reverse Transcriptase according to the manufacturer’s protocol. Add either pooled 48 RT primers for parallel Access Array amplification or random hexamers for single-PCR amplicons.

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Suggested controls: Include RT minus controls for each sample as the PCR primers are not necessarily designed to span exon-exon junctions. In the controls, use 1μL of H2O instead of reverse transcriptase. 3. After the reaction is complete, dilute the cDNAs 1:10 in ultrapure H2O for PCR amplification. 3.6 Individual PCR Amplification, Quantification, and Pooling

1. For a 10μL PCR, add 0.2μL of KAPA HiFi DNA Polymerase, 2μL of 5 HiFi Fidelity buffer (with MgCl2), 0.3μL of 10 mM dNTP, 0.4μL of 10μM forward primer (CS1_GSS), 0.4μL of 10μM reverse primer (CS2_GSS), 1μL of diluted cDNA, and H2O to a final volume of 10μL. Perform PCR for each amplicon in triplicate. 2. Gently finger vortex, briefly centrifuge, and place into a preheated thermal cycler. 3. Perform a two-step thermal cycling PCR program. See Table 1 for more details. 4. Pool the triplicates and perform an AMPure bead cleanup at a ratio of 1.8:1 to remove unincorporated primers and primer dimers. Repeat this step (see Notes 13 and 14). 5. Assess PCR amplicon size and concentration after separation on a Shimadzu Microchip Electrophoresis System MCE®-202 MultiNA. 6. Normalize the concentration of each amplicon in the experiment by dilution with H2O to a concentration in the range of 0.5–5 ng/μL. 7. Perform the barcoding and Illumina adapter addition PCR. In a 10μL PCR, add 0.2μL of KAPA HiFi DNA Polymerase, 2μL of 5 HiFi Fidelity buffer (with MgCl2), 0.3μL of 10 mM dNTP, 1μL of 10μM forward primer (P5_CS1), 1μL of 10μM reverse primer (P7_CS2), 2μL of diluted PCR amplicon, and H2O to a final volume of 10μL. 8. Gently finger vortex, briefly centrifuge, and place into a preheated thermal cycler. 9. Perform a two-step thermal cycling PCR program. See Table 2 for more details. 10. Assess PCR amplicon size and concentration after separation on a Shimadzu Microchip Electrophoresis System MCE®-202 MultiNA. 11. Pool the amplicons in equimolar concentration and purify them using AMPure beads according to the manufacturer’s protocol. Use a ratio of beads to pooled amplicons of 0.9:1 to ensure binding of amplicons and not primer dimers or unincorporated primers.

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Table 1 Two-step thermal cycling conditions for the amplification of individual amplicons Stage

Temperature ( C)

Time (s)

Initial denaturation

98

15

Denaturation

94

10

Annealing

60

30

Extension

72

15

Denaturation

94

10

Annealing

55

30

Extension

72

15

Final extension

72

60

Hold

4

Forever

Step I (10 cycles)

Step II (20 cycles)

Table 2 One-step thermal cycling conditions for the addition of barcodes and Illumina adapters Stage

Temperature ( C)

Time (s)

Initial denaturation

98

15

Denaturation

94

10

Annealing

63

30

Extension

72

30

Final extension

72

120

Hold

4

Forever

One step (12 cycles)

12. First estimate the DNA concentration using a Qubit dsDNA Broad Range Assay Kit according to the manufacturer’s protocol. Then accurately assess the DNA concentration by using KAPA Library Quantification Kit for Illumina® Platforms. Perform serial dilution of the pooled amplicons such that they fall into the dynamic range of the assay of 5.5–0.000055 pg/μL. 3.7 Parallel PCR Amplification Using a Fluidigm Access Array Integrated Fluidic Circuit (IFC)

1. Prime the Access Array according to the manufacturer’s protocol. 2. Pre-warm the 20 Access Array loading reagent to room temperature before use. Prepare the pooled 48-oligonucleotide

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primer mix by mixing 2.0μL of 50μM CS1-GS forward, 2.0μL of 50μM CS1-GS reverse, 5.0μL of 20 Access Array loading reagent, and 91μL of H2O to a final volume of 100μL. 3. Finger vortex the mix and centrifuge to spin the contents to the bottom of the tube. 4. Prepare the sample premix solution by mixing 30μL 10 FastStart High Fidelity Reaction Buffer (without MgCl2), 54μL 25 mM MgCl2, 15μL DMSO, 6.0μL 10 mM dNTP mix, 3.0μL FastStart High Fidelity Enzyme Blend, 15.0μL 20 Access Array Loading Reagent, and 57μL H2O. 5. Finger vortex the mix and centrifuge to spin the contents to the bottom of the tube. 6. Prepare the sample mix solutions, 48 in total, in a 96-well plate. Mix 3.0μL sample premix, 1.0μL cDNA, and 1.0μL Access Array Barcode library primers. 7. Thoroughly vortex the solutions for at least 30 s and then centrifuge to spin down the contents to the bottom of the plate. N.B.: Each well should receive a uniquely barcoded primer pair. 8. Load 4.0μL of the primer solution and 4.0μL of the sample mix solution into the primer and sample inlets of the Access Array by using an 8-channel pipette. 9. Load the Access Array into the Pre-PCR IFC Controller AX according to the manufacturer’s protocol. 10. Place the Access Array onto the FC1 Cycler and start thermal cycling by selecting the protocol AA 48  48 Standard v1. The thermal cycling conditions are presented in Table 3. 11. To harvest the PCR products from the Access Array follow the manufacturer’s protocol. Once the final step is completed, eject the Access Array. 12. Collect the harvested PCR products into a labeled PCR 96-well plate. Carefully transfer 10μL of harvested PCR products from each of the sample inlets into columns 1–6 of the labeled 96-well plate by using an 8-channel pipette. 13. Assess PCR amplicon size and concentration after separation on a Shimadzu Microchip Electrophoresis System MCE®-202 MultiNA. 14. Pool the amplicons in equimolar concentration and purify them using AMPure beads according to the manufacturer’s protocol. Use a ratio of beads to pooled amplicons of 0.9:1 to ensure binding of amplicons and not primer dimers or unincorporated primers (see Note 14). 15. First estimate the DNA concentration using a Qubit dsDNA Broad Range Assay Kit according to the manufacturer’s

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Table 3 Multistep thermal cycling conditions for the Access Array Temperature ( C)

Time (s)

Number of cycles

50

120

1

70

1200

1

95

600

1

95 60 72

15 30 60

10

95 80 60 72

15 30 30 60

2

95 60 72

15 30 60

8

95 80 60 72

15 30 30 60

2

95 60 72

15 30 60

8

95 80 60 72

15 30 30 60

5

protocol. Then accurately assess the DNA concentration by using KAPA Library Quantification Kit for Illumina® Platforms. Perform serial dilution of the pooled amplicons such that they fall into the dynamic range of the assay of 5.5–0.000055 pg/μL. 3.8 MiSeq Sequencing

1. Prepare the sample sheet using the Illumina Experiment Manager by following the manufacturer’s protocol (see Note 15). 2. Dilute the library to 10 nM in EBT buffer based on the concentrations determined by the qPCR. From this point, keep the libraries on ice. 3. Dilute the PhiX control library to 2 nM by adding 8μL EBT buffer to 2μL of the 10 nM PhiX control library (see Note 16). 4. Denature the pooled libraries and PhiX control library separately by adding 10μL of 0.2 M NaOH to 10μL of the 2 nM libraries (see Note 17).

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5. Vortex thoroughly to mix and centrifuge at 1000  g for 30 s. Incubate at room temperature for 5 min. 6. Dilute the denatured pooled libraries and PhiX control library separately to 20 pM by adding 980μL pre-chilled HT1 to 20μL denatured libraries. 7. Dilute the 20 pM pooled libraries and PhiX control library separately to 10 pM by adding 500μL pre-chilled HT1 to 500μL 20 pM libraries. 8. Combine 100μL of the 10 pM PhiX control library with 900μL of the 10 pM pooled libraries and vortex to mix (see Note 18). 9. Load 600μL of the final sample into the cartridge. Ensure that air bubbles are removed by gently tapping the cartridge. 10. Perform the sequencing run according to the manufacturer’s protocol. 3.9 Bioinformatics Analysis of Data

1. To trim the Illumina adaptor sequences that were incorporated into the amplicons to permit sequencing of the 150 bp pairedend reads, use Trimmomatic in palindromic mode [15]. 2. Sequencing reads can be aligned with meRanTK by using Bowtie2 internally [16]. Assemble reference sequences for the alignment by using the segments of RNA interrogated by sequencing prior to bisulfite conversion. 3. Extract the methylation state of individual cytosines from bisulfite-read alignments by using meRanCall. The number of reads can be extracted from the aligned sequencing reads in order to determine read coverage at a given cytosine. 4. To call differentially methylated cytosines use meRanCompare. The number of reads can be extracted from the aligned sequencing reads in order to determine read coverage at a given cytosine (Fig. 4).

4

Notes 1. Slowly add 10 M NaOH dropwise to the sodium bisulfite solution while mixing. Slightly less than 1 mL is required to adjust the pH to 5.1. 2. The MiSeq Reagent Kit v3 (150- or 600-cycle) provides 1  150 bp or the 600-cycle kit allows combinations of cycles that add to 600, for example 200 and 400 cycles. 3. Do not machine or finger vortex the RNA as this will increase the risk of RNA loss. 4. Air-drying the samples in a sterile laminar flow hood is recommended. Do not allow the RNA to completely dry as this will cause difficulties in resuspending the RNA.

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Fig. 4 Representative analysis of an Illumina MiSeq amplicon sequencing of negative and positive controls. (a) A region of the MGFP spiked-in in vitro control transcript showing even coverage and all cytosines are converted (no methylation). The y-axis shows the read depth and the x-axis shows the cytosines (numbers) in the sequenced region. (b) A region of the Mag5 gene that shows converted and non-converted cytosine, C3349. Cytosines flanking C339 are completely converted, demonstrating that bisulfite conversion was very efficient. The heatmaps display the cytosine non-conversion percentage

5. As the in vitro MGFP transcript will most likely be at a high concentration, it is good practice to perform a serial dilution in H2O such that the estimated concentrations are in the range of 5–50 ng/μL. Prepare and run three dilutions on the RNA Nano chip.

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6. Tilt the 1.5 mL tube at a 45 angle and then slowly pipette the mineral oil directly on top of the RNA-bisulfite reaction mixture. 7. Emptying of the Micro Bio-Spin gel column takes about 2 min. If the gel column does not empty by gravity, place the lid back onto the column and remove again. 8. Gently pipette the reaction mixture onto the gel bed and avoid disturbing the gel bed. Minimize the transfer of mineral oil to the column although there will be traces which is unavoidable. 9. About 25% of the RNA is lost during the procedure, and we find that 10μL of H2O/2μg RNA used in the bisulfite conversion reaction results in concentrations of ~150 ng/μL. 10. Bisulfite treatment of the RNA causes significant shearing and we have observed that shorter amplicons are preferentially amplified over longer amplicons. Longer PCR amplicons increase the tendency of detecting non-converted cytosines in RNA exhibiting strong secondary structure. 11. Inefficient bisulfite conversion may result in unconverted cytosines, so it is important to ensure that the PCR primers are not biasing the amplification toward converted cytosines. 12. Occasionally, not all triplicates successfully amplify and it may be necessary to optimize the PCR. 13. We elute the purified PCR products in 10–30μL depending on the amount of amplified PCR products. 14. After purification of the amplicons, residual ethanol may remain in the purified amplicons. We find that concentrating down the pooled amplicons even if there is Output_File_Read1_trimmed.fastq 4. Use the following awk command in order to discard the sequences shorter than 10 nt: awk ’BEGIN {FS ¼ "\t" ; OFS ¼ "\n"} {header ¼ $0 ; getline seq ; getline qheader ; getline qseq ; if (length(seq) >¼ 10) {print header, seq, qheader, qseq}}’ Output_File_Read1_trimmed. fastq > Length_Filtered.fastq 3.4.2 Mapping of the Trimmed Sequences

1. Download the database corresponding to the organism of interest from miRBase as follows. Go to http://www.mirbase. org/ftp.shtml and download the “mature.fa” file. Note that the sequences are indicated in RNA notation. Replace the U residues by T with the following command (see Note 3): sed -i ’/^>/! s/U/T/g’ mature.fa 2. Select the miRNA sequences of your organism of interest with the following command: awk ’/name_of_the_organism/{print; nr[NR+1]; next}; NR in nr’ mature.fa > mature_name_of_the_organism_mirs.fa 3. Map the reads to the above-created file using Bowtie2 [21] (version 2.3.0) allowing no mismatches. First, build an index for your file with the following command: bowtie2-build mature_name_of_the_organism_mirs.fa mature_name_of_the_organism_mirs 4. Align the sequencing reads to the database, requiring that a read map entirely to a miRNA of the database, without any mismatches. To this end, use the following tool (see Note 4):

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bowtie2 -N 0 -L 10 --score-min C,0,0 --end-to-end --time -x mature_name_of_the_organism_mirs -U Length_Filtered. fastq -S Length_Filtered_ALIGNMENT.sam 5. To discard the reads that did not align, use the following command (see Note 5): samtools view -F 4 Length_Filtered_ALIGNMENT.sam > Reads_aligned_to_Mirs.sam

4

Notes 1. Small amounts of gel may remain in the 0.5 mL tube; carefully transfer with a pipette tip. 2. We typically perform 11 cycles, but this should be optimized by the user. Try to perform the smallest possible number of PCR cycles. 3. This command will yield a complete list of all miRNAs in miRBase, originating from a variety of organisms. 4. The option --score-min C,0,0 ensures that alignment is without any mismatches. For an explanation of the various parameters in the tool, please visit the following website: http://bowtiebio.sourceforge.net/bowtie2/manual.shtml. 5. As a result of these steps, you should now have obtained the aligned reads, corresponding to miRNAs.

Acknowledgments This work was supported by the National Center for Scientific Research (CNRS), the French Alternative Energies and Atomic Energy Commission (CEA), and Paris-Sud University. The members of the I2BC Next-Generation Sequencing service are acknowledged for critical reading of the manuscript and helpful suggestions. References 1. Ghildiyal M, Zamore PD (2009) Small silencing RNAs: an expanding universe. Nat Rev Genet 10(2):94–108. https://doi.org/10. 1038/nrg2504 2. Chang TC, Mendell JT (2007) microRNAs in vertebrate physiology and human disease. Annu Rev Genomics Hum Genet 8:215–239. https://doi.org/10.1146/annurev.genom.8. 080706.092351 3. Zhuang F, Fuchs RT, Robb GB (2012) Small RNA expression profiling by high-throughput

sequencing: implications of enzymatic manipulation. J Nucleic Acids 2012:360358. https:// doi.org/10.1155/2012/360358 4. van Dijk EL, Jaszczyszyn Y, Thermes C (2014) Library preparation methods for nextgeneration sequencing: tone down the bias. Exp Cell Res 322(1):12–20. https://doi.org/ 10.1016/j.yexcr.2014.01.008 5. Munafo DB, Robb GB (2010) Optimization of enzymatic reaction conditions for generating representative pools of cDNA from small

Improved Capture of 20 -O-Methyl RNAs by Small RNA-seq RNA. RNA 16(12):2537–2552. https://doi. org/10.1261/rna.2242610 6. Hafner M, Renwick N, Brown M, Mihailovic A, Holoch D, Lin C, Pena JT, Nusbaum JD, Morozov P, Ludwig J, Ojo T, Luo S, Schroth G, Tuschl T (2011) RNA-ligasedependent biases in miRNA representation in deep-sequenced small RNA cDNA libraries. RNA 17(9):1697–1712. https://doi.org/10. 1261/rna.2799511 7. Sorefan K, Pais H, Hall AE, Kozomara A, Griffiths-Jones S, Moulton V, Dalmay T (2012) Reducing ligation bias of small RNAs in libraries for next generation sequencing. Silence 3(1):4. https://doi.org/10.1186/ 1758-907X-3-4 8. Sun G, Wu X, Wang J, Li H, Li X, Gao H, Rossi J, Yen Y (2011) A bias-reducing strategy in profiling small RNAs using Solexa. RNA 17 (12):2256–2262. https://doi.org/10.1261/ rna.028621.111 9. Jayaprakash AD, Jabado O, Brown BD, Sachidanandam R (2011) Identification and remediation of biases in the activity of RNA ligases in small-RNA deep sequencing. Nucleic Acids Res 39(21):e141. https://doi.org/10.1093/ nar/gkr693 10. Zhuang F, Fuchs RT, Sun Z, Zheng Y, Robb GB (2012) Structural bias in T4 RNA ligasemediated 30 -adapter ligation. Nucleic Acids Res 40(7):e54. https://doi.org/10.1093/ nar/gkr1263 11. Fuchs RT, Sun Z, Zhuang F, Robb GB (2015) Bias in ligation-based small RNA sequencing library construction is determined by adaptor and RNA structure. PLoS One 10(5): e0126049. https://doi.org/10.1371/journal. pone.0126049 12. Dard-Dascot C, Naquin D, d’AubentonCarafa Y, Alix K, Thermes C, van Dijk E (2018) Systematic comparison of small RNA library preparation protocols for nextgeneration sequencing. BMC Genomics 19 (1):118. https://doi.org/10.1186/s12864018-4491-6 13. Van Nieuwerburgh F, Soetaert S, Podshivalova K, Ay-Lin Wang E, Schaffer L, Deforce D, Salomon DR, Head SR,

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Ordoukhanian P (2011) Quantitative bias in Illumina TruSeq and a novel post amplification barcoding strategy for multiplexed DNA and small RNA deep sequencing. PLoS One 6(10): e26969. https://doi.org/10.1371/journal. pone.0026969 14. Harrison B, Zimmerman SB (1984) Polymerstimulated ligation: enhanced ligation of oligoand polynucleotides by T4 RNA ligase in polymer solutions. Nucleic Acids Res 12 (21):8235–8251 15. Song Y, Liu KJ, Wang TH (2014) Elimination of ligation dependent artifacts in T4 RNA ligase to achieve high efficiency and low bias microRNA capture. PLoS One 9(4):e94619. https://doi.org/10.1371/journal.pone. 0094619 16. Zhang Z, Lee JE, Riemondy K, Anderson EM, Yi R (2013) High-efficiency RNA cloning enables accurate quantification of miRNA expression by deep sequencing. Genome Biol 14(10):R109. https://doi.org/10.1186/gb2013-14-10-r109 17. Barberan-Soler S, Vo JM, Hogans RE, Dallas A, Johnston BH, Kazakov SA (2018) Decreasing miRNA sequencing bias using a single adapter and circularization approach. Genome Biol 19(1):105. https://doi.org/10. 1186/s13059-018-1488-z 18. van Dijk EL, Eleftheriou E, Thermes C (2019) Improving small RNA-seq: less bias and better detection of 2’-O-methyl RNAs. J Vis Exp (151). https://doi.org/10.3791/60056 19. Chen YR, Zheng Y, Liu B, Zhong S, Giovannoni J, Fei Z (2012) A cost-effective method for Illumina small RNA-Seq library preparation using T4 RNA ligase 1 adenylated adapters. Plant Methods 8(1):41. https://doi. org/10.1186/1746-4811-8-41 20. Martin M (2011) Cutadapt removes adapter sequences from high-throughput sequencing reads. EMBnet. https://doi.org/10.14806/ ej.17.1.200 21. Langmead B, Trapnell C, Pop M, Salzberg SL (2009) Ultrafast and memory-efficient alignment of short DNA sequences to the human genome. Genome Biol 10(3):R25. https:// doi.org/10.1186/gb-2009-10-3-r25

Part IV Assessing RNA Modifications Using qPCR- and Molecular Biology-Based Methods

Chapter 11 Assessing 20 -O-Methylation of mRNA Using Quantitative PCR Brittany A. Elliott and Christopher L. Holley Abstract 20 -O-methylation (Nm) is an RNA modification commonly found on rRNA and snRNA, and at the mRNA 50 -cap, but has more recently been found internally on mRNA. The study of internal Nm modifications on mRNA is in the early stages, but we have reported that this sort of Nm modification can regulate mRNA abundance and translation. Although there are many methods to determine the presence of Nm on rRNA, detecting Nm on specific mRNA transcripts is technically difficult because they are much less abundant than rRNA. Some of these methods rely on the fact that Nm modification of RNA disrupts reverse transcription reactions when performed at low dNTP concentrations. In this chapter, we describe our approach to using quantitative PCR in conjunction with reverse transcription at low dNTPs, which is sensitive enough to detect changes to Nm modification of mRNA. Key words 20 -O-methylation, RNA modifications, Reverse transcription, Low dNTP, qPCR

1

Introduction 20 -O-methylation is a posttranscriptional ribose modification that can occur in conjunction with any RNA base (Nm, see Fig. 1). Nm is commonly found on noncoding RNAs such as ribosomal RNA (rRNA), snRNA, and tRNA, as well as the first and second nucleotides at the 50 -cap of mRNA [1–3]. More recently, transcriptome-wide mapping has suggested that Nm sites are also present internally on mRNA and pre-mRNA [4]. Validating these novel sites on an individual basis has been challenging, given the low abundance of gene-specific transcripts. Using genetic models, we have recently demonstrated snoRNA-guided Nm modification of Pxdn mRNA and shown that this modification inhibits translation in vivo—which is consistent with other in vitro work [5– 8]. Our findings demonstrate that at least some mRNA Nm sites

Supplementary Information The online version of this chapter (https://doi.org/10.1007/978-1-0716-13740_11) contains supplementary material, which is available to authorized users. Mary McMahon (ed.), RNA Modifications: Methods and Protocols, Methods in Molecular Biology, vol. 2298, https://doi.org/10.1007/978-1-0716-1374-0_11, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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“Nm”

“N”

Base

Base

2’-O-methyltransferase + SAM Fig. 1 20 -O-methylation (Nm) adds a methyl group to any nucleoside ribose (red arrow). The 20 -O-methyltransferase (i.e., fibrillarin and others) uses S-adenosyl methionine (SAM) as the methyl donor

are guided by snoRNAs and catalyzed by the enzyme fibrillarin (FBL), which is the same mechanism used for Nm modification of rRNA and snRNA. Although there are no antibodies that can specifically bind and detect Nm sites, Nm sites can be detected by mass spectrometry, resistance to site-specific cleavage, oxidation-elimination chemistry, and interference with reverse transcription [9–13]. Mass spectrometry has the advantage of being quantitative and exquisitely specific for detecting nucleoside modifications, but most applications involve the digestion of RNA into single nucleosides, thereby losing positional information about the modification(s). The chemistry of Nm sites can also be exploited for detection and mapping. Nm sites are resistant to alkaline hydrolysis (by preventing 20 -OH nucleophile attack on the 30 -phosphate backbone), and this property has been exploited for mapping them with primer extension techniques and newer high-throughput mapping methods such as RiboMeth-seq [11]. Nm sites are also resistant to oxidationelimination chemistry, which is the basis of the RNA-seq-based Nm mapping methods known as Nm-seq and RibOxi-seq [4, 12]. Unfortunately, these methods are limited by the requirement of a large amount of purified starting material or high-depth RNA-sequencing and are not practical for routine detection of Nm modifications on less abundant RNA molecules such as mRNA. The most commonly used method for detecting Nm on RNA relies on the fact that reverse transcription (RT) is inhibited by Nm modifications, if the RT reaction is carried out with low amounts of dNTPs. The presence of limiting dNTPs generally reduces processivity of the reverse transcriptase, which results in pausing or stoppage at Nm and other points of steric hindrance (such as highly structured regions) [14]. One of the earliest applications was to map Nm sites on rRNA and snRNAs using radiolabeled primers and primer extension assay at low dNTPs. This approach was very successful and a mainstay of the field for several decades [15]. There has also been more recent work done to develop RT enzymes that

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A.

Gm1328

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B.

Um1326 Gm1328

Fig. 2 Detection of Nm sites by primer extension. (a) SuperScript III Reverse Transcriptase (SSIII RT) or (b) an engineered 20 -O-methyl-sensitive DNA polymerase (Klen Taq V669L; KTQ) was used to detect Nm on 18S rRNA by fluorescently labeled primer extension. RNAs from hearts of WT and Rpl13a snoRNA KO mice (lacking U32A, U33, U34, and U35A) were processed with (a) SSIII RT and low dNTP or (b) Nm-sensitive KTQ. Rpl13a snoRNA KO mice lack U33 that guides 18S Um1326 and they also lack U32A, which is one of the two guides for 18S Gm1328 (U32B is a redundant snoRNA with the same guide sequence). Reduction of Gm1328 can be clearly seen in KO mice from samples processed with SSIII with low dNTPs (a, arrow), but nearby Um1326 is unable to be detected in WT or KO RNA (a). In samples processed with Nm-sensitive KTQ, both Um1326 and Gm1328 are resolved in WT mice (b, arrows). In KO mice, loss of Um1326 is observed as well as partial loss of methylation of Gm1328. Loss of Gm1328 is incomplete due to the redundant U32B snoRNA

are inhibited by Nm at high (normal) dNTP concentrations. One such enzyme is an Nm-sensitive DNA polymerase with RT activity: Klen Taq V669L [16]. To avoid radioisotopes, we have used fluorescently labeled RT primers to detect Nm on abundant rRNA using both SuperScript III at low dNTPs and Klen Taq V669L at high dNTPs, in conjunction with a genetic model that lacks snoRNAs guiding the modifications in question. In the presence of low dNTPs, presence and reduction of 18S Gm1328 were detected with RT SuperScript III (Fig. 2a). However, detection of Nm was even better using Klen Taq V669L. Using this approach, a “stop” product from both 18S Gm1328 and nearby Um1326 could be easily visualized, and loss or reduction in these sites could be seen in animals lacking the Rpl13a snoRNAs that guide them (Fig. 2b). Reverse transcription “low dNTP” methods and engineered enzymes suffer from several weaknesses that must be considered, including lack of specificity, need to know approximate position of Nm site, and poor sensitivity for low-abundance transcripts with primer extension. One contributor to the lack of specificity is that RNA modifications other than Nm can also disrupt RT to a lesser extent, and that inherently structured regions of RNA can strongly inhibit RT at low dNTPs [14, 15]. It is therefore essential to have proper controls, such as unmodified synthetic RNA or genetic models that lack the modification being studied.

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A.

RT

m 5’ Fu

FD

B. RT primer(s) 3’ RNA target PCR primers

Methylated Transcript (WT) m 1. mRNA

OR

RT odT

AAAAAA 3’

5’ F

R

C.

RT GSP m

R

RT primer(s) mRNA target qPCR primers

Unmethylated Transcript (snoRNA KO or FBL KD)

2. RT High dNTP m

3. qPCR

Low dNTP mx

m

mx

m

m

High dNTP

Low dNTP

Less Nm

More Nm

Fig. 3 Detecting snoRNA-guided 20 -O-methylation of mRNA using reverse transcription under low-dNTP conditions followed by qPCR (RTL-P). (a) Schematic of primer design for RTL-P as described in Dong et al., 2012. (b) Schematic of primer design sensitive to the presence or absence of 20 -O-methylation on mRNA. Oligo-dT or gene-specific primers (GSP) for target and housekeeping genes are used as an RT primer. qPCR primers (F & R) are designed to be upstream of the putative Nm site. (c) Schematic of expected results at each step of the assay. (1) Total RNA is extracted from control and fibrillarin (Fbl) or snoRNA KO cells or tissue. (2) Under high-dNTP conditions, RT will read through Nm and methylated transcripts will be indistinguishable from unmethylated ones. In low-dNTP conditions, RT will frequently pause at sites of Nm, resulting in truncated transcripts that will not be amplified by PCR. (3) RT products are amplified by qPCR. In this example, loss of Nm modification due to snoRNA KO leads to increased read-through during RT, which leads to increased qPCR product

To overcome sensitivity limitations with primer extension, the low dNTP/primer extension approach has been combined with PCR amplification, allowing for the use of very little starting material (see “Reverse Transcription Under Low dNTP Conditions Followed by PCR (RTL-P)” [17]). In this method, RT is performed at both high (standard) and low dNTP conditions, followed by PCR amplification of the cDNA (Fig. 3). In its original form (Fig. 3a, c), the products of endpoint PCR are run on an agarose gel, quantified by imaging, and the relative inhibition of RT by Nm is calculated by comparing the amount of product in the low dNTP condition to the standard reaction. A forward primer 50 to the methylation (Nm sensitive, Fu) will generate a longer product than a forward primer 30 to the point of the methylation (Nm unsensitive, FD). Following endpoint PCR, the amplified reactions are run on an electrophoresis gel and analyzed using densitometry of a fluorescent nucleotide reporter dye. In conditions where an Nm is present, there will be less observed longer

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product compared with short product (FU/FD) in the low-dNTP condition, whereas the ratio will be nearly equal in high dNTP (Fig. 3a, c). Although the sensitivity of RTL-P assay for detecting RT disruption from Nm is exponentially higher than a simple primer extension assay, there is a trade-off in the loss of exact positional information. That is, with labeled primer extension, it is possible to determine the exact position of an Nm site relative to the primer by running a high-resolution gel, as in Fig. 2 (though general knowledge of the Nm site location is also necessary, since primer extension assays are typically limited to regions of 50-200 nucleotides upstream of the RT primer). Using the RTL-P approach, the Nm site needs only to be between the RT primer and the PCR amplification region, and these two elements can be much farther apart. As above, use of genetic knockouts or unmodified RNA controls is necessary to distinguish Nm sites from other places where RNA may be otherwise modified or heavily structured and cause a similar disruption to RT reactions. Careful placement of RT primers 30 - to the suspected mRNA Nm site has also been described as a way to “map” the Nm site, but this has variable efficacy in our experience when analyzing less abundant RNAs. Another factor to consider when designing an RTL-P assay for mRNA is that while rRNA is typically consistent among experimental conditions, the abundance of mRNA transcripts may be highly variable. In fact, we recently reported that the loss of 1 Nm on Pxdn mRNA led to a 50% decrease in Pxdn transcript abundance [8]. Normalization to the amount of PCR product at normal (high) levels of dNTP accounts for this in the RTL-P approach. We have modified RTL-P to more accurately quantitate snoRNA-guided Nm modifications on an mRNA target, as described below. First, we use genetic models to perturb the presence of Nm, since inherently structured regions of RNA can also block RT at low dNTPs. For snoRNA-guided methylations, this is achieved by knockdown of FBL or a genetic knockout of the snoRNA guide. Next, we frequently use oligo-dT as an RT primer for mRNA, which is surprisingly effective for detecting Nm sites that are even several kilobases away from the polyA tail. Alternatively, a gene-specific primer (GSP) can be used but requires an analogous GSP for the housekeeping gene so that accurate quantification of the target can be calculated between genetic conditions. While the original RTL-P method used standard PCR as a readout, we use real-time quantitative PCR (RT-qPCR) to allow for even more sensitive detection of Nm between conditions. In our approach, qPCR primers are designed 50 to the Nm being evaluated. A schematic of our approach is shown in Fig. 3b, with the downstream analysis again shown in Fig. 3c.

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Materials

2.1 Cell Culture and RNA Isolation

1. Cell line of interest, human or animal tissue: Here, we use 293 T cells (ATCC® CRL-3216™). 2. Dulbecco’s modified Eagle’s medium (DMEM). 3. Fetal bovine serum (heat inactivated). 4. L-Glutamine. 5. PBS. 6. TRIzol reagent. 7. 1-Bromo-3-chloropropane. 8. Glycogen (optional). 9. Isopropanol. 10. Ethanol (75% and 100%). 11. Nanodrop or Qubit. 12. Bioanalyzer (optional). 13. RNase-free 1.5 mL polypropylene tubes. 14. RNase-free water.

2.2 Reverse Transcription and qPCR

1. PCR instrument (Bio-Rad or equivalent). 2. Real-time PCR instrument (StepOne Plus or equivalent). 3. RNase-free 0.65 mL thin-wall qPCR tubes. 4. qPCR plate and optical film. 5. SuperScript III 10 Buffer. 6. Reverse transcriptase enzyme (SuperScript III or enzyme of choice). 7. dNTPs (10 mM and 10μM). 8. 25 mM Magnesium chloride. 9. 0.1 M DTT. 10. RNaseOUT. 11. RNase-free water. 12. Oligo-dT or alternate gene-specific primer (GSP) as reverse transcription primer. 13. Target-specific qPCR primers: Examples of qPCR primers used for RTL-P can be found in Elliott et al. 2019 [8]. 14. Housekeeping (HK) gene qPCR primers. 15. Power SYBR (or alternate PCR master mix).

2.3

Software

1. StepOne Plus (or alternate qPCR software). 2. Microsoft Excel or similar.

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Methods RTL-P can be performed on a variety of RNA types from different sources including cultured cells, animal tissues, and synthetic oligos. While the method was traditionally developed to confirm Nm in rRNA [17], the instructions outlined here will focus on detection of snoRNA-guided Nm in mRNA from cultured cells. RNA integrity can be volatile but good-quality RNA is required for this assay. Ensure that all consumables are RNase free and that tools and work surfaces have been decontaminated with RNase-eliminating reagents.

3.1 Cell Cultivation and Collection

1. Subculture each 293 T cell line 1:3–1:8 into 10 cm2 culture dishes in 10–15 mL DMEM complete with 10% FBS and 2 mM L-glutamine. Grow cells until 80–90% confluent, changing medium every 2–3 days. 2. Prior to harvest, aspirate medium, wash briefly with PBS, and add 1 mL TRIzol reagent. 3. Scrape the plates using a cell scraper and transfer the TRIzol/ cell lysate into an RNase-free 1.5 mL tube using an RNase-free pipet filter tip. 4. Incubate the tube for 5 min at room temperature (RT) to allow for the dissociation of nucleoprotein complexes and proceed to RNA extraction.

3.2

RNA Extraction

1. Add 200μL 1-bromo-3-chloropropane to each tube and shake vigorously by hand for 15 s. Do not vortex. 2. Incubate for 5 min at RT. 3. Centrifuge tubes at 12,000  g at 4  C for 15 min. 4. Carefully collect the clear, upper aqueous phase (~0.5 mL) and transfer it to a new tube. It is important not to disturb the interphase and red organic phase below that contains DNA and protein that could interfere with downstream steps. 5. (Optional) Add glycogen to the aqueous phase, as an RNA carrier to maximize yield. 6. Add an equivalent volume of isopropanol (~0.5 mL) to the aqueous phase and mix. 7. Incubate at RT for 10 min. 8. Centrifuge tubes at 12,000  g at 4  C for 10 min. Precipitated RNA will appear as a white pellet at the bottom of the tube. 9. Discard the supernatant. 10. Wash pellet with 1 mL 75% ethanol. 11. Vortex briefly to release pellet from the bottom of the tube.

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12. Centrifuge tubes at 7500  g at 4  C for 5 min. 13. (Optional) Repeat steps 9–12 twice to reduce contaminates carried over from the TRIzol. 14. Discard the supernatant, being careful to remove excess ethanol surrounding the pellet by pipetting or careful vacuum aspiration. 15. Allow the pellet to air-dry for 5–10 min at RT until the edges of the pellet become translucent. Do not overdry the pellet. 16. Resuspend the RNA pellet in RNase-free water to ~100–500 ng/μL final concentration (typically 10–100μL of water, depending on the amount of starting material). 17. Heat tubes for 5–10 min at 50  C using a heat block to facilitate resuspension. 18. Vortex and/or pipet up/down to ensure resuspension. 19. Quantitate RNA using a Nanodrop or Qubit spectrophotometer. Ideally, A260/280 and A260/230 should both be >1.8. 20. (Optional) RNA can be assessed for integrity and lack of DNA contamination using an Agilent Bioanalyzer. 21. Store RNA at 80  C until ready to use. 3.3 Reverse Transcription Under High- and Low-dNTP Conditions (See Notes 1–4)

1. Ensure that each sample to be processed has 1000 ng of total RNA in 16μL or less volume. 2. Thaw RNA sample, RT primers, dNTPs, SuperScript III 10 buffer, 25 mM magnesium, and 0.1 M DTT on ice. Keep enzymes (SuperScript III and RNaseOUT) in a freezer or enzyme cooler until ready to use. 3. For each sample, label two 0.65 mL polypropylene tubes for high and low dNTPs. 4. Add 500 ng of RNA sample into each tube and adjust the volume to 8μL with RNase-free water. 5. Prepare two master mixes, one for high and low dNTPs as outlined below, in 1.5 mL RNase-free tubes. Vortex each master mix prior to use.

Master Mix 1

1 (μL)

Sample number (N)  (μL)

10 mM (high) OR 10 uM (low) dNTP

1

1N

Oligo-dT (50μM) and/or GSP RT primer 1 (2 pmol)

1N

Total volume to pipet per reaction

–

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6. Pipet 2μL of each master mix (high or low dNTP) into the respectively labeled RNA sample tubes. Vortex or pipet up and down to mix. Each tube should now contain 10μL of RNA, RT primer, and dNTPs. 7. Move tubes to the thermocycler, heat at 65  C for 5 min, and hold at 25  C. 8. Prepare “Master Mix 2” in a 1.5 mL RNase-free tube while the samples are being heated as outlined below: Master Mix 2

1 (μL)

N (μL)

10 RT buffer

2

2N

25 mM MgCl2

4

4N

0.1 M DTT

2

2N

RNase-free water

1

1N

RNaseOUT

0.5

0.5  N

SuperScript III

0.5

0.5  N

Total volume to pipet

10

–

9. Add 10μL of Master Mix 2 into each tube, pipetting up and down to mix. 10. Incubate tubes using the following program on the thermocycler: 25  C for 5 min, 50  C for 30 min, and 85  C for 5 min, and hold at 4  C. 11. Proceed to qPCR processing or freeze until ready to use. 3.4

qPCR

1. Thaw cDNA samples, Power SYBR, and target-specific qPCR primers on ice. 2. Prepare “Master Mix 3” for qPCR analysis of samples in a 1.5 mL RNase-free tube as outlined below: Master Mix 3

1 (μL)

N (μL)

Forward primer (10μM)

1.1

1.1  N

Reverse primer (10μM)

1.1

1.1  N

RNase-free water

17.8

17.8  N

2X power SYBR

22

22  N

Total volume to pipet

42

–

Volume cDNA

2

–

Total volume mix

44

–

Total volume to pipet per qPCR reaction

20

–

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3. Add 42μL of Master Mix 3 into a 0.65 mL polypropylene tube per cDNA to be assayed for each target primer set. 4. Pipet 2μL of cDNA sample into tube and vortex to thoroughly mix. 5. Add 20μL of sample + qPCR mix into one sample well on a 96-well qPCR plate in duplicate. 6. After all samples are loaded onto qPCR plate, seal with optical film. 7. Centrifuge plates at 5000  g and ensure that no bubbles are present in sample wells. 8. Load qPCR plate into thermocycler. 9. Assay the plate with the following cycle program: 95  C for 10 min (1), 95  C for 15 s, 60  C for 1 min (40), and 1 dissociation curve. 10. After assay is complete, collect Ct values for quantification. 3.5 Quantification (See Note 5)

1. Calculate the average Ct from technical duplicates for each sample, condition, and target gene. We have provided a templated Excel file for calculations (Electronic Supplemental Material, available on link.springer.com). See Fig. 4 and Table 1 for an example using summary data (or single samples). 2. Calculate the delta Ct (dCt): (a) For high-dNTP reactions, subtract the average Ct of the housekeeping (HK) gene from the average Ct of the target gene for each condition. This normalizes the results of the target to the housekeeping gene, which should have similar expression across the biological conditions (such as WT and KO) and therefore serves as a loading control for the amount of input RNA in each reaction. (b) For the low-dNTP reaction, which is the same RNA and PCR primers but under low dNTPs, we are simply interested in the product difference between the two experimental conditions (here, WT and KO): subtract each KO (or experimental) Ct from the WT (or control) Ct. The high-dNTP reactions serve to normalize for differences in the overall expression of the target gene being studied. 3. Next, calculate the delta-delta Ct (ddCt) for the high-dNTP samples, as you usually would: subtract each dCt from the WT (or control) dCt. This step provides a relative quantification of the high-dNTP PCR products produced in the two biological conditions. See Fig. 4 for a schematic of how these values are representative of the raw data. 4. Now calculate the RQ for each target sample (at both high and low dNTP), using estimated or calculated primer efficiency

RFU

qPCR Quantification of mRNA 20 -O-methylation

A,B

C

D

F

181

E

G: dCt (C-A) H: dCt (D-B)

Low dCt: E-F ddCt: G-H

Ct

Fig. 4 Schematic of RTL-P quantification (as detailed in Table 1). RFU ¼ relative fluorescence units; Ct ¼ cycle threshold. (a, b) Housekeeping (HK) genes should be essentially overlapping at high dNTP. For high-dNTP reactions, subtract the average Ct of the HK gene from the average Ct of the target gene for each condition to calculate its delta Ct (G and H, black dotted lines). Calculate the delta-delta CT (ddCT) in the high-dNTP samples by subtracting each dCT condition from the WT (blue dotted line). For the low-dNTP reactions, subtract each KO (or experimental) Ct from the WT/control (red dotted line)

(estimated ¼ 2), to the power of each sample’s (d)dCt. Our example shows less target expression in the KO condition (measured at high dNTP), yet greater product from the KO sample under low dNTP (from loss of Nm). 5. Calculate the relative RT efficiency by dividing the RQ of low-dNTP samples by the respective RQ of the same sample under high-dNTP conditions. This final step normalizes the relative RT efficiency based on the measured target gene expression. Higher RT efficiency relative to the control sample indicates less Nm; lower values indicate more Nm.

4

Notes 1. Concentrations of RNA used for reverse transcription may need to be determined empirically. Transcript abundance and processing of sample (e.g., DNase treatment) may necessitate more or less RNA input. 2. With rare or heavily modified transcripts, qPCR on cDNAs generated under low-dNTP conditions may fail. To

Low L WT Target E ¼ 33.09 E–E ¼ 0 dNTP L KO Target F ¼ 31.20 E–F ¼ 1.89 1 3.71

RQ (high ¼ 2^ddCt; low ¼ 2^dCt)

A ¼ 15.08 B ¼ 15.09 C ¼ 22.04 C–A ¼ 6.96 ¼ G G–G ¼ 0 1 D ¼ 23.08 D–B ¼ 7.98 ¼ H G–H ¼ 1.03 0.49

HK HK Target Target

ddCt

High WT dNTP KO WT KO

dCt

Average Ct

Quantification of RT efficiency for target of interest Gene

1/1 ¼ 1 3.71/0.49 ¼ 7.57

RT efficiency (low dNTP RQ/high dNTP RQ, per genotype or condition)

Table 1 Example of quantification of sample RT efficiency of experimental RNA samples, using WT and snoRNA KO samples

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troubleshoot this, more RNA may be needed for cDNA reactions. Alternatively, water may be decreased in PCR reactions and more cDNA added. 3. Different RT or qPCR primer sets may also be needed for the assay in the event that RNA structure is impeding RT too heavily for the transcript to be detected. Examples of qPCR primers used for RTL-P can be found in Elliott et al., 2019 [8]. 4. Alternative concentrations of dNTPs can also be used in the case that 1 uM dNTP is too restrictive. For instance, 1 mM of dNTP could cause enough RT pausing to visualize a difference in Nm between genetic conditions, whereas 1 uM is too restrictive. Similarly, analysis of rRNA is more efficient with even lower dNTP (200 nM), due to its abundance. 5. RTL-P is most useful for relative quantification of Nm modification for a given target mRNA under conditions that change the Nm modification (such as fibrillarin knockdown, snoRNA KO, synthetic unmodified control). Without an appropriate control, it is difficult to say whether an isolated RT efficiency (or stopping) in low dNTPs is due to the presence of Nm or some other determinant (such as complex secondary structure).

Acknowledgments This work was funded in part by NIH R01 GM135383 (to CLH) and NIH T32 HL007101 (to BAE). References 1. Baskin F, Dekker CA (1967) A rapid and specific assay for sugar methylation in ribonucleic acid. J Biol Chem 242:5447–5449 2. Wei CM, Gershowitz A, Moss B (1975) Methylated nucleotides block 50 terminus of HeLa cell messenger RNA. Cell 4:379–386. https://doi.org/10.1016/0092-8674(75) 90158-0 3. Furuichi Y, Morgan M, Shatkin AJ, Jelinek W, Salditt-Georgieff M, Darnell JE (1975) Methylated, blocked 5 termini in HeLa cell mRNA. Proc Natl Acad Sci U S A 72:1904–1908. https://doi.org/10.1073/pnas.72.5.1904 4. Dai Q, Moshitch-Moshkovitz S, Han D, Kol N, Amariglio N, Rechavi G, Dominissini D, He C (2017) Nm-seq maps 20 -O-methylation sites in human mRNA with base precision. Nat Methods 14:695–698. https://doi.org/10.1038/nmeth.4294

5. Hoernes TP, Clementi N, Faserl K, Glasner H, Breuker K, Lindner H, Hu¨ttenhofer A, Erlacher MD (2016) Nucleotide modifications within bacterial messenger RNAs regulate their translation and are able to rewire the genetic code. Nucleic Acids Res 44:852–862. https://doi.org/10.1093/nar/gkv1182 6. Choi J, Indrisiunaite G, DeMirci H, Ieong K-W, Wang J, Petrov A, Prabhakar A, Rechavi G, Dominissini D, He C, Ehrenberg M, Puglisi JD (2018) 20 -O-methylation in mRNA disrupts tRNA decoding during translation elongation. Nat Struct Mol Biol 25:208–216. https://doi.org/10.1038/ s41594-018-0030-z 7. Hoernes TP, Heimdo¨rfer D, Ko¨stner D, Faserl K, Nußbaumer F, Plangger R, Kreutz C, Lindner H, Erlacher MD (2019) Eukaryotic translation elongation is modulated by single natural nucleotide

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derivatives in the coding sequences of mRNAs. Genes (Basel) 10. https://doi.org/10.3390/ genes10020084 8. Elliott BA, Ho H-T, Ranganathan SV, Vangaveti S, Ilkayeva O, Abou Assi H, Choi AK, Agris PF, Holley CL (2019) Modification of messenger RNA by 20 -O-methylation regulates gene expression in vivo. Nat Commun 10. https://doi.org/10.1038/s41467-01911375-7 9. Yu YT, Shu MD, Steitz JA (1997) A new method for detecting sites of 2’-O-methylation in RNA molecules. RNA 3:324–331 10. Rose RE, Pazos MA, Curcio MJ, Fabris D (2016) Global Epitranscriptomics profiling of RNA post-transcriptional modifications as an effective tool for investigating the epitranscriptomics of stress response. Mol Cell Proteomics 15:932–944. https://doi.org/10.1074/mcp. M115.054718 11. Marchand V, Blanloeil-Oillo F, Helm M, Motorin Y (2016) Illumina-based RiboMethSeq approach for mapping of 2’-O-me residues in RNA. Nucleic Acids Res 44:e135. https:// doi.org/10.1093/nar/gkw547 12. Zhu Y, Pirnie SP, Carmichael GG (2017) High-throughput and site-specific

identification of 20 - O -methylation sites using ribose oxidation sequencing (RibOxi-seq). RNA 23:1303–1314. https://doi.org/10. 1261/rna.061549.117 13. Motorin Y, Marchand V (2018) Detection and analysis of RNA ribose 2’-O-methylations: challenges and solutions. Genes (Basel) 9. https://doi.org/10.3390/genes9120642 14. Motorin Y, Muller S, Behm-Ansmant I, Branlant C (2007) Identification of modified residues in RNAs by reverse transcription-based methods. Meth Enzymol 425:21–53. https:// doi.org/10.1016/S0076-6879(07)25002-5 15. Maden BE (2001) Mapping 2’-O-methyl groups in ribosomal RNA. Methods 25:374–382. https://doi.org/10.1006/ meth.2001.1250 16. Aschenbrenner J, Marx A (2016) Direct and site-specific quantification of RNA 2’-O-methylation by PCR with an engineered DNA polymerase. Nucleic Acids Res 44:3495–3502. https://doi.org/10.1093/nar/gkw200 17. Dong Z-W, Shao P, Diao L-T, Zhou H, Yu C-H, Qu L-H (2012) RTL-P: a sensitive approach for detecting sites of 2’-O-methylation in RNA molecules. Nucleic Acids Res 40: e157. https://doi.org/10.1093/nar/gks698

Chapter 12 Relative Quantification of Residue-Specific m6A RNA Methylation Using m6A-RT-QPCR Ane Olazagoitia-Garmendia and Ainara Castellanos-Rubio Abstract Technological advances in high-throughput sequencing in combination with antibody enrichment and/or induced nucleotide-specific chemical modifications have accelerated the mapping of epitranscriptomic modifications. However, site-specific detection and quantification of m6A are still technically challenging. Here, we describe a simple RT-QPCR-based approach for the relative quantification of candidate m6A regions that takes advantage of the diminished capacity of BstI enzyme to retrotranscribe m6A residues. Key words RNA methylation, m6A, Retrotranscription, QPCR

1

Introduction N6-methyladenosine (m6A) is the most common internal RNA modification in eukaryotic mRNAs and noncoding RNAs. M6A readers bind to m6A-methylated RNAs and regulate downstream effects as mRNA stability, splicing, cellular localization of RNAs, or translation efficiency, the extent of which is closely related to the levels of methylation of each RNA [1–3]. It is known that m6A modifications are dynamic and can be regulated upon different stimuli and thus the analysis of m6A methylation levels is gaining interest in the field of gene regulation [4–6]. Being involved in such diverse cellular functions, m6A methylation has rapidly been described to play key roles in different biological processes, including pathogenesis of a wide range of diseases as cancer and immune disorders [3, 7–10]. A growing interest into deciphering the mechanisms regulated by m6A modifications, together with their involvement in disease development, highlights the need of identifying m6A-methylated sites. In addition, techniques that give quantitative information of the methylation status of individual sites would be of particular interest in order to compare m6A levels between disease stages,

Mary McMahon (ed.), RNA Modifications: Methods and Protocols, Methods in Molecular Biology, vol. 2298, https://doi.org/10.1007/978-1-0716-1374-0_12, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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among different cellular processes and/or in response to different stimuli. To this aim, different techniques have been developed, such as m6A-seq and MERIP-seq (m6A-specific methylated RNA immunoprecipitation (IP) with next-generation sequencing) [11, 12], PA-m6A-seq (photo-crosslinking-assisted m6A sequencing) [13], m6A-CLIP/IP and miCLIP (m6A individual-nucleotideresolution cross-linking and immunoprecipitation) [14, 15] or SCARLET (site-specific cleavage and radioactive labeling followed by ligation-assisted extraction and thin-layer chromatography) [16], and m6A-LAIC-seq (m6A-level and isoform-characterization sequencing) [17]. All of these techniques require long and tedious protocols that are technically challenging. In addition, most of them require high-throughput sequencing and/or the use of a m6A antibodies or chemical modifications that may not be available and affordable to every laboratory. Alternative protocols based on the reduced ability of some polymerases (i.e., BstI) to retrotranscribe m6A-modified adenosines located adjacent to the priming oligo have also been described [18, 19]. When using these enzymes, the conformational freedom between the complex formed by the primer, the template, and the enzyme itself is decreased in the presence of an RNA template containing m6A residues, reducing the flexibility needed by the enzyme for successful retrotranscription (RT) [19, 20]. These enzymes have previously been used for single-nucleotide primer extension followed by polyacrylamide gel electrophoresis of the extension products to quantify the m6A methylation levels of specific sites, but again these experiments are highly time consuming [5, 18, 19]. In the protocol described herein, we take advantage of the diminished capability of BstI enzyme to retrotranscribe methylated RNAs followed by a more sensitive and accurate quantitative PCR (QPCR)-based quantification method. We describe a new, affordable, and easy-to-use method that is able to relatively quantify m6A methylation sites in RNA samples derived from human tissues or cell cultures (Fig. 1). This method is also useful to quantify relative m6A changes in response to stimuli or in different disease stages [21].

2

Materials

2.1 RNA Extraction and Quantification

1. Biological material (cells, tissue specimens, etc.) (see Note 1). 2. Nucleospin RNA extraction kit including all accompanying buffers and reagents (see Notes 2 and 3). 3. rDNAse I. 4. Beta-mercaptoethanol. 5. 70% Ethanol.

RT-QPCR Based m6A Quantification Non-methylated Processed RNA transcript

A

187

Methylated A

A m6A

5’ Primer(+)

MRT

Primer(-) Primer(-)

BstI

Primer(+)

3’ Primer(+)

MRT

Primer(-) BstI

Primer(-) Primer(+)

Fig. 1 Schematic representation of m6A-RT-QPCR. When the candidate A is not methylated (left) the difference in retrotranscription between the two enzymes (MRT and BstI) will be equal, independently of the primer used. When the candidate A is methylated (right) BstI will lose retrotranscription capability, augmenting the difference between both enzymes when using a primer (+) adjacent to the m6A residue Table 1 Primer sequences used for the TUG1 positive and HPRT negative controls Primer name

Sequence (50 –30 )

HPRTF

ACCAGTCAACAGGGGACATAA

HPRT-

CTTCGTGGGGTCCTTTTCACC

HPRT+

CCTCCTACAACAAACTTGTCTGGAATT

TUG1F

ATTCCACGACCATGGTTGTC

TUG1-

ATTCACCACCAACCACACAGCC

TUG1+

TTCCAGTGAGCCCGCTTGCTAAAAG

6. Microcentrifuge. 7. Micropipettes and filter tips (see Note 4). 8. Disposable plastic tubes (1.5 mL) (see Note 4). 9. Nanodrop. 10. Nuclease-free H2O. 2.2 Retrotranscription

1. m6A (+) and m6A (
) primers: See Table 1 for primer sequences of targets described in this protocol. 2. NEB BstI enzyme. 3. 10 ThermoPol reaction buffer pack (see Note 5). 4. Maxima RT (MRT) enzyme and accompanying buffer. 5. dNTPs. 6. Micropipettes and filter tips (see Note 4). 7. Disposable plastic tubes for master mix preparation (1.5 mL) (see Note 4). 8. PCR tubes and caps (see Note 4). 9. Mini-centrifuge for PCR tubes. 10. Thermocycler.

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QPCR Analysis

1. QPCR forward and reverse primers: See Table 1 for primer sequences of targets described in this protocol. 2. SYBR-based master mix. 3. Micropipettes and filter tips (see Note 4). 4. Plastic tubes for reagent storage and master mix preparation (1.5 mL) (see Note 4). 5. Optical plates and films. 6. Plate centrifuge. 7. Real-time thermocycler.

3 3.1

Methods Primer Design

3.1.1 Retrotranscription (RT) Primer Design

A schematic representation of the primer location for the TUG1 transcript (methylated positive control) is shown in Fig. 2a. RT primer sequences for positive and negative controls described here are shown in Table 1. 1. Design a reverse positive (+) primer that will measure the relative methylation of your candidate methylated adenosine (m6A) within a m6A motif (DR-A-CH) [22, 23]. The last nucleotide of the 30 end of primer will be adjacent to the candidate m6A (see Note 6). 2. Design a reverse negative (
) primer located close to adenosine (A) (not within a motif), which does not overlap with the (+) primer-covered region. The last nucleotide of the 30 end of primer will be adjacent to the non-methylated A (see Note 6). 3. Primer GC content should be between 40% and 60%. 4. Optimal primer length is between 20 and 30 nucleotides (see Note 7).

3.1.2 QPCR Primer Design

For QPCR primer design, it should be taken into account that the BstI enzyme has a lower retrotranscription ability than the MRT enzyme, which results in shorter cDNA products. For this reason, the closer the QPCR primers are located to the RT primers the better. A schematic representation of QPCR primer design options is shown in Fig. 2b. QPCR primer sequences for positive and negative controls described here are shown in Table 1. 1. Select a 200 bp cDNA sequence upstream of the RT primers (see Note 8). 2. Use the selected sequence for QPCR primer design with the tool of your choice (e.g., primer3plus: https://primer3plus. com) (see Note 9).

RT-QPCR Based m6A Quantification

A

189

TUG1 m6A positive control: chr22:30,970,683-30,970,787 5’GTCTAGGCTGTGTGGTTGGTGGTGAATAGGCTTCTTTTTACATGGTGCTGCCAGCCCAGCTAATTAATGGTGCACGTGGACTTTTAGCAAGCGGGCTCACTGGAA 3’ ,5’ ,5’ TTCCAGTGAGCCCGCTTGCTAAAAG

ATTCACCACCAACCACACAGCC

Primer (-)

Primer (+)

B

Fig. 2 Schematic representation of primer design for m6A-RT-QPCR. (a) RT primer design for TUG1 mRNA is shown—positive control. (b) Schematic representation of QPCR primer design. In option 1, the RT primer (
) is also used as the reverse primer for the QPCR step; in option 2 a new QPCR reverse primer, independent of the one used in RT, can be designed and used for QPCR

3. Alternatively, in order to reduce the length of the retrotranscription, RT (
) primer can be used as the reverse primer for QPCR (see Note 10). 4. Check QPCR primer pairs for cross-hybridization with other regions of the transcriptome using NCBI/Primer-BLAST (https://www.ncbi.nlm.nih.gov/tools/primer-blast/). 3.2 RNA Extraction and Quantification

1. Using the Nucleospin RNA extraction kit, add 350 μL of RA1 lysis buffer and 3.5 μL beta-mercaptoethanol to the cell pellet and vortex vigorously (see Note 11). 2. Place the violet ring column (Nucleospin filter) in a collection tube, apply the sample from step 1, and centrifuge for 1 min at 11,000  g to clear the lysate. 3. Discard the violet ring column and add 350 μL of 70% ethanol to the homogenized lysate to adjust the RNA binding conditions. Mix by pipetting up and down. 4. Load the mixed lysate to the blue ring (Nucleospin RNA) column. Centrifuge for 30 s at 11,000  g and place the column in a new collection tube. 5. Add 350 μL of the membrane desalting buffer (MDB buffer) and centrifuge at 11,000  g for 1 min to desalt the membrane. 6. On-column DNA digestion: Prepare DNase reaction mixture with 10 μL reconstituted rDNase I and 90 μL rDNase reaction buffer. Mix by flicking and apply 95 μL DNase reaction mix onto the membrane. Incubate at room temperature for 15 min.

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7. Add 200 μL of buffer RAW2 to the blue ring column. Centrifuge for 30 s at 11,000  g and place the column into a new collection tube. 8. Add 600 μL buffer RA3 to the blue ring column. Centrifuge for 30 s at 11,000  g. Discard the flow through and place the column back into the same collection tube. 9. Add 250 μL buffer RA3 to the blue ring column. Centrifuge for 30 s at 11,000  g. Discard flow through and place the column back into the same collection tube. 10. Centrifuge for 1 min at 11,000  g to completely dry the membrane. Place the column into a nuclease-free collection tube (1.5 mL). 11. Add 20–40 μL of RNase-free H2O to the membrane, incubate for 1 min at room temperature, and centrifuge at 11,000  g for 1 min (see Note 12). 12. Measure the RNA concentration with a Nanodrop using 2 μL of the eluted RNA sample. 3.3 Retrotranscription (RT)

A schematic representation of the RT reaction setup is presented in Fig. 3a. 1. Prepare the primer solutions by making a working primer solution with all the positive (+) primers and an independent solution with all the negative (
) primers. Prepare the working primer solution at 1 μM. 2. Adjust the RNA concentration (40 ng–150 ng/μL) so all the reactions will start from the same RNA amount. Prepare 10 μL for each sample (see Note 13). 3. Pipette 2 μL of each RNA into four different PCR tubes. 4. Add 2 μL of the (+) or (
) working primer mix to the corresponding tubes. 5. Make two separate RT mixes, one for each enzyme. Include an extra sample to account for pipetting errors: (a) BstI mix: 50 μM dNTPs, 0.1 U enzyme, 1 μL 10 ThermoPol buffer, up to 6 μL H2O. (b) MRT mix: 50 μM dNTPs, 0.8 U enzyme, 2 μL 5 RT buffer, up to 6 μL H2O. 6. Add 6 μL of the corresponding mix to the tubes to get a reaction volume of 10 μL. 7. Spin PCR strip. 8. Run RT reaction using the following conditions on a thermocycler: 50  C for 15 min, 85  C for 3 min, 4 1.

sample x sample y primer (+) primer (-) MM BstI MM MRT

+ + + -

+ + + -

+ + +

+ + +

+ + + -

+ + + -

+ + +

+ + +

mo tif

ctr

l

191

tes t

B

HP m 6 RT An eg.

A

TU G1 m6 Ap os.

ctr

l

RT-QPCR Based m6A Quantification

sample x Bst (+) sample x Bst (-) sample x MRT (+) sample x MRT (-) sample y Bst (+) sample y Bst (-) sample y MRT (+) sample y MRT (-)

Fig. 3 Schematic representation of the setup of the RT reaction. (a) Sample x and sample y represent two conditions (e.g., treated vs. non-treated). (b) Schematic representation of the QPCR experiment setup. TUG1 and HPRT1 are used as m6A positive and negative controls, respectively 3.4

QPCR

A schematic representation of the QPCR reaction setup is presented in Fig. 3b. 1. Dilute RT reactions two times by adding 10 μL of sterile H2O (see Note 14). 2. Prepare a 10 μM QPCR working primer solution by mixing 10 μL of 100 μM stock of each forward and reverse primer in 80 μL sterile H2O. 3. Prepare the master mix for each of the genes of interest to be analyzed by mixing 5 μL SYBR green, 0.2 μL primer mix (F and R), and 2.8 μL sterile water per reaction. Make calculations considering that samples will be run in duplicate. Add two wells for NTC and three extra samples for the pipetting errors. 4. Pipette 2 μL of the diluted RT reaction in the bottom of the wells. 5. Pipette 8 μL of the master mix (step 3) in the top of the wells. 6. Seal the plate using the optical cover. Make sure that corners are properly sealed to avoid evaporation of the samples. 7. Spin the plate in a centrifuge at 1200  g for 1 min. 8. Run QPCR reaction using the following conditions: 95  C for 30 s, 50 cycles  (95  C for 15 s, 60  C for 30 s) followed by the melting curve analysis using the following conditions: 95  C for 10 s and 65–95  C with a 0.5  C increment.

192

Ane Olazagoitia-Garmendia and Ainara Castellanos-Rubio Optimal melting curve

Non-optimal melting curve

Melt Peak

Melt Peak 120 100

-d(RFU)/dT

-d(RFU)/dT

300 200

80 60 40

100

20 0

0 65

70

75 80 85 Temperature, Celsius

90

95

65

70

75 80 85 Temperature, Celsius

90

95

Fig. 4 Melting curve analysis after QPCR. The figure shows one example of an optimal melting curve, indicating successful RT and QPCR (left panel) and one example of a nonoptimal melting curve, indicating nonspecific amplification (right panel) 3.5

Data Analysis

1. Analyze melting curves. Examples of optimal and nonoptimal melting curves are shown in Fig. 4. 2. Calculate the difference of Cts between the two enzymes using primer (+): Delta ðþÞ ¼ Ct BstI ðþÞ
Ct MRT ðþÞ: 3. Calculate the difference of Cts between the two enzymes using primer (
): Delta ð
Þ ¼ Ct BstI ð
Þ
Ct MRT ð
Þ 4. If Delta (+) > Delta (
), the candidate m6A is considered methylated. 5. If Delta (+)  Delta (
), the candidate m6A is considered non-methylated. 6. To calculate the relative m6A levels the following formula can be used: Relative m6A ¼ 2 –((Ct Bst (
) - Ct MRT (
))/(Ct Bst (+) - Ct MRT (+)), where values above 0.5 are considered positive for methylation and below 0.5 are considered as negative for methylation (see Note 15). 7. TUG1 gene primers can be used as a positive control of m6A, and HPRT gene primers can be used as a negative control of a non-methylated A (primer sequences for these controls are shown in Table 1). Relative m6A results for TUG1 and HPRT in different cell lines and tissue samples are shown in Fig. 5a, b.

RT-QPCR Based m6A Quantification

1

* ** *

0,8

**

TUG1 HPRT

0,6 0,4 0,2

HCT15

HCT116

HEK293

C

Human intestinal biopsies 1,4 1,2

*

1 0,8

**

0,6 0,4 0,2 0

Disease

Control

TUG1 HPRT

Ct Bst-Ct MRT

B

Cell lines 1,2

Relative m6A (RT -QPCR)

Relative m6A (RT -QPCR)

A

193

Bstl v3 vs MRT 12 10 8 6 4 2 0 -2 -4 -6 -8

TUG1 HPRT

primer(+)

primer(-)

Fig. 5 Quantification of m6A on candidate transcripts in biological samples using m6A-RT-QPCR. (a) Relative m6A quantification of TUG1 positive and HPRT negative controls in three different cell lines: HCT15 and HCT116 human intestinal and HEK293 human embryonic kidney cells. (b) Relative m6A quantification of TUG1 positive and HPRT negative controls in intestinal biopsies of celiac disease patients and controls. (c) Delta Ct values (Ct Bst
Ct MRT) of primer (+) and primer (
) in the TUG1 positive and HPRT1 negative controls when using BstI v3 enzyme. Values are represented as the mean and standard error from three independent experiments. *p < 0.05; **p < 0.01 based on Students t-test

4

Notes 1. RNA can be extracted from cell cultures treated with different agents or from tissue sections in which the relative methylation level of a certain motif needs to be compared (e.g., disease versus healthy tissue). 2. RNA extraction can be performed with alternative kits or methods. 3. Make sure to add ethanol to buffer RA3 prior to the first use, as stated in the kit protocol. 4. Gloves should be worn at all times and all materials should be RNase free and sterile in order to avoid RNA degradation. 5. ThermoPol buffer contains 20 mM MgSO4. Additional MgSO4 is not added to the reaction. 6. For RT reverse primer design and selection of the As (+ and
), confirm whether the gene of interest is transcribed from the sense or antisense strand of the DNA. 7. The 30 end of the primer cannot be modified as it has to be adjacent to the test A, and length can be variable toward the 50 end. 8. If no QPCR primers can be designed in the first 200 bp of the cDNA, the region can be extended up to 300 bp. However, some changes in the primer concentration or BstI amount will be needed in order to get the retrotranscription to achieve a longer cDNA. 9. When possible, design the QPCR F and R primers in different exons in order to avoid amplification of any remaining DNA.

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10. If the RT (
) primer is used as the R primer for the QPCR, it should be confirmed that it has an adequate melting temperature for QPCR. 11. When using tissue samples, tissue should be homogenized with a pellet pestle after the addition of the RA1 buffer containing beta-mercaptoethanol. If the tissue cannot be homogenized using a pellet pestle, a 20-gauge syringe can be used. 12. Higher concentrations of RNA are preferred; if the starting material is limited (

tRNAHis

G> ox> Q>

tRNAAsn

G> ox> tRNATyr Q/G> ox> tRNAAsp Q/G> ox>

Fig. 1 Analysis of Q-tRNA modifications in mouse cells. Mouse embryonic stem cells (mESCs) and mESC-differentiated cells are grown either in a standard medium containing FBS as the source of Q or in 2i medium without any source of Q. 5 μg of total RNA harvested from various conditions is subjected to APB Northern blot analysis revealing differences in the level of Q-tRNAHis and Q-tRNAAsn modifications. No separate Q- and G-bands are detected for tRNAAsp and tRNATyr. (a) Nucleic acid detection by SYBR Gold staining. (b) After hybridization and detection with each probe, the membrane is stripped and probed with a new probe. The slower migration of tRNAs is eliminated by oxidizing the ribose with periodate, producing a single, faster migrating band (ox). Data information: 2i: serum-free medium containing GSK3b and Mek 1/2 inhibitors, FBS: serum-supplemented medium, EB: embryoid bodies cultured in serum-supplemented medium, Ad: mESC-derived adipocytes cultured in serumsupplemented medium, ox: periodate-oxidized control sample. Q: Q-tRNA, G: G-tRNA, ox: periodate-oxidized control

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2. Immediately insert a gel comb without introducing any air bubbles and allow the gel to polymerize for at least 45 min at room temperature. 3. Remove the comb, gently wash the plates using tap water to remove gel debris, assemble the electrophoresis unit, and fill up the tank with pre-chilled 1 TAE (see Note 12). 4. Denature control and sample RNAs containing 2 RNA loading dye at 72  C for 3 min and store on ice until loading. 5. Wash the gel wells with a syringe to remove the urea and immediately load the samples (see Note 13). 6. Run the gel using Bio-Rad short plates at 85 V for 30 min and at 140 V for 2–3 h at 4  C or until the xylene cyanol marker reaches 2–3 cm from the bottom of the gel (see Note 14). 7. Following the electrophoresis, separate the gel from the glass plates using a spatula or similar (see Note 15). 8. Stain the nucleic acids with SYBR Gold dye staining solution in a plastic container for 15 min shaking/tumbling at room temperature. 9. Visualize the nucleic acids on a transilluminator (see Note 16). 3.3 Northern Blot and Hybridization

3.3.1 Semidry Transfer of RNAs to the Membrane

Northern blot is a valuable tool for probing individual tRNA species. Sequence-specific hybridizing probes are usually labeled with radioisotopes, such as 32P, or fluorophores to detect the RNA of interest. Although radiolabeled probes are advantageous for improving detection sensitivity and assay quality, the usage of radioactive substances should be minimized [15]. 1. Place three equally sized Whatman filter papers on the positively charged side of the semidry blotting system and moist the filter papers with 1 TAE. 2. Place a positively charged nylon membrane on the wet filter papers (see Note 17). 3. Gently put the gel onto the membrane and cover with three equally sized filter papers (see Note 18). 4. Pour 1 TAE on the gel-membrane sandwich until it gets wet and remove any air bubbles using a cylinder-shaped tool. 5. Start the transfer at 5 V/gel for 40 min at room temperature. 6. To fix RNA targets to the membrane surface, UV cross-link the membrane using a calibrated UV cross-linker with the RNA surface facing up. Expose membrane to shortwave UV light (254 nm, the 1200 mJ auto-cross-linking setting in a Stratalinker) for 1 min (see Note 19). Membrane can be stored at 20  C for up to a year.

Queuosine Detection by APB Northern 3.3.2 50 -End Labeling

225

Before proceeding, complementary oligonucleotide probes for the tRNA species of interest should be designed. Oligos can be ordered from a standard DNA synthesizing company. Oligonucleotide probes used to detect human and mouse Q-modified tRNAs (see Note 20) are listed in Subheading 2.3. T4 polynucleotide kinase is used to catalyze the reaction for the transfer and exchange of the radiolabeled terminal (γ) phosphate of ATP to the 50 -end of an oligonucleotide. 1. Experiments involving radioactivity must be conducted in a protected room after contacting a radioactivity safety officer (see Note 21). Prepare the labeling buffer in a tube and add 5 μL γ-32P-ATP (10 μCi/μL) and 1 μL T4 polynucleotide kinase. Vortex and spin down the mix and incubate at 37  C on a heat block for 30–60 min. 2. Inactivate the reaction for 5 min at 72  C. 3. Place the reaction on ice and add 100 μL dH2O. 4. Purify the probes using Micro Bio-Spin P-30 chromatography columns according to the manufacturer’s instructions. 5. Quantify the 32P incorporation using a conventional scintillation counter. 6. The probe is ready to use. Alternatively, it can be stored at 20  C for a few days considering the short half-life of 32P.

3.3.3 Hybridization and Detection

Pre-hybridization and hybridization are carried out in a glass hybridization bottle rotating in a dry hybridization oven. The radioactive signals are captured and documented on X-ray films. Alternatively, a phosphor imager could be used. 1. Pre-warm the pre-hybridization buffer at 42  C until it turns to liquid. 2. Put the membrane in a glass hybridization bottle and add preheated pre-hybridization buffer. 3. Rotate the bottle in a hybridization oven at 42  C for at least 1 h. 4. Denature the radiolabeled probe (106 cpm/mL) at 72  C for 10 min and add into the hybridization bottle (see Note 22). 5. Rotate the bottle in a hybridization oven at 42  C overnight. 6. Discard the hybridization buffer and wash the membrane two times with 2 SSC/5% SDS for 15 min each wash. 7. Wash the membrane with 1 SSC/1% SDS for another 15 min. 8. Remove the membrane from the bottle and put it into a plastic bag. Seal the bag to avoid leakage and drying of the membrane.

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9. Place the membrane into an X-ray cassette with the RNA side facing up and place an X-ray film on top in the dark (see Note 23). Expose for 2 h, 4 h, and 24 h using an automatic film processor (X-ray film developer). If necessary, expose for shorter or longer time to obtain clear signals. 3.3.4 Membrane Stripping

Membranes can be re-probed several times. Pay attention to verify the background signal before each hybridization. 1. Incubate the membrane in stripping buffer at 80  C for 2 h (see Note 24). 2. Rinse the membrane twice in 2 SSC for 15 min. The membrane is ready to use for probing and downstream analysis. The membrane can also be stored at 20  C after stripping for up to a year.

3.3.5 Quantification

If the method is performed correctly, two separate bands corresponding to G- and Q-tRNA should be seen (Figs. 1 and 2) (see Note 25). If periodate-oxidized controls are used, a distinct single, faster migrating band will appear due to oxidation of the cisdiol groups (Fig. 1) [6, 11]. Membrane images can be quantified using an image processing program such as ImageJ (Fig. 2). 1. Scan the exposed film. 2. Measure the area of the detected bands using ImageJ. 3. All signals in one column represent total RNA detected in a sample. 4. Invert the membrane (ImageJ > Edit > Invert).

color

on

ImageJ

5. In order to eliminate the background signal (blank), select an empty area on the membrane and subtract the measured signal intensity from the signal intensity detected in Q- and G-bands (ImageJ > Draw a rectangle that could cover the biggest band on the membrane > Analyze > Measure). 6. Measure the areas by dragging the rectangle on desired areas containing signals (Drag rectangle > Analyze > Measure). Repeat this until all the bands are measured. Pay attention to use the same sized rectangle when measuring all the areas on one membrane. 7. Calculate Q-tRNA modification levels by dividing Q-band signals to total Q- plus G-band signals [(Q band-blank)/ ((Q band-blank) + (G band-blank))].

Queuosine Detection by APB Northern

a q 20nM sample

227

b 1

2

3

+ 4

+ 5

+ 6 100

Q> G>

80 % of Q-tRNAHis

tRNAHis

60 40 20 0

1

2

3

4

5

6

Fig. 2 Detection and quantification of Q-tRNAHis modification in human HCT116 cells. (a) APB Northern blot analysis reveals differences in the level of Q-tRNAHis modification. (b) The modification level is quantified using ImageJ software. Data information: Cells grown in serum-free medium are indicated as samples 1, 2, and 3. Cells grown in serum-free medium supplemented with 20 nM queuine (q), which is the base of queuosine (Q), are shown as samples 4, 5, and 6. Q: Q-tRNA, G: G-tRNA

4

Notes 1. As little as 1 μg of total RNA can be used as starting material. We strongly recommend not to use more than 10 μg of total RNA. Excess RNA may cause diffuse bands. 2. It is recommended to first use concentrated acid solutions to narrow down the pH. Once the required pH is close, use a series of solution with lower strength for precision. 3. Glucose can get contaminated very easily. Therefore, it should be prepared freshly prior to use. 4. A larger amount of gel solution can be prepared and stored at 4  C for up to a month before APS and TEMED are added. One gel consists of ca. 12 mL of gel solution and the gel volume can be adapted to the dimension of the gel plates. 5. Do not autoclave the SDS solution. It will irreversibly precipitate. 6. Cells can be collected using enzymatic or mechanical methods. If using an enzymatic method, be sure to use an enzyme, such as accutase, that does not contain any Q source.

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7. Be careful when separating the aqueous phase from the organic phase to avoid any contamination. 8. Pay attention to the 260/280 and 260/230 ratios. Abnormal ratios may indicate protein, phenol/chloroform, or other chemical contaminations, which might negatively affect downstream analyses. 9. Quick protocol for sample preparation: Incubate 1–10 μg RNA in 100 mM of Tris–HCl, pH 9, for 30 min at 37  C. Reaction mix: 1–10 μg of RNA, 2 μL of 1 M Tris–HCl, and up to 20 μL of dH2O. Add 20 μL of 2 RNA loading dye to the reaction mix. Sample can be stored at 80  C. 10. We find it helpful to incubate the precipitation mix at 80  C overnight when the initial material is as little as 1 μg. 11. The experiment can be paused after adding the loading dye to the samples. 2 RNA dye containing formamide helps to avoid degradation. The samples can then be stored at 80  C before loading. 12. We use 1 mm glass plates and an electrophoresis apparatus normally used for Western blots like the Mini-PROTEAN® handcast system provided by Bio-Rad. Alternatively, cast systems with gel dimensions of 70  100  1 mm are also suitable. 13. Load the samples immediately after flushing the gel wells. To ease the loading of the samples, use long pipette tips. 14. We find that it is important to keep the chamber on an even surface for equal running of samples. If a larger separation between G- and Q-tRNA is desired, run the gel for longer time. Pay attention not to lose the target of interest. 15. When separating the gel between the glass plates, it may stick to one of the plates. To keep the gel intact, be sure that the gel is wet enough. If necessary, use additional 1 TAE and lift the gel with a help of a wet spatula or similar. Cut one edge of the gel in a way that the orientation of the samples can be recognized. 16. Do not let the gel dry while visualizing. Do not expose the gel to UV for longer time periods to avoid RNA degradation. It is useful to take a picture of the loaded gel to document the quality of the RNA and equal loading (Fig. 1a). 17. Cut one edge of the membrane in a way that the orientation of the samples can be recognized. We find it helpful to label the membrane on a side. 18. Mark the position of the wells on the membrane with a pencil prior to blotting.

Queuosine Detection by APB Northern

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19. Move the membrane 90 to increase cross-linking efficiency after the first cross-linking. As an alternative to the Stratalinker, a 254 nm UV transilluminator can be used, exposing the membrane with the RNA surface facing down against the glass surface, or bake the membrane in a vacuum oven for 1 h at 80  C. 20. tRNA sequences can be found in the Genomic tRNA database: http://gtrnadb.ucsc.edu/. 21. Wear a lab coat, gloves, googles, and a dosimeter. Survey yourself, materials, and work area for contamination before and after each experiment. Decontaminate if necessary. 22. Dilute the labeled probe with ~100 μL of 1 PBS prior to adding it into the hybridization bottle to ensure accurate pipetting. Do not pipette the probe directly onto the membrane. 23. We find it helpful to cut or mark one side of the X-ray film in a way that the order of the samples can be recognized. 24. Renewing the stripping solution after 1 h increases the stripping efficiency. 25. Using APB Northern analysis, we could detect Q-tRNA modification with probes specific to tRNAHis and tRNAAsn in human and mouse samples, whereas no separate Q- and G-bands could be detected with tRNAAsp and tRNATyr probes, which is most likely due to a secondary mannosyl modification of Q-tRNAAsp and galactosyl modification of Q-tRNATyr in these organisms [14, 16]. Recently, a method based on the charging of the secondary amine group of galQ/manQ allowed the detection of those modifications by slowing their migration in acid-denaturing gels commonly used for the tRNA charging studies [15]. References 1. McCown PJ, Ruszkowska A, Kunkler CN, Breger K, Hulewicz JP, Wang MC, Springer NA, Brown JA (2020) Naturally occurring modified ribonucleosides. Wiley Interdiscip Rev RNA:e1595. https://doi.org/10.1002/ wrna.1595 2. Boccaletto P, Machnicka MA, Purta E, Piatkowski P, Baginski B, Wirecki TK, de Crecy-Lagard V, Ross R, Limbach PA, Kotter A, Helm M, Bujnicki JM (2018) MODOMICS: a database of RNA modification pathways. 2017 update. Nucleic Acids Res 46(D1):D303–D307. https://doi.org/ 10.1093/nar/gkx1030 3. Tuorto F, Lyko F (2016) Genome recoding by tRNA modifications. Open Biol 6(12). https://doi.org/10.1098/rsob.160287

4. Harada F, Nishimura S (1972) Possible anticodon sequences of tRNA His, tRNA Asn, and tRNA Asp from Escherichia coli B. Universal presence of nucleoside Q in the first postion of the anticondons of these transfer ribonucleic acids. Biochemistry 11(2):301–308. https:// doi.org/10.1021/bi00752a024 5. Fergus C, Barnes D, Alqasem MA, Kelly VP (2015) The queuine micronutrient: charting a course from microbe to man. Nutrients 7 (4):2897–2929. https://doi.org/10.3390/ nu7042897 6. Tuorto F, Legrand C, Cirzi C, Federico G, Liebers R, Muller M, Ehrenhofer-Murray AE, Dittmar G, Grone HJ, Lyko F (2018) Queuosine-modified tRNAs confer nutritional control of protein translation. EMBO J 37

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(18). https://doi.org/10.15252/embj. 201899777 7. Helm M, Lyko F, Motorin Y (2019) Limited antibody specificity compromises epitranscriptomic analyses. Nat Commun 10(1):5669. https://doi.org/10.1038/s41467-01913684-3 8. Thuring K, Schmid K, Keller P, Helm M (2016) Analysis of RNA modifications by liquid chromatography-tandem mass spectrometry. Methods 107:48–56. https://doi.org/10. 1016/j.ymeth.2016.03.019 9. Cao X, Limbach PA (2015) Enhanced detection of post-transcriptional modifications using a mass-exclusion list strategy for RNA modification mapping by LC-MS/MS. Anal Chem 87(16):8433–8440. https://doi.org/10. 1021/acs.analchem.5b01826 10. Singhal RP, Kopper RA, Nishimura S, ShindoOkada N (1981) Modification of guanine to queuine in transfer RNAs during development and aging. Biochem Biophys Res Commun 99 (1):120–126. https://doi.org/10.1016/ 0006-291x(81)91721-6 11. Igloi GL, Kossel H (1985) Affinity electrophoresis for monitoring terminal phosphorylation and the presence of queuosine in RNA. Application of polyacrylamide containing a covalently bound boronic acid. Nucleic Acids Res 13(19):6881–6898 12. Yuan Y, Zallot R, Grove TL, Payan DJ, MartinVerstraete I, Sepic S, Balamkundu S,

Neelakandan R, Gadi VK, Liu CF, Swairjo MA, Dedon PC, Almo SC, Gerlt JA, de Crecy-Lagard V (2019) Discovery of novel bacterial queuine salvage enzymes and pathways in human pathogens. Proc Natl Acad Sci U S A 116(38):19126–19135. https://doi. org/10.1073/pnas.1909604116 13. Muller M, Legrand C, Tuorto F, Kelly VP, Atlasi Y, Lyko F, Ehrenhofer-Murray AE (2019) Queuine links translational control in eukaryotes to a micronutrient from bacteria. Nucleic Acids Res 47(7):3711–3727. https:// doi.org/10.1093/nar/gkz063 14. Zaborske JM, DuMont VL, Wallace EW, Pan T, Aquadro CF, Drummond DA (2014) A nutrient-driven tRNA modification alters translational fidelity and genome-wide protein coding across an animal genus. PLoS Biol 12 (12):e1002015. https://doi.org/10.1371/ journal.pbio.1002015 15. Zhang W, Xu R, Matuszek Z, Cai Z, Pan T (2020) Detection and quantification of glycosylated queuosine modified tRNAs by acid denaturing and APB gels. RNA. https://doi. org/10.1261/rna.075556.120 16. Kasai H, Nakanishi K, Macfarlane RD, Torgerson DF, Ohashi Z, McCloskey JA, Gross HJ, Nishimura S (1976) Letter: the structure of Q* nucleoside isolated from rabbit liver transfer ribonucleic acid. J Am Chem Soc 98 (16):5044–5046

Chapter 15 Detecting ADP-Ribosylation in RNA Deeksha Munnur and Ivan Ahel Abstract ADP-ribosylation is a widespread reversible chemical modification of macromolecular targets. Protein ADP-ribosylation has been widely studied and plays a vital role in the regulation of several biological processes. In recent years there has been increasing interest in alternative ADP-ribosylation targets such as nucleic acids—DNA and RNA. Here we report different methods to detect ADP-ribosylation of RNA substrates. Key words RNA, ADP-ribosylation, PARPs, TRPT1

1

Introduction The chemical process of transferring single or poly ADP-ribose unit onto macromolecular targets by utilizing β-nicotinamide adenine dinucleotide (NAD+) is known as ADP-ribosylation. ADP-ribosylation plays an important role in regulating a number of cellular processes such as DNA repair, cell cycle regulation, host-virus interaction, transcription, cellular stress responses, and apoptosis [1–4]. Enzymes that catalyze ADP-ribosylation are known as ADPribosyltransferases (ARTs) and belong to different protein families, namely diphtheria-toxin-like ARTs (also known as poly ADP-ribose polymerase, PARPs), cholera-toxin-like transferases (ARTCs), and sirtuins [5–8]. PARPs are the most widely studied ADP-ribosyltransferases [9–11]. ADP-ribosylation is a reversible process wherein the enzyme that removes either the poly or the mono ADP-ribose unit is termed as an ADP-ribosylhydrolase enzyme. Some of the better characterized ADP-ribosylhydrolases in humans are PARG, TARG1, MACROD1, MACROD2, ARH1, ARH2, ARH3, NUDT16, and ENPP1 [12–22]. Traditionally, ADP-ribosylation has been studied as a post translational modification process targeting proteins. There was some early evidence of DNA ADP-ribosylation by ARTC family proteins like pierisins and CARP-1 [23–25]. Pierisins were shown to

Mary McMahon (ed.), RNA Modifications: Methods and Protocols, Methods in Molecular Biology, vol. 2298, https://doi.org/10.1007/978-1-0716-1374-0_15, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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irreversibly modify DNA on guanine nucleotide [23]. However, in the recent years there has been increasing evidence of signaling by reversible DNA ADP-ribosylation. The bacterial toxin-antitoxin system, DarT-DarG, was shown to reversibly mono ADP-ribosylate DNA on thymidine residues in a sequence-specific manner [26]. DarTG system is present in many bacterial species including Mycobacterium tuberculosis, pathogenic E. coli, and many extremophilic species where it has been suggested to control dormancy/persistence by stalling replication [26, 27]. More recently, human DNA repair PARPs—PARP1, PARP2, and PARP3—were shown to modify phosphorylated DNA ends [28–31] with efficient reversal by PARG, MACROD1, MACROD2, ARH3, and TARG1 [28, 29, 32]. In addition to proteins and DNA, one of the newly characterized macromolecular ADP-ribosylation targets is RNA. PARP-like orthologous proteins called TRPT1/Tpt1/KptA from bacterial and fungal origin were shown to ADP-ribosylate RNA and DNA ends on the phosphate group at the 50 end [33, 34]. Human TRPT1 ortholog as well as human PARP10, PARP11, and PARP15 can also ADP-ribosylate phosphorylated RNA ends [34]. RNA ADP-ribosylation can be reversed (hydrolyzed) by several human, viral, and bacterial ADP-ribosylhydrolases [34]. The biological function of ADP-ribosylation of phosphorylated RNA ends is not yet known, but there has been growing interest in understanding this modification of nucleic acid in general. In this chapter, we report different ways to detect RNA ADP-ribosylation and removal of ADP-ribosylation. RNA ADP-ribosylation can be detected by gel-based assay with or without the use of radioactivity. The same techniques mentioned herein can also be used to study DNA ADP-ribosylation.

2

Materials

2.1 Denaturing Urea Polyacrylamide Gel

1. Denaturing urea PAGE gel: 20% (w/v) Acrylamide/bis 19:1, 8 M urea, and 1 TBE. 2. 10% APS. 3. TEMED. 4. Apparatus to cast mini gel of 1 mm thickness. 5. 8  8 cm mini gel cassette of 1 mm thickness. 6. Gel-running apparatus. 7. 10 TBE buffer: 1 M Tris base, 1 M boric acid, and 0.02 M EDTA (or purchase commercially).

ADP-Ribosylation of RNA Substrates

2.2 RNA ADP-Ribosylation Using Radioactivity

233

1. Single-stranded RNA (ssRNA) oligos. 5P ssRNA

50 -[Phos] GUGGCGCGGAGACUUAGAGAA-30

3P ssRNA

50 -GUGGCGCGGAGACUUAGAGAA [Phos]-30

noP ssRNA

50 -GUGGCGCGGAGACUUAGAGAA-30

2. NAD+, [32P]-800 Ci/mmol 5 mCi/mL. 3. ATP, [γ 32P]-3000 Ci/mmol 10 mCi/mL. 4. T4 Polynucleotide Kinase (PNK) 30 -phosphatase minus with buffer. 5. illustra™ MicroSpin G-25 columns. 6. Whatman™ 3 MM Chromatography paper. 7. Cling film. 8. Gel dryer (preferably with functionality to increase temperature 1  C per min). 9. BIMAX Maximum Sensitivity Film (Kodak). 10. Purified ADP-ribosyltransferase protein (such as PARP10, TRPT1, or similar): For PARP10, the catalytic domain (818–1025aa) protein sequence was cloned in pGEX-4 T1 vector, expressed in Rosetta DE3 competent cells, and induced with 0.5 mM IPTG. Cells were lysed using BugBuster (Merck) and purified using 20 mM reduced glutathione (see Note 1). For TRPT1, TRPT1 protein sequence was codon optimized with 6 His tag and cloned in pET28a vector. Plasmid was expressed in Rosetta DE3 competent cells and induced with IPTG. Cells were lysed with BugBuster and purified using Ni-NTA resin (Qiagen). 11. DNase/RNase-free water. 12. 5 Reaction buffer: 100 mM HEPES-KOH pH 7.6, 250 mM KCl, 25 mM MgCl2, and 5 mM DTT made in DNase/RNasefree water (see Note 2). 13. RNA buffer: 20 mM HEPES-KOH pH 7.6 and 50 mM KCl made in DNase/RNase-free water (see Note 2). 14. Protein dilution buffer: 25 mM HEPES pH 8.0, 150 mM NaCl, 1 mM EDTA, and 0.1 mM TCEP (see Note 2). 15. 20 mM NAD+ from Trevigen. 16. Thermoblock. 17. 20 mg/mL Proteinase K. 18. 10% w/v SDS solution. 19. Benzonase (Sigma Aldrich) (optional).

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20. Alkaline phosphatase, calf intestinal (CIP) (optional). 21. 2 TBE urea loading dye: 8 M Urea, 20 μM EDTA pH 8.0, 2 μM Tris pH 7.5, and bromophenol blue. 22. 10 TBE buffer: 1 M Tris base, 1 M boric acid, and 0.02 M EDTA (or purchase commercially). 2.3 RNA ADP-Ribosylation Using Nonradioactive Cyanine-Tagged Oligos

1. Cyanine-tagged ssRNA oligo: 5P ssRNA 3Cy3

50 -[Phos] GUGGCGCGGAGACUUAGAGAA [Cy3]-30

noP ssRNA 3Cy3

50 -GUGGCGCGGAGACUUAGAGAA [Cy3]-30

3P ssRNA 5Cy3

50 -[Cy3] GUGGCGCGGAGACUUAGAGAA [Phos]-30

noP ssRNA 5Cy3

50 -[Cy3] GUGGCGCGGAGACUUAGAGAA-30

2. Apparatus to detect cyanine3 fluorophore—in this study we used Molecular Imager PharosFX systems. 3. Purified ADP-ribosyltransferase protein (such as PARP10, TRPT1, or similar): As described in Subheading 2.2 above. 4. DNase/RNase-free water. 5. 5 Reaction buffer: 100 mM HEPES-KOH (pH 7.6), 250 mM KCl, 25 mM MgCl2, and 5 mM DTT made in DNase/RNase-free water (see Note 2). 6. RNA buffer: 20 mM HEPES-KOH (pH 7.6) and 50 mM KCl made in DNase/RNase-free water (see Note 2). 7. Protein dilution buffer: 25 mM HEPES pH 8.0, 150 mM NaCl, 1 mM EDTA, and 0.1 mM TCEP (see Note 2). 8. 20 mM NAD+ from Trevigen. 9. Thermoblock. 10. 2 TBE urea loading dye: 8 M Urea, 20 μM EDTA pH 8.0, 2 μM Tris pH 7.5, and bromophenol blue. 11. 10 TBE buffer: 1 M Tris base, 1 M boric acid, and 0.02 M EDTA or commercially from Sigma Aldrich. 2.4 Hydrolysis of RNA ADP-Ribosylation

1. 3-Aminobenzamide (3-ABA), PARP inhibitor. 2. ADP-ribosylhydrolase enzyme. 3. 2 TBE urea loading dye: 8 M Urea, 20 μM EDTA pH 8.0, 2 μM Tris pH 7.5, and bromophenol blue.

ADP-Ribosylation of RNA Substrates

3

235

Methods

3.1 Denaturing Urea Polyacrylamide Gel Preparation and Pre-Run

1. Prepare 10 mL denaturing urea gel solution to contain final concentrations of 20% (w/v) acrylamide/bis 19:1, 8 M urea, and 1 TBE. Heat solution for 10 s in a microwave until the solution is warm to aid dissolution of the urea. Add 99 μL 10% APS and 1 μL TEMED to the gel solution. Pour into 8  8 cm mini gel cassette of 1 mm thickness. Insert gel comb and allow to set for 40–60 min. 2. Rinse each well with 0.5 TBE buffer to ensure that the wells are free from urea. Pre-run gel in 0.5 TBE running buffer for 30 min at 7 W/gel to heat the gel up and remove any excess urea. Ensure that the pre-run is complete just before reaction samples are ready to load. 3. Re-rinse the wells with 0.5 TBE buffer before loading samples to run the gel.

3.2 Preparing RNA Oligo

1. Dissolve single-stranded RNA oligo in RNA buffer to a final stock concentration of 100 μM. 2. Perform radioactive labeling of non-phosphorylated RNA oligo (noP ssRNA) according to T4 PNK enzyme manufacturer’s protocol. Briefly, set up a 25 μL reaction containing 25 μM noP ssRNA, 1 T4 PNK buffer, 25 μM of [γ-32P] ATP, and 20 units of T4 PNK (30 -phosphatase minus) enzyme made in DNase/RNase-free water. Incubate reaction at 37  C for 30 min. Heat inactivate reaction by incubating at 65  C for 20 min. 3. Purify radioactively labeled oligo using an illustra™ MicroSpin G-25 column according to the manufacturer’s protocol to remove any unincorporated labeled nucleotide (see Note 3).

3.3 RNA ADP-Ribosylation Using Radioactive NAD+

1. Calculate the amount of PARP10 protein (or similar) needed to make a 10 μM stock solution using protein dilution buffer. 2. Prepare a master mix solution of reaction buffer calculated to a final 1 concentration and 50 kBq 32P NAD+. Calculate enough volume for 4 reactions plus an extra reaction to account for pipetting error. [For 5 reactions, master mix would contain 10 μL of reaction buffer and 0.27 μL 32P NAD+.]

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3. In four Eppendorf tubes, prepare a 10 μL ADP-ribosylation reaction including a negative control reaction (see Note 4). Prepare all reactions on ice as outlined below: Final concentration Volume per reaction

Stock concentration

Reagent

5

Reaction buffer

1

2 μL

9.25 MBq (calculate radioactive decay)

32

50 kBq

0.054 μL (calculate radioactive decay)

100 μM

RNA oligo

10 μM

1 μL

10 μM

Protein/ buffer

2 μM

2 μL

P NAD+

Adjust to 9 μL

Water 500 μM

+

NAD

50 μM

1 μL Total volume 10 μL

4. Add the required volume of DNase/RNase-free water to make final reaction volume of 9 μL. Then add PARP10 protein (or other ARTs) to a final concentration of 2 μM (protein dilution buffer added to negative control reactions), RNA oligo to a final concentration of 10 μM (phosphorylated or non-phosphorylated), and, lastly, an appropriate volume of master mix per reaction. 5. Just before starting the reaction add NAD+ to a final concentration of 50 μM. 6. Incubate the reaction at 37  C for 30 min. 7. Put the Eppendorf tubes back on ice. Calculate and add a volume of proteinase K and SDS required to have final reaction concentrations of 50 ng/μL and 0.15%, respectively. Incubate the reaction at 50  C for 30 min. This step degrades the PARP10 protein and thereby removes any contamination caused by automodification of protein (see Note 5). 8. Stop the reaction by adding 2 TBE urea loading dye. Calculate the appropriate volume of dye required to achieve 1 final concentration. 9. Incubate at 95  C for 3 min (see Note 6). 10. Place the reaction back on ice to avoid gradual annealing of the oligos which could lead to the formation of secondary structures. 11. Load half the volume of the reaction onto a pre-run urea polyacrylamide denaturing gel prepared in Subheading 3.1 (see Note 7).

ADP-Ribosylation of RNA Substrates

M

No protein

PARP10

No protein

noP ssRNA

PARP10

5P ssRNA

237

1

2

3

4

5

Fig. 1 RNA ADP-ribosylation using radioactive NAD+. In this assay only the reaction product is visualized. The reaction can be catalyzed by PARP10 and several other ARTs on 50 phosphorylated, but not on non-phosphorylated, RNA oligo substrate. Control reactions are indicated as no protein lanes. Lane 1 represents 32P-labeled ssRNA that is used as a size marker

12. Additionally, load radioactively labeled non-phosphorylated RNA oligo (from Subheading 3.2) as a size marker. 13. Run gel at 7 W/gel for 40–60 min or until the dye front has run off the bottom of the gel. 14. Remove the gel from the cassette and wash with 0.5 TBE buffer to get rid of any free urea. Carry out two further washes at room temperature with gentle agitating for 5–10 min each. 15. Place the gel on two sheets of 3 MM Whatman paper and cover with cling film before drying. 16. Dry the gel using a gel dryer by raising the temperature 1  C/ min until it reaches 80  C. 17. Expose the gel to high-sensitivity X-ray film (Fig. 1). 3.4 RNA ADP-Ribosylation Using Radioactively Labeled RNA Oligos

1. Calculate the appropriate amount of 32P-labeled RNA oligo needed to make a 2 μM stock solution using RNA buffer. Radioactive labeling of the non-phosphorylated RNA oligo (noP ssRNA) can be performed as described in Subheading 3.2. 2. Calculate the volume of the reagents required to achieve a final concentration of 1 reaction buffer, 0.2 μM 32P-labeled RNA oligo, and 1 μM protein, PARP10 (or other ARTs). Include a reaction with no protein to serve as negative control and marker. 3. Prepare four distinct ADP-ribosylation reactions each with a total volume of 10 μL as outline below:

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Stock concentration

Reagent

Final concentration

Volume per reaction

5

Reaction buffer

1

2 μL

2 μM

32

0.2 μM

1 μL

10 μM

Protein/buffer 1 μM

1 μL

Water

Adjust to 9 μL

5 mM

P-labeled RNA

NAD+

500 μM

1 μL Total volume 10 μL

4. Add DNase/RNase-free water to each tube to make the final volume of 9 μL. 5. Start the reaction by adding 1 μL NAD+ at a final concentration of 0.5 mM to each tube. 6. Incubate at 37  C for 30 min. 7. To tube no. 1, which is the control tube (no protein), add 2 μL DNase/RNase-free water and incubate at 50  C for a further 30 min. 8. To tube no. 2, add proteinase K and SDS to a final concentration of 50 ng/μL and 0.15%, respectively, and incubate at 50  C for 30 min. 9. To tube nos. 3 and 4 add calf intestinal phosphatase (CIP) and benzonase, respectively. Incubate at 37  C for 30 min. 10. Add 10 μL of 2 TBE urea loading dye to all reactions and carry out a final incubation at 95  C for 3 min (see Note 6). 11. Place the reactions back on ice to avoid gradual annealing of oligos and the potential formation of secondary structures. 12. Load half the reaction on a pre-run denaturing urea polyacrylamide gel prepared in Subheading 3.1 (see Note 7). Run the gel at 7 W/gel until the dye front runs off. 13. Wash gel twice with 0.5 TBE buffer for 5–10 min each. 14. Place the gel on two sheets of 3 MM Whatman paper and cover with cling film for drying. 15. Dry the gel by gradually increasing the temperature at 1  C/ min until it reaches 80  C using a gel dryer. 16. Expose the gel to high-sensitivity X-ray film (Fig. 2).

ADP-Ribosylation of RNA Substrates

B

labelled ssRNA

β-NAD+

Protein

M

Benzonase

5ʹ-32P

Phosphatase

+

P

5ʹ 32P ssRNA P Protein Proteinase K

A

239

Nicotinamide ADPr

ADPr

P

P

P

ADP-ribosylated RNA

1

2

3

4

Fig. 2 Schematic representation of the RNA ADP-ribosylation assay that allows visualization of both the substrate and the product of the reaction using radioactively labeled RNA oligos. (a) RNA ADP-ribosylation reaction mediated by ADP-ribosyltransferase (ARTs) protein in the presence of NAD+. (b) 32P-labeled RNA oligo is modified by ARTs. ADP-ribosylation leads to a mobility shift in denaturing urea gels and protection from hydrolysis by the alkaline calf intestinal phosphatase (lane 3). As further controls, reactions can be treated with proteinase K (lane 2) or benzonase (lane 4). Lane 1 represents the unmodified 32P-labeled RNA oligo that is used as both size marker and no-protein control reaction 3.5 RNA ADP-Ribosylation Using Nonradioactive Cyanine-Tagged RNA Oligos

1. Prepare Eppendorf tubes on ice for RNA ADP-ribosylation reactions of 10 μL each as outlined below: Stock concentration

Reagent

Final concentration

Volume per reaction

1 μM

1 μL

10 μM

Cy3-labeled RNA

5

Reaction buffer 1

2 μL

10 μM

Protein/buffer 2 μM

2 μL

Water

Adjust to 9 μL

5 mM

NAD

+

500 μM

1 μL Total volume 10 μL

2. Calculate the amount of Cy3-labeled RNA oligo (Subheading 2.3) required for a final concentration of 1 μM and add it to individual Eppendorf tubes.

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3. Prepare a master mix for the planned number of reactions including an additional reaction volume to account for pipetting error. Calculate the volume of reaction buffer at final 1 concentration, protein (PARP10 or other ARTs) 2 μM, and required volume of DNase/RNase-free water to make the final master mix volume (as outlined above). 4. Add the master mix to individual tubes. 5. Finally, add NAD+ at 500 μM final concentration to each tube. 6. Incubate reaction at 37  C for 30 min. 7. Add 10 μL 2 TBE urea loading dye to each tube (see Note 8). 8. Incubate reaction at 95  C for 3 min (see Note 6). 9. Place tube back on ice to avoid gradual annealing of oligos and the potential formation of secondary structures. 10. Load half the volume of each reaction on a pre-run denaturing urea polyacrylamide gel prepared in Subheading 3.1 and run at 7 W/gel for 40–60 min or until the dye front reaches the end (see Note 7). 11. Briefly wash the gel twice for 1 min each with distilled water to remove any free urea. 12. Visualize the gel using equipment that can detect cyanine fluorophore. We used Molecular Imager PharosFX systems with laser excitation for cyanine3 fluorophore at 532 nM wavelength. 3.6 Hydrolysis of RNA ADP-Ribosylation

1. Prepare RNA ADP-ribosylated substrate as mentioned in Subheadings 3.3, 3.4, or 3.5, by adding NAD+ and incubating the reaction at 37  C for 30 min. 2. Stop PARP10-mediated ADP-ribosylation process by addition of equivalent amount of PARP inhibitor such as 3-ABA (see Note 9). 3. Calculate the volume of ADP-ribosylhydrolase enzyme required to achieve 2 μM final concentration and add it to the reaction. 4. Incubate reaction at 30  C for 30 min. 5. Add TBE urea loading dye at 1 final concentration and heat reaction at 95  C for 3 min (see Note 6). 6. Load half the volume of reaction on a pre-run urea polyacrylamide gel and run at 7 W/gel (see Note 7). 7. Further process the gel as mentioned in Subheadings 3.3, 3.4, or 3.5 depending on the method used to prepare ADP-ribosylated substrate.

ADP-Ribosylation of RNA Substrates

4

241

Notes 1. pGEX-4 T1 PARP10 catalytic domain (818–1025aa) plasmid was a gift from Bernhard Lu¨scher (RWTH Aachen University) [35]. 2. All buffers prepared in DNase/RNase-free water were filter sterilized, aliquoted, and frozen at 20  C to avoid contamination by DNase or RNase that could cause degradation of RNA oligos. 3. Radioactively labeled RNA oligo can be stored at 20  C. 4. Do not add protein to the negative control reaction. Instead add the same volume of protein dilution buffer. 5. This step can be modified by adding benzonase or calf intestinal phosphatase and incubating the reaction at 37  C for 30 min to degrade nucleic acid or to remove the phosphate group, respectively. 6. After heating the reaction at 95  C, it can be frozen at 20  C to be processed later, if required. 7. The other half of the reaction can be frozen at 20  C to be processed later, if required. 8. The additional step of treating the reaction with proteinase K/SDS or benzonase or phosphatase can be performed before addition of 2 TBE urea loading dye as described earlier and if necessary. 9. ADP-ribosylation reaction can also be stopped by heat inactivation at 80  C.

Acknowledgments This work was supported by the Wellcome Trust (101794 and 210634), Biotechnology and Biological Sciences Research Council [BB/R007195/1], and Cancer Research United Kingdom [C35050/A22284]. References 1. Gibson BA, Kraus WL (2012) New insights into the molecular and cellular functions of poly(ADP-ribose) and PARPs. Nat Rev Mol Cell Biol 13:411–424 2. Vyas S, Matic I, Uchima L, Rood J, Zaja R, Hay RT, Ahel I, Chang P (2014) Family-wide analysis of poly(ADP-ribose) polymerase activity. Nat Commun 5:4426

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5. Gupte R, Liu Z, Kraus WL (2017) PARPs and ADP-ribosylation: recent advances linking molecular functions to biological outcomes. Genes Dev 31:101–126 6. Palazzo L, Mikocˇ A, Ahel I (2017) ADP-ribosylation: new facets of an ancient modification. FEBS J 284:2932–2946 7. Rack JGM, Morra R, Barkauskaite E, Kraehenbuehl R, Ariza A, Qu Y, Ortmayer M, Leidecker O, Cameron DR, Matic I et al (2015) Identification of a class of protein ADP-ribosylating sirtuins in microbial pathogens. Mol Cell 59:309–320 8. Choi J-E, Mostoslavsky R (2014) Sirtuins, metabolism, and DNA repair. Curr Opin Genet Dev 26:24–32 9. Barkauskaite E, Jankevicius G, Ahel I (2015) Structures and mechanisms of enzymes employed in the synthesis and degradation of PARP-dependent protein ADP-ribosylation. Mol Cell 58:935–946 10. Eisemann T, Pascal JM (2020) Poly (ADP-ribose) polymerase enzymes and the maintenance of genome integrity. Cell Mol Life Sci 77:19–33 11. Kim D-S, Challa S, Jones A, Kraus WL (2020) PARPs and ADP-ribosylation in RNA biology: from RNA expression and processing to protein translation and proteostasis. Genes Dev 34:302–320 12. Rack JGM, Palazzo L, Ahel I (2020) (ADP-ribosyl)hydrolases: structure, function, and biology. Genes Dev 34:263–284 13. Slade D, Dunstan MS, Barkauskaite E, Weston R, Lafite P, Dixon N, Ahel M, Leys D, Ahel I (2011) The structure and catalytic mechanism of a poly(ADP-ribose) glycohydrolase. Nature 477:616–620 14. Oka S, Kato J, Moss J (2006) Identification and characterization of a mammalian 39-kDa poly(ADP-ribose) glycohydrolase. J Biol Chem 281:705–713 15. Lin W, Ame´ J-C, Aboul-Ela N, Jacobson EL, Jacobson MK (1997) Isolation and characterization of the cDNA encoding bovine poly (ADP-ribose) glycohydrolase. J Biol Chem 272:11895–11901 16. Sharifi R, Morra R, Appel CD, Tallis M, Chioza B, Jankevicius G, Simpson MA, Matic I, Ozkan E, Golia B et al (2013) Deficiency of terminal ADP-ribose protein glycohydrolase TARG1/C6orf130 in neurodegenerative disease. EMBO J 32:1225–1237 17. Jankevicius G, Hassler M, Golia B, Rybin V, Zacharias M, Timinszky G, Ladurner AG

(2013) A family of macrodomain proteins reverses cellular mono-ADP-ribosylation. Nat Struct Mol Biol 20:508–514 18. Chen D, Vollmar M, Rossi MN, Phillips C, Kraehenbuehl R, Slade D, Mehrotra PV, von Delft F, Crosthwaite SK, Gileadi O et al (2011) Identification of macrodomain proteins as novel O-acetyl-ADP-ribose deacetylases. J Biol Chem 286:13261–13271 19. Ono T, Kasamatsu A, Oka S, Moss J (2006) The 39-kDa poly(ADP-ribose) glycohydrolase ARH3 hydrolyzes O-acetyl-ADP-ribose, a product of the Sir2 family of acetyl-histone deacetylases. Proc Natl Acad Sci U S A 103:16687–16691 20. Kato J, Zhu J, Liu C, Moss J (2007) Enhanced sensitivity to cholera toxin in ADP-ribosylarginine hydrolase-deficient mice. Mol Cell Biol 27:5534–5543 21. Palazzo L, Thomas B, Jemth A-S, Colby T, Leidecker O, Feijs KLH, Zaja R, Loseva O, Puigvert JC, Matic I et al (2015) Processing of protein ADP-ribosylation by Nudix hydrolases. Biochem J 468:293–301 22. Palazzo L, Daniels CM, Nettleship JE, Rahman N, McPherson RL, Ong S-E, Kato K, Nureki O, Leung AKL, Ahel I (2016) ENPP1 processes protein ADP-ribosylation in vitro. FEBS J 283:3371–3388 23. Takamura-Enya T, Watanabe M, Totsuka Y, Kanazawa T, Matsushima-Hibiya Y, Koyama K, Sugimura T, Wakabayashi K (2001) Mono(ADP-ribosyl)ation of 2’deoxyguanosine residue in DNA by an apoptosis-inducing protein, pierisin-1, from cabbage butterfly. Proc Natl Acad Sci U S A 98:12414–12419 24. Nakano T, Takahashi-Nakaguchi A, Yamamoto M, Watanabe M (2015) In: KochNolte F (ed) Endogenous ADP-ribosylation. Springer, Cham, pp 127–149 25. Nakano T, Matsushima-Hibiya Y, Yamamoto M, Enomoto S, Matsumoto Y, Totsuka Y, Watanabe M, Sugimura T, Wakabayashi K (2006) Purification and molecular cloning of a DNA ADP-ribosylating protein, CARP-1, from the edible clam Meretrix lamarckii. Proc Natl Acad Sci U S A 103:13652–13657 26. Jankevicius G, Ariza A, Ahel M, Ahel I (2016) The toxin-antitoxin system DarTG catalyzes reversible ADP-ribosylation of DNA. Mol Cell 64:1109–1116 27. Laware´e E, Jankevicius G, Cooper C, Ahel I, Uphoff S, Tang CM (2020) DNA ADP-ribosylation stalls replication and is

ADP-Ribosylation of RNA Substrates reversed by RecF-mediated homologous recombination and nucleotide excision repair. Cell Rep 30:1373–1384 28. Talhaoui I, Lebedeva NA, Zarkovic G, SaintPierre C, Kutuzov MM, Sukhanova MV, Matkarimov BT, Gasparutto D, Saparbaev MK, Lavrik OI et al (2016) Poly(ADP-ribose) polymerases covalently modify strand break termini in DNA fragments in vitro. Nucleic Acids Res 44:9279–9295 29. Munnur D, Ahel I (2017) Reversible monoADP-ribosylation of DNA breaks. FEBS J 284:4002–4016 30. Zarkovic G, Belousova EA, Talhaoui I, SaintPierre C, Kutuzov MM, Matkarimov BT, Biard D, Gasparutto D, Lavrik OI, Ishchenko AA (2018) Characterization of DNA ADP-ribosyltransferase activities of PARP2 and PARP3: new insights into DNA ADP-ribosylation. Nucleic Acids Res 46:2417–2431

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31. Belousova EA, Ishchenko A, Lavrik OI (2018) DNA is a new target of Parp3. Sci Rep 8:4176–4176 32. Agnew T, Munnur D, Crawford K, Palazzo L, Mikocˇ A, Ahel I (2018) MacroD1 is a promiscuous ADP-Ribosyl hydrolase localized to mitochondria. Front Microbiol 9:20–20 33. Munir A, Banerjee A, Shuman S (2018) NAD +-dependent synthesis of a 5’-phospho-ADPribosylated RNA/DNA cap by RNA 2’phosphotransferase Tpt1. Nucleic Acids Res 46:9617–9624 34. Munnur D, Bartlett E, Mikolcˇevic´ P, Kirby IT, Matthias Rack JG, Mikocˇ A, Cohen MS, Ahel I (2019) Reversible ADP-ribosylation of RNA. Nucleic Acids Res 47:5658–5669 35. Kleine H, Poreba E, Lesniewicz K, Hassa PO, Hottiger MO, Litchfield DW, Shilton BH, Lu¨scher B (2008) Substrate-assisted catalysis by PARP10 limits its activity to mono-ADPRibosylation. Mol Cell 32:57–69

Part V Mass Spectrometry- and NMR-Based Methods for RNA Modifications Analysis

Chapter 16 Detecting Internal N7-Methylguanosine mRNA Modifications by Differential Enzymatic Digestion Coupled with Mass Spectrometry Analysis Xue-Jiao You and Bi-Feng Yuan Abstract The recent discovery of reversible chemical modifications on mRNA has opened a new era of post-transcriptional gene regulation in eukaryotes. Among these modifications identified in eukaryotic mRNA, N7-methylguanosine (m7G) is unique owing to its presence in the 50 cap structure. Recently, it has been reported that m7G also exists internally in mRNA. Here, we describe a protocol of combining differential enzymatic digestion with liquid chromatography-electrospray ionization-tandem mass spectrometry (LC-ESI-MS/MS) analysis to detect internal m7G modification in mRNA. This protocol can also be used to quantify the level of m7G at both the 50 cap and internal positions of mRNA. Key words mRNA modification, N7-methylguanosine, Differential enzymatic digestion, Mass spectrometry

1

Introduction In view of the rich epigenetic regulatory layer resulting from DNA and histone modifications, reversible RNA modifications are now considered to function as another layer for biological regulation [1, 2]. To date, more than 150 structurally distinct modifications have been identified in various RNA species [3–6]. Most of these chemical modifications are found in highly abundant rRNA and tRNA, which contribute to RNA folding, structural stability, and molecular recognition of these RNA species [7, 8]. A series of modifications are also present in mRNA, such as N6-methyladenosine [9], N1-methyladenosine [10–12], 5-methylcytidine [13–15], N3-methylcytidine [16, 17], N6, 20 -O-dimethyladenosine [18], 20 -O-methylation [19], pseudouridine [20], inosine [21], N4-acetylcytidine [22], and 5-methyluridine [23]. These modifications are

Mary McMahon (ed.), RNA Modifications: Methods and Protocols, Methods in Molecular Biology, vol. 2298, https://doi.org/10.1007/978-1-0716-1374-0_16, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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reversible and dynamic, and play key roles in regulating gene expression and modulating various physiological processes [24, 25]. N7-methylguanosine (m7G) is an evolutionarily conserved type of modification that is known as a ubiquitous mRNA cap [26]. During transcription initiation, m7G is installed at the 50 cap, forming the first co-transcriptional 50 -terminal modification of nascent mRNA [26]. The 50 -m7G-cap structure is essential for the stability of transcripts and regulation of mRNA life, including transcription elongation, pre-mRNA splicing, polyadenylation, nuclear export, and translation [26]. In addition to being present as part of the cap structure, m7G is also an internal modification in tRNA [26] and rRNA [27]. However, for years it remained unknown if m7G existed internally in mRNA of mammals, which is largely attributed to the lack of an appropriate analytical method to differentiate internal m7G in mRNA from that in the 50 cap. Mass spectrometry-based analysis can enable sensitive detection of RNA modifications due to its superior capability for identification and quantification of compounds [28–35]. In this respect, we recently demonstrated the presence of internal m7G in mammalian mRNA via mass spectrometry analysis [36]. Later, He’s group confirmed our findings and performed mapping analysis of internal m7G in mammalian mRNA [37]. In this chapter, we describe a detailed protocol to detect internal m7G in mRNA by differential enzymatic digestion coupled with liquid chromatography-tandem mass spectrometry (LC-ESI-MS/MS) analysis (Fig. 1).

2

Materials All solvents and chemicals are of analytical grade, and all solutions are prepared using Milli-Q water (Milli-Q apparatus, Millipore).

2.1 Nucleoside Standards

1. 4 mM Cytidine (C) in water. 2. 4 mM Guanosine (G) in water. 3. 4 mM Adenosine (A) in water. 4. 4 mM Uridine (U) in water. 5. 4 mM N7-methylguanosine (m7G) in water.

2.2

Oligonucleotides

1. 10 μM Forward primer (27-mer DNA): 50 - GAATTAATAC GACTCACTATAGGGAGA-30 (prepare in water). 2. 10 μM Reverse primer (22-mer DNA): 50 -AGCCGTTTCTG TAATGAAGGAG-30 (prepare in water). 3. Plasmid containing DNA template (Table 1).

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Fig. 1 Schematic illustration for the use of differential enzymatic digestion to detect internal and external 50 cap m7G in mRNA. S1 nuclease allows for the release of internal m7G from mRNA but displays almost no activity in liberating m7G from the 50 cap of mRNA. Phosphodiesterase I releases both internal m7G and m7G in the 50 cap of mRNA (reprinted with permission from [36], American Chemical Society) 2.3 Chemicals and Reagents

1. Chloroform. 2. Solvent A: 2 mM ammonium bicarbonate, pH 7.0, in water. 3. Solvent B: Chromatographic grade methanol. 4. 2.5 mM dNTPs. 5. Nuclease-free water. 6. 180 units/μL S1 nuclease. 7. 0.001 units/μL Phosphodiesterase I. 8. 30 units/μL Alkaline phosphatase (CIAP). 9. 5 units/μL Ex Taq DNA polymerase. 10. 10 S1 nuclease buffer: 300 mM CH3COONa, pH 4.6, 2800 mM NaCl, 10 mM ZnSO4.

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Table 1 The sequence of the DNA template sense strand for use in the in vitro transcription reaction and the sequence of the in vitro-transcribed 50 -m7G-capped RNA (reprinted with permission from [36], American Chemical Society) Name

Sequence (50 ! 30 )

Sense strand of GAATTAATACGACTCACTATAGGGAGACAGACTAAACTGGCTGACGGAATT DNA TATGCCTCTTCCGACCATCAAGCATTTTATCCGTACTCCTGATGATGCAT template GGTTACTCACCACTGCGATCCCCGGGAAAACAGCATTCCAGGTATTAGA AGAATATCCTGATTCAGGTGAAAATATTGTTGATGCGCTGGCAGTGTTC CTGCGCCGGTTGCATTCGATTCCTGTTTGTAATTGTCCTTTTAACAGCG ATCGCGTATTTCGTCTCGCTCAGGCGCAATCACGAATGAATAACGGTTT GGTTGATGCGAGTGATTTTGATGACGAGCGTAATGGCTGGCCTGTTGAA CAAGTCTGGAAAGAAATGCATAAACTTTTGCCATTCTCACCGGATTCAGT CGTCACTCATGGTGATTTCTCACTTGATAACCTTATTTTTGACGAGGGGA AATTAATAGGTTGTATTGATGTTGGACGAGTCGGAATCGCAGACCGATAC CAGGATCTTGCCATCCTATGGAACTGCCTCGGTGAGTTTTCTCCTTCATT ACAGAAACGGCT In vitrom7G(50 )ppp(50 )GGGAGACAGACUAAACUGGCUGACGGAAUUUAUGCCUCU transcribed UCCGACCAUCAAGCAUUUUAUCCGUACUCCUGAUGAUGCAUGGUUA 50 CUCACCACUGCGAUCCCCGGGAAAACAGCAUUCCAGGUAUUAGAAGA AUAUCCUGAUUCAGGUGAAAAUAUUGUUGAUGCGCUGGCAGUGUUC m7G-capped CUGCGCCGGUUGCAUUCGAUUCCUGUUUGUAAUUGUCCUUUUAAC RNA AGCGAUCGCGUAUUUCGUCUCGCUCAGGCGCAAUCACGAAUGAAUA ACGGUUUGGUUGAUGCGAGUGAUUUUGAUGACGAGCGUAAUGGCU GGCCUGUUGAACAAGUCUGGAAAGAAAUGCAUAAACUUUUGCCAUU CUCACCGGAUUCAGUCGUCACUCAUGGUGAUUUCUCACUUGAUAA CCUUAUUUUUGACGAGGGGAAAUUAAUAGGUUGUAUUGAUGUUGG ACGAGUCGGAAUCGCAGACCGAUACCAGGAUCUUGCCAUCCUAUGG AACUGCCUCGGUGAGUUUUCUCCUUCAUUACAGAAACGGCU Highlighted in red is the T7 promoter sequence. Underlined is the first base for the in vitro transcription. “p” represents the phosphate

11. 10 Alkaline phosphatase buffer: 500 mM Tris–HCl, 10 mM MgCl2, pH 9.0. 12. 10  Ex Taq buffer. 13. mMESSAGE mMACHINE T7 kit. 14. E.Z.N.A. Cycle-Pure kit. 15. E.Z.N.A. HP Total RNA kit. 16. E.Z.N.A. Tissue RNA kit. 17. TRIzol reagent. 18. Promega PolyATtract mRNA Isolation System. 2.4 Biological Samples

1. Human embryonic kidney (HEK293T) cells. 2. Human breast cancer (MCF-7) cells. 3. Human cervical cancer (HeLa) cells.

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4. Liver tissue of a male Sprague-Dawley rat (4 weeks old). 5. Shoot tissues of plants (rice, Arabidopsis thaliana, tobacco, and cotton). 2.5

Equipment

1. AB 3200 QTRAP mass spectrometer (Applied Biosystems) with an electrospray ionization source (Turbo Ionspray). 2. Shimadzu LC-20AD HPLC with two LC-20AD pumps, a SIL-20A autosampler, a CTO-20AC thermostated column compartment, and a DGU-20A3 degasser. 3. B-500 spectrophotometer (Metash Instruments Co.). 4. Inertsil ODS-3 column (2.1  250 mm i.d., 5 μm, GL Sciences).

3

Methods

3.1 Synthesis of 50 m7G-Capped RNA

1. Perform PCR amplification to generate a 558 bp duplex DNA containing a T7 promoter that will be used as a template for in vitro transcription. The PCR reaction mixture contains 0.25 μL of Ex Taq DNA polymerase, 4 μL of dNTPs, 1 μL of forward primer, 1 μL of reverse primer, 5 μL of 10 Ex Taq buffer, 5 μL of plasmid containing the T7 promoter and DNA template (see Table 1) and 33.75 μL of nuclease-free water. PCR reactions are performed as follows: 94  C for 2 min followed by 30 cycles at 94  C for 30 s, 56  C for 30 s, and 72  C for 30 s, and finally at 72  C for 2 min. 2. Purify the PCR products using the E.Z.N.A. Cycle-Pure kit according to the manufacturer’s recommended protocol. 3. Synthesize the 50 -m7G-capped RNA by in vitro transcription using a mMESSAGE mMACHINE T7 kit according to the manufacturer’s recommended protocol. 4. Purify the transcript by chloroform extraction and isopropanol precipitation. The sequence of the resulting 50 -m7G-capped RNA is presented in Table 1.

3.2 Isolation of mRNA from Biological Samples

1. Total RNA of HEK293T, HeLa, and MCF-7 cells is extracted using the commercially available E.Z.N.A. HP Total RNA kit according to the manufacturer’s recommended protocol. 2. Total RNA of rat live tissue is extracted using an E.Z.N.A. Tissue RNA kit according to the manufacturer’s recommended protocol. 3. Total RNA of the plant shoot is extracted using TRIzol reagent according to the manufacturer’s recommended protocol. 4. mRNA of different cells and tissues can be isolated from the corresponding total RNA by three successive polyA+-based

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purification using a PolyATtract mRNA Isolation System. Purified mRNA is dissolved in nuclease-free water. 5. Determine the concentrations of the purified RNA using a B-500 spectrophotometer or similar (see Note 1). 3.3 Enzymatic Digestion of RNA 3.3.1 S1 Nuclease Digestion

1. Denature a defined amount (0.1–2 μg) of RNA (dissolved in 8.5 μL of H2O) per sample by heating at 95  C for 5 min and then chilling on ice for 2 min (see Note 2). 2. Add 1 μL of 10 S1 nuclease buffer and 0.5 μL of S1 nuclease. Incubate the mixture (10 μL) at 37  C for 1 h. 3. Add 4 μL of alkaline phosphatase buffer, 0.3 μL of alkaline phosphatase, and 25.7 μL of H2O to the mixture. 4. Continue the incubation at 37  C for an additional 30 min followed by adding 160 μL of H2O (see Note 3). The total volume of the mixture is 200 μL. 5. Extract the digested nucleosides by adding 200 μL of chloroform. Shake vigorously and let it sit for 2 min before spinning at 12,000  g for 15 min at 4  C. Remove the aqueous layer to a new tube. 6. Repeat step 5 three times to ensure complete removal of proteins (see Note 4). 7. Collect the resulting aqueous layer and dry at 37  C under vacuum. 8. Redissolve the digested nucleosides in 65 μL of water for LC-ESI-MS/MS analysis.

3.3.2 Phosphodiesterase I Digestion

1. Denature a defined amount (0.1–2 μg) of RNA (dissolved in 7 μL of H2O) per sample by heating at 95  C for 5 min and then chilling on ice for 2 min. 2. Add 1 μL of 10 alkaline phosphatase buffer and 2 μL of phosphodiesterase I. Incubate the mixture (10 μL) at 37  C for 1 h. 3. Add 4 μL of alkaline phosphatase buffer, 0.3 μL of alkaline phosphatase, and 25.7 μL of H2O to the resulting solution. 4. Continue the incubation at 37  C for an additional 30 min followed by adding 160 μL of H2O. The total volume of the mixture is 200 μL. 5. Extract the digested nucleosides by adding 200 μL of chloroform. Shake vigorously and let it sit for 2 min before spinning at 12,000  g for 15 min at 4  C. Remove the aqueous layer to a new tube. 6. Repeat step 5 three times to ensure complete removal of proteins.

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7. Collect the resulting aqueous layer and dry at 37  C under vacuum. 8. Redissolve the digested nucleosides in 65 μL of water for LC-ESI-MS/MS analysis. 3.4 Analysis of Nucleosides by LC-ESI-MS/MS

1. Analysis of nucleosides is performed on the LC-ESI-MS/MS system consisting of an AB 3200 QTRAP mass spectrometer with an electrospray ionization source and a Shimadzu LC-20AD HPLC. 2. The HPLC separation of nucleosides is performed on an Inertsil ODS-3 column with a flow rate of 0.2 mL/min at 35  C (see Note 5). 3. Solvent A and solvent B are employed as mobile phases. A gradient of 0–5 min 5% B, 5–25 min from 5% B to 80% B, 25–28 min 80% B, 28–30 min from 80% B to 5% B, and 30–40 min 5% B for the separation is used. 4. Mass spectrometric detection is performed under positive electrospray ionization mode. 5. Monitor the nucleosides by multiple reaction monitoring (MRM) mode with the mass transitions (precursor ions ! product ions) of C (244.1 ! 112.1), U (245.1 ! 113.1), A (268.1 ! 136.1), G (284.1 ! 152.1), and m7G (298.1 ! 166.1). 6. The MRM parameters of all nucleosides are optimized to achieve maximal detection sensitivity. The optimized parameters are listed in Table 2. 7. Apply the AB SCIEX Analyst 1.5 Software for data acquisition and processing.

Table 2 The MRM transitions and optimal parameters for analysis of the nucleosides by LC-ESI-MS/MS (reprinted with permission from [36], American Chemical Society) Analytes

Precursor ion

Product ion

DP (V)

C

244.1

112.1

20.0

U

245.1

113.1

A

268.1

CEP (V)

CE (V)

CXP (V)

8.0

10.0

20.0

2.5

25.0

6.0

13.0

15.0

3.0

136.1

15.0

5.0

15.0

23.0

2.0

284.1

152.1

25.0

5.0

10.0

23.0

3.0

m G

298.1

166.1

20.0

10.0

15.0

20.0

3.0

DHzR

354.1

222.1

25.0

8.0

10.0

25.0

2.5

G 7

EP (V)

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Table 3 Evaluation of the rate of S1 nuclease-mediated release of m7G from the 50 cap of an in vitrotranscribed 50 -m7G-capped RNA (reprinted with permission from [36], American Chemical Society) In vitro-transcribed 50 -m7G-capped RNA (μg)

Measured ratio of m7G (S1 nuclease/ phosphodiesterase I)

0.2 0.5 1.0

1.9  0.03% 2.0  0.02% 1.9  0.04%

3.5 Detection of Internal m7G in mRNA

Average 1.9  0.06%

1. Evaluate the performance of the two enzymatic digestion (S1 nuclease digestion and phosphodiesterase I digestion) conditions toward m7G at the 50 cap of mRNA using the in vitrotranscribed 50 -m7G-capped mRNA by LC-ESI-MS/MS analysis. The results are listed in Table 3. 2. Based on the above experiments, we derived the following formulas: Pm7G ¼ Capm7G þ Internalm7G

ð1Þ

Sm7G ¼ 1:9%  Capm7G þ Internalm7G

ð2Þ

where Pm7G, Capm7G, and Internalm7G represent the measured amount of m7G by phosphodiesterase I digestion, m7G from the 50 cap, and internal positions of mRNA, respectively. Sm7G designates the measured amount of m7G by S1 nuclease digestion; 1.9%  Capm7G and Internalm7G represent the amounts of nonspecifically digested m7G from the 50 cap and internal m7G of mRNA, respectively. 3. According to formulas (1) and (2) above, the content of the internal m7G in mRNA can be calculated with the following formula:
ð3Þ Internalm7G ¼ Sm7G  1:9%  Capm7G =98:1% 4. The content of m7G in the 50 cap of mRNA could be calculated using the following formula: Capm7G ¼ Pm7G  Internalm7G 3.6 Validation of the Analytical Method

ð4Þ

1. Construct a calibration curve of m7G by plotting the mean peak area ratio of m7G/G versus the mean molar ratio of m7G/G (ranging from 0.0002 to 0.01% and from 0.01 to 0.2% of m7G/G) based on data obtained from triplicate LC-ESI-MS/MS measurements (Table 4).

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Table 4 Linearity of m7G by LC-ESI-MS/MS (reprinted with permission from [36], American Chemical Society) Regression line Analyte

Linear range (m7G/G, %)

Slope

Intercept

m7G

0.0002–0.01

17.13

7.11  105

7

m G

3.7 Analysis of Internal m7G in mRNA from Eukaryotic Cells and Tissues

0.01–0.2

35.21

8.40  10

R2 5

0.999 0.999

1. Use the developed method to analyze internal m7G in mRNA of eukaryotic cells and tissues. Shown in Fig. 2a, b are the typical extracted-ion chromatograms and the product-ion spectra for the detection of m7G in mRNA of rice. 2. Calculate the amount of m7G in mRNA of eukaryotic cells and tissues, including cultured human cells (HeLa, HEK293T, MCF-7), rat liver, and shoot tissues of plants (rice, Arabidopsis thaliana, tobacco, and cotton). Figure 3a, b shows quantification of m7G in mRNA from different cells and tissues. All quantitative results are based on three independent sample processing and measurements.

4

Notes 1. High purity of mRNA is essential for qualitative and quantitative analysis of modifications in mRNA. Since the proportion of mRNA in total RNA is low, contamination of other types of RNA (such as tRNA and rRNA) may result in false-positive and inaccurate results. 2. Complete digestion of RNA is essential for accurate quantitative analysis of nucleosides. Heating at 95  C for 5 min and then chilling on ice for 2 min can effectively destroy the secondary structure of RNA, thereby promoting the full digestion of RNA by S1 nuclease and phosphodiesterase I. 3. Extraction of the digested RNA with chloroform may cause loss of sample when collecting the upper aqueous solution. Adding a certain volume of H2O to the digested RNA will reduce sample loss. 4. Removal of the protein in the digested nucleoside mixture before LC-ESI-MS/MS analysis can avoid blockage of the chromatographic column. 5. Salts are introduced into the samples during enzymatic digestion, which may contaminate the mass spectrometer. A valveswitch procedure should be used to cut the first few minutes of elution into the waste solution because the salts are usually eluted at the dead time.

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Fig. 2 Analysis of internal m7G in mRNA of rice sample. (a) Extracted-ion chromatograms of the m7G standard, m7G liberated from rice mRNA by digestion with S1 nuclease or phosphodiesterase I, and a blank control (without mRNA) treated with S1 nuclease or phosphodiesterase I. (b) The product-ion spectra of m7G standard, m7G released from rice mRNA by digestion with S1 nuclease or phosphodiesterase I (reprinted with permission from [36], American Chemical Society)

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Fig. 3 Quantification of m7G RNA modifications in biological samples. Quantification of internal m7G (a) and m7G in the 50 cap structure (b) of mRNA in different cells and tissues (reprinted with permission from [36], American Chemical Society)

Acknowledgments The work is supported by the National Key R&D Program of China (2017YFC0906800) and the National Natural Science Foundation of China (21672166, 21635006, 21721005).

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13. Squires JE, Patel HR, Nousch M, Sibbritt T, Humphreys DT, Parker BJ, Suter CM, Preiss T (2012) Widespread occurrence of 5-methylcytosine in human coding and non-coding RNA. Nucleic Acids Res 40:5023–5033 14. Qi CB, Jiang HP, Xiong J, Yuan BF, Feng YQ (2019) On-line trapping/capillary hydrophilic-interaction liquid chromatography/mass spectrometry for sensitive determination of RNA modifications from human blood. Chin Chem Lett 30:553–557 15. Huang W, Lan MD, Qi CB, Zheng SJ, Wei SZ, Yuan BF, Feng YQ (2016) Formation and determination of the oxidation products of 5-methylcytosine in RNA. Chem Sci 7:5495–5502 16. Xu L, Liu X, Sheng N, Oo KS, Liang J, Chionh YH, Xu J, Ye F, Gao YG, Dedon PC, Fu XY (2017) Three distinct 3-methylcytidine (m3C) methyltransferases modify tRNA and mRNA in mice and humans. J Biol Chem 292:14695–14703 17. Ma CJ, Ding JH, Ye TT, Yuan BF, Feng YQ (2019) AlkB homologue 1 demethylates N(3)methylcytidine in mRNA of mammals. ACS Chem Biol 14:1418–1425 18. Mauer J, Sindelar M, Despic V, Guez T, Hawley BR, Vasseur JJ, Rentmeister A, Gross SS, Pellizzoni L, Debart F, Goodarzi H, Jaffrey SR (2019) FTO controls reversible m(6)Am RNA methylation during snRNA biogenesis. Nat Chem Biol 15:340–347 19. Lan MD, Xiong J, You XJ, Weng XC, Zhou X, Yuan BF, Feng YQ (2018) Existence of diverse modifications in small-RNA species composed of 16-28 nucleotides. Chem Eur J 24:9949–9956 20. Schwartz S, Bernstein DA, Mumbach MR, Jovanovic M, Herbst RH, Leon-Ricardo BX, Engreitz JM, Guttman M, Satija R, Lander ES, Fink G, Regev A (2014) Transcriptome-wide mapping reveals widespread dynamic-regulated pseudouridylation of ncRNA and mRNA. Cell 159:148–162 21. Eisenberg E, Levanon EY (2018) A-to-I RNA editing - immune protector and transcriptome diversifier. Nat Rev Genet 19:473–490 22. Arango D, Sturgill D, Alhusaini N, Dillman AA, Sweet TJ, Hanson G, Hosogane M, Sinclair WR, Nanan KK, Mandler MD, Fox SD, Zengeya TT, Andresson T, Meier JL, Coller J, Oberdoerffer S (2018) Acetylation of cytidine in mRNA promotes translation efficiency. Cell 175:1872–1886

Detecting Internal N7-Methylguanosine 23. Cheng QY, Xiong J, Ma CJ, Dai Y, Ding JH, Liu FL, Yuan BF, Feng YQ (2020) Chemical tagging for sensitive determination of uridine modifications in RNA. Chem Sci 11:1878–1891 24. Frye M, Harada BT, Behm M, He C (2018) RNA modifications modulate gene expression during development. Science 361:1346–1349 25. Qi CB, Ding JH, Yuan BF, Feng YQ (2019) Analytical methods for locating modifications in nucleic acids. Chin Chem Lett 30:1618–1626 26. Ramanathan A, Robb GB, Chan SH (2016) mRNA capping: biological functions and applications. Nucleic Acids Res 44:7511–7526 27. Zorbas C, Nicolas E, Wacheul L, Huvelle E, Heurgue-Hamard V, Lafontaine DL (2015) The human 18S rRNA base methyltransferases DIMT1L and WBSCR22-TRMT112 but not rRNA modification are required for ribosome biogenesis. Mol Biol Cell 26:2080–2095 28. Feng Y, Ma CJ, Ding JH, Qi CB, Xu XJ, Yuan BF, Feng YQ (2020) Chemical labeling assisted mass spectrometry analysis for sensitive detection of cytidine dual modifications in RNA of mammals. Anal Chim Acta 1098:56–65 29. You XJ, Liu T, Ma CJ, Qi CB, Tong Y, Zhao X, Yuan BF, Feng YQ (2019) Determination of RNA hydroxylmethylation in mammals by mass spectrometry analysis. Anal Chem 91:10477–10483 30. Xiong J, Yuan BF, Feng YQ (2019) Mass spectrometry for investigating the effects of toxic

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Chapter 17 A General LC-MS-Based Method for Direct and De Novo Sequencing of RNA Mixtures Containing both Canonical and Modified Nucleotides Ning Zhang, Shundi Shi, Xiaohong Yuan, Wenhao Ni, Xuanting Wang, Barney Yoo, Tony Z. Jia, Wenjia Li, and Shenglong Zhang Abstract Mass spectrometry (MS)-based sequencing has advantages in direct sequencing of RNA, compared to cDNA-based RNA sequencing methods, as it is completely independent of enzymes and base complementarity errors in sample preparation. In addition, it allows for sequencing of different RNA modifications in a single study, rather than just one specific modification type per study. However, many technical challenges remain in de novo MS sequencing of RNA, making it difficult to MS sequence mixed RNAs or to differentiate isomeric modifications such as pseudouridine (Ψ) from uridine (U). Our recent study incorporates a two-dimensional hydrophobic end labeling strategy into MS-based sequencing (2D-HELS MS Seq) to systematically address the current challenges in MS sequencing of RNA, making it possible to directly and de novo sequence purified single RNA and mixed RNA containing both canonical and modified nucleotides. Here, we describe the method to sequence representative single-RNA and mixedRNA oligonucleotides, each with a different sequence and/or containing modified nucleotides such as Ψ and 5-methylcytosine (m5C), using 2D-HELS MS Seq. Key words Direct RNA sequencing, Two-dimensional retention time-mass ladders, Hydrophobic end-labeling strategy, RNA modification sequencing, Single-base resolution

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Introduction Mass spectrometry (MS)-based ladder sequencing is an essential tool for studying posttranslational modifications of peptides in proteomics, where the peptides are fragmentated into mass ladders to reveal the identity and location of their chemical modifications [1–3]. However, a comparable method is not available for RNA

Supplementary Information The online version of this chapter (https://doi.org/10.1007/978-1-0716-13740_17) contains supplementary material, which is available to authorized users. Mary McMahon (ed.), RNA Modifications: Methods and Protocols, Methods in Molecular Biology, vol. 2298, https://doi.org/10.1007/978-1-0716-1374-0_17, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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modifications, despite the increasing interest in the new field of epitranscriptomics. There are several major technical challenges for MS ladder sequencing of RNA. First, it is not trivial technically to generate a complete set of MS ladders needed for ladder sequencing, i.e., ladder components/ fragments corresponding to all nucleotides from the first to the last in an RNA strand. Ideally, ladder cleavage must be highly uniform with exactly one random and unbiased cut on each RNA strand [4]. RNA can be in situ fragmented, such as what MS/MS accomplishes, into mass ladders to be directly inputted into a mass spectrometer. However, there are multiple theoretically possible fragmentation sites on any given RNA strand; for example, each phosphodiester bond often results in multiple different mass ladders from an RNA strand with significant overlap with other mass ladders, which complicates downstream data analysis for sequencing. To better control ladder fragmentation, chemical and/or enzymatic approaches have been developed to generate MS ladders in advance before inputting into the mass spectrometer [5– 10]. Encouragingly, conditions of RNA acid hydrolysis, including temperatures and reaction durations, can be well controlled to ensure a single random cut in most RNA strands via balancing the yield of single-cut fragments versus the unwanted appearance of multiple-cut fragments [4]. Unfortunately, even with optimal fragmentation techniques under well-controlled conditions, the ladder sequences generated may not always be of satisfactory quality and purity, as such ladders are often mixed with undesired fragments resulting from multiple cuts on each RNA strand (which is not avoidable 100% of the time) and subsequent metal adduct formation, complicating downstream data analysis. The second challenge in MS ladder sequencing of RNA is interpretation of the acquired MS data, which is often complicated by the issues raised above, so as to identify the complete set of MS ladder components/fragments required for sequencing. Significant efforts have been made to address this challenge through developments in data analysis techniques, e.g., via developing robust computational approaches to handle acquired complex MS/MS data [11]. In addition to the mass value of each fragment, an additional parameter, retention time (tR) of each fragment resulting from the chromatography step, has recently been included in data analysis as a second dimension to assist in sequence ladder identification from complex liquid chromatography-mass spectrometry (LC-MS) data [4, 12]. The tR is unique to each RNA fragment as lower mass RNA fragments are typically eluted from the LC column earlier (i.e., a shorter tR) and longer mass fragments are eluted from the LC column later (i.e., a longer tR). An ideal set of mass ladders would be visualized as a sigmoidal curve in a two-dimensional (2D) plot of tR vs. mass [4, 12]. By generating this curve, undesired data points that would otherwise overlap with the desired sequence

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ladder fragments in traditional one-dimensional (1D) (mass only) data are easily separated out in the 2D data analysis due to differences in their tRs. Therefore, this 2D method makes the identification of fragments that contribute to the sequence ladder much more straightforward than in traditional 1D MS sequencing methods, thus enabling streamlined data analysis for de novo sequencing of RNAs and their modifications. The third challenge of MS-based RNA sequencing is differentiation of isomeric nucleotides that share an identical mass. It is impossible to differentiate pseudouridine (ψ) from its identical uridine (U) isomer by mass alone; both ψ and U share an identical mass but are structurally different. Since MS-based sequencing techniques rely on a unique mass value for identifying and locating each nucleotide in MS sequencing of RNA, structural isomers with an identical mass bring extra difficulty in MS sequencing of ψ-containing RNAs. Similarly, it is also difficult to differentiate all isomeric methylated nucleotides by mass alone. The methyl group can be varied at different positions within the nucleotide, such as in 5-methylcytosine (m5C) and 3-methylcytosine (m3C), and this variation does not introduce a mass change to these methylated nucleotides. Although MS can be used to locate each methylated nucleotide site-specifically, it is challenging for MS to differentiate them and identify which isomeric nucleotide is at the specified position. Last but not least, sequence determination of RNA mixtures with multiple distinct RNA strands is very difficult [13]; this is probably one of the most formidable challenges in MS ladder sequencing of RNA. This challenge is deeply related to previously mentioned outstanding issues in MS-based RNA sequencing—how to generate an ideal MS ladder and how to interpret complex MS data. An ideal fragmentation method used in MS sequencing results in a single cut on each RNA strand, and every RNA molecule is cleaved into two fragments: one containing the original 50 -end of the RNA, and the other containing the original 30 -end of the RNA (Fig. 1). Therefore, without the second dimension, tR, the mass of each fragment generated within the two ladders often overlaps with each other in traditional 1D MS sequencing when one ladder is measured from the 50 -end (50 -ladder) while the other is measured from the 30 -end (30 -ladder) [13], resulting in the possibility of an incorrect association of a fragment to an incorrect mass ladder and subsequent incorrect base-calling. Such overlaps resulting in incorrect base-calling become more problematic if the RNA sample is a mixture with several different RNAs, due to the generation of a large number of mass-overlapping sequence ladders, significantly complicating downstream data analysis. As such, traditional 1D MS has so far only typically been used to sequence a purified single short RNA oligonucleotide for its sequence confirmation. Even though the 2D LC-MS-based method is capable of de novo and

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Fig. 1 RNA fragmentation and MS sequencing. (a) Each cleavage of an RNA phosphodiester bond by acidmediated hydrolysis generates two fragments, one containing the original 50 -hydroxyl (OH) and a newly formed phosphate at the 30 -end (in red), and the other containing the original 30 -OH and a newly formed OH at the 50 -end (in red). (b) A schematic representation using a short oligonucleotide 50 -HO-ACGUAC-OH-30 as an example to illustrate the potential overlap of mass peaks of ladder fragments that contribute to the formation of 50 -ladder (in black) and 30 -ladders (in blue) in traditional 1D MS sequencing

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direct sequencing of RNA, this method is still limited to a singleRNA sequence, because the two ladders (50 - and 30 -ladders) still exist closely to each other in the 2D mass-tR plot. Indeed, the increasing number of overlapping mass-tR ladder fragments upon sequencing of multiple RNA sequences simultaneously results in stepwise increases in difficulty for ladder identification and sequencing [12]. These technical barriers are significant and challenging, and overcoming them requires innovation beyond incremental advances, which would ultimately result in more robust and accurate MS-based sequencing of RNA. Recently, we have introduced a novel hydrophobic end-labeling strategy into the 2D LC-MS-based sequencing (termed 2D-HELS MS Seq) [12]. Together with wellcontrolled acid degradation, this method offers a strategy to systematically address the abovementioned challenges and allows for direct and de novo sequencing of both single- and mixed-RNA sequences containing both canonical and modified nucleotides [12]. To assist the RNA community in applying 2D-HELS MS Seq for their own studies, here, we describe the detailed experimental procedures for this direct sequencing approach, including all essential and crucial steps required when applied toward direct sequencing of RNA samples. Although this approach has already been used to sequence biological tRNA samples by utilizing an advanced anchor-based algorithm [14], we aim to illustrate the sequencing process in this chapter where it is still possible to manually read out sequences of RNA samples without such advanced base-calling algorithms (Fig. 2a). Specifically, we use both a synthetic single RNA sequence and a mixture of five synthetic RNAs as a representative example to illustrate the sequencing technique. We select ψ as a representative non-mass-altered nucleotide modification and m5C as a representative mass-altered nucleotide modification to demonstrate how to sequence purified RNAs as well as RNA mixtures with different nucleotide modifications. Currently, the minimal loading amount for sequencing a purified short RNA sample less than 35 nt is ~100 pmol per run on a standard high-resolution LC-MS (40 K mass resolution). However, more material is required (up to 400 pmol per RNA sample) when additional experiments must be conducted, e.g., to distinguish isomeric base modifications that share identical masses. With improvements in sequencing algorithms and instrument sensitivity and resolution, we anticipate that our method will eventually be applicable for direct sequencing of a broader range of cellular RNAs containing both canonical and modified nucleotides [14].

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Fig. 2 Workflow for 2D-HELS MS Seq (two-dimensional Hydrophobic End-Labeling Strategy to MS-based sequencing). (a) The major steps include hydrophobic tag labeling of RNA to be sequenced, acid hydrolysis, LC-MS measurement, and sequence generation by a computational algorithm. (b) The chemical structure of a hydrophobic tag, AppCp-biotin

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Materials Prepare all solutions using nuclease-free, diethyl pyrocarbonate (DEPC)-treated water (expressed as DEPC-treated H2O unless otherwise indicated). All reagents are of analytical grade and are used as received without further purification. Use RNase-free microcentrifuge tubes and pipette tips and use RNaseZap™ to wipe RNases off surfaces of lab equipment or apparatuses to avoid possible RNA sample degradation. Stock solutions are stored long term at 20  C unless otherwise indicated, and are allowed to equilibrate to the appropriate temperatures, as indicated, immediately prior to use.

2.1 Synthetic RNA Oligonucleotides

1. Design six short synthetic RNA oligonucleotides with different lengths (19 nt, 20 nt, and 21 nt). These RNA oligonucleotides are randomly selected as representative sequences to demonstrate how to use our sequencing method. RNA #6 contains both canonical and modified nucleotides. Similarly, ψ is employed as a representative non-mass-altering modification having an identical mass to U; m5C is selected as a representative mass-altering modification to demonstrate the robustness

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of our approach. The following RNA oligonucleotides are obtained from IDT (Integrated DNA Technologies) and used without further purification: RNA #1: 50 -HO-CGCAUCUGACUGACCAAAA-OH-30 . RNA #2: 50 -HO-AUAGCCCAGUCAGUCUACGC-OH-30 . RNA #3: 50 -HO-AAACCGUUACCAUUACUGAG-OH-30 . RNA #4: 50 -HO-GCGUACAUCUUCCCCUUUAU-OH-30 . RNA #5: 50 -HO-GCGGAUUUAGCUCAGUUGGGA-OH-30 . RNA #6: 50 -HO-AAACCGUψACCAUUAm5CUGAG-OH-30 . 2. Dissolve each synthetic RNA in nuclease-free, DEPC-treated water to obtain respective RNA stock solutions with a concentration of 100 μM (based on the amount provided by IDT). Store at 20  C. Thaw the reagents in a water bath at room temperature and mix well by vortexing and centrifuging before use. 2.2 Labeling the 30 -End of RNA

1. 100 μM Biotinylated cytidine bisphosphate (pCp-biotin, TriLink Bio Technologies) (used for the two-step 30 -end labeling protocol): Add 1.3 mL DEPC-treated H2O to 0.1 mg pCp-biotin and mix well by vortexing and centrifuging. Store at 20  C. 2. 150 μM Adenosine-50 -50 -diphosphate-{50 -(cytidine-20 -Omethyl-30 -phosphate-TEG)}-biotin (AppCp-biotin (ChemGenes)) (Fig. 2b) (used for the one-step 30 -end labeling protocol): Add 2.7 mL DEPC-treated H2O to 0.5 mg AppCp-biotin and mix well by vortexing and centrifuging. Store at 20  C. 3. 1 mM ATP (Sigma-Aldrich). 4. 50 μM Mth RNA ligase and 10 adenylation buffer (SigmaAldrich). 5. DMSO (anhydrous dimethyl sulfoxide, 99.9%) (SigmaAldrich). 6. T4 RNA ligase 1 (10 units/μL) with 10 ligation buffer (Sigma-Aldrich). 7. Zymo Research Oligo Clean & Concentrator (Zymo Research).

2.3 Biotin/ Streptavidin Capture and Release

1. Streptavidin beads (Thermo Fisher Scientific) (10 mg/mL, ~7–10  109 beads/mL) in PBS buffer, pH 7.4, 0.01% Tween™ 20, and 0.09% sodium azide. Store at 4  C.

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2. Binding and washing (B&W) buffer (2): 10 mM Tris–HCl, pH 7.5, 1 mM EDTA, 2 M NaCl. Add 0.5 mL 1 M Tris–HCl buffer to 49.4 mL DEPC-treated H2O. Add 0.1 mL 0.5 M EDTA. Add 5.844 g NaCl and mix well by vortexing. Dilute 2 B&W buffer to 1 B&W buffer by adding 25 mL 2 B&W buffer into 25 mL DEPC-treated H2O. Store at 4  C. 3. Solution A: DEPC-treated 0.1 M NaOH and DEPC-treated 0.05 M NaCl. Weigh 0.2 g NaOH and 0.15 g NaCl, add to 50 mL DEPC-treated H2O, and mix well by vortexing. Store at 4  C. 4. Solution B: DEPC-treated 0.1 M NaCl. Weigh 0.3 g NaCl, add to 50 mL DEPC-treated H2O, and mix well by vortexing. Store at 4  C. 5. 95% Formamide containing 10 mM EDTA, pH 8.2. Store at 4 oC. 6. Formic acid (98–100%). 2.4

CMC Conversion

1. CMC (N-cyclohexyl-N0 -(2-morpholinoethyl)-carbodiimide metho-p-toluenesulfonate (Sigma-Aldrich)): Weigh 0.0141 g CMC in a 1.5 mL RNase-free microcentrifuge tube. Store at 20  C. 2. Urea: Weigh 0.07 g urea in a 1.5 mL RNase-free microcentrifuge tube. Store at 4  C. 3. Bicine buffer: 1 M Bicine, pH 8.3. Weigh 1.6317 g bicine in a 15 mL RNase-free centrifuge tube and add 8 mL DEPCtreated H2O. Adjust solution to pH 8.3 with 10 N NaOH. Make up to 10 mL total volume with DEPC-treated H2O. Store at 4  C. 4. Sodium acetate (NaOAc) solution: 1.5 M NaOAc, pH 5.6. Add 500 μL 3 M NaOAc to 499 μL DEPC-treated H2O. Then add 1 μL 0.5 M EDTA and mix well by vortexing. Store at 4  C. 5. Sodium bicarbonate (Na2CO3) buffer: 0.1 M Na2CO3, pH 10.4. Weigh 1.992 g Na2CO3 and 8.086 g sodium carbonate (anhydrous) in a 15 mL RNase-free centrifuge tube and add 8 mL DEPC-treated H2O. Make up to 10 mL total volume with DEPC-treated H2O. Store at 4  C.

2.5 LC-MS Elution Buffers

1. Mobile phase A: 25 mM Hexafluoro-2-propanol (HFIP) with 10 mM diisopropylamine (DIPA) in LC-MS-grade water. Add 2.6 mL HFIP into 996 mL LC-MS-grade water and mix well by shaking. Add 1.4 mL DIPA (1.0 g) and mix well. Store at room temperature. 2. Mobile phase B: LC-MS-grade methanol.

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Methods Perform all experimental procedures at room temperature unless otherwise specified.

3.1 RNA 30 -End Labeling with Biotin (See Note 1)

1. Add 1 μL 10 adenylation reaction buffer, 1 μL 1 mM ATP, 1 μL 100 μM pCp-biotin, 1 μL 50 μM Mth RNA ligase, and 6 μL DEPC-treated H2O (total volume of 10 μL) in an RNase-free, thin-walled 0.2 mL PCR tube. Incubate the reaction in a PCR machine (GeneAmp™ PCR System 9700) at 65  C for 1 h and inactivate the enzyme by incubation at 85  C for 5 min (see Note 2). 2. Conduct the ligation step by adding the 10 μL reaction solution from the previous step to 3 μL 10 ligation buffer, 1.5 μL 100 μM RNA sample to be sequenced (for example, RNA #1), 3 μL anhydrous DMSO to reach 10% (v/v), 1 μL T4 RNA ligase (10 units), and 11.5 μL DEPC-treated H2O (total volume of 30 μL). Add the reaction components at room temperature due to the high freezing point of DMSO (18.45  C). Incubate the reaction in a PCR machine overnight (~16 h) at 16  C. 3. Quench and purify the reaction by column purification to remove enzymes and free pCp-biotin using Oligo Clean & Concentrator according to the manufacturer’s instructions. To elute the RNA, add 15 μL DEPC-treated H2O to the column and centrifuge at 10,000  g for 30 s. Store at 20  C. 4. Replace pCp-biotin with AppCp-biotin (see Note 3) by performing a one-step ligation reaction containing 2 μL 150 μM AppCp-biotin, 3 μL 10 ligase reaction buffer, 1.5 μL 100 μM RNA sample to be sequenced, 3 μL anhydrous DMSO (to reach 10% (v/v)), 1 μL T4 RNA ligase (10 units), and 19.5 μL DEPC-treated H2O (total volume of 30 μL). Incubate the reaction overnight (~16 h) at 16  C. Perform column purification as described above to elute the 30 -biotinylated RNA sample with 15 μL DEPC-treated H2O in a 1.5 mL RNase-free microcentrifuge tube.

3.2 Streptavidin Beads for Physical Separation of Biotinylated RNA (See Note 4)

1. Activate streptavidin beads by adding 200 μL 1 B&W buffer to 200 μL streptavidin beads. Vortex this solution for 30 s, place it on a magnetic stand for 2 min, and then discard the supernatant. 2. Wash the beads twice with 200 μL Solution A and once in 200 μL Solution B. For each wash step, vortex the solution for 30 s, place it on a magnetic stand for 2 min, and then discard the supernatant. Finally, after all wash steps, add 100 μL 2 B&W buffer to the washed beads.

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3. Add 1 B&W buffer to the biotinylated RNA sample to a final volume of 100 μL. Then add this solution to the washed beads stored in 100 μL 2 B&W buffer. Incubate for 30 min at room temperature on a rocking platform shaker at 300 rpm. Place the tube on a magnetic stand for 2–3 min and discard the supernatant. 4. Wash the biotin-coated beads three times in 1 B&W buffer and measure the RNA concentration in the supernatant during each wash step by Nanodrop for recovery analysis to confirm that the biotinylated RNAs remain on the beads (see Note 5). 5. Incubate the beads with 95% formamide containing 10 mM EDTA, pH 8.2, in a PCR machine at 65  C for 5 min. Put the tube on the magnetic stand for 2 min and collect the supernatant using a pipette, carefully avoiding the beads. The supernatant contains the biotinylated RNAs released from the streptavidin beads. Measure the final RNA concentration in the supernatant using a Nanodrop. 3.3 Generation of MS Sequence Ladders by Controlled Acid Degradation of RNA

1. Divide the collected biotinylated RNA sample into three equal aliquots in RNase-free, thin-walled 0.2 mL PCR tubes. For instance, divide an RNA sample with a volume of 15 μL into 3  5 μL aliquots. 2. Add an equal volume of formic acid (98–100%) to achieve 50% (v/v) formic acid in each reaction tube (see Note 6). 3. Incubate the reaction at 40  C in a PCR machine, with one reaction for 2 min, one for 5 min, and one for 15 min. 4. Immediately freeze the sample on dry ice after each specified time interval to quench the acid degradation reaction. Use a centrifugal vacuum concentrator to dry the sample. The sample is typically completely dry within 30 min. 5. Resuspend each dried sample in 20 μL DEPC-treated H2O and combine all the samples in a LC-MS sample vial for LC-MS measurement.

3.4 Sequencing a Mixed RNA Sample (See Note 7)

1. A mixture of five different RNA sequences (RNA #1 to #5) are used here as an example to demonstrate the experimental procedures. Mix 15 μL 10 ligase reaction buffer, 1.5 μL of each RNA strand (100 μM stock of RNA #1 to #5, respectively, for a total volume of 7.5 μL), 10 μL 150 μM AppCp-biotin (one-step protocol), 15 μL anhydrous DMSO, 5 μL T4 RNA ligase (10 units/μL), and 97.5 μL DEPC-treated H2O to produce a reaction solution with a total volume of 150 μL in a 1.5 mL RNase-free microcentrifuge tube. Distribute the reaction solution into five equal-volume aliquots; each microcentrifuge tube now contains 30 μL reaction solution.

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2. Incubate the reaction overnight (~16 h) at 16  C as described in Subheading 3.1, step 2. Conduct column purification according to the procedure described above in Subheading 3.1, step 3, using Oligo Clean & Concentrator. A mixed sample of 30 -biotinylated RNA sequences (RNA #1 to #5) should be eluted with 15 μL DEPC-treated H2O in a 1.5 mL RNase-free microcentrifuge tube. 3. Combine the purified mixture samples from each of the five tubes into one 1.5 mL RNase-free microcentrifuge tube. Perform formic acid degradation (50% (v/v)) according to the procedure described above in Subheading 3.3 to generate MS ladders for sequencing. 3.5 CMC Conversion for Identifying and Locating Pseudouridine (See Notes 8 and 9)

1. Add 80 μL DEPC-treated H2O to a 1.5 mL RNase-free microcentrifuge tube containing 0.0141 g CMC and 0.07 g urea. Then add 10 μL of the RNA (100 μM) to be sequenced, 8 μL bicine buffer, and 1.28 μL 0.5 M EDTA. Bring the total reaction volume to 160 μL by adding 60.72 μL DEPC-treated H2O. The final concentrations of CMC, urea, EDTA, and bicine (pH 8.3) are 0.17 M, 7 M, 4 mM, and 50 mM, respectively [15]. 2. Divide the 160 μL reaction solution into four equal aliquots of 40 μL each in RNase-free, thin-walled 0.2 mL PCR tubes and incubate in a PCR machine at 37  C for 20 min. The maximum reaction volume is 50 μL per tube based on the PCR machine used in this procedure. 3. Add 10 μL sodium acetate solution to quench each reaction. 4. Perform column purification with four parallel spin columns provided by Oligo Clean & Concentrator to remove excessive reactants according to the procedure as described above in Subheading 3.1, step 3. 5. Transfer the purified product to four RNase-free, thin-walled 0.2 mL PCR tubes. To each 15 μL purified product, add 20 μL 0.1 M Na2CO3 buffer (pH 10.4) and make up the volume to 40 μL with 5 μL DEPC-treated H2O. Incubate these four reaction tubes in a PCR machine at 37  C for 2 h. 6. Use four parallel spin columns provided by Oligo Clean & Concentrator to purify the reaction products. The CMC-ψ-converted product should be eluted with 15 μL DEPC-treated H2O in a 1.5 mL RNase-free microcentrifuge tube. 7. Transfer the purified CMC-ψ-converted sample to four RNasefree, thin-walled 0.2 mL PCR tubes. Add an equal volume of formic acid to achieve 50% (v/v) formic acid in each reaction tube. Perform acid degradation according to the procedures as described above in Subheading 3.3 to generate MS ladders for sequencing.

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Fig. 3 2D mass-tR plots of sequencing of representative RNA samples. (a) Sequencing of RNA #1 (19 nt). The 30 -end is biotin-labeled during sample preparation before acid degradation. All of the 30 -ladder fragments are well

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1. Transfer the RNA samples, stored in DEPC-treated H2O prior to LC-MS analysis, to a conical bottom micro-insert (250 μL) in a 2 mL glass HPLC sample vial for analysis. The maximum injection volume for each sample is 20 μL containing 100–400 pmol RNA.

3.6 LC-MS Measurement and Analysis of RNA Samples

2. Use the following LC conditions: a column temperature of 35  C and flow rate of 0.3 mL/min as well as a linear gradient from 2% to 20% mobile phase B over 15 min followed by a 2-min wash step with 90% mobile phase B (see Note 10). 3. Set the MS analysis for data recording with the following settings: negative ion mode; range, 350–3200 m/z; scan rate, 2 spectra/s; drying gas flow, 17 L/min; drying gas temperature, 250  C; nebulizer pressure, 30 psig; capillary voltage, 3500 V; and fragmentor voltage, 365 V (see Note 11). 4. Extract data files with MassHunter acquisition software provided by Agilent Technologies. Use the molecular feature extraction (MFE) algorithm to export compound information to an Excel spreadsheet file, which includes mass, retention time, volume (the MFE abundance for the respective ion species), and quality score. The MFE settings are as follows: “centroid data format, small molecules (chromatographic), peak with height  100, up to a maximum of 1000, quality score  50” (see Note 12).

ä Fig. 3 (continued) separated from the unlabeled 50 -ladder fragments and other undesired fragments in the 2D plot due to a systematic increase in their tRs. The sequences are automatically generated by an anchor-based computational algorithm. (b) Sequencing of a mixture of RNA containing five different RNA sequences (RNAs #1–#5). A biotin tag is used to label each RNA at the 30 -end, and tRs of each RNA ladder are normalized to begin at 7-min intervals for ease of visualization. All base-calls are performed manually by calculating the mass differences of two adjacent ladder components and matching them with the theoretical mass differences in the RNA nucleotide and modification database. With base-by-base base-calling, all sequences of the five RNAs are correctly read out. (c) Sequencing of RNA #6, which contains one ψ. The increase in mass and hydrophobicity caused by conversion of the ψ to the CMC-ψ adduct (ψ*) results in a systematic mass-tR shift on all ψ*-containing ladder fragments beginning at the ψ position. This site-specific shift indicates that a ψ is at position 8 in the RNA sequence. The other modification, m5C, can be simultaneously identified and located at position 16 based on its unique mass. The sequences are acquired by an anchor-based computational algorithm. All three 2D plots are reconstructed by OriginLab based on sequences read out by the anchor-based algorithm or manual calculation

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3.7 Generate RNA Sequence by an Anchor-Based Computational Algorithm (See Note 13)

1. Use a slightly revised version of a previously published anchorbased algorithm [14] to process the MFE files of RNA #1 and CMC-converted RNA #6, respectively. Reconstruct 2D masstR plots for better visualization for each sequence as shown in Fig. 3a and c using OriginLab, based on the sequence read out by the algorithm (Electronic Supplementary Material, Tables S1–S4). The observed mass, tR, volume, and quality score for each ladder fragment are reported in the MFE file obtained in Subheading 3.6, step 4. Related MFE data and a revised version of anchor-based algorithm (including both the Web-based sequencing application and the source code) are available upon request and are also uploaded to GitHub (https://github.com/directRNASeq/anchorseqapp). 2. Manually calculate the mass differences between the two adjacent ladder components for base-calling to confirm the order of each nucleotide in each algorithm-reported sequence. The structures of RNA modifications can be found in RNA modification databases [4], and their corresponding theoretical masses are obtained by ChemDraw. Calculate the PPM (parts per million) mass difference to compare the observed mass to the theoretical mass for a specific ladder component, and a value less than 10 PPM is considered a good match for basecalling [4, 12] (see Note 14). Manually verify each nucleotide in each RNA sequence using base-by-base manual calculation.

3.8 Manually Reading Sequences in an RNA Sample Mixture (Fig. 3b) (See Note 15)

4

Perform all base-calling procedures manually as described above in Subheading 3.7, step 2, and match well with the theoretical bases in the RNA nucleotide and modification database [4]. The matched bases with a mass PPM “Actions” > “Copy Calibration Level” in the method adjustment window. This will activate a script which transfers the respective peak areas of the unlabeled calibration compound to each isotopologue which is in the respective compound group (also see Table 4). From this timepoint on it is crucial to only quantify the batch if needed. Activating “Analyze batch” results in reversing of the used script to copy calibration curves to each isotopologue.

Acknowledgments This study was funded through the Deutsche Forschungsgemeinschaft (KE1943/3-1, KE1943/4-1–SPP1784, and ProjectID 325871075–SFB 1309). We are grateful to Prof. Peter Dedon, Prof. Mark Helm, and Prof. Thomas Carell for generous donation of synthetic standards of modified nucleosides.

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References 1. Heiss M, Kellner S (2016) Detection of nucleic acid modifications by chemical reagents. RNA Biol 0. https://doi.org/10.1080/15476286. 2016.1261788 2. Motorin Y, Helm M (2019) Methods for RNA modification mapping using deep sequencing: established and new emerging technologies. Genes (Basel) 10(1). https://doi.org/10. 3390/genes10010035 3. Helm M, Lyko F, Motorin Y (2019) Limited antibody specificity compromises epitranscriptomic analyses. Nat Commun 10(1):5669. https://doi.org/10.1038/s41467-01913684-3 4. Brandmayr C, Wagner M, Bruckl T, Globisch D, Pearson D, Kneuttinger AC, Reiter V, Hienzsch A, Koch S, Thoma I, Thumbs P, Michalakis S, Muller M, Biel M, Carell T (2012) Isotope-based analysis of modified tRNA nucleosides correlates modification density with translational efficiency. Angew Chem Int Ed Engl 51(44):11162–11165. https://doi.org/10.1002/anie.201203769 5. Kellner S, Ochel A, Thuring K, Spenkuch F, Neumann J, Sharma S, Entian KD, Schneider D, Helm M (2014) Absolute and relative quantification of RNA modifications via biosynthetic isotopomers. Nucleic Acids Res 42(18):e142. https://doi.org/10.1093/ nar/gku733 6. Thuring K, Schmid K, Keller P, Helm M (2017) LC-MS analysis of methylated RNA. Methods Mol Biol 1562:3–18. https://doi. org/10.1007/978-1-4939-6807-7_1 7. Borland K, Diesend J, Ito-Kureha T, Heissmeyer V, Hammann C, Buck AH, Michalakis S, Kellner S (2019) Production and application of stable isotope-labeled internal standards for RNA modification analysis. Genes (Basel) 10(1). https://doi.org/10. 3390/genes10010026 8. Reichle VF, Petrov DP, Weber V, Jung K, Kellner S (2019) NAIL-MS reveals the repair of 2-methylthiocytidine by AlkB in E. coli. Nat Commun 10(1):5600. https://doi.org/10. 1038/s41467-019-13565-9

9. Heiss M, Reichle VF, Kellner S (2017) Observing the fate of tRNA and its modifications by nucleic acid isotope labeling mass spectrometry: NAIL-MS. RNA Biol 14(9):1260–1268. https://doi.org/10.1080/15476286.2017. 1325063 10. Heiss M, Hagelskamp F, Kellner S (2020) Cell culture NAIL-MS allows insight into human RNA modification dynamics in vivo. bioRxiv. https://doi.org/10.1101/2020.04.28. 067314 11. Thumbs P, Ensfelder TT, Hillmeier M, Wagner M, Heiss M, Scheel C, Schon A, Muller M, Michalakis S, Kellner S, Carell T (2020) Synthesis of galactosyl-queuosine and distribution of hypermodified Q-nucleosides in mouse tissues. Angew Chem Int Ed Engl. https://doi.org/10.1002/anie.202002295 12. Chionh YH, Ho CH, Pruksakorn D, Ramesh Babu I, Ng CS, Hia F, McBee ME, Su D, Pang YL, Gu C, Dong H, Prestwich EG, Shi PY, Preiser PR, Alonso S, Dedon PC (2013) A multidimensional platform for the purification of non-coding RNA species. Nucleic Acids Res 41(17):e168. https://doi.org/10.1093/nar/ gkt668 13. Hagelskamp F, Borland K, Ramos J, Hendrick AG, Fu D, Kellner S (2020) Broadly applicable oligonucleotide mass spectrometry for the analysis of RNA writers and erasers in vitro. Nucleic Acids Res 48(7):e41. https://doi. org/10.1093/nar/gkaa091 14. Hauenschild R, Tserovski L, Schmid K, Thuring K, Winz ML, Sharma S, Entian KD, Wacheul L, Lafontaine DL, Anderson J, Alfonzo J, Hildebrandt A, Jaschke A, Motorin Y, Helm M (2015) The reverse transcription signature of N-1-methyladenosine in RNA-Seq is sequence dependent. Nucleic Acids Res 43(20):9950–9964. https://doi. org/10.1093/nar/gkv895 15. Collart MA, Oliviero S (2001) Preparation of yeast RNA. Curr Protoc Mol Biol Chapter 13: Unit13 12. https://doi.org/10.1002/ 0471142727.mb1312s23

Chapter 19 A Method to Monitor the Introduction of Posttranscriptional Modifications in tRNAs with NMR Spectroscopy Alexandre Gato, Marjorie Catala, Carine Tisne´, and Pierre Barraud Abstract During their biosynthesis, transfer RNAs (tRNAs) are decorated with a large number of posttranscriptional chemical modifications. Methods to directly detect the introduction of posttranscriptional modifications during tRNA maturation are rare and do not provide information on the temporality of modification events. Here, we report a methodology, using NMR as a tool to monitor tRNA maturation in a nondisruptive and continuous fashion in cellular extracts. This method requires the production of substrate tRNA transcripts devoid of modifications and active cell extracts containing the necessary cellular enzymatic activities to modify RNA. The present protocol describes these different aspects of our method and reports the time-resolved NMR monitoring of the yeast tRNAPhe maturation as an example. The NMR-based methodology presented here could be adapted to investigate diverse features in tRNA maturation. Key words Transfer RNA, tRNA, Posttranscriptional modifications, RNA modifications, Yeast, RNA transcription, NMR spectroscopy, Yeast extract, Time-resolved NMR, In extract NMR

1

Introduction Synthesis and maturation of transfer RNAs (tRNAs) involve posttranscriptional chemical modifications of their nucleotides. These modifications occur at specific sites in a tightly controlled manner, which ensures that the tRNA biogenesis process effectively leads to the formation of mature and functional tRNAs [1–4]. Currently, over 140 posttranscriptional modifications are reported in RNAs, the vast majority being found in tRNAs [5, 6]. This family of RNAs displays not only the largest variety of posttranscriptional decoration, but also the highest density of modification per RNA molecule, with ~8–25% of modified nucleotides in tRNAs depending on the organisms [7, 8]. Notably, all tRNA functions within cells are affected by modifications. In particular, modifications in and around the anticodon are implicated in the decoding process

Mary McMahon (ed.), RNA Modifications: Methods and Protocols, Methods in Molecular Biology, vol. 2298, https://doi.org/10.1007/978-1-0716-1374-0_19, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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Fig. 1 Schematic overview of the NMR method used to monitor RNA modifications. (a) Step 1: tRNA transcription with 15N-labeled nucleotides. A 15N-labeled tRNA (represented as red dots) is produced by T7 in vitro transcription. The

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[1, 9, 10]. In addition, some modifications control cleavages producing tRNA fragments [11, 12]. And finally, cellular stress can result in changes in tRNA modifications, which can lead to translational reprogramming [13, 14]. Even though modifications are central in tRNA biology, methods to directly detect their introduction during tRNA biosynthesis are rare and do not provide information on the temporality of modification events. With the aim of filling this gap, we have developed a methodology, using NMR as a tool to monitor tRNA maturation in a nondisruptive and continuous manner [15]. Briefly, introducing isotope-labeled tRNAs into unlabeled cell extracts containing the cellular enzymatic activities required to modify tRNA, combined with the use of isotope filters in NMR experiments, enables the detection of a tRNA of interest within the complex cell extract environment (Fig. 1). RNA modification events are directly monitored in a time-resolved fashion, by measuring successive NMR experiments on a single sample directly incubated in the NMR spectrometer. In this chapter, we describe the detailed procedure for monitoring tRNA maturation in cell extracts with NMR spectroscopy. We demonstrate our methodology with the maturation of the yeast tRNAPhe in yeast extracts. Our methodology consists of three steps (Fig. 1). In the first step, a 15N-labeled tRNA sample is produced by in vitro transcription with 15N-labeled nucleotides and purified by ion-exchange chromatography (Fig. 1; step 1). In the second step, an unlabeled cellular extract is produced from the lysis of a cell culture (Fig. 1; step 2). To preserve as far as possible the cellular enzymatic activities in the extract, cell lysis is performed under gentle conditions and in the presence of anti-proteases. After ultracentrifugation of the cell debris, active unlabeled cellular extracts ä Fig. 1 (continued) transcription reaction is purified by ion-exchange chromatography (MonoQ column), and the purified 15N-labeled tRNA is dialyzed and concentrated for use in step 3. (b) Step 2: unlabeled cell extract preparation. A cell culture is performed in an Erlenmeyer flask with unlabeled growth media. After cell lysis, cell debris are removed by ultracentrifugation, leading to the production of unlabeled cellular extract subsequently used in step 3. (c) Step 3: NMR monitoring of tRNA maturation. The 15N-labeled tRNA produced in step 1 is introduced in the unlabeled cellular extract produced in step 2. This mix is transferred to an NMR tube to yield an in extract NMR sample. The tRNA maturation sample is incubated directly in the NMR spectrometer. Successive NMR measurements are performed to provide time-resolved information on the tRNA maturation process. NMR chemical shift changes (indicated with arrows on the NMR spectra) are identified and linked to specific modification events. The acquisition of NMR spectra in a continuous and time-resolved fashion enables the identification of a sequential order in the introduction of posttranscriptional modifications

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are obtained. In the third step, the NMR monitoring of tRNA maturation is performed (Fig. 1; step 3). An in extract NMR sample is produced by mixing the 15N-labeled tRNA sample obtained in step 1 with the unlabeled cell extract obtained in step 2. This sample is incubated directly in the NMR spectrometer and a series of NMR experiments are measured in order to monitor tRNA maturation events in a time-resolved fashion (Fig. 1; step 3). In the NMR spectra, the progressive appearance of new signals and the correlated disappearance of signals from the unmodified tRNA sample are the signature of chemical modifications being introduced in the initial transcript. Since NMR spectra are measured as a time-course series, our methodology enables the identification of early and later modifications events. Overall, NMR spectroscopy provides the means to observe sequential orders in the introduction of modifications along the tRNA maturation pathway.

2

Materials All solutions are prepared using reagents of the highest available purity and with ultrapure deionized water. Particular care should be taken to avoid RNase contamination during preparation of reagents that are required for the RNA transcription reaction (see Note 1). The use of ultrapure deionized water is sufficient to assure RNasefree condition, provided that the water purification system is well maintained. All reagents required for RNA transcription and cellular extract preparation are stored at
20  C, unless otherwise indicated (see Note 2).

2.1 tRNA Transcription and Purification

1. T7 RNA polymerase (stock at 1 mg/mL stored at
20  C in 20 mM Na-phosphate pH 7.7, 100 mM NaCl, 10 mM DTT, 1 mM EDTA, 50% (v/v) glycerol). Commercial or in-house produced (see Note 3). 2. In vitro T7 transcription buffer concentrated 20 times (TB20x): 400 mM Tris–HCl pH 8.0, 10 mM spermidine, 50 mM DTT, 0.1% (v/v) Triton X-100 in water. Store at
20  C. 3. 100 mM Stock solution of unlabeled ATP and CTP, pH 8. Store at
20  C. 4. 100 mM Stock solution of 15N-labeled UTP and GTP (Cambridge Isotope Laboratories). Store at
20  C. 5. 100 μM DNA oligonucleotide (of high purity and dissolved in water) for use as template for the yeast tRNAPhe transcription (see Note 4). For yeast tRNAPhe, use an oligonucleotide with the following sequence:

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50 TGG TGC GAA TTC TGT GGA TCG AAC ACA GGA CCT CCA GAT CTT CAG TCT GGC GCT CTC CCA ACT GAG CTA AAT CCG CTA TAG TGA GTC GTA TTA 30 . 6. 100 μM DNA oligonucleotide for use as T7 promotor primer (see Note 5). For example: 50 TAA TAC GAC TCA CTA TAG 30 . 7. 280 mM MgCl2 in water. Sterilize by filtration with a 0.22 μm filter and store at
20  C. 8. 80 mM GMP, pH 7.5 in water. Sterilize by filtration with a 0.22 μm filter and store at
20  C. 9. Water bath with temperature control. 10. 0.5 M EDTA, pH 8.0. Store at room temperature. ¨ KTA Purifier (GE Healthcare) or similar chromatographic 11. A system. 12. MonoQ 10/100 GL column (GE Healthcare). 13. 0.5 M NaOH. 14. Purification buffer A: 25 mM Na-phosphate pH 6.5, 50 mM NaCl, 5 mM MgSO4 (filtrate with a 0.22 μm filter). 15. Purification buffer B: 25 mM Na-phosphate pH 6.5, 1 M NaCl (filtrate with a 0.22 μm filter). 16. NMR buffer stock solutions: 500 mM Na-phosphate pH 6.5, 1 M MgCl2. 17. Dialysis sacks with 3.5 kDa MWCO (e.g., Spectra/Por 3 Dialysis Tubing Spectra/Por). 18. Standard closures for dialysis tubing. 19. Concentrators Amicon Ultra-15 MWCO 10 kDa (Millipore/ Merck). 20. Ultrapure deionized water. 2.2 Cellular Extract Preparation

1. 100 mg/mL Ampicillin stock solution in water. Store at
20  C. 2. YEPD-ampicillin agar plates: Dissolve 10 g of yeast extract, 20 g of peptone, and 20 g of agar in 950 mL of water and sterilize by autoclaving. Prepare a solution of 40% (w/v) of glucose in water and sterilize by filtration with 0.22 μm filter. Add 50 mL of the sterile-filtered glucose solution in the autoclaved medium to obtain 1 L of YEPD agar medium containing 2% (w/v) glucose. Allow the solution to cool to ~50  C and add ampicillin at a final concentration of 100 μg/mL. Mix well and pour ~25 mL of the solution per Petri dish. Plates can be stored upside down at 4  C for several weeks.

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3. YEPD medium: Dissolve 10 g of yeast extract and 20 g of peptone in 950 mL of water and sterilize by autoclaving. Prepare a solution of 40% (w/v) of glucose in water and sterilize by filtration with 0.22 μm filter. Solutions can be stored for months at room temperature. Add 50 mL of the sterile-filtered glucose solution in the autoclaved medium to obtain 1 L of YEPD medium containing 2% (w/v) of glucose. 4. Glycerol stock of c13-ABYS-86 S. cerevisiae strain stored at
80  C (see Note 6). 5. Static and shaking incubators at 30  C. Sterile 100 mL and 2 L Erlenmeyer flasks. 6. Spectrophotometer to measure the absorbance at 600 nm (OD600). 7. Centrifugation bottles for cell harvesting and S30 cell extract preparation and corresponding rotors (e.g., Beckman Coulter JA-10 and JA-20). Ultracentrifugation bottles for S100 cell extract preparation and corresponding rotor (e.g., Beckman Coulter Type 70.1 Ti). 8. Centrifuge and ultracentrifuge with temperature control that allows cooling to 4  C. 9. Mechanical device to disrupt yeast cells such as an Eaton press or a French press (see Note 7). 10. Lysis buffer: 25 mM KH2PO4/Na2HPO4 pH 7.0, 10 mM MgCl2, 0.1 mM EDTA. Sterilize by filtration with a 0.22 μm filter and divide into aliquots of 10 mL. Store at
20  C. 11. 1 M DTT stock solution: Dissolve 3.085 g of 1,4-dithio-DLthreitol into 20 mL of 10 mM sodium acetate pH 5.2. Sterilize by filtration with a 0.22 μm filter and divide into aliquots of 1 mL. Store at
20  C. 12. 100 mM Phenylmethanesulfonyl fluoride (PMSF) solution in isopropanol. Store at 4  C. 13. 500 mM Benzamidine stock solution: Dissolve 437 mg of benzamidine hydrochloride into 5 mL of water. Sterilize by filtration with a 0.22 μm filter and divide into aliquots of 500 μL. Store at
20  C. 14. Antiprotease stock solutions concentrated 5000: Dissolve 5 mg of leupeptin and 5 mg of antipain into 1 mL of water and divide into aliquots of 200 μL. Separately, dissolve 5 mg of pepstatin A and 5 mg of chymostatin into 1 mL of DMSO and divide into aliquots of 200 μL. Store both antiprotease stock solutions at
20  C. 15. Liquid nitrogen (see Note 8). 16. Ultrapure deionized water.

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1. Maturation buffer ten times concentrated (MB10): 1.5 M NaH2PO4/K2HPO4 pH 7.5, 50 mM NH4Cl, 20 mM DTT, 1 mM EDTA in water (see Note 9). Sterilize by filtration with a 0.22 μm filter and divide into aliquots of 500 μL. Store at
20  C. 2. 1 M MgCl2 in water: Sterilize by filtration with a 0.22 μm filter and divide into aliquots of 500 μL. Store at
20  C. 3. 100 mM S-adenosyl-L-methionine (SAM) stock solution (see Note 10) in 5 mM H2SO4 (see Note 11). Divide into aliquots of 25 μL. Store at
20  C for up to 4–6 months. 4. 100 mM ATP stock solution, pH 8.0. Store at
20  C. 5. 100 mM β-Nicotinamide adenine dinucleotide 20 -phosphate reduced (NADPH) stock solution (see Note 12) pH 8 in water (see Note 13). Divide into aliquots of 25 μL. Store at
20  C for up to 1–2 months. 6. Deuterium oxide 99.96% (D2O) (e.g., Eurisotop, catalog number: D215B). 7. 5 mm Shigemi NMR tubes (see Note 14). 8. Manual centrifuge equipped with a swing-out rotor and 15 mL conical tube adaptors for the centrifugation of NMR tubes (e.g., Hettich, catalog numbers: HET-1011 and HET-1014). 9. Solution-state NMR spectrometer operating at 600 MHz or higher, equipped with a cryogenically cooled triple-resonance probe with z-axis gradients. 10. RNase-free water.

3

Methods

3.1 tRNA Transcription and Purification

The optimal T7 in vitro transcription conditions depend greatly on the MgCl2 and NTP concentrations. The [MgCl2]/[NTP] ratio should be adjusted to optimize the yield of the RNA of interest. We recommend testing small-scale reactions (e.g., 40 μL) before largescale production of RNAs (5–20 mL) (see Note 15). For large-scale in vitro transcription, scale up the best conditions determined with small-scale reactions. In our hands, large amount of unmodified yeast tRNAPhe 15N-labeled on Us and Gs (see Note 16), sufficient for monitoring its maturation with NMR, can be obtained from a 10 mL transcription reaction as described here. 1. Prepare the large-scale transcription reaction of 10 mL in a 50 mL conical tube as follows: 40 μL of DNA template premixed with the T7 promotor primer at a 1:1 ratio (stock at 50 μM), 500 μL of TB20 transcription buffer, 500 μL of non-labeled ATP (stock at 100 mM), 500 μL of non-labeled CTP (stock at 100 mM), 500 μL of 15N-labeled UTP (stock at

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100 mM), 500 μL of 15N-labeled GTP (stock at 100 mM), 1300 μL of MgCl2 (stock at 280 mM), 500 μL of GMP (stock at 80 mM), 250 μL of T7 RNA polymerase (stock at 1 mg/mL), and complete with ultrapure deionized water to 10 mL. 2. Incubate in a water bath for ~3–4 h at 37  C (see Note 17). 3. Stop the reaction by adding 1.0–1.5 mL of 0.5 M EDTA pH 8.0 (see Note 18). The transcription reaction can be stored for a few days at
20  C until purification or it can be purified directly as described below. ¨ KTA Purifier or equivalent 4. Wash the purification system (A FPLC purification system) and the column (MonoQ 10/100 GL) with 0.5 M NaOH and rinse them extensively with water (see Note 19). Then, equilibrate the MonoQ column in purification buffer A. 5. Filtrate the large-scale transcription sample with a 0.22 μm filter and load up to 5–6 mL of the transcription sample (see Note 20) on the MonoQ column previously equilibrated with purification buffer A at a flow rate of 2 mL/min. 6. Elute the tRNA transcript from the MonoQ column with a 20–80% gradient of purification buffer B over 75 mL (~10 column volumes) at a flow rate of 3 mL/min. Collect 1 mL fractions. Transcribed yeast tRNAPhe is usually eluted between 50% and 55% of buffer B (see Note 21). 7. Repeat steps 5 and 6 with the remaining of the 10 mL largescale transcription, and pool the fractions containing the tRNA of interest from both purifications. 8. Dialyze extensively the pooled fractions of yeast tRNAPhe in a dialysis sack against 1 mM Na-phosphate pH 6.5 (see Note 22). 9. Recover the desalted tRNAPhe in a 50 mL conical plastic tube and refold it by heating the sample at 95  C for 5 min and letting it cool down slowly at room temperature. Add NMR buffer stock solutions (e.g., 500 mM Na-phosphate pH 6.5 and 1 M MgCl2) to place the refolded sample in the final NMR buffer, namely 10 mM Na-phosphate pH 6.5 and 10 mM MgCl2 (see Note 23). 10. Concentrate the yeast tRNAPhe sample in the NMR buffer up to ~1.5–2.0 mM using Amicon concentrators of 10 kDa. Typically, a 10 mL transcription with 15N-labeled Us and Gs provides a final NMR sample of 300 μL at ~1.8 mM, corresponding to 0.5–0.6 μmol of purified tRNA. Store the concentrated yeast tRNAPhe NMR sample at
20  C until needed (see Note 24).

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All steps after cell harvest are performed on ice, unless indicated otherwise. Precool centrifugation bottles, rotors, and centrifuges before usage. 1. Streak yeast c13-ABYS-86 cells from glycerol stock on YEPDampicillin agar plate and incubate the plate in a static incubator at 30  C for ~2 days until the colonies appear. 2. Start a preculture by transferring an individual colony with a sterile inoculation loop into a 100 mL Erlenmeyer containing 25 mL of YEPD medium supplemented with 100 μg/mL of ampicillin. Incubate the preculture in a shaking incubator at 30  C and 200 rpm for 20 h. 3. Measure the OD600 of the preculture. Start a culture with an OD600 of 0.15 by transferring the appropriate volume of preculture into a 2 L Erlenmeyer containing 500 mL of YEPD medium. Incubate the culture in a shaking incubator at 30  C and 200 rpm for 24 h. 4. Harvest the cells by centrifugation at 3500  g for 20 min at room temperature in 500 mL centrifugation bottles. Resuspend the cell pellet in 40 mL of cold water and transfer the suspension into a 50 mL conical tube. Centrifuge cells at 3500  g for 20 min at 4  C. Weight the cell pellet (see Note 25). Store the cells at
80  C until needed for cellular extract preparation. 5. Add one equivalent volume of lysis buffer (i.e., 1 mL of lysis buffer per g of yeast cells) to the frozen cell pellet on ice and resuspend cells gently. Add DTT at a final concentration of 2 mM, PMSF, and benzamidine both at a final concentration of 1 mM, and leupeptin, antipain, pepstatin A, and chymostatin each at a final concentration of 1 μg/mL. 6. Lyse the cells by two sequential passages in a precooled Eaton pressure chamber at
80  C operated via a hydraulic press with a working pressure of 20,000–30,000 psi. Alternatively, cells may be lysed by two sequential passages in a precooled French press at 4  C operating at a pressure of 10,000–20,000 psi (see Note 7). 7. Evaluate the volume of lysed cell suspension recovered after lysis and add additional DTT, PMSF, benzamidine, leupeptin, antipain, pepstatin A, and chymostatin to double the previously mentioned final concentrations (see Note 26). 8. Centrifuge the disrupted cells at 30,000  g for 1 h at 8  C. Carefully recover the supernatant and transfer it into a clean ultracentrifugation bottle. Ultracentrifuge the yeast S30 cell extract at 100,000  g for 1 h at 8  C. Transfer 250 μL aliquots of the supernatant into 1.5 mL safe-lock tubes. Snap freeze the S100 yeast extract aliquots into liquid nitrogen (see Note 8) and store the cellular extracts at
80  C until needed (see Note 27).

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NMR Monitoring of tRNA Maturation tRNA maturation can simply be monitored by adding the unmodified 15N-labeled tRNA prepared in step 1 into the unlabeled cellular extract prepared in step 2 (Fig. 1). In our hands, using yeast tRNAPhe, sufficient signal-to-noise ratio in NMR measurements can be achieved with a concentration of 15N-labeled tRNA of 40 μM in the cellular extract (see Note 28). Additionally, cofactors of modification enzymes should be added to the reaction to ensure efficient activity of the cell extract. In practice, we add S-adenosyl-Lmethionine (SAM), the almost universal methyl donor used by RNA methyltransferases, and reduce nicotinamide adenine dinucleotide phosphate (NADPH), a hydride donor implicated in the formation of dihydrouridines. All components of the maturation reaction should be prepared and mixed together on ice. The 15 N-labeled tRNA must be added last and the delay from mixing to the starting of the NMR experiments must be as quick as possible and kept shorter than ~5–10 min (see Note 29). The incubation is done directly in the NMR spectrometer, and a series of NMR experiments are measured in a time-resolved fashion in order to visualize the sequential order of tRNA modification events (Fig. 2). Although the observation of NMR chemical shift changes that correspond to modification events is easily performed at this step, the interpretation of these changes in terms of posttranscriptional modifications is not necessarily straightforward and might necessitate in-depth NMR studies (see Note 30). 1. Prepare the tRNA maturation reaction of 280 μL in a 1.5 mL tube on ice as follows: 185 μL of yeast cellular extract prepared in Subheading 3.2, 28 μL of MB10 maturation buffer, 1.4 μL of MgCl2 (stock at 1 M; 5 mM final), 11.2 μL of SAM (stock at 100 mM; 4 mM final), 11.2 μL of ATP (stock at 100 mM; 4 mM final), 11.2 μL of NADPH (stock at 100 mM; 4 mM final), 14 μL of D2O (5% (v/v) final), 6.3 μL of 15N-labeled yeast tRNAPhe prepared in Subheading 3.1 (stock at 1.8 mM; 40 μM final), and complete with RNase-free water to 280 μL (see Note 31). 2. Quickly transfer the tRNA maturation mix to a 5 mm Shigemi tube. Spin down the reaction mix using a manual centrifuge adapted for NMR tube centrifugation. Put the Shigemi plunger in place and maintain it in position with sealing plastic film (e.g., Parafilm). Insert the NMR tube into the NMR spectrometer probe previously equilibrated at 30  C. 3. Perform a rapid adjustment of the shims and quickly start the series of 2D (1H, 15N) correlation spectra with NMR parameters determined on a reference sample of similar composition (see Note 29). In our hands, with the yeast tRNAPhe,

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Fig. 2 Time-resolved NMR monitoring of RNA modifications in yeast tRNAPhe. (a) Imino (1H, 15N) correlation spectrum of a 15N-labeled yeast tRNAPhe measured in vitro to provide a reference spectrum (top left spectrum) and in a time-resolved fashion during a continuous incubation at 30  C in yeast cellular extract over 18 h (remaining five spectra). Each NMR spectrum measurement spreads over a 2-h time period (incubation time indicated on each spectrum). Modifications occurring at the different steps are reported with arrows. (b) Schematic view of the sequential order of the introduction of modifications in yeast tRNAPhe as observed using NMR

high-quality NMR spectra were measured with 2D (1H, 15N)BEST-TROSY experiments [16] with 210 transients and interscan delays of 200 ms. Sweep widths were set to 24.0 and 26.0 ppm and 3072  96 complex points were measured for the 1H and 15N dimensions, respectively. With these settings, a 2D NMR spectra is recorded in ~120 min. Record a series of 12 identical spectra in a continuous fashion, for monitoring tRNA maturation during 24 h.

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4. Process the series of spectra with cosine-bell apodization functions and zero filling to 4096 and 512 data points in the 1H and 15 N dimensions, respectively. Other more complicated apodization schemes may also be used to achieve better S/N ratios or resolution. 5. Analyze the series of NMR spectra and identify the NMR chemical shift changes that correspond to modification events. For accurate interpretations of the changes appearing upon incubation, an in-depth NMR study may be needed (see Note 30). The acquisition of NMR spectra in a continuous and timeresolved fashion enables the identification of a potential sequential order in the introduction of posttranscriptional modifications. In the case of the yeast tRNAPhe, some modifications appear early in the modification process, such as Ψ55, m7G46, and T54, and some appear late, such as m1A58 (Fig. 2).

5

Notes 1. The introduction of undesired RNase activity into the in vitro transcription reaction can significantly reduce the efficiency of the reaction and therefore the overall yield of tRNA production by T7 in vitro transcription. 2. Storage at
20  C significantly prolongs the lifetimes of hydrolysis-sensitive reagents. We usually do not observe any significant deterioration of stored reagents within 1 year at
20  C, unless indicated otherwise. Yeast cellular extracts are more sensitive and should be stored at
80  C. 3. Protocol for the expression and purification of T7 RNA polymerase in E. coli is not detailed here, but has been described comprehensively elsewhere, and active T7 RNA polymerase can be easily prepared following published protocols [17, 18]. 4. We usually order the oligonucleotide at 1 μmol of synthesis scale with a PAGE purification. Design the correct sequence to produce other tRNAs of interest for studying their maturation with NMR. 5. Efficient transcription is typically obtained from an 18-nucleotide-long T7 promotor primer: 50 TAA TAC GAC TCA CTA TAG 30 [19]. We usually order the oligonucleotide at 0.2 μmol of synthesis scale purified with a simple desalting step. 6. The multiple protease-deficient strain c13-ABYS-86 (genotype MATα ura3Δ5 leu2-3112 his3 pra1-1 prb1-1 prc1-1 cps1-3) [20] lacks the four major nonspecific proteases, namely the two endoproteinases A and B and the two carboxypeptidases Y and S. Cell extracts prepared from this strain have a much

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reduced ability to degrade unspecific protein substrates, namely ~1% of the remaining protease activity as compared to a wildtype strain harboring these enzymes [21]. 7. Several methods for lysing yeast cells are available, such as pressure chambers (e.g., Eaton press and French press), abrasives (glass bead vortexing), and enzymatic lysis (e.g., zymolyase). In our hands, the best enzymatic activity is achieved using mechanical lysis of frozen cells in an Eaton pressure chamber [22]. If an Eaton press is not available in the laboratory, we recommend using a French press for cell lysis. 8. Cell extracts may also be snap frozen using dry ice or liquid nitrogen. 9. The composition of the maturation buffer aims to achieve cellular conditions while at the same time remaining compatible with NMR spectroscopy [23]. 10. The precise concentration of the SAM stock solution should be calculated from the millimolar extinction coefficient of SAM at 260 nm (ε260(SAM) ¼ 15.4 mM
1 cm
1) and measurement of the absorbance at 260 nm of a suitable dilution of the solution. 11. We recommend preparing SAM stock solutions from SAM p-toluenesulfonate salt. In addition, SAM is not stable in the presence of nucleophiles, and is therefore not stable in basic conditions. Sulfuric acid H2SO4 is a strong acid with a conjugate base SO42
being a poor nucleophile. SAM solutions are quite stable in these conditions, if stored at
20  C. Note that the pH of a 5 mM H2SO4 solution is ~2, which may be checked by putting a drop of the SAM stock solution on a pH indicator paper. 12. The precise concentration of the NADPH stock solution should be calculated from the millimolar extinction coefficient of NADPH at 340 nm (ε340(NADPH) ¼ 6.3 mM
1 cm
1) and measurement of the absorbance at 340 nm of a suitable dilution of the solution. 13. We recommend preparing NADPH stock solution from NADPH tetrasodium salt. In addition, NADPH is not stable at acidic pH; therefore adjust the pH of the stock solution to pH ~8 by adding a moderately concentrated base (e.g., NaOH 3–5 M) drop by drop with a micropipette. pH should be checked by putting a drop of the NADPH stock solution on a pH indicator paper. 14. Other types of NMR tubes, such as standard 5 mm or 3 mm tubes, may also be used, but volumes of tRNA maturation reactions should be adjusted accordingly. 15. We recommend testing small-scale reactions of 40 μL before large-scale production of tRNAs. Transcriptions are performed

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with each of the four NTPs at 5 mM (total concentration of 20 mM). Concentration of MgCl2 is varied in small steps in order to evaluate the efficiency of transcription with the [MgCl2]/[NTPs] ratio ranging from 0.3 to 3.0. Concentration of T7 RNA polymerase and DNA template and the addition of GMP at 2–6 mM also need to be tested in a second step after determination of the best MgCl2 concentration. The setup and the analysis of small-scale transcriptions with denaturing urea-PAGE can be performed as described in [24]. 16. Our methodology to monitor posttranscriptional modifications in tRNAs with NMR relies on the fact that imino signals are very sensitive to their chemical environment. Imino groups are carried by uridines and guanosines, and can be observed in (1H, 15N) correlation spectra on condition that the imino proton is protected from exchange with the solvent by hydrogen bonding. Since adenosines and cytosines do not carry imino groups, it is not necessary to produce a sample that is 15 N-labeled on As and Cs. This has the advantage of drastically reducing the cost of sample production. 17. After ~1 h of incubation, a white precipitate of magnesium pyrophosphate may become visible and is indicative of an efficient transcription. 18. After addition of EDTA and a few minutes of incubation, the solution should turn clear. Magnesium ions are chelated by EDTA, which displaces the equilibrium of magnesium pyrophosphate precipitation. In case of remaining turbidity, add EDTA 0.5 M progressively, by increments of 0.1 mL until the solution becomes clear. 19. Transcribed RNAs are sensitive to RNases; one should therefore be careful to operate in RNase-free conditions. Washing the purification system and the column with 0.5 M NaOH is sufficient to remove most sources of RNase contamination. In addition, we recommend running the purification at 4  C in a cold room or refrigerated cabinet, but purification may also be performed at room temperature. 20. It is not recommended to load more than 5–6 mL of transcription onto the column. With larger quantities of tRNAs, the purification becomes difficult due to a broadening of the elution peaks. Repeat steps 5 and 6 as many times as needed for large-scale transcriptions of more than 5 mL. 21. Elution fractions can be analyzed with denaturing urea-PAGE. Yeast tRNAPhe usually elutes between 50 and 55% of buffer B in five fractions of 1 mL each. Do not add fractions of tRNAPhe eluted after 55% of buffer B, since they usually contain longer RNA transcripts originating from non-templated nucleotide addition at the 3’-end of the transcript.

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22. For an efficient and extensive dialysis, we usually dialyze the sample against a 2 L dialysis solution for at least 1–2 h and change it with a fresh solution three times. 23. We recommend performing the refolding step on diluted tRNAs at a low salt concentration. In the case of the yeast tRNAPhe, we have observed that refolding at high salt and/or in the presence of MgCl2 was less suitable for obtaining tRNAs of high-quality NMR spectra. Refolding with high salt most likely leads to increased misfolding. However, the situation might be different for the refolding of other tRNAs and should be tested. 24. The yeast tRNAPhe NMR sample can be stored at
20  C for at least 12–18 months without observable degradation or alteration of its NMR spectral properties. 25. The weight of the cell pellets can be evaluated by subtracting the weight of an empty conical tube. 26. Final concentrations in the yeast cell extract are as follows: 4 mM DTT, 2 mM PMSF, 2 mM benzamidine, and leupeptin, antipain, pepstatin A, and chymostatin each at 2 μg/mL. 27. Yeast cellular extracts can be stored at
80  C for at least 12 months without a significant reduction of activity as monitored using NMR of posttranscriptional tRNA modifications. 28. For maturation studies, we selected a 15N-labeled tRNA concentration of 40 μM as a compromise to achieve sufficient signal-to-noise ratio in NMR measurements while seeking to approach cellular tRNA concentrations. As a comparison, the concentration of total tRNAs has been estimated to be 100–200 μM in yeast and 200–350 μM in E. coli, with typical concentrations of individual tRNAs of 2–15 μM [25, 26]. 29. To keep the delay between mixing of the reaction components and start of the NMR experiments as short as possible, we recommend setting up NMR experiments on a reference sample of similar composition. Specifically, shimming, tuning, and matching of the probe and pulse calibrations are performed in advance on this reference sample. In these conditions, tRNA maturation experiments can be quickly started, since only a rapid adjustment of the shims on the real sample is needed. 30. One of the main challenges of NMR monitoring of tRNA maturation is to identify the NMR signature of individual modifications and thus to associate a particular chemical shift change on the NMR spectra with a specific tRNA modification event. For the yeast tRNAPhe, we assigned its imino groups in three forms differing in their modification content [24, 27]. From the analysis of the differences between their (1H, 15N) chemical shifts, we could identify the NMR

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signature of individual modifications [15, 27]. The requirements for such a detailed analysis might be system dependent, but we anticipate that for any system, the complete NMR analysis of at least two tRNA samples, with and without modifications, would be crucial. 31. In cases where the solution stocks of SAM, NADPH, and 15 N-labeled tRNA cannot be prepared at the mentioned concentration, please adjust the volumes to reach a final concentration of SAM and NADPH of 4 mM and a final concentration of 15N-labeled tRNA of 40 μM.

Acknowledgments The authors are grateful to Henri Grosjean for protocols and stimulating discussions about RNA modifications, Sylvie Auxilien (I2BC) for the c13-ABYS-86 yeast strain, Bruno Sargueil for guidance regarding the Eaton press implementation, and Christel Le Bon for ensuring the best performance of the NMR infrastructure at the IBPC. The authors acknowledge access to the biomolecular NMR platform of the IBPC that is supported by the CNRS, the Labex DYNAMO (ANR-11-LABX-0011), the Equipex CACSICE (ANR-11-EQPX-0008), and the Conseil Re´gional d’Iˆle-de-France (SESAME grant). This work was supported by grant ANR-14CE09-0012 from the ANR. References 1. El Yacoubi B, Bailly M, de Cre´cy-Lagard V (2012) Biosynthesis and function of posttranscriptional modifications of transfer RNAs. Annu Rev Genet 46:69–95. https://doi.org/ 10.1146/annurev-genet-110711-155641 2. Phizicky EM, Hopper AK (2010) tRNA biology charges to the front. Genes Dev 24:1832–1860. https://doi.org/10.1101/ gad.1956510 3. Hopper AK (2013) Transfer RNA posttranscriptional processing, turnover, and subcellular dynamics in the yeast Saccharomyces cerevisiae. Genetics 194:43–67. https://doi. org/10.1534/genetics.112.147470 4. Barraud P, Tisne´ C (2019) To be or not to be modified: miscellaneous aspects influencing nucleotide modifications in tRNAs. IUBMB Life 71:1126–1140. https://doi.org/10. 1002/iub.2041 5. Boccaletto P, Machnicka MA, Purta E, Piatkowski P, Baginski B, Wirecki TK, de Cre´cy-Lagard V, Ross R, Limbach PA,

Kotter A, Helm M, Bujnicki JM (2018) MODOMICS: a database of RNA modification pathways. 2017 update. Nucleic Acids Res 46:D303–D307. https://doi.org/10. 1093/nar/gkx1030 6. Helm M, Alfonzo JD (2014) Posttranscriptional RNA modifications: playing metabolic games in a cell’s chemical Legoland. Chem Biol 21:174–185. https://doi.org/10.1016/ j.chembiol.2013.10.015 7. Jackman JE, Alfonzo JD (2013) Transfer RNA modifications: nature’s combinatorial chemistry playground. Wiley Interdiscip Rev RNA 4:35–48. https://doi.org/10.1002/wrna. 1144 8. Machnicka MA, Olchowik A, Grosjean H, Bujnicki JM (2014) Distribution and frequencies of post-transcriptional modifications in tRNAs. RNA Biol 11:1619–1629. https://doi.org/ 10.4161/15476286.2014.992273 9. Agris PF, Vendeix FAP, Graham WD (2007) tRNA’s wobble decoding of the genome:

Monitoring the Introduction of Modifications in tRNAs with NMR 40 years of modification. J Mol Biol 366:1–13. https://doi.org/10.1016/j.jmb.2006.11.046 10. Grosjean H, de Cre´cy-Lagard V, Marck C (2010) Deciphering synonymous codons in the three domains of life: co-evolution with specific tRNA modification enzymes. FEBS Lett 584:252–264. https://doi.org/10. 1016/j.febslet.2009.11.052 11. Oberbauer V, Schaefer MR (2018) tRNAderived small RNAs: biogenesis, modification, function and potential impact on human disease development. Genes (Basel) 9:607. https://doi.org/10.3390/genes9120607 12. Lyons SM, Fay MM, Ivanov P (2018) The role of RNA modifications in the regulation of tRNA cleavage. FEBS Lett 592:2828–2844. https://doi.org/10.1002/1873-3468.13205 13. Chan CTY, Pang YLJ, Deng W, Babu IR, Dyavaiah M, Begley TJ, Dedon PC (2012) Reprogramming of tRNA modifications controls the oxidative stress response by codonbiased translation of proteins. Nat Commun 3:937. https://doi.org/10.1038/ ncomms1938 14. Huber SM, Leonardi A, Dedon PC, Begley TJ (2019) The versatile roles of the tRNA epitranscriptome during cellular responses to toxic exposures and environmental stress. Toxics 7:17. https://doi.org/10.3390/ toxics7010017 15. Barraud P, Gato A, Heiss M, Catala M, Kellner S, Tisne´ C (2019) Time-resolved NMR monitoring of tRNA maturation. Nat Commun 10:3373. https://doi.org/10. 1038/s41467-019-11356-w 16. Farjon J, Boisbouvier J, Schanda P, Pardi A, Simorre J-P, Brutscher B (2009) Longitudinalrelaxation-enhanced NMR experiments for the study of nucleic acids in solution. J Am Chem Soc 131:8571–8577. https://doi.org/10. 1021/ja901633y 17. Rio DC (2013) Expression and purification of active recombinant T7 RNA polymerase from E. coli. Cold Spring Harb Protoc 2013:pdb. prot078527. https://doi.org/10.1101/pdb. prot078527 18. De´gut C, Monod A, Brachet F, Cre´pin T, Tisne´ C (2016) In vitro/in vivo production of tRNA

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for X-ray studies. Methods Mol Biol 1320:37–57. https://doi.org/10.1007/9781-4939-2763-0_4 19. Milligan JF, Groebe DR, Witherell GW, Uhlenbeck OC (1987) Oligoribonucleotide synthesis using T7 RNA polymerase and synthetic DNA templates. Nucleic Acids Res 15:8783–8798. https://doi.org/10.1093/ nar/15.21.8783 20. Heinemeyer W, Kleinschmidt JA, Saidowsky J, Escher C, Wolf DH (1991) Proteinase yscE, the yeast proteasome/multicatalyticmultifunctional proteinase: mutants unravel its function in stress induced proteolysis and uncover its necessity for cell survival. EMBO J 10:555–562. https://doi.org/10.1002/j. 1460-2075.1991.tb07982.x 21. Achstetter T, Emter O, Ehmann C, Wolf DH (1984) Proteolysis in eukaryotic cells. Identification of multiple proteolytic enzymes in yeast. J Biol Chem 259:13334–13343 22. Eaton NR (1962) New press for disruption of microorganisms. J Bacteriol 83:1359–1360 23. Freund J, Kalbitzer HR (1995) Physiological buffers for NMR spectroscopy. J Biomol NMR 5:321–322. https://doi.org/10.1007/ BF00211760 24. Catala M, Gato A, Tisne´ C, Barraud P (2020) Preparation of yeast tRNA sample for NMR spectroscopy. Bio-protocol 10:e3646. https:// doi.org/10.21769/BioProtoc.3646 25. Waldron C, Lacroute F (1975) Effect of growth rate on the amounts of ribosomal and transfer ribonucleic acids in yeast. J Bacteriol 122:855–865 26. Dong H, Nilsson L, Kurland CG (1996) Co-variation of tRNA abundance and codon usage in Escherichia coli at different growth rates. J Mol Biol 260:649–663. https://doi. org/10.1006/jmbi.1996.0428 27. Catala M, Gato A, Tisne´ C, Barraud P (2020) 1H, 15N chemical shift assignments of the imino groups of yeast tRNAPhe: influence of the post-transcriptional modifications. Biomol NMR Assign 14(2):169–174. https://doi. org/10.1007/s12104-020-09939-6

Part VI Approaches to Assess Kinetics, Determinants, and Functions of RNA Modifications

Chapter 20 Effects of mRNA Modifications on Translation: An Overview Bijoyita Roy Abstract The mRNA epitranscriptome imparts diversity to gene expression by installing chemical modifications. Advances in detection methods have identified chemical modifications in eukaryotic, bacterial, and viral messenger RNAs (mRNAs). The biological functions of modifications in mRNAs still remain to be understood. Chemical modifications are introduced in synthetic mRNAs meant for therapeutic applications to maximize expression from the synthetic mRNAs and to evade the host immune response. This overview provides a background of chemical modifications found in mRNAs, with an emphasis on pseudouridine and its known effects on the mRNA life cycle, its potential applications in synthetic mRNA, and the methods used to assess its effects on mRNA translation. Key words mRNA modification, Translation, Synthetic mRNA, Therapeutic mRNAs, Pseudouridine

1

Introduction The emerging field of epitranscriptomics has uncovered an additional layer of complexity in the regulation of gene expression that stems from the addition of RNA modifications posttranscriptionally. More than 150 modifications have been identified in highly abundant noncoding RNAs (ncRNAs, such as ribosomal RNAs (rRNAs), transfer RNAs (tRNAs), and small nuclear RNAs (snRNAs)) [1, 2]. It was thought for a long time that posttranscriptional chemical modifications of RNA that affected the RNA’s biogenesis, function, and stability were limited to ncRNAs. Additionally, the only modifications that were reported for proteincoding messenger RNAs (mRNAs) were those in the mRNA 50 cap (N7-methylguanosine (m7G)) and 30 poly(A) tail, internal inosine (I) modifications, and modifications of internal adenosines to N6methyladenosine (m6A) [1]. With advances in next-generation sequencing technologies and in analytical chemistry, a wide range of RNA modifications have now been identified in mRNAs, albeit at lower levels. Among the modifications that have been identified in mRNAs, m6A and pseudouridine (Ψ) occur most frequently.

Mary McMahon (ed.), RNA Modifications: Methods and Protocols, Methods in Molecular Biology, vol. 2298, https://doi.org/10.1007/978-1-0716-1374-0_20, © The Author(s) 2021

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Although the first chemical modifications on mRNA were identified more than 40 years ago, their functional significance has only just begun to be uncovered. The localization of modifications throughout the body of the mRNAs suggests that modifications could potentially alter protein production by multiple mechanisms, and it is becoming increasingly clear that these modifications play diverse roles in a variety of physiological and disease settings. Here, the effects of mRNA modifications on the life cycle of an mRNA, with a specific emphasis on the process of translation, are discussed. The discovery of mRNA modifications, and the pros and cons of the various detection methods, has been covered extensively in the literature. Because m6A has been discussed in detail in other chapters in this issue, I will focus on Ψ, the second most abundant mRNA modification. I also summarize the current state of knowledge of mRNA modifications, highlighting their known effects on protein expression, and also discuss the applications of mRNA modifications in the synthesis of functional mRNAs. Finally, I provide an overview of the methods used to assess the biological functions of mRNA modifications.

2

Overview of mRNA Life Cycle Gene expression in eukaryotes is a complex and highly regulated process, and mRNAs are subjected to multiple regulatory mechanisms throughout the course of their life cycle (Fig. 1) [3]. Eukaryotic pre-mRNAs are processed by the capping, splicing, and polyadenylation machineries in the nucleus. The mature mRNA, which is now ready for export, travels to the nuclear envelope and is translocated into the cytoplasm, where it is accessed by either the translation machinery to begin the process of protein synthesis or the RNA degradation machinery. Under certain circumstances, some mRNA species also undergo an additional localization process that enables the migration of translationally silenced mRNAs to specific cellular destinations before subsequent translation. In the cytosol, the number of protein molecules produced from each mRNA molecule is a combinatorial effect of the mRNA’s translational efficiency and its half-life. Early notions of the interplay between translation and decay were simplistic and based on the idea that mRNAs undergoing translation were protected from decay [3, 4]. It is now well understood that the decay of individual mRNAs can be either accelerated or antagonized by loss of translation efficiency, and changes in mRNA half-life can further alter translational fidelity [4]. A complex set of protein–mRNA and RNA–RNA interactions determine the fate of the mRNA, and perturbation to this set of interactions can have consequences on the mRNA’s life cycle. The presence of chemical tags on the mRNA is one such factor.

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Translation

Transcription

Cap binding complex

Pol II

Nucleus

Decapping Cytosol

Modifications Deadenylation

Nascent peptide Uridylation mRNA cap

AAAAAA Poly(A)

tail

Fig. 1 A simplified illustration showing the different steps in the life cycle of an mRNA. The pre-mRNA undergoes 50 - and 30 -processing, splicing, and polyadenylation, and the mature mRNA is then exported to the cytosol where it undergoes either translation or decay. Chemical modifications can influence all of the steps in mRNA metabolism, including splicing, maturation, translation, and decay 2.1 Chemical Modifications of mRNA

It is well documented that chemical modifications in ncRNAs can alter the charge, sterics, and conformation of the target RNA molecule, thus modulating the RNA’s biogenesis, stability, and function. Therefore, it is not surprising that the presence of chemical modifications in the mRNA can have similar effects in altering gene expression (Fig. 1). Previous studies have shown that the levels of different mRNA modifications vary widely and that there can be >1000-fold differences in the levels of different modifications [2]. Across organisms, the m6A modification has been found to be the most prevalent, and the abundances of Ψ, N4-acetylcytidine (ac4C), Cm, and Gm have been observed to be approaching those of m6A [2]. On the other hand, 3-methylcytidine (m3C), 1-methylguanine (m1G), and 5-hydroxymethylcytidine (hm5C) are rare, with levels at least 500-fold lower than that of m6A, making accurate detection of these modifications more difficult [2]. In general, the absence of high-quality quantitative mRNA modification maps makes it difficult to correlate the effects of modifications with biological phenotypes, and thus, to understand their physiological relevance, especially with respect to translation and mRNA

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B.

Cap binding complex

A.

Modification

Cap binding complex

Nascent peptide mRNA cap

AAAAAA Poly(A)

Premature termination

tail

RNA-binding protein

Aberrant protein pool (truncated protein)

Normal protein pool

Cap binding complex

C.

Translation recoding

D.

Cap binding complex

Altered ribosome loading and ribosome transit

Aberrant protein pool (miscoded protein)

Aberrant protein pool (misfolded protein, low protein abundance)

Fig. 2 A simplified illustration showing the consequences of mRNA chemical modifications on translation. In contrast to an mRNA without any modification (a), the presence of a modification can result in aberrant translation termination, resulting in a protein pool with truncated peptides as observed for N1-m-pseudouridine (b). The presence of pseudouridine can also result in translation recoding and misincorporation of amino acids in the growing peptide (c). Modification-induced changes in secondary structure or RNA-protein interactions can alter ribosome recycling and ribosome transit time (as observed for N1-m-pseudouridine), subsequently affecting mRNA stability or even protein folding (d). All of these can lead to a change in the contents of the protein pool

stability. Furthermore, another critical factor to consider when dissecting the functional consequences of a specific modification is the relative position of the modified nucleotide in the mRNA. On the one hand, the presence of a modification in the coding sequence (CDS) of an mRNA has the potential to alter translation fidelity, decoding, and ribosome transit (Fig. 2). On the other hand, the presence of a modification in the untranslated regions (UTRs) can result in changes to the stability of the RNA structure, thereby modulating its ability to form RNA–protein interactions, which can consequently impact mRNA maturation, translation, and decay through pathways dependent on these interactions. Understanding the effects of modified nucleotides in mRNAs is further complicated by the fact that some of these modifications are reversible, whereas others are irreversible, suggesting that modifications have the potential to dynamically control gene expression.

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Table 1 Summary of modifications present in the body of the mRNA

Chemical structure

Distribution in mRNA Erasers Role in translation

Writers

Readers

N -methyladenosine (m6A)

METTL3/1, WTAP, KIAA1429, METTL16

ALKBH, IGF2BP, FMRP, FTO G3BP1, PRRC2A, HuR YTHDF1/2/3, YTHDC2

30 UTR (near stop codon), 50 UTR

Pseudouridine (Ψ)

PUSs, box h/ACA snoRNAs

Not known

Not known

50 UTR, CDS, 30 UTR

N1-methyladenosine (m1A)

TRMT6/TRMT61A

Not known

ALKBH3, 50 UTR CDS, 30 UTR FTO (mitochondrial mRNAs)

N7-methylguanosine (m7G)

METTL1

Not known

Not known

50 UTR, CDS, 30 UTR

5-Methylcytosine (m5C) NSUN2

YBX1

–

50 UTR, CDS, 30 UTR

3-Methylcytosine (m3C) METLL8

Not known

ALKBH1

Not known

N -acetylcytidine (ac C) NAT10

Not known

Not known

50 UTR, CDS, 30 UTR

5-Methyluridine (m5U) TRMA

Not known

Not known

Not known

20 -O-Me

Fibrillarin

Not known

Not known

CDS

8-Oxo-7,8dihydroguanosine (8-oxo-G)

In response to reactive YBX1, AUF1 oxygen species

Not known

CDS

5-Hydroxy methylcytosine (hm5C)

TET/ABH1

Not known

CDS

6

4

4

Not known

This form of dynamic control of gene expression can be viewed as a coordinated effort between writer proteins (RNA-modifying enzymes), reader (RNA-binding) proteins, and eraser proteins (which remove the modification from the RNA) (Table 1). The interactions of the reader and eraser proteins with the m6A-containing transcripts have been critical in dissecting the effects of m6A on the mRNA life cycle [5, 6]. In addition, positional information of m6A in the mRNA has greatly contributed to a functional understanding of m6A in biological contexts; depending on its location along the mRNA, m6A can affect fundamental processes

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of cell metabolism—such as splicing, transport, stability, and translation of the target mRNA—by providing interaction platforms for specific reader proteins [7]. Transcriptome-wide studies in conjunction with genetic and biochemical investigations have helped to elucidate how the presence of m6A in an mRNA affects the processes of mRNA degradation, transcript maturation, and translation [5, 8, 9]. Mounting evidence has now shown the involvement of m6A methylation status in embryonic development, stem cell regulation, adipogenesis, development of obesity and pathogenesis of type 2 diabetes, immunological processes, and carcinogenesis [10–13]. In the subsequent sections, chemical modifications (other than m6A) that have been identified in mRNAs and their potential effects on mRNA stability and translation are discussed.

3

Modifications at the Ends of the mRNA

3.1 Modifications of mRNA 50 Termini

The mRNA 50 cap is a well-studied and well-documented structure that is post-transcriptionally added to the pre-mRNA. It consists of an untemplated guanosine that is then methylated to form 7-methylguanosine (m7G) via three enzymatic steps involving an RNA triphosphatase (TPase), an RNA guanylyltransferase (GTase), and a guanine-N7 methyltransferase (guanine-N7 MTase) [14]. m7G is linked to the first transcribed nucleotide via a 50 –50 triphosphate bond to form the minimal Cap 0 structure (m7GpppRNA) in lower eukaryotes. mRNAs in higher eukaryotes can further have an additional methyl group on the ribose 20 -O position of the first transcribed nucleotide (20 -O-methylated versions of A, U, C, G) to form Cap 1 RNA, and the adjoining nucleotide can also be subsequently methylated to yield the Cap 2 structure [15]. Furthermore, certain vertebrate mRNAs have either an m6A or a 20 -O-dimethyladenosine (m6Am) as the first nucleotide [5, 8, 16, 17]. The modifications at the 50 end of the mRNA act as a platform for multiple mRNA–protein interactions. The m7G cap is first recognized by the nuclear cap-binding complex (CBC), which facilitates mRNA export from the nucleus and is later replaced by the translation initiation factor eIF4E. Furthermore, the Cap 1 and Cap 2 structures (presence of the 20 -O methylation) have been shown to be pivotal for distinction between self and foreign RNA via altered recognition by RIG-I, MDA5, and IFIT-1 (cellular sensors that induce type I interferon (IFN) signaling) [18–21]. Binding of IFIT-1 to the mRNAs at the extended cap structure can also result in suppression of translation by competing for binding by eIF4E [18]. As for a direct effect on translation and mRNA stability, a recent study has shown that presence of the 20 -Omethylation of the first transcribed nucleotide blocks the decapping and 50 –30 -exoribonuclease activity of DXO (a noncanonical decapping enzyme) on RNA substrates [22].

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The m6Am modification is formed by methylation of an adenosine nucleotide that is adjacent to the m7G cap (denoted as “Am”) by a unique writer protein, phosphorylated CTD-interacting factor 1 (renamed as CAPAM—cap-specific adenosine N6-methyltransferase) [16, 23]. The reversibility of m6Am has been observed only very recently, and FTO (originally identified as a demethylase for internal m6A) has been identified as the demethylase that acts on m6Am in snRNAs [24]. Whether or not the m6Am modification is reversible in mRNAs is still unknown. Some studies have shown an increased half-life for an m6Am-containing mRNA versus mRNAs that do not contain this modification, which can be attributed to an increased resistance of the modified mRNA to the action of the DCP2 decapping enzyme [17]. The molecular mechanisms of these altered interactions of DCP2 with m6Am-containing mRNAs are not well understood and need to be explored further. A direct correlation between the presence of an m6Am cap and translation of the m6Am-containing mRNA is missing, even though knockout of CAPAM has shown decreased translation for a subset of mRNAs [16]. It has been reported that the presence of a methylated nucleotide adjacent to the 50 cap alters the binding of eIF4E to the mRNA cap [25]. Whether or not a methylated nucleotide adjacent to the mRNA cap affects the binding of other components of the translation initiation complex or the cytoplasmic CBC to the mRNA is unclear; this information is needed to further our understanding of how this particular modification might differentially affect translation of a subset of mRNAs. Furthermore, it remains a possibility that understanding the interaction of the cytosolic CBC and its interaction with the m6Am cap could help us understand more about eraser proteins. In addition to the classical m7G cap found at the 50 ends of the majority of eukaryotic mRNAs, nicotinamide adenine dinucleotide (NAD+), NADH, and dpCoA caps have been identified in eukaryotic mRNAs [26, 27]. In contrast to the m7G cap, the NAD+ cap cannot support translation in human cells and destabilizes mRNAs through the deNADding activity of the DXO proteins [28]. The discovery of alternative types of cap structures and the elucidation of their functions are certainly exciting areas of research, and the development of tools for cap metabolism will be pivotal for our understanding of the biological consequences of modifications at the 50 termini of mRNAs. 3.2 Modifications of mRNA 30 Termini

Because of the impact of an RNA’s ends on the stability of the RNA, post-transcriptional modifications at the ends of mRNA are tightly controlled. 30 -end processing generates a mature 30 terminus, which is typically polyadenylated by canonical poly (A) polymerases. The poly(A) tail is bound by poly(A)-binding proteins (PABPs), which facilitates mRNA export from the nucleus,

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enhances protein synthesis through interactions with translation initiation factors, and prevents exoribonucleolytic decay of the mRNA [3]. Shortening a poly(A) tail in the cytoplasm to fewer than 15–20 nt destabilizes its interaction with the PABP, and release of the last PABP makes the mRNA translationally inactive and susceptible to degradation either through the decapping and 50 -to-30 decay pathway or by the 30 -to-50 decay pathway [3]. Deadenylated mRNAs can also be uridylated with terminal uridylyltransferases (TUTases) or noncanonical poly(A) polymerases [29]. Uridylation of the 30 end is diverse and has physiological consequences, illustrating functional diversity of mRNA 30 modifications in controlling RNA stability and degradation. Our understanding of the complex regulatory networks of protein–protein and protein–RNA interactions involved in capping/decapping, polyadenylation/deadenylation, and uridylation is expanding and so is the knowledge of the modifications decorating the 50 and 30 extremities of an RNA.

4 4.1

Modifications in the Body of the mRNA Inosine

Inosine is a widespread modification found in RNA and DNA, and it is formed by the hydrolytic deamination at the C6 position of adenosine. Originally discovered as a precursor of the purine synthesis pathway, inosine is introduced in RNA either nonenzymatically by spontaneous hydrolysis or enzymatically by deaminases [30]. Adenosine-to-inosine (A-to-I) editing sites have been mapped extensively, and most of the sites have been mapped to noncoding regions of transcripts such as introns or UTRs [31, 32]. However, more than 1000 sites have been located in the CDSs of transcripts. When present in RNA, inosine can base pair to A, C, or U, thus allowing inosine to function as a wobble base [32]. The presence of inosine in the RNA has the potential to affect multiple stages in the RNA life cycle because the cellular machinery (splicing and translation factors) recognizes inosines as a G. Not surprisingly, inosine has been demonstrated to alter mRNA stability, splicing, and translational recoding [33, 34]. Ato-I editing in the coding region of the mRNA results in translation recoding and amino acid misincorporation mediated in part by the destabilization of the codon-anticodon interactions [34]. It has been reported that even though inosine is primarily interpreted as guanosine, it can also be decoded, in a context-dependent way, as either adenosine or uracil. Ribosome profiling data have demonstrated inosine-dependent ribosome stalling (up to nine ribosomes) in vivo and premature peptide termination upstream of the inosinecontaining codon. Recoding is rare, most likely because the consequences of recoding are dire, and therefore, it is not surprising that predominant A-to-I editing events occur in the UTRs of mRNAs.

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4.2 N1-Methyladenosine (m1A)

Recent studies implementing transcriptome-wide analyses have suggested the widespread presence of m1A at internal sites of mRNAs with frequencies of 0.007–0.054% (modified/main base) in mammals [35–37]. m1A in the 50 UTR of cytosolic mRNAs has been associated with increased translation efficiency. Additionally, m1A has been reported to be prevalent in the CDSs and 30 UTRs of mitochondrion-encoded transcripts [35, 37]. Manipulation of m1A levels using the mitochondrion-localized m1A methyltransferase TRMT61B was found to result in alteration of translation, thus highlighting a link between m1A and translation regulation [38]. The presence of m1A can disrupt the U-A Watson–Crick base pairing, and the increased efficiency of translation initiation, early elongation, and enhanced translation rates could be a consequence of the altered base-pair interactions. A tRNA T-loop-like structure has been identified to have an increased number of m1A marks, which is dependent on the TRMT6/TRMT61A methyltransferase complex [37]. Even though the altered activity of the writer, reader, and eraser proteins of m1A has been implicated in certain diseases [39], the exact molecular function of m1A is not known, and whether or not there is any direct pathological outcome related to m1A status is unclear.

4.3 N7-Methylguanosine (m7G)

m7G is one of the most abundant modifications present in the mRNA 50 cap, and it is also abundant in tRNAs and rRNAs. m7G sites have now been observed in internal sequences in mRNAs and ncRNAs, and mass spectrometry (MS)-based quantification analyses have revealed the presence of internal m7Gs in decapped mRNAs with a frequency of 0.02–0.05% (modified/main base) [40]. Internal m7G in mRNA has been mapped to singlenucleotide resolution, which has revealed enrichment of this modification in CDSs, 30 UTRs, some 50 UTRs, and AG-rich contexts (AGm7GA, GGm7GAA, AUCGm7GA, and UUm7GAU), a feature that has been observed to be well conserved across different cell lines and tissues [41, 42]. Knockdown of METTL1 (methyltransferase complex for m7G) expression has shown a decrease in the translation efficiency of a subset of transcripts with hypomethylated sites [41, 42]. The exact mechanism of how methylation affects translation is not understood, and future mechanistic studies are required to understand whether internal m7Gs also act as a platform for the binding of translation factors, as has been described for the m7G at the 50 end of the mRNA. Methylation can affect the binding of protein partners and can also alter the stability, or structure, of its target RNAs via steric effects. Furthermore, dynamic regulation of the internal m7G modification has been reported with increased methylation of the m7G sites and subsequent increased translation from the m7G-containing mRNAs in response to hydrogen peroxide and heat-shock treatments. Internal METLL1-mediated m7G marks have been identified in miRNAs as well and N7 methylation

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of the guanosine can regulate the biogenesis and structure of the miRNA [41]. It will be interesting to test if any of the effects of m7G on translation is mediated via miRNAs. 4.4 5-Methylcytosine (m5C)

m5C is a prevalent base modification that is present in tRNAs and rRNA; it has also been mapped broadly in the transcriptome, including in mRNAs [43, 44]. Although the presence of m5C in mRNA was first observed in the 1970s, its complete transcriptomewide mapping has been performed only recently. Initial MS-based transcriptome-wide m5C mapping has suggested high levels of m5C in mRNAs; however, recent work has suggested that the m5C frequency in the transcriptome might be overrepresented due to contamination with m5C from rRNA and tRNAs [45]. Bisulfite sequencing has mapped m5C sites in mRNAs to the 50 UTR; however, a caveat to these findings is that incomplete conversion of cytidine and m5C during bisulfite treatment could have potentially affected the precise mapping of m5C [44, 46, 47]. Overall, 1000 m5C sites have been identified in the 50 UTRs of mRNAs from human cell lines such as HeLa cells. Polysome profiling of HeLa cell lysates followed by bisulfite conversion and RNA sequencing has demonstrated the prevalence of m5C sites near the start codons in mRNAs and has also identified sequence contexts, similar to the tRNA variable loop, where m5C sites are present [48]. Furthermore, the levels of modification were inversely related to ribosome association, suggesting that the presence of m5C results in translation repression. When present in the CDS, m5C has been observed to be enriched in the first and second positions of a few specific codons, suggesting site-specific functions in translation regulation, which could be mediated by the reader proteins that might either perturb ribosome loading during translation initiation or represent an obstacle in the transit of elongating ribosomes [48]. m5C may also have the potential to affect decoding. The presence of m5C in the second position of the codon altered decoding in a bacterial in vitro translation system; however, whether or not m5C can have a same effect in vivo is not understood and will require more in-depth studies [49]. m5C regulation in mRNAs can be mediated by a specific set of writer, reader, and eraser proteins. An indirect effect of m5C on transcript stability can occur via the m5C reader protein YBX1 and its interaction with PABP [50]. Identification of hydroxymethylcytosine (hm5C), 5-formylcytosine, and 5-carboxylcytosine in RNA further highlights potential mechanisms to “erase” m5C from RNA.

4.5 3-Methylcytosine (m3C)

m3C was originally identified in tRNAs and plant mRNAs, but stringent purification of mRNA and quantification by LC-MS/ MS have now shown that it also occurs in mammalian mRNA. Quantification has demonstrated that the abundance of m3C (0.0006–0.0038%, modified/main base) is similar to that of m5C

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and m1A but is much lower than that of m6A [2, 51]. How m3C affects an mRNA’s fate is still unclear. m3C has been found to be installed on mRNAs via METTL8 in murine and human cell lines [51]. Mettl8 KO mice had lower m3C levels in mRNAs, although considerable changes in the m3C levels in tRNAs were also observed [51]. Mettl8 knockout in mice also decreased the ratio of polysomes to monosomes, suggesting that Mettl8 might affect translation. Whether or not the reduced polysome status is a direct effect of the m3C levels in mRNAs needs to be investigated. 4.6 N4-Acetylcytidine (ac4C)

In HeLa cells, ac4C has been detected (0.11%, modified/main base) in more than 4000 regions of the human transcriptome, and these marks have been mapped predominantly in the CDSs, with ac4C content gradually decreasing from the 50 to 30 end of the gene transcript [52]. mRNAs enriched in ac4C have been observed to have a longer half-life, and the effect of ac4C on mRNA life cycle is further highlighted by the observation that ac4C was enriched in a wobble site C codon and that it significantly promoted translation efficiency in a luciferase reporter assay [52]. It has been hypothesized that the presence of ac4C in the wobble position promotes translation by facilitating tRNA decoding and that ac4C increases translation efficiency by enhancing the thermal stability of baseassociated guanosine, thus altering the tRNA–mRNA interaction during translation. The cytidine acetyltransferase NAT10 has been determined to catalyze the formation of ac4C in a variety of mRNAs, because knocking out NAT10 reduces the level of ac4C on the RNA [52]. Analysis of ac4C position on the acetylated transcripts indicated that transcripts with ac4C sites in the CDS or 30 UTR are preferentially downregulated by loss of NAT10. Furthermore, Ribo-seq analyses have demonstrated that acetylated mRNAs have increased translation efficiency compared to non-acetylated mRNAs, and loss of NAT10 decreases the translation efficiency of acetylated mRNAs [52]. It remains to be seen whether the ac4C mRNA modifications exist in other eukaryotes and/or whether this modification is seen in bacteria or archaea. No known reader proteins or a dedicated deacetylation process has been identified for ac4C mRNAs.

4.7 5-Methyluridine (m5U)

m5U is a common modification in the tRNAs of bacteria and eukaryotes, and its formation is catalyzed by TrmA in E. coli. A recent high-throughput sequencing approach with increased sensitivity led to the identification of m5U in mRNAs of various mammalian cells and tissues [53]. The measured contents of m5U in the mRNAs of various types of human cells and mouse tissues range from 0.001% to 0.0059% (m5U/U), comparable to that of m6Am and m3C in mRNAs of mammals. The potential role of m5U in mRNA remains elusive, but it is plausible that m5U in mRNA might affect interaction of the modified codon with the ribosome

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or lead to the alteration of RNA secondary structures and recruitment of m5U-specific binding proteins. Future work on transcriptome-wide mapping of m5U will further uncover the functional roles of m5U in the mRNAs of mammals. 4.8

2’-O-Methylation

While the existence of 20 -O-methyl modifications has been documented since the 1960s, much of the function of 20 -O-methyl modifications still remains unknown. 20 -O-methyl methylation in CDSs induces ribosome stalling by disrupting codon reading [54]. Moreover, knockdown of the methyltransferase fibrillarin was found to result in a decrease in ribosome 20 -O-methylation and a corresponding increase in translational infidelity [55]. 20 -Omethylation also modulates translation of internal ribosome entry site-containing mRNAs [55]. Whether or not this is mediated by the rRNA modification alterations or alteration of distinct IREScontaining mRNA modification is unclear. Apart from a direct function in translation, self- and non-self-identification of mRNAs is dependent on the presence or absence, respectively, of 20 -Omethyl marks in humans [21].

4.9 8-Oxo-7, 8-Dihydroguanosine (8-Oxo-G)

Modifications are introduced into RNAs not just by nucleotidespecific modifying enzymes but also in response to environmental stress conditions (e.g., oxidative stress, heat shock, and UV irradiation), which can cause chemical damage [56]. Reactive oxygen species can oxidize RNA bases and generate 8-oxo-G, 8-dihydroadenosine, 5-hydroxyuridine, 5-hydroxycytidine, and cytosine glycol. 8-Oxo-G, the most abundant form of the oxidized base, has been detected in purified mRNA. Incorporation of 8-oxo-G in luciferase mRNAs leads to the presence of truncated proteins, suggesting that 8-oxo-G can alter decoding [57]. The presence of 8-oxo-G in the CDS has also been reported to alter peptide bond formation and subsequent triggering of an mRNA surveillance pathway, no-go decay (NGD), that targets stalled ribosomes during translation elongation [57]. 8-Oxo-G-containing mRNAs have been reported to be degraded with the involvement of the reader proteins YBX1, AUF1 (also known as hnRNP D), and PCBP1 [58, 59]. The exact mechanism by which the reader proteins trigger the degradation of 8-oxo-G-containing mRNAs, and the interactions that lead to decay factor recruitment, is not well understood. Whether or not 8-oxo-G is reversible or whether this modification preferentially forms in specific mRNAs is not known.

4.10 5-Hydroxymethylcytosine (hm5C)

hm5C can be generated in RNA by the oxidization of m5C via deoxygenases of the ten-eleven translocation (Tet) family. Transcriptome-wide approaches to map hm5C (hydroxymethylated RNA immunoprecipitation followed by sequencing; hMeRIP-seq) demonstrated enrichment of hm5C sites in the CDS [60]. It has been postulated that hm5C acts as an eraser of m5C and can restore

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mRNA translation in mRNAs whose translation is otherwise inhibited by the presence of m5C [61]. Furthermore, m5C and hm5C are thought to affect mRNA secondary structure and thereby regulate target mRNA recognition by RNA-binding proteins. The relative abundance of hm5C in mRNAs is debated because hm5C has not been detected in highly purified mRNA preparations.

5

Pseudouridine in mRNAs Pseudouridine, one of the first post-transcriptional RNA modifications discovered, is now recognized as one of the most abundant types of RNA modifications. Although Ψ was first discovered in rRNAs, tRNAs, and snRNAs, recent transcriptome-wide analysis of Ψ profiles in humans and yeast has now revealed that hundreds of mRNAs contain this modification [2, 62–65]. The ratio of Ψ/ uridine in mammalian mRNA, as measured by LCMS/MS, has been observed to be about 0.2–0.6%, a frequency comparable to that of m6A [66]. Pseudouridine is generated by the C–C glycosidic isomerization of a uridine base, and there are two distinct mechanisms that have been identified for conversion of uridine into Ψ in vivo [67, 68]. The first is an RNA-independent mechanism in which Ψ is deposited by various Ψ synthases (PUSs) that have different substrate specificities and subcellular localizations; the second involves an RNA-dependent mechanism that is guided and catalyzed by Box H/ACA small nucleolar RNAs (snoRNAs). The enzymatic cores of the PUS enzymes are conserved, and substrate RNA recognition is mediated via the various protein domains [69]. However, the Ψ synthase NAP57/DKC1, also known as dyskerin, uses H/ACA snoRNAs as guides to position a uridine at the active site of the enzyme, which then installs the Ψ marks in rRNAs and snRNAs [70]. Interestingly, studies have shown that knocking down the expression of individual PUSs does not drastically alter Ψ levels in mRNA, suggesting that there is functional redundancy among the PUS proteins. The mechanism by which PUS proteins regulate pseudouridylation in specific mRNAs is an intriguing and outstanding question in the field. Recent work combining in silico RNA-folding predictions with systematic substrate mutagenesis and quantitative kinetic analyses has provided insights into the structural features in mRNA that might help determine pseudouridylation by yeast PUS1 and has demonstrated that, in vitro, PUS1 can recognize substrates in a structuredependent manner [71]. These studies suggest that PUS1mediated pseudouridylation in vivo requires the substrates to be structured; however, because mRNA structures are likely to be dynamic in vivo, the presence of cellular RNA-binding proteins probably influences the pseudouridylation process. By extension, the findings also indicate that modulation of an mRNA’s structure

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might be a critical checkpoint for the Ψ-mediated regulation of gene expression; additional studies to understand mRNA structural dynamics in vivo and their role in the process of pseudouridylation will help shed more light on this aspect. In humans, DKC1 has been reported to catalyze the isomerization of a subset of uridines in mRNAs [63]. The chemical properties of Ψ differ from those of uridine, and Ψ has an altered strength of base-pairing with adenine because the phosphodiester backbone is more rigid. The presence of Ψ55 in tRNA has been proposed to stabilize the tertiary structure of tRNAs, particularly under extreme thermal stress. The Ψ modification has also been observed to be in locations crucial for intermolecular RNA–RNA and RNA–protein interactions such as the peptidyl transferase center (for peptide bond formation), in tRNAs and rRNAs (to provide structural stabilization), and in major and minor spliceosomal snRNA complexes. In mRNAs, Ψ has not been found to have any positional bias and has been detected in 50 UTRs, CDSs, and 30 UTRs [62]. Because Ψ can alter base-pairing interactions, it is plausible that Ψ could directly or indirectly influence mRNA stability and translation (Fig. 3). Early evidence for this hypothesis came from in vitro studies showing that targeted conversion of U to Ψ in translation termination codons (UAA, UGA, and UAG) converted these stop codons into missense codons, and U-to-Ψ conversion of premature termination codons inhibited the accelerated degradation of these mRNAs triggered by the process of nonsense-mediated mRNA decay (NMD) [72]. Multiple lines of evidence have demonstrated a role for Ψ in maintaining mRNA stability, although the effects have been dependent on the system in which this was evaluated. In vitro-transcribed mRNAs containing Ψ have been observed to have higher stability than unmodified mRNAs in cell-based assays. Deletion of Pus7 in yeast resulted in reduction of Ψ-containing mRNAs, suggesting that the presence of Ψ can stabilize mRNAs [63]. Contradictory to these observations, in Toxoplasma gondii it has been reported that PUS1targeted mRNAs have an increased half-life in PUS1 mutant [73]. These examples highlight that Ψ can have a multifold effect on mRNA structure and function, and a better understanding of Ψ’s effect on mRNA stability will require a greater comprehension of how the other coupled mRNA processes, especially translation, are affected by the presence of Ψ. There is a growing body of evidence showing that Ψ can affect both the rate and fidelity of translation. Structural studies of the 70S ribosome with ΨUU in the A site and tRNAPhe bound to the ribosome have demonstrated that the 30 -CCA end of tRNAPhe is not properly positioned in the peptidyl-transferase center in the presence of Ψ despite correct mRNA: tRNA base-pairing interactions [74]. Kinetic data have further revealed that replacing a single uridine with Ψ impedes amino acid incorporation in vitro; this misincorporation of amino

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Modification

A. Alter RNA:protein interacon Nascent peptide

AAAA

mRNA cap

AAAAAA Poly(A)

tail

RNA-binding protein AAAA

B.

Influence RNA secondary structure AAAA

AAAA

C.

Influence decoding and terminaon

AAAA

AAAA

AA AAAA

Fig. 3 Pseudouridine affects mRNA in multiple ways. (a) The presence of Ψ provides rigidity to the mRNA backbone and can alter RNA–protein interactions. (b) Secondary structures in the mRNA can be stabilized in the presence of Ψ. (c) Presence of Ψ in a sense codon can lead to translation recoding, and in a nonsense codon can lead to nonsense suppression

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acids in presence of Ψ has been recapitulated in vivo using cellbased assays where complete substitution of U with Ψ in synthetic mRNA showed misincorporation of amino acids, albeit at low frequencies. Taken together, these studies indicate that Ψ-containing codons can slow the elongating ribosome. The magnitude by which the presence of Ψ in an mRNA can affect translation will likely depend on several parameters, including sequence context and mRNA secondary structures. The sequence context of the Ψ-containing codon can affect the interaction between the ribosome and the codon, as has been observed for inosinecontaining codons. Translation of the Ψ-containing mRNAs can also be affected by the indirect effect of Ψ on mRNA secondary structure, and RNA secondary structure mapping studies (e.g., using DMS or SHAPE reagents) comparing mRNAs with modified or unmodified nucleotides have now provided additional insights into how mRNA structure affects ribosome transit time. Complete substitution of uridines in an mRNA with N1-methyl-pseudouridine (m1Ψ) resulted in m1Ψ-containing mRNAs that have a higher translation efficiency that could, in part, be attributed to a highly structured CDS that extended the half-life of the mRNA [75]. Additionally, increased ribosome occupancy on m1Ψ-containing mRNAs can further facilitate translation initiation or ribosome recycling, resulting in enhanced overall translation from the mRNA [76]. The effect of modifications on the translation can thus be imparted by multiple mechanisms. Identifying the consequences of Ψ in mRNAs is complicated in cells because the enzymes that incorporate Ψ into mRNAs also catalyze Ψ addition in ncRNA species. Furthermore, the impact of Ψ on mRNA stability can be difficult to deconvolute from its effects on translation in cells. It is also unknown whether readers for Ψ exist or whether Ψ can be reverted back to U in vivo; the identification of Ψ reader and eraser proteins is needed to further our functional understanding of Ψ marks in mRNA. Similar to the other modifications discussed in this review, Ψ has been implicated in various human diseases, although a direct association of the modification with the disease state has not been elucidated.

6

Applications of RNA Modifications As has been discussed, chemical tags in RNAs impart an additional level of gene expression regulation, and modified nucleotides (or modified RNAs, per se) are being explored for a number of different applications, including extending their use to antisense oligonucleotides and miRNA derivatives to enhance the therapeutic potential of these RNA species. Because chemical modifications have the potential to alter RNA–RNA and RNA–protein interactions, chemically modified synthetic RNA-based biosensors are

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being evaluated for improved interactions between the modified RNA and the target molecule. Modified ribonucleosides or their chemical analogs (e.g., ribavirin, acyclovir, and valacyclovir) are being used as antiviral agents or as cancer therapies. Although the mechanism of action of these analogs varies, they typically mimic the canonical nucleosides in terms of uptake and incorporation into the host nucleic acids, which results in chain termination of the newly synthesized DNA, leading to incomplete replication and viral or cell death. 6.1 Synthetic mRNA Vaccines

A prominent application of chemical modifications in RNA is in synthetic mRNA-based therapeutics. mRNA-based therapeutics is based on the idea that a synthetic mRNA can be delivered in vivo and the mRNA can utilize the cellular machinery to generate the protein of interest. The field of RNA-based therapeutics is emerging as a promising new alternative to conventional proteinbased vaccine approaches [77, 78]. Use of mRNA as a therapeutic modality is advantageous because the molecule is noninfectious and the mRNA does not get integrated into the genome. Furthermore, the mRNA’s half-life can be regulated by the cellular machineries, thus making expression from the mRNA transient and amenable to regulation. In addition, the manufacturing process to produce an mRNA vaccine is rapid, inexpensive, and scalable, making it a lucrative platform. Even though the first report for the successful in vivo use of synthetic mRNA was published in the early 1990s, the idea failed to gain traction until more recently, mainly due to concerns associated with mRNA instability in vivo, low translation efficiency, immunogenicity concerns, and inefficient in vivo delivery. The pharmacokinetic (PK) and pharmacodynamic (PD) properties of mRNA-based therapeutics are affected by the half-life of the synthetic mRNA, which subsequently affects the efficacy of the drug. Several approaches have been used to increase the efficiency of protein synthesis from a synthetic mRNA, including optimization of the 50 UTR sequences to minimize secondary structures that could impair ribosome recruitment or transit, removal of alternate initiation codons in the 50 leader to maximize translation initiation from the main CDS, addition of stabilizer elements in the 30 UTR, elimination of miRNA-binding sites from the 30 UTR, and optimization of poly-A tail length and codon sequences (which affect translation efficiency and mRNA stability) [79–82]. Over the past decade, technological innovations together with an increased understanding of the interconnected mRNA processes have enabled advancement of synthetic mRNAs for therapeutics and vaccines—use of chemically modified mRNAs for therapeutics being at the forefront to address the issues of mRNA instability in vivo and immunogenicity of synthetic mRNAs [83].

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6.2 Modifications of the mRNA Termini

As discussed in previous sections, the mRNA 50 cap structure not only promotes translation via its interaction with the translation initiation factor 4E (EIF4E), but also modulates mRNA decay via interactions with the DCP1/DCP2 complex (Fig. 1) [3]. Synthetic mRNAs delivered in vivo interact with the same cytosolic proteins as an endogenous mRNA, and therefore, the interaction of the 50 cap with either the translation or the decay machinery is pivotal in determining the therapeutic efficacy of a synthetic mRNA molecule. An added level of complication arises from the fact that a synthetic mRNA can also encounter cytosolic immune receptors that recognize viral RNAs [19–21]. Translation efficiency and immune evasion can be enhanced by the use of cap analogs during mRNA synthesis that have chemical modifications, and several mimics of the 50 cap have been evaluated for these purposes [84– 88]. The most common cap analogs used are the anti-reverse cap analogs (ARCAs), which have modifications within the ribose moiety of the m7G cap [87]. ARCA possesses a 30 -O-methyl group on the sugar adjacent to the m7G, which prevents incorrect cap incorporation during the mRNA synthesis process, resulting in a higher percentage of capped mRNA species and an increased population of mRNAs that can be translated in vivo. A single phosphorothioate (O-to-S) substitution in the triphosphate bridge of the ARCA cap analog has been reported to increase the affinity of this cap structure for EIF4E, thereby reducing an mRNA’s susceptibility to DCP1/DCP2-mediated decapping and subsequent decay [86]. Because the presence of a 20 -O-methylation modification in Cap 1 prevents IFIT-1-induced translation inhibition, cap analogs that confer such structures are also routinely used to reduce binding to IFIT-1 and to subsequently prevent activation of an immune response. Systematic analyses of methylation status of the first nucleotide using cap analogs have further demonstrated that the identity of the first nucleotide and its methylation status in an mRNA influences the expression in a cell-type-specific manner [25]. Whether or not this increased translation is due to a direct effect on the binding of eIF4E or an indirect effect of reduced IFIT1 binding is not clear. Modifications at the 30 end of the mRNA also affect mRNA translation and stability (Fig. 1), and poly(A) tails of optimum length are added to synthetic mRNAs for efficient translation. For certain synthetic mRNAs, the combination of a cap modification with a poly(A) tail length of 100 residues has been shown to increase translation in dendritic cells. The correlation between poly(A) tail length and translation is not clear, but in naturally occurring mRNAs, regulation of poly(A) tail length is critical for maintaining specific biological behaviors. During eukaryotic mRNA degradation, deadenylation by 30 -exonucleases such as the poly(A)-specific ribonuclease (PARN) selectively deadenylates single-stranded mRNA via recognition of the 20 ,30 -adenosine diol

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intermediate [89]. Chemical modification of the end 20 ,30 -diol group in a synthetic Luciferase mRNA resulted in a threefold increase in protein synthesis due to an increase in transcript stability [90]. These findings suggest that chemical modification of the mRNA’s 30 end might interfere with the cellular interactions that lead to decay. Even though the mRNA half-life can be extended, whether or not these modifications alter the stable RNP formation or affect translation indirectly needs to be investigated. 6.3 Internal Modifications of the Synthetic mRNA

In vivo studies of synthetic mRNAs modified with Ψ have demonstrated increased translational rates and protein expression compared with mRNAs lacking Ψ [82, 91–95]. For example, injection of Ψ-modified erythropoietin (EPO) mRNAs into mice led to 10to 100-fold higher levels of expression from the modified mRNAs than from unmodified mRNAs, making this modification very attractive for the purposes of mRNA therapeutics. However, the effects of Ψ on protein expression have been quite varied, and translation enhancement was not observed in other systems. The discrepancies in the effects of Ψ on expression can be explained by differences in the sequences tested and differences in the innate immune responses in the target cells. In vitro, Ψ has been shown to repress translation when introduced at all three nucleotide positions in a codon. m1Ψ has been shown to enhance translation by increasing ribosome pausing, and consequently ribosome density, on synthetic mRNAs [76]. A correlation between protein expression and structure of the mRNA has also been demonstrated for synthetic mRNAs modified with m1Ψ, with the modified mRNAs having an increased functional half-life [75]. An mRNA with a combination of Ψ, m5U, and m1Ψ is currently being evaluated in preclinical research. The effect of Ψ on translation could also be indirect and mediated by interference with global protein synthesis shutdown during a cellular immune response. Uridine-containing mRNAs have been reported to activate protein kinase R (PKR)mediated phosphorylation of the α subunit of eukaryotic translation initiation factor 2 (eIF2α), thus leading to global downregulation of protein synthesis. In contrast, reduced phosphorylation of eIF2α has been observed with Ψ-containing mRNAs, suggesting that the increased translation is imparted by reducing the activation of PKR [96]. Reduced activation of 20 –50 -oligoadenylate synthetase (OAS) by Ψ-containing mRNAs and reduced 20 –50 -oligoadenylatemediated activation of RNase-L have also been observed. Increased resistance of the Ψ-containing mRNAs to RNase-L-mediated degradation further results in increased stability of the mRNA and, subsequently, in increased protein synthesis. Another modification that is currently being evaluated for its therapeutic potential is m5C, because earlier studies found that m5C incorporated in synthetic mRNAs stimulated expression from these transcripts in vivo.

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Immunostimulatory effects of synthetic mRNAs are imparted by two different structural elements—the cap structure (as discussed in the previous section) as well as the nucleotide content of the synthetic RNA. Synthetic RNA is recognized in nonimmune cells through the RIG-I receptor and in immune cells through the Toll-like receptors (TLRs). It has been observed that the presence of U-rich sequences in the RNA is a trigger for TLRs, and thus reduction of the U-content of the mRNA, as well as the substitution of uridine with modified uridine in the mRNA, is a strategy that has been implemented to evade recognition by the TLRs. Replacement of uridine with ψ, m1Ψ, m5U, 2-thiouridine (s2U), or 5-methoxyuridine (5moU), and replacing the natural cytidine with m5C, has demonstrated a reduction in the immune responses to synthetic mRNAs. It has also been hypothesized that the effect of uridine can be counterbalanced by altering the poly (A) tail to generate mRNAs with low immunogenicity because the U content decreases or is shielded in the sequence. In light of these findings, chemical modifications of mRNA are applied to not only modulate translation efficiency but also alter mRNA recognition by the innate immune system. Understanding these mechanisms underlying how modifications affect interrelated mRNA processes can aid in the rationalized design of efficacious synthetic mRNA molecules. Modifications can be introduced to affect the mRNA secondary structure without affecting the codon when required, and rationalized incorporation of the modifications can be done when minimal secondary structure changes are desirable. In addition, modifications can also be introduced sitespecifically to affect immunogenicity without affecting decoding. In a cellular context, destabilization of the local secondary structure (as seen with m6A) alters the accessibility of RNA-binding proteins to specific sequence elements; the same can be achieved for synthetic mRNAs. Decoupling structural stability from primary sequence changes can be instrumental in altering the expression from the synthetic mRNA.

7

Approaches to Understand the Effects of mRNA Modifications on Translation As we continue to identify modifications in mRNAs, the next question that needs to be addressed is how these modifications affect the mRNA life cycle. The effects of modifications on the mRNA life cycle are complicated to tease out in a biological context, i.e., in vivo, because the modifications are installed not only in the mRNA but also in other noncoding mRNAs by the same set of writer proteins. Therefore, cell-based assays using synthetic modified mRNAs are routinely implemented to evaluate the effect of mRNA modifications. Protein expression from the transfected mRNAs in cell-based assays is measured a few hours after

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transfection, making it difficult to study the direct effects of the modifications; also, the kinetics of protein synthesis are difficult to analyze. Transfection of synthetic mRNAs also results in degradation of a significant proportion of the mRNAs in endosomes, and the final readout does not reflect the actual efficiency of protein expression. Differences in delivery modes, cellular uptake, and endosomal targeting can affect the mRNA pool for translation. Furthermore, deconvoluting the effects of mRNA modifications on translation versus their effects on mRNA stability is not trivial. Because chemical modifications of mRNAs have been shown to affect mRNA translation and stability, it is critical that cell-based assays be combined with in vitro assays that can help tease out the function of the mRNA modification. 7.1 mRNA Termini Modifications

The discovery of diverse modifications in endogenous mRNA caps has provided versatility to mRNA function, and because the cap structure helps distinguish endogenous mRNAs from viral RNAs, the 50 termini of synthetic mRNAs are routinely modified to recapitulate these endogenous structures [14, 15]. The biological significance of the 50 cap variations is routinely evaluated using in vitro-synthesized RNA in either cell-based or biochemical assays to understand the effects of the modifications in enhancing translation and evading immune responses [25, 88, 97, 98]. There are some considerations that should be taken into account when utilizing cell-based assays in combination with synthetic modified mRNAs. Current methods involve using cap analogs during in vitro transcription to introduce the cap modifications at the 50 end of the mRNA. However, the incorporation of the cap analog is not always efficient, and the resulting RNA population can have a mix of capped and uncapped RNAs [86, 87]. Furthermore, the purity of the in vitro-transcribed RNA can also affect the immune response generated against the mRNA preparation, which can subsequently affect its translation efficiency [99, 100]. There are multiple parameters that can play a role in expression from an exogenously delivered mRNA. Interaction with cellular immune receptors (RIG-I, MDA5, IFIT-1) is dependent on the nature and the modification status of the 50 cap and the presence of impurities in the synthetic mRNA can also activate PKR and 20 -50 OAS [96]. PKR-mediated phosphorylation of eIF2α represses translation initiation and 20 -50 OAS-mediated activation RNase-L can lead to degradation of cellular RNAs and a subsequent shutdown of global protein biosynthesis [96]. Teasing out the direct effects of the 50 cap modification in translation (via binding to the cytoplasmic CBC, specifically, eIF4E) versus the indirect effects due to global protein synthesis shutdown is critical. Not choosing the right cell type for cell-based assays can also bias the results; in a recent study evaluating the effect of modifications in the first

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transcribed nucleotide in the RNA, it was observed that the effect of the methylation status of the nucleotide after the cap is strongly dependent on the type of cultured cells [25]. Biochemical assays to determine the binding kinetics allow determination of apparent KD; however, they do not give insights into the kinetics of IFIT1–RNA interactions. Biophysical assays allowing direct mRNA–protein binding have also been used in combination with cell-based assays. Binding of the 50 ends of mRNA with either the cap-binding protein eIF4E, the cytosolic receptors (IFIT1), or the decapping enzyme (DcpS) has been evaluated to determine the kinetics of these interactions and deduce a role of the 50 cap modification [25, 97]. 7.2 Internal Modifications

Cell-free in vitro translation systems, such as those derived from Krebs cells, HeLa cells, and rabbit reticulocyte lysates, have been used to understand the effects of RNA modifications on translation and to recapitulate the increase in protein expression seen using the modified mRNAs in cell-based assays [34, 74, 76, 83, 93– 96]. There are discrepancies in the studies using synthetic modified mRNAs, and an emerging body of evidence is making clear that the translation systems and the cell types used for evaluating the effects of modifications can have an impact on the final outcome. The effect of m5C on translation yield has been found to be quite different when tested in rabbit reticulocyte extracts, wheat germ extracts, or bacterial translation systems [49, 76, 101]. Discrepancies may also arise because of differences in sequence contexts of the modifications in a synthetic mRNA. A concerted evaluation of multiple mRNAs—including mRNAs of varied length, structure, and sequence—would be required to get a comprehensive idea of the effects of modifications in vitro. Current cell-based or in vitro assays have utilized approaches where the entire body of the mRNA has been modified with a specific analog, which does not reflect the modification status of a naturally occurring mRNA, and it should not be surprising that the consequences of modifying the entire mRNA will be quite distinct from the consequences of introducing a modification at a single position in a codon. In order to circumvent the unnatural representation of modifications in these synthetic mRNAs, systematic modification of the first, second, or third position of the codon has been performed, and the peptide products of the corresponding mRNAs have been analyzed [101]. Site-specific introduction of modifications in the CDS of the mRNA followed by translation in HEK293 cells revealed a strong position-dependent effect of modifications on translation efficiency and accuracy. Whereas single m5C or Ψ modifications did not have any effect on product formation, m1A impeded translation, and a codon-position-dependent effect on translation was observed for 20 -O-methylated nucleotides and m6A. Miscoding in

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I. In vitro transcription of modified mRNA AAAA

II. Transfection into HEK293 cells

Modification

FLAG HA

III. Translation in HEK293 cells followed by purification of full-length protein

*

IV. Identification of amino acid misincorporation by MS analyses

mRNA cap AAAAAA Poly(A)

tail

* Protein of choice

Fig. 4 Experimental procedure to test translation recoding from modified mRNAs (containing pseudouridine) in vivo. The modification is incorporated during in vitro transcription, and the mRNA is transfected into HEK293 cells. After optimal expression, the protein is purified using an N-terminal hemagglutinin (HA) and a C-terminal FLAG tag, ensuring enrichment of full-length protein (normal and recoded) and not prematurely terminated products. The purified protein is then subjected to digestion and mass spectrometry (MS) analyses to identify amino acid misincorporation

the presence of modification can be measured either in vitro or using cell-based assays. Eukaryotic translation is highly accurate, and miscoding in the presence of modifications seems unlikely. However, in a recent study, in vitro biochemical and structural studies were combined together with cell-based assays to demonstrate miscoding in the presence of Ψ (Fig. 4, [101]). Misincorporation in the presence of m5C has also been reported in bacterial systems [49]. However, in vivo experiments in human cells have yielded conflicting results for miscoding that can be attributed to the sensitivity of the methods implemented, as well as to the sequence contexts of the codons. Understanding how mRNA modifications impact translation is critical, but it is also important to gain quantitative insights into how mRNA–protein interactions affect the mRNA life cycle. Alteration of protein–mRNA interactions has been investigated for m6A using a combination of immunoprecipitation and pull-down assays, mass spectrometry, and RNA-seq approaches. m6A can both weaken and enhance RNA–protein interactions. The YTH family of m6A reader proteins differentially interact with methylated and unmethylated transcripts, a discrimination more prominent than for other RNA-binding proteins [102]. Unlike m6A, reader proteins for other mRNA modifications are not known yet, and the plausible interactions between the translation and mRNA decay machineries have not yet been mapped.

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Future Perspectives The identification of modifications in mRNA has unveiled an uncharted layer of regulation of RNA stability and translation. The study of mRNA modifications is bound to grow rapidly as researchers seek to understand the influence of these modifications on human health and biology. Advancements in analytical chemistry and deep sequencing-based methods have furthered our understanding of RNA modifications. Future work combining the quantitative transcriptome-wide mapping of modifications with in-depth characterization of the writer, reader, and eraser proteins using structural biology and in vitro biochemistry will be pivotal in getting a comprehensive landscape of mRNA modifications and their function in the regulation of gene expression. Structural and biochemical analyses to unveil modification-induced changes in mRNAs, and the factors driving the selection of modification sites and the stoichiometry of the modifications, are all outstanding questions in the field, and their answers will enable a better understanding of how cellular machineries encounter the modifications and the consequences of such an event. Most of the current studies have been performed using single types of modifications, and the combinatorial effect of different types of modifications when present in the same mRNA will be of interest. How mRNA modifications contribute to a specific disease state can only be addressed when a comprehensive understanding of the molecular mechanisms is available. A better understanding of the correlation between mRNA modifications and the regulation of factors involved in the dynamic control of these modifications will help screening of inhibitors of mRNA modification regulators that will open up new therapeutic avenues. It is not surprising that new mechanisms of mRNA modification will be discovered as the analyses of modifications are refined and technological advances are made.

Acknowledgments I would like to thank Monica Z. Wu for help with the illustrations and New England Biolabs Inc. for supporting the basic research on mRNA modifications. Funding for open access was provided by New England Biolabs Inc.

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Chapter 21 Assaying the Molecular Determinants and Kinetics of RNA Pseudouridylation by H/ACA snoRNPs and Stand-Alone Pseudouridine Synthases Dominic P. Czekay, Sarah K. Schultz, and Ute Kothe Abstract Posttranscriptional modifications of RNA play an important role in promoting the maturation and functional diversity of many RNA species. Accordingly, understanding the enzymes and mechanisms that underlie RNA modifications is an important aspect in advancing our knowledge of the continually expanding RNA modification field. However, of the more than 160 currently identified RNA modifications, a large portion remains without quantitative detection assays for their biochemical characterization. Here, we describe the tritium release assay as a convenient tool allowing for the quantitative assessment of in vitro RNA pseudouridylation by stand-alone or box H/ACA RNA-guided pseudouridine synthases. This assay enables quantification of RNA pseudouridylation over a time course to effectively compare pseudouridylation activity between different substrates and/or different recombinant enzymes as well as to determine kinetic parameters. With the help of a quench-flow apparatus, the tritium release assay can be adapted for rapid kinetic measurements under single-turnover conditions to dissect reaction mechanisms. As examples, we show the formation of pseudouridines by a reconstituted Saccharomyces cerevisiae H/ACA small ribonucleoprotein (snoRNP) and an Escherichia coli stand-alone pseudouridine synthase. Key words Pseudouridine, RNA modification tritium release assay, Pseudouridine synthase, H/ACA small ribonucleoprotein, Enzyme kinetics

1

Introduction Pseudouridine is an abundant modification of uridine occurring in all domains of life [1]. It is found in all RNA species including transfer RNA (tRNA), small nuclear RNA (snRNA), ribosomal RNA (rRNA), and messenger RNA (mRNA) [2–5]. Pseudouridine is a structural isomer of uridine, containing a noncanonical C-C glycosidic bond (instead of C-N) as well as an additional imino group at position 5 of the modified nucleobase (Fig. 1). The formation of pseudouridine is catalyzed by either stand-alone enzymes or H/ACA small nucleolar ribonucleoprotein (snoRNP)

Mary McMahon (ed.), RNA Modifications: Methods and Protocols, Methods in Molecular Biology, vol. 2298, https://doi.org/10.1007/978-1-0716-1374-0_21, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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Fig. 1 Isomerization of uridine to pseudouridine. Pseudouridine synthases catalyze the isomerization of uridine to pseudouridine. For the tritium release assay, the uridine is radiolabeled at position C5 with a tritium (3H, highlighted in red) which is released into solution upon formation of the new C10 –C5 glycosidic bond

complexes, which recognize a target RNA sequence by base-pairing with a box H/ACA RNA [6–8]. Following target selection, pseudouridylation begins with a universally conserved aspartate residue in the pseudouridine synthase attacking the C20 position of the ribose sugar [9]. This attack breaks the C-N glycosidic bond and forms a glycal intermediate [10]. After reorienting the nucleobase and forming a new C10 –C5 glycosidic bond, the resulting pseudouridine nucleobase appears as mirrored across the N3–C6 plane when compared to uridine (Fig. 1). The tritium release assay exploits the fact that during pseudouridylation, a hydrogen attached to the C5 of uridine is released into solution. Replacing hydrogen at this position with the radioactive isotope tritium (Fig. 1, highlighted in red) forms the foundation of the tritium release assay as each released tritium corresponds exactly to the formation of one pseudouridine. Following the release of tritium, radioactively labeled RNAs and proteins are removed by absorption to activated charcoal, which is then separated from any released tritium in solution via centrifugation. The tritium content in the supernatant is quantified with a liquid scintillation analyzer providing a direct measurement of target RNA pseudouridylation (Fig. 2). Importantly, the tritium release assay effectively allows for temporal quantification of RNA pseudouridylation as several time points can be easily processed in parallel. The tritium release assay can be used under multiple-turnover (substrate excess) conditions in order to determine the initial velocity of the reaction. When measuring the initial velocity at increasing concentrations of substrate RNA, Michaelis-Menten kinetic parameters can be determined to quantitively compare, for example, different substrate

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Fig. 2 Overview of the tritium release assay. As an example, pseudouridine formation by an S. cerevisiae H/ACA snoRNP complex is depicted. First, the H/ACA snoRNP is reconstituted from purified proteins and a refolded H/ACA RNA. Second, incubation of the H/ACA snoRNP with [3H-5]-uridine-containing RNA results in pseudouridine formation in vitro. Third, the reaction is stopped with HCl, and the radiolabeled RNAs and proteins are extracted by absorption to charcoal and centrifugation. Fourth, the tritium, that is released during pseudouridylation, remains in the supernatant and can be quantified by liquid scintillation counting such that the time course of pseudouridylation can be determined

RNAs or different enzyme variants [11–13]. Alternatively, the tritium release assay allows for measurement of single-turnover reactions (enzyme excess) in order to identify and characterize individual reaction steps of a pseudouridine synthase [13, 14]. Under these conditions, the reaction is often very fast such that time courses need to be followed in the milliseconds-toseconds range which can be achieved using a quench-flow apparatus. In brief, a quench-flow apparatus is a rapid-mixing device where substrate and enzyme are combined within less than three milliseconds, the reaction is allowed to proceed for a desired time in the millisecond-to-second range, and subsequently the reaction is stopped by rapid mixing with a quenching solution. For example, this approach often allows direct determination of the rate constant of the catalytic step independent of substrate RNA binding to the enzyme. This provides direct insight into the reaction mechanism and characterization of active-site enzyme variants.

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As with every assay, the tritium release assay has some limitations. Most importantly, the tritium release assay cannot determine the position at which pseudouridylation occurs within an RNA strand. This information must be known prior to the experiment or needs to be verified in complementary experiments, e.g. the suspected target uridine can be mutated to cytosine leading to a loss of tritium release. Moreover, the tritium release assay is inherently a radioactive enzyme assay requiring skilled workers and a safe work environment where additional safety measures and documentation are needed. As further explained in Subheading 3, another limitation of the tritium release assay is that it depends on the preparation of [C5-3H]U-labeled RNA through in vitro transcription. Conveniently, RNA that has been specifically prepared for the tritium release assay can be used over a long time period due to the long half-life of tritium (12.5 years).

2

Materials Prepare reagents using ultrapure RNase-free water as well as RNA-grade chemical reagents.

2.1 In Vitro Transcription of [5-3H]-Substrate RNA

1. 5 Transcription buffer: 200 mM Tris–HCl, pH 7.5, 75 mM MgCl2, 10 mM spermidine, 50 mM NaCl. To prepare, add 2.42 g of Tris, 1.52 g of MgCl2·6H2O, 0.145 g of spermidine, and 0.292 g of NaCl to 80 mL of RNase-free water in a graduated cylinder. Mix to dissolve. Adjust pH to 7.5 with HCl. Add RNase-free water to a final volume of 0.1 L. Syringe filter through 0.22 μm pore size into 1 mL aliquots and store at 20  C. 2. 100 mM DL-dithiothreitol (DTT): Dissolve 0.154 g of DTT in 10 mL of RNase-free water and syringe filter through 0.2 μm pore size generating 1 mL aliquots. 3. 100 mM Adenosine-50 -triphosphate: Dissolve 1 g of ATP disodium salt hydrate in 3 mL of 500 mM magnesium acetate pH 7.5, adjust the pH to 7.5 using KOH, then dilute to 100 mM (based on A260 measurements), and store in 100 μL aliquots. 4. 100 mM Cytosine-50 -triphosphate (CTP): Dissolve 1 g of CTP disodium salt in 3 mL of 500 mM magnesium acetate pH 7.5, adjust the pH to 7.5 using KOH acid, then dilute to 100 mM (based on A260 measurements), and store in 100 μL aliquots. 5. 100 mM Guanosine-50 -triphosphate (GTP): Dissolve 1 g of GTP sodium salt hydrate in 3 mL of RNase-free water, adjust the pH to 7.5 using KOH, then dilute to 100 mM (based on A260 measurements), and store in 100 μL aliquots.

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6. 100 mM Uridine-50 -triphosphate (UTP): Dissolve 1 g of UTP trisodium salt hydrate in 3 mL of 500 mM magnesium acetate pH 7.5, adjust the pH to 7.5 using KOH, then dilute to 100 mM (based on A260 measurements), and store in 100 μL aliquots. 7. [5-3H]-uridine-50 -triphosphate tetraammonium salt (Moravek Radiochemicals)—supplied as a 1 μCi/μL solution in 50% ethanol. 8. 100 mM Guanosine-50 -monophosphate (GMP): Dissolve 0.4072 g of guanosine-50 -monophosphate disodium salt in 10 mL of RNase-free water and syringe filter through 0.2 μm pore size, preparing 1 mL aliquots. 9. 0.5 U/μL Inorganic pyrophosphatase (IPPase) from S. cerevisiae: Add 200 μL of 5 transcription buffer, 280 μL of RNasefree water, 20 μL of 100 mM DTT, and 500 μL of glycerol to 500 units of IPPase from baker’s yeast to dissolve the enzyme. 10. DNA template for RNA to be analyzed: See Table 1 for sequences of DNA templates used to generate a substrate RNA within S. cerevisiae 25S rRNA. 11. T7 RNA polymerase. 12. 40 U/μL Ribolock RNase inhibitor. 13. RNase-free DNase I. 14. RNase-free 1.5 mL/2 mL tubes. 15. Heat block. 16. Vortex. 17. Temperature-controlled water bath. 18. Microcentrifuge.

Table 1 DNA oligo sequences used as in vitro transcription templates to generate S. cerevisiae 25S rRNA fragments Primer

Sequence (50 –30 )

snR34 30 hairpin sub_sense

GCTAATACGACTCACTATAGGGGACAACTGGCTTGTGGCTGCCT

snR34 30 hairpin sub_antisense

mAmGGCAGCCACAAGCCAGTTGTCCCCTATAGTGAGTCGTATTAGC

0

snR34 5 hairpin sub_sense

GCTAATACGACTCACTATAGGGACGTCGGCTCTTCCTATCATACC

snR34 50 hairpin sub_antisense

mGmGTATGATAGGAAGAGCCGACGTCCCTATAGTGAGTCGTATTAGC

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2.2 Crush and Soak Purification of [5-3H]-Labeled Substrate RNA

1. 2 RNA loading dye: 95% Formamide, 0.025% bromophenol blue, 0.025% xylene cyanol, 0.025% sodium dodecyl sulfate (SDS), 0.5 mM EDTA. To 900 μL of formamide add 25 μL each of 1% (w/v) bromophenol blue in formamide and 1% (w/v) xylene cyanol in formamide, 2.5 μL of 10% SDS, 1.0 μL of 0.5 M EDTA, and 46.5 μL of RNase-free water. Mix well by vortexing. Store at room temperature (see Note 1). 2. 10 TBE buffer: 900 mM Tris, pH 8.3, 900 mM boric acid, 10 mM EDTA. In a 1 L graduated cylinder, dissolve 108 g of Tris base, 55 g of boric acid, and 3.36 g of EDTA disodium salt in 800 mL of RNase-free water. Adjust pH to 8.3 as needed. Filter sterilize through a 0.22 μm pore size into a 1 L glass bottle and store at 4  C. 3. 15% Polyacrylamide gel: Combine 0.6 mL of 10 TBE, 2.25 mL of 40% acrylamide:bis-acrylamide (19:1), 2.88 g of urea, RNase-free water to 6 mL, 25 μL of 10% APS, and 5 μL of TEMED. 4. Polyacrylamide electrophoresis equipment (gel chamber, lid, running module, glass plates, power supply, etc.). 5. Isopropanol. 6. 3 M Sodium acetate, pH 5.2. 7. Heat block. 8. Syringe. 9. Gel loading pipette tips. 10. Plastic (Saran) wrap. 11. Handheld UV lamp. 12. Thin-layer chromatography plate—silica gel 60 F254 (EMD Millipore). 13. Scalpel or razor. 14. 15 mL Tubes. 15. Tabletop centrifuge (for 15 mL tubes). 16. Rocking platform. 17. Phenol, pH 4.5. 18. Chloroform. 19. UV-visible spectroscopy spectrophotometer.

2.3 Tritium Release Assay

cuvette

and

absorbance

1. 5 Tritium release reaction buffer: 100 mM HEPES-KOH, pH 7.4, 750 mM NaCl, 0.5 mM EDTA, 7.5 mM MgCl2, 50% (v/v) glycerol, 3.75 mM DTT. Mix 250 mL of glycerol and 150 mL of RNase-free water in a graduated cylinder. Add 11.9 g of HEPES, 21.9 g of NaCl, 0.5 mL of 0.5 M EDTA solution, 0.762 g of MgCl2·6H2O, and 0.289 g of DTT. Mix

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to dissolve. Adjust pH to 7.4 with KOH. Add RNase-free water to a final volume of 0.5 L. Vacuum filter through 0.22 μm pore size. Store at 20  C (see Note 2). 2. 0.1 M HCl, 5% (w/v) activated charcoal: In a 1 L glass bottle, mix together approximately 700 mL of RNase-free water with 8.3 mL of 12 M HCl. Add 50 g of activated charcoal Norit® using adequate ventilation when handling activated charcoal Norit®. Add RNase-free water to the 1 L mark on the bottle. Store at room temperature (see Note 3). 3. Glass wool: Handle with gloves, lab coat, and adequate ventilation. 4. 6 mL Liquid scintillation vials. 5. EcoLite(+)™ Liquid Scintillation Cocktail (MP Biomedicals). 6. Liquid scintillation analyzer. 7. Purified pseudouridine synthase of interest (stand-alone enzyme or H/ACA snoRNP). 2.4

Quench Flow

1. Quench-flow (KinTek model RQF-3). 2. Circulating water bath (VWR 1156D). 3. Peristaltic pump (GE Pump P-1). 4. 0.1 M HCl. 5. 1.5 mL Microcentrifuge tubes. 6. Single-use 10 mL Luer-Lock syringes. 7. Single-use 1 mL Slip-Tip syringes. 8. Hypodermic needles. 9. Two 100 mL beakers.

3

Methods Unless otherwise stated, keep samples on ice. Change gloves regularly to reduce the likelihood of RNase contaminations. Safety considerations: When working with radioactive [5-3H]labeled UTP and RNA, appropriate safety procedures must be followed including wearing personnel protective equipment, detecting contaminations, documenting the use of radioactive material, and following guidelines for disposal of radioactive waste. Inform yourself of the institutional requirements and adhere closely to them.

3.1 In Vitro Transcription of [5-3H]-Substrate RNA

1. Template DNA for in vitro transcription of RNAs shorter than 40 nt in length is prepared by annealing complementary DNA oligos to one another as follows: boil (100  C, 5 min) and then passively cool to room temperature equivalent amounts

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(in pmole) of sense and antisense DNA oligos in 1 transcription buffer at a concentration of 1 μM each. Annealed DNA oligos are added directly into the in vitro transcription reaction. For substrate RNAs up to 100 nt in length, template DNA is assembled by first annealing partially complementary DNA oligos (as described above) and then performing a polymerase chain reaction (PCR) to extend the oligos followed by PCR cleanup. Template DNA for RNAs longer than 100 nt is generated from a suitable plasmid by PCR amplification followed by PCR cleanup. Exemplary oligonucleotides for the generation of two short S. cerevisiae 25S rRNA substrates are shown in Table 1 (see Note 4 for details on oligonucleotide design including information on the T7 promoter sequence). 2. For the in vitro transcription reaction, combine the following components to the indicated final concentrations in a 2 mL tube: 1 transcription buffer; 10 mM DTT (added from a 100 mM stock; see Note 5); 3 mM each ATP, CTP, and GTP (see Note 5); 0.1 mM of [5-3H]UTP/UTP mixture (see Note 6); 5 mM guanosine monophosphate (see Note 5); 0.01 U/μL inorganic pyrophosphatase from baker’s yeast (added from a 0.5 U/μL stock; see Note 7); 0.3 μM T7 RNA polymerase (see Note 8); 0.2 U/μL RiboLock RNase Inhibitor; and 1–2 ng/μL template DNA (from step 1 above). Mix gently until homogenous. The recommended reaction volume is 1 mL. 3. Incubate the in vitro transcription reaction at 37  C for 1–4 h with intermittent gentle mixing every 30 min. Add 2 U DNase I per 1000 μL reaction and incubate for an additional 1 h at 37  C to digest the DNA template. Remove excess inorganic pyrophosphate by centrifuging at 10,000  g for 10 min, and then pipette the supernatant into a new RNase-free 2 mL tube (see Note 9). 3.2 Purification of [5-3H]-Substrate RNA via Crush and Soak PAGE Purification

Here, a convenient and widely used method for gel extraction of RNA is described. Any other purification method that efficiently removes free nucleotides is equally suitable such as anion-exchange chromatography (e.g., disposable columns) or size-exclusion chromatography (e.g., spin columns) [13]. 1. To facilitate gel loading, reduce the volume of the in vitro transcription sample by performing an RNA precipitation as follows: Add 1/10 volume of 3 M sodium acetate and 1 volume of isopropanol (or 2.5 volumes of 100% ethanol). Mix and incubate overnight at 4  C. 2. Collect RNA by centrifuging at 10,000  g for 20 min. Remove supernatant with a pipette being careful not to disrupt the pellet (see Note 10). Add 1 mL of 70% ethanol to the pellet and vortex until resuspended. Repeat the centrifuge step as

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before, remove as much supernatant as possible with a pipette, and let the pellet air-dry until no visible liquid is left (see Note 11). Resuspend the pellet by adding 100 μL of RNase-free water and add 100 μL of 2 RNA loading dye. If required to help dissolve the pellet, add an additional 50 μL of RNase-free water and 50 μL of 2 RNA loading dye. Avoid exceeding a total volume of 300 μL. 3. Cast a 1 mm thick denaturing polyacrylamide gel in 1 TBE with a single flat well by pouring the gel solution until it reaches 1 cm below the height of the small plate and layering 250 μL of 100% isopropanol on the top during polymerization. For RNAs shorter than 40 nt, a 15% acrylamide concentration is ideal. 4. Once the gel has polymerized, wash out the isopropanol with RNase-free water, and then assemble the polymerized gel in the electrophoresis apparatus. Add a total volume of 600 mL of pre-warmed 1 TBE buffer in the upper and lower chambers of the system. Warm the gel by running at 300 V for at least 30 min prior to sample loading. 5. Heat the in vitro transcription sample containing loading buffer (step 2) to 70  C for 10 min, and snap cool immediately on ice for 3 min. Double-check to ensure that there is no precipitate forming in the RNA sample. Use a syringe or 1 mL pipette to clean the wells removing air bubbles and residual buffer containing urea; this helps greatly with sample loading. Use a gel loading tip to carefully pipette the entire in vitro transcription sample evenly across the gel surface. 6. Run the gel at a constant voltage of 300 V (37.5 V/cm) for an appropriate amount of time based on the gel percentage and RNA size. For 20–50 nt RNAs, a denaturing 15% acrylamide gel for 30 min at 300 V should center the RNA in the middle of the gel. Gently pull the plates apart using a wedge and sandwich the gel between two clear sheets of plastic (Saran) wrap. 7. Place the wrapped gel directly over an F254 silica gel thin-layer chromatography plate and expose to a 254 nm UV wavelength light source (see Note 12). By absorbing the UV light, the RNA should generate a visible dark shadow on the TLC plate, effectively allowing for RNA to be visualized without the use of stains. Using a permanent marker, draw a line on the plastic wrap to indicate where the RNA band is located. 8. Turn off the UV source to minimize damage of the RNA by UV light. Flip the gel to the other side and remove the top layer of plastic wrap being careful not to shift the position of the gel relative to the marked plastic wrap underneath the gel. Using a sterile scalpel or razor, excise the gel piece containing RNA by

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cutting along the permanent marker line. Slice the resulting long strip of gel into small segments and transfer to a 15 mL tube. 9. Clean the shaft and tip ejector of a P1000 pipette using ethanol. With a P1000 pipette tip, pulverize the gel pieces inside the 15 mL falcon tube into a fine powder. Do NOT discard the pipette tip at this step as it will have collected small pieces of gel on the inside. 10. Add 4 mL of filter-sterilized 1 TBE and 200 U of RiboLock RNase Inhibitor to a final concentration of 0.05 U/μl to the crushed gel pieces and pipette up and down a few times to flush the gel pieces from the P1000 pipette tip into solution before discarding the pipette tip. Incubate the content of the tube overnight at room temperature on a platform rocker. The RNA inside the gel will passively diffuse out of the gel and into the supernatant. 11. The next day, pellet the gel pieces by centrifuging at 10,000  g for 15 min. Transfer the entire supernatant to a small syringe attached to a 0.2 μm pore size filter. Push the contents through the filter into a clean, sterile 15 mL tube to remove small pieces of gel. 12. Perform an RNA phenol:chloroform extraction as follows: create a 1:1 mixture of acidic phenol:chloroform and add 1 volume to the filtered RNA sample in a 15 mL tube. Vortex for 15 s, and then centrifuge at 10,000  g for 2 min at room temperature. Carefully remove the tube from the centrifuge to avoid mixing the organic and aqueous phases. Use a pipette to transfer the upper (aqueous) phase into a fresh 15 mL tube. 13. Extract the RNA once more with 1:1 acidic phenol:chloroform and then once with chloroform alone, following the steps described above. Be careful to avoid transfer of chloroform during the final aqueous-phase removal. 14. Perform an isopropanol precipitation of RNA as described in Subheading 3.2, steps 1 and 2. Resuspend the RNA pellet into 20–100 μL RNase-free water (see Note 13). 15. Determine the specific activity of the purified [5-3H]-substrate RNA as follows: perform triplicate absorbance measurements at a wavelength of 260 nm and calculate the concentration (c) using the extinction coefficient (ε260) of the RNA (calculated using IDT OligoAnalyzer Tool) and Lambert-Beer law: A260 ¼ ε260 * c * d where d is the path length. 16. After each measurement, quantitatively transfer the measured RNA from the cuvette into a 6 mL scintillation vial and determine its radioactivity (in disintegrations per minute, dpm) using a liquid scintillation analyzer. Using the RNA

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concentration and activity of each measured RNA sample, calculate the specific activity of [5-3H]-substrate RNA as dpm/pmole∙RNA (see Note 14). Then divide the specific activity by the number of uridines in the specific RNA sequence to obtain a specific activity of dpm/pmole uridine. 3.3 Tritium Release Assay

The following protocol describes quantifying pseudouridylation of an rRNA fragment by a S. cerevisiae H/ACA snoRNP. As indicated, other stand-alone pseudouridine synthases and different [5-3H]labeled substrate RNAs such as tRNAs can be used in this assay [12–16]. 1. Prepare folded H/ACA snoRNA as follows: dilute purified box H/ACA snoRNA into 1 reaction buffer to a final concentration of 1.0 μM. Unfold diluted box H/ACA guide RNA by heating at 65  C for 5 min on a heat block. Remove from heat block and slowly cool for 15 min at room temperature to allow for refolding of box H/ACA guide RNA. 2. Allow formation of the reconstituted H/ACA snoRNP by combining the folded H/ACA snoRNA with a 2.2-fold excess of Cbf5 (dyskerin), Nop10, Gar1, and Nhp2 proteins in 1 reaction buffer into a 1.5 mL microcentrifuge tube and mix gently by pipetting (see Note 15). Incubate the H/ACA snoRNP for 10 min at 30  C in a water bath to allow formation of the H/ACA snoRNP. For stand-alone enzymes, prepare a dilution of the enzyme in 1 reaction buffer. As a negative control, include a sample that lacks the pseudouridine synthase, but is processed identically otherwise. 3. Prepare [5-3H]-substrate RNA by diluting into 1 reaction buffer to a final concentration of 1.0 μM. Unfold [5-3H]substrate RNA by heating at 65  C for 5 min in a heat block. Remove from heat block and slowly cool for 10 min at room temperature to allow RNA refolding. 4. To initiate the pseudouridylation reaction, add [5-3H]-substrate RNA to each sample tube containing the H/ACA snoRNP or a stand-alone pseudouridine synthase (see Note 16). Mix gently by pipette and continue incubating at 30  C for enzymes from S. cerevisiae and 37  C for enzymes from E. coli or Homo sapiens. 5. During the reaction, remove samples and quench by adding 1 mL of pre-aliquoted 0.1 M HCl and 5% (w/v) activated charcoal (see Notes 17 and 18). Vortex the quenched samples and incubate for at least 10 min at room temperature. Following the incubation, centrifuge the quenched samples at 10,000  g for 2 min to pellet the charcoal.

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6. Using a micropipette, transfer 800 μL of the supernatant into 500 μL of pre-aliquoted 0.1 M HCl and 5% (w/v) activated charcoal. Vortex well and incubate for 10 min at room temperature. Pellet charcoal as described previously. 7. Remove and then filter the entire supernatant through a glass wool plug into a fresh 1.5 mL tube (see Note 19). Centrifuge filtrate at 10,000  g for 2 min to pellet any charcoal that was not removed during filtration through the glass wool. Be careful to avoid any pelleted charcoal and transfer 800 μL of clear supernatant into a clean 6 mL liquid scintillation vial. 8. As a control, remove 2 μL of the pseudouridylation reaction directly to determine the total radioactivity present in the reaction by scintillation counting. This should be done in triplicate. 9. Add 4 mL of EcoLite(+)™ Liquid Scintillation Cocktail (MP Biomedicals) to all samples, vortex for 30 s, and then measure the radioactivity of the released tritium (in disintegrations per minute, dpm) using a liquid scintillation analyzer. 3.4

Data Analysis

1. Tritium release assay results can be represented as the concentration of pseudouridine formed or as the percentage of pseudouridine formation (Y) versus reaction time (X) (Fig. 3). The calculation of percentage of pseudouridine formation from a measured radioactivity (dpm) is explained here. After processing of the sample (charcoal extraction, filtration), only a fraction (F) of 0.492 of each original sample volume is subjected to

Fig. 3 Multiple-turnover pseudouridine formation of an excess of substrate RNA by S. cerevisiae H/ACA snoRNP. The tritium release assay was used to monitor pseudouridylation of two different short substrate RNAs that comprise different fragments of 25S rRNA by in vitro-reconstituted yeast snR34 H/ACA snoRNP. The two different substrate RNAs bind to the 30 and 50 hairpins of the snR34 H/ACA snoRNA, respectively [15, 16]

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liquid scintillation counting (see Note 20). Using the sample radioactivity (A in dpm) measured by the liquid scintillation analyzer, the sample volume (V0), and the fraction analyzed (F), the tritium released in dpm/μL of reaction volume (A[3H released]) is then calculated:   A 3 H released ¼ A=ðF ∗V 0 Þ Next, the concentration of released tritium, i.e., the concentration of pseudouridine formed (c[Ψ ] in μM), is determined with the help of the specific activity of the [5-3H]substrate RNA (aRNA in dpm/pmole uridine):   c ½Ψ  ¼ A 3 H released =a RNA If desired, the percentage of pseudouridine formation can be obtained by dividing the concentration of pseudouridine formed by the RNA concentration in the reaction (see Note 21). 2. The initial velocity of the reaction (v0) is the slope of the pseudouridine formation time course. To calculate the initial velocity, plot the concentration of pseudouridine formation versus time. Select only the data points where less than 30% of pseudouridines are formed and determine the slope by linear regression. Determining the initial velocity at different substrate RNA concentrations [S] allows generating a MichaelisMenten plot of v0 versus [S] which can be fit to the MichaelisMenten equation: v0 ¼ vmax ∗½S=ðK M þ ½SÞ to obtain the Michaelis constant (KM) and the maximal velocity, vmax. Dividing vmax by the enzyme concentration [E] will yield the catalytic constant kcat. 3.5 Adaptation to Quench Flow

Here, we focus on adapting the tritium release assay to singleturnover pseudouridylation measurements using a quench-flow apparatus. For details of operating the quench-flow apparatus, please consult the instrument manual. We provide only specific advice on select steps in Subheading 4. 1. Prepare a dilution of the pseudouridine synthase in 1 reaction buffer at a concentration twofold higher than the desired final concentration to account for the twofold dilution upon mixing in the quench-flow apparatus (see Note 22). Store the enzyme on ice until ready to use. 2. Dilute the tritium-labeled substrate RNA in 1 reaction buffer at a twofold higher concentration than the final reaction concentration (see Note 22). Fold the substrate RNA by heating to 65  C for 5 min and allow to passively cool to room temperature for at least 10 min.

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3. Fill the Drive Syringe C with 0.1 M HCl, which will be used as the quencher for the reaction. Subsequently wash all other lines in the quench-flow apparatus with reaction buffer. 4. Use a single-use 1 mL syringe with attached needle to carefully draw the pseudouridine synthase solution into the syringe (see Note 23). Remove needle, attach syringe to Sample Load Port A, and inject until the solution reaches the Sample Load Valve. Repeat the same procedure with the substrate RNA solution to load it into Sample Load Port B (Fig. 4a). 5. For the measurement of each desired time point, the following steps should be repeated: (a) Load the pseudouridine synthase and the RNA into the reaction loops up to the Central Mixer (see Note 24). (b) Enter the desired reaction time in the quench-flow controller and select the required reaction loop as indicated by the controller. Hold a labeled microcentrifuge tube under the exit line (see Note 25) and initiate the reaction on the controller (see Note 26). (c) Wash and dry the reaction loop. (d) Repeat sequence a–c for each time point to be measured (see Note 27). 6. Process each time point collected from the quench-flow apparatus through the tritium release assay similarly as described in Subheading 3.3, steps 5–9, with some deviations as follows. The volume of the quenched reaction obtained from the quench-flow apparatus will differ as it is dependent on the reaction loop. For example, a volume of ~220 μL is generally obtained when using reaction loop 7. Of this sample, first 2 μL is counted directly in 2 mL EcoLite(+) scintillation cocktail to determine the concentration of [5-3H]-substrate RNA present in the quenched reaction. Second, a constant volume (such as 200 μL) from each quenched reaction is added to 1 mL of pre-aliquoted 5% (w/v) activated charcoal in 0.1 M HCl and then treated as described in Subheading 3.3, steps 5–9. 3.6 Quench-Flow Data Analysis

1. The concentration of pseudouridine present in the quenched sample is determined as described in Subheading 3.4. Note that the dilution factor for processing of the sample should be adjusted as a large sample volume is used (see Note 28 for an example). In short, counts from each extracted sample are adjusted for the dilutions that occurred throughout the tritium release assay. 2. Once the concentration of pseudouridine present in each quenched sample is known, this value must be compared to the RNA concentration in the quenched sample to obtain the

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Fig. 4 Single-turnover pseudouridine formation in tRNA by an excess of a bacterial stand-alone pseudouridine synthase. (a) Rapid pseudouridylation under single-turnover conditions can be measured using a rapid-mixing quench-flow apparatus. Here, a step motor rapidly mixes the pseudouridine synthase and the substrate RNA within a few milliseconds. Pseudouridylation occurs while the mixture flows through the reaction loop, and the reaction is stopped when the mixture is combined with a quench solution (from Drive Syringe C). The quenched sample is collected and analyzed with the tritium release assay to quantify pseudouridine formation. (b) Exemplary time course of a quench-flow experiment measuring tRNA pseudouridylation by the E. coli stand-alone pseudouridine synthase TruB. Note that the time is plotted logarithmically on the x-axis. The smooth line is the result of one-exponential fitting of the data to obtain the apparent rate constant, kapp (here 0.5  0.1 s1) [13]

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percentage of pseudouridine formation. To determine the concentration of RNA in each quenched sample (in μM), the activity (in dpm) determined for 2 μL of the quenched sample is divided by the specific activity (dpm/pmole) and divided by the volume (2 μL). Now, the percentage of pseudouridine formation can be calculated by dividing the concentration of pseudouridine formed in the quenched reaction by the RNA concentration in the quenched reaction sample. This analysis must be repeated for each reaction time point. 3. In order to determine the apparent rate (kapp) of the reaction, the percentage of pseudouridine formed is plotted as a function of time and fit with a one-exponential equation using GraphPad Prism software:
Y ¼ Y 0 þ ðY 1  Y 0 Þ  exp kapp  t where Y is the percentage of pseudouridine formation, Y0 is the initial level of pseudouridine formation (see Note 29), and Y1 is the end level of pseudouridine formation. 4. Determining kapp at increasing concentrations of pseudouridine synthase will reveal whether this value is concentration dependent or not. If kapp is not concentration dependent, then it should not be limited by enzyme substrate binding and will likely directly reflect the rate of pseudouridylation (kΨ) (see Note 30).

4

Notes 1. Although having a colored loading dye is helpful in judging the migration of samples during electrophoresis, it may prove problematic when excising a target RNA from the gel during PAGE purification. If either the bromophenol blue or the xylene cyanol dye front overlaps with the desired RNA location following electrophoresis, we recommend removing this dye from the loading buffer. This will help visualize the RNA during UV shadowing and also ensure that the excised RNA sample is free of dyes. 2. The reaction buffer recommended here enables a high pseudouridylation activity of yeast H/ACA snoRNPs. For standalone enzymes, we have also successfully used a 1 TAKEM4 buffer: 50 mM Tris–HCl pH 7.5, 70 mM NH4Cl, 30 mM KCl, 1 mM EDTA, and 4 mM MgCl2. In general, the buffer conditions may have to be optimized to the respective enzyme used in the tritium release assay. 3. The charcoal in this solution settles. Therefore, it is important that the solution is mixed constantly with a stir bar to evenly distribute the charcoal when aliquoting samples prior to performing an assay.

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4. Sense DNA oligos must include a T7 promoter sequence (50 GCTAATACGACTCACTATAGGG-30 ) immediately 50 of the substrate RNA sequence. Additionally, the two 50 -most bases of the antisense DNA oligo should be methylated to reduce unwanted additions to the 30 end of the transcribed RNA. See Table 1 for examples. 5. 100 mM DTT, 100 mM stocks of each NTP, and 100 mM GMP stocks are stored at 20  C and can be freeze-thawed several times without loss of activity during in vitro transcription. 6. For in vitro transcription reactions, prepare a 0.3 mM UTP/[5-3H]UTP mixture with an activity of 2000 dpm/ pmole by mixing [5-3H]UTP with nonradioactive UTP. For example, with a 1 μCi/μL stock of [5-3H]UTP at an activity of 23 Ci/mmole (or 51,060 dpm/pmole), a 200 μL UTP mixture (0.3 mM) would be created by combining 54.05 μL of [5-3H]UTP (1.2 * 108 dpm, 2350 pmole), 2.31 μL of 25 mM cold UTP (57,650 pmole), and 143.64 μL of RNase-free water. The final concentration of ethanol in the UTP mixture should not exceed 15% (v/v) such that no more than 5% (v/v) ethanol is present in the in vitro transcription reaction. 7. Aliquots of 100 μL inorganic pyrophosphatase will not freeze at the 20  C storage temperature and should always be handled for minimum amounts of time outside the freezer and on ice. 8. T7 RNA polymerase can be purchased or expressed and purified in-house. For example, we express hexahistidine-tagged T7-RNA polymerase in E. coli BL21(DE3) followed by Ni2+sepharose affinity purification. Purified T7 RNA polymerase is rebuffered into a storage buffer (20 mM K2HPO4, pH 7.5, 1 mM DTT, 1 mM EDTA, 50% (v/v) glycerol) and concentrated to 4 μM. T7 RNA polymerase retains in vitro transcription activity after years of storage at 20  C. Aliquots are always handled for minimum amounts of time outside the freezer and on ice. Particular care should be taken to use RNase-free tubes and reagents during purification of T7 RNA polymerase to avoid RNase contaminations. 9. If inorganic pyrophosphatase is working well, there may not be a visible pellet formed during this step. 10. Often, a large white salt pellet will co-precipitate along with the transcribed RNA; this will not affect the downstream steps. 11. If the pellet is very small, an effective way to remove the supernatant completely is to push the pipette tip tightly against the bottom of the tube ensuring that it does not make direct contact with the pellet, which is usually located slightly to the side of the tube. Once liquid is removed, small pellets will

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air-dry in 10 min while larger pellets will take significantly longer since the pellet will trap some water that cannot be removed easily with a pipette. Large pellets could take up to 30 min to air-dry once you remove the supernatant. Avoid overdrying the RNA as it will be difficult to resuspend. 12. (A) Performing this step in a darkened room greatly helps to see the shadow cast onto the thin-layer chromatography plate. (B) Reduce the amount of time you expose RNA to strong UV light to prevent damage to the RNA. (C) It is typical to see a very strong shadow cast near the bottom of the gel; this is likely due to excess free nucleotide triphosphates left over from the in vitro transcription reaction, and not RNA. There may also be a signal at the very top of the gel immediately below the wells; this may be due to proteins from the in vitro transcription reaction and should not be excised. The shadow from the transcribed RNA should appear as a much weaker signal that will be seen above the height of the NTP shadow. 13. If the RNA pellet is clearly visible, use 100 μL to resuspend the pellet. If little to no pellet is visible, dissolve the RNA in 20–50 μL to ensure that the RNA is of a reasonable concentration (e.g., 10–50 μM) to be quantified with absorbance measurements and visualized with electrophoresis. 14. A specific activity of 750–2000 dpm/pmole∙U for [5-3H]substrate RNA is useful for tritium release. [5-3H]-substrate RNAs with specific activities of 250–750 dpm/pmole∙U can also be used if the reaction volumes for each time point are scaled up accordingly (see Note 16). 15. In the tritium release assay, substrate RNA and protein should be combined under multiple turnover conditions where substrate RNA is in at least threefold excess over the enzyme: 1 reaction buffer, 45 nM H/ACA guide RNA, 100 nM proteins (Cbf5-Nop10-Gar1-Nhp2), and typically 500 nM [5-3H]substrate RNA (150–1000 nM). Alternatively, 5–50 nM of stand-alone enzyme can be used together with 100 nM to 10 μM [5-3H]-substrate RNA. To determine MichaelisMenten kinetic parameters, the initial velocity v0 of pseudouridylation must be determined at increasing substrate RNA concentrations while keeping the enzyme concentration constant. 16. The reaction volume processed at each time point should be selected such that a tritium release of at least 500 dpm at the earliest time point is obtained to allow accurate determination of the extent of pseudouridine formation. For example, for a substrate RNA with a specific activity of 2000 dpm/pmole uridine, and assuming 10% pseudouridine formation at the first time point, the reaction volume sampled at each time

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point should contain at least 2.5 pmole RNA. Therefore, the reaction volume may need to be increased when detecting pseudouridylation of a [5-3H]-substrate RNA that has a low specific activity. 17. Time points should be carefully chosen and optimized for each enzyme. To accurately measure the initial velocity (v0) of the reaction, several time points should be recorded in the linear range of the reaction where less than 30% of the substrate RNA is pseudouridylated. 18. Immediately prior to removing each sample, vortex the reaction tube on low speed for a few seconds. If there is condensation occurring under the lid, briefly spin the tube to collect the contents of the tube at the bottom. 19. A glass wool plug can be created by using tweezers to pack glass wool into a P1000 pipette tip. A glass wool plug should be 1–2 cm in size. 20. Calculating the percentage of sample counted: in the first charcoal extraction, a fraction of 0.8 of the sample is retained (800 μL of originally 1000 μL). Note that this assumes that the sample volume itself is low (less than 50 μL) and negligible compared to the 1000 μL of 0.1 M HCl and 5% (w/v) activated charcoal. In the second charcoal extraction and glass wool filtration, a fraction of 0.615 of sample is kept (800 μL of originally 800 μL + 500 μL ¼ 1300 μL). Consequently, overall a fraction of 0.492 of the original sample is used in liquid scintillation counting (0.8 * 0.615). When deviating from volumes suggested in the tritium release assay (steps 5 and 6), the fraction of sample that is counted must be recalculated accordingly. 21. In our experience, the actual substrate RNA concentration in the reaction often differs from the intended substrate RNA concentration. By directly counting 2 μL of the pseudouridylation reaction as explained in step 7, the actual RNA concentration can be calculated with the help of the specific activity of the substrate RNA (aRNA in dpm/pmole uridine). It is then best to use this experimentally determined substrate RNA concentration, which is unique to each individual reaction, to calculate the concentration of percentage of pseudouridine formation. 22. To ensure pre-steady-state conditions facilitating data analysis, the pseudouridine synthase enzyme concentration must be at least threefold higher than the substrate RNA concentration. Similar as described for multiple-turnover reactions, the concentration of [5-3H]-substrate RNA must be high enough to yield counts of at least 500 dpm in each reaction sample analyzed from the tritium release assay. As explained, for a substrate RNA with a specific activity of 2000 dpm/pmole uridine,

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the reaction sample taken at each time point should contain at least 2.5 pmole of substrate RNA (see Note 16). Note that the reaction volume in the quench-flow apparatus is approximately 15 μL in each sample loop (i.e., 30 μL in total); that is, a final substrate RNA concentration of at least 0.083 μM (0.165 μM in the original sample) should be used. To further increase the signal, we typically use higher concentrations such as 5 μM pseudouridine synthase and 1 μM [5-3H]-substrate RNA [13]. For a time course of more than ten time points, we recommend the total volume of pseudouridine synthase and RNA sample to be 250–300 μL each. 23. Prior to loading the samples into the disposable 1 mL syringe, insert a small volume (~50 μL) of air prior to attaching the needle. Carefully draw sample into the syringe, leaving the small pocket of air between the sample and the plunger. Remove the needle, and carefully displace the air on top of the sample. Try to leave the air pocket between the sample and the plunger undisturbed, as it is necessary for pushing the last several microliters of sample to the central mixer. Also, ensure that no air bubbles are present within the sample as these will lead to mixing of inaccurate volumes. 24. Be careful to ensure that the sample is loaded completely up to the central mixer but stop before any sample enters the mixer. This is critical for maintaining proper reaction volumes. 25. Insert the exit line halfway into a microcentrifuge tube and close the lid as much as possible to avoid splashing. 26. Note that for longer reaction times, the motor will move twice: first to move samples into the reaction loop and second to move samples through the exit line with quencher. Ensure that you do not move the microcentrifuge tube until the reaction is complete. 27. As time courses will be analyzed by fitting to an exponential equation, time points should be distributed logarithmically rather than linearly. Also, time points may need to be optimized based on the rate of the reaction. We often use time points between 0.01 and 60 s when characterizing bacterial stand-alone pseudouridine synthases (Fig. 4b, [13]). 28. For a quenched sample volume of 200 μL, the dilution factor is 0.406: here, in the first charcoal extraction, a fraction of 0.66 of the sample is retained (800 μL of originally 1200 μL ¼ 1000 μL 0.1 M HCL, activated charcoal +200 μL sample). In the second charcoal extraction and glass wool filtration, a fraction of 0.615 of sample is kept, and overall a fraction of 0.406 of the original sample is used in liquid scintillation counting.

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29. Theoretically, the initial level of pseudouridine formation (Y0) should be zero. However, due to hydrogen/tritium exchange with water, it is possible that some tritium was already released from the RNA prior to modification by a pseudouridine synthase. This background value should always be determined and typically increases with the age of a [5-3H]-substrate RNA preparation. 30. For more details on the kinetic analysis of pre-steady-state reaction kinetics, an appropriate textbook should be consulted [17–19].

Acknowledgments This work was supported by the Natural Sciences and Engineering Research Council of Canada [Discovery Grant RGPIN-201405954] and Alberta Innovates [Strategic Research Chair 2015]. References 1. Spenkuch F, Motorin Y, Helm M (2014) Pseudouridine: still mysterious, but never a fake (uridine)! RNA Biol 11(12):1540–1554 2. Madison JT, Everett GA, Kung H (1966) Nucleotide sequence of a yeast tyrosine transfer RNA. Science 153(3735):531–534 3. Kato N, Harada F (1984) Nucleotide sequence of nuclear 5.4 S RNA of mouse cells. Biochim Biophys Acta 782(2):127–131 4. Brand RC, Klootwijk J, Sibum CP, Planta RJ (1979) Pseudouridylation of yeast ribosomal precursor RNA. Nucleic Acids Res 7 (1):121–134 5. Carlile TM, Rojas-Duran MF, Zinshteyn B, Shin H, Bartoli KM, Gilbert WV (2014) Pseudouridine profiling reveals regulated mRNA pseudouridylation in yeast and human cells. Nature 515(7525):143–146 6. Hamma T, Ferre-D’Amare AR (2006) Pseudouridine synthases. Chem Biol 13 (11):1125–1135 7. Rintala-Dempsey AC, Kothe U (2017) Eukaryotic stand-alone pseudouridine synthases - RNA modifying enzymes and emerging regulators of gene expression? RNA Biol 14(9):1185–1196 8. Watkins NJ, Bohnsack MT (2012) The box C/D and H/ACA snoRNPs: key players in the modification, processing and the dynamic folding of ribosomal RNA. Wiley Interdiscip Rev RNA 3(3):397–414

9. Huang L, Pookanjanatavip M, Gu X, Santi DV (1998) A conserved aspartate of tRNA pseudouridine synthase is essential for activity and a probable nucleophilic catalyst. Biochemistry 37(1):344–351 10. Veerareddygari GR, Singh SK, Mueller EG (2016) The Pseudouridine synthases proceed through a glycal intermediate. J Am Chem Soc 138(25):7852–7855 11. Kamalampeta R, Keffer-Wilkes LC, Kothe U (2013) tRNA binding, positioning, and modification by the pseudouridine synthase Pus10. J Mol Biol 425(20):3863–3874 12. Kamalampeta R, Kothe U (2012) Archaeal proteins Nop10 and Gar1 increase the catalytic activity of Cbf5 in pseudouridylating tRNA. Sci Rep 2:663 13. Wright JR, Keffer-Wilkes LC, Dobing SR, Kothe U (2011) Pre-steady-state kinetic analysis of the three Escherichia coli pseudouridine synthases TruB, TruA, and RluA reveals uniformly slow catalysis. RNA 17(12):2074–2084 14. Tillault AS, Schultz SK, Wieden HJ, Kothe U (2018) Molecular determinants for 23S rRNA recognition and modification by the E. coli Pseudouridine synthase RluE. J Mol Biol 430 (9):1284–1294 15. Caton EA, Kelly EK, Kamalampeta R, Kothe U (2018) Efficient RNA pseudouridylation by eukaryotic H/ACA ribonucleoproteins requires high affinity binding and correct

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positioning of guide RNA. Nucleic Acids Res 46(2):905–916 16. Kelly EK, Czekay DP, Kothe U (2019) Basepairing interactions between substrate RNA and H/ACA guide RNA modulate the kinetics of pseudouridylation, but not the affinity of substrate binding by H/ACA small nucleolar ribonucleoproteins. RNA 25(10):1393–1404

17. Bernasconi C (1976) Relaxation kinetics. Academic Press, New York 18. Cook PC, Cleland WW (2012) Enzyme kinetics and mechanism. Garland Science, New York 19. Fersht A (1999) Structure and mechanism in protein science: guide to enzyme catalysis and protein folding, 3rd Revised edn ed. W.H. Freeman & Co Ltd, San Francisco

Chapter 22 Investigating Pseudouridylation Mechanisms by High-Throughput in Vitro RNA Pseudouridylation and Sequencing Nicole M. Martinez and Wendy V. Gilbert Abstract Pseudouridine profiling has revealed many previously unknown sites of the RNA modification pseudouridine (Ψ) in cellular RNAs. All organisms express multiple pseudouridine synthases (PUS) whose RNA targets and mechanisms of targeting remain to be elucidated. Here, we describe a high-throughput in vitro pseudouridylation assay to interrogate pseudouridine status upon incubation with recombinant pseudouridine synthases (PUS) at thousands of RNA sequences of interest in parallel. This approach allows validation of sites provisionally identified in cells, identification of the direct targets of individual PUS, and interrogation of the determinants of target recognition including primary sequence and RNA secondary structure. Key words Pseudouridine, RNA modification, mRNA modification, PUS, Pseudouridine synthase, Pseudouridylation, Pseudo-seq

1

Introduction Recent advances in transcriptome-wide sequencing methods allowed the discovery of pseudouridine [1–3] among a growing repertoire of modified nucleotides in mRNAs [4]. Validation of sites called from transcriptome-wide sequencing experiments is an important step in establishing a high confidence map of pseudouridylation in biological samples of interest. Multiple PUS could potentially modify any given site; thus identifying the direct targets of individual enzymes is critical for interrogating the function of mRNA modifications in gene regulation. How specificity is achieved by RNA-modifying enzymes and why some sequences are targeted for modification, but not others, are not completely understood. Methods to validate sites, identify the enzymes, and dissect the RNA sequence and structures that direct modification are needed to address these questions.

Mary McMahon (ed.), RNA Modifications: Methods and Protocols, Methods in Molecular Biology, vol. 2298, https://doi.org/10.1007/978-1-0716-1374-0_22, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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Here we present a detailed method for parallel testing of thousands of RNA sequences for pseudouridylation by individual recombinant pseudouridine synthases (PUS) using high-throughput in vitro pseudouridylation and sequencing (Fig. 1). Pseudouridines installed by recombinant PUS are identified based on carbodiimide (CMC) derivatization to produce reverse transcriptase stops at modified sites [1]. Using this approach, we determined that multiple human pseudouridine synthases modify mRNA sequences and defined a sequence and structural motif that is necessary and sufficient for pseudouridylation by yeast PUS1 [5]. Importantly, our strategy allows high sequencing coverage at sites of interest even in mRNA sequences that are lowly expressed in cells. The approach that we developed for the study of pseudouridine in mRNA sequences can be adapted to investigate other RNA modifications of interest.

2

Materials

2.1 DNA Pool and In Vitro Transcription

1. Pool of DNA sequences of targets. Each sequence should include the following: T7 sequence to include at 50 end of DNA oligos: GCTAATAC GACTCACTATAGGG 30 Adapter to append to 30 end of DNA oligo: CACTCGGG CACCAAGGAC 2. 10 mM Tris·Cl, pH 8.0. 3. Phusion High-Fidelity (HF) DNA polymerase and HF buffer. 4. Pool PCR F primer: GCTAATACGACTCACTATAGGG 5. Pool PCR R primer: GTCCTTGGTGCCCGAGTG 6. 10 mM dNTPs. 7. DMSO. 8. 8% Non-denaturing TBE-polyacrylamide. 9. DNA loading dye (6): 30% (v/v) Glycerol, 0.025% (w/v) bromophenol blue, 0.025% (w/v) xylene cyanol FF. 10. 10 bp DNA ladder (Invitrogen). 11. 0.5 TBE. 12. SYBR Gold. 13. DNA elution buffer: 300 mM NaCl, 10 mM Tris–Cl, pH 8.0. 14. Isopropanol. 15. GlycoBlue. 16. 70% Ethanol. 17. Deionized distilled H2O. 18. MEGAshortscript T7 Transcription kit (Ambion).

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sequence flanking Ψ site Ψ

T7 promoter

3’ adapter

Ψ

Pool of >1,000 of DNA oligos

Ψ

in vitro transcribe RNA

PUS7

RNA sequences

wildtype

recombinant PUS

PUS1

+

TRUB1

mutants to test impact of structure on Ψ

in vitro pseudouridylation

Ψ wildtype

Ψ mutants to test impact of structure on Ψ

Pseudo-seq

PUS1

Ψ

Assign Ψ to PUS & Identify RNA elements required for Ψ

Fig. 1 Overview schematic of high-throughput in vitro RNA pseudouridylation and sequencing. Design a pool of DNA sequences that includes candidate pseudouridines and flanking sequence. Include wild-type and mutant sequences to interrogate the sequence and structural features that direct Ψ. The DNA oligos should include a T7 promoter sequence and a 30 adapter for PCR amplification and in vitro transcription. In vitro transcribe RNA sequences from the DNA pool template. In vitro pseudouridylate by incubating the folded pool of RNA sequences with recombinant pseudouridine synthases (PUS). Perform pseudo-seq to detect Ψ installed by specific PUS and to determine the effects of mutations on Ψ. Figure adapted from Carlile et al. 2019 Nat Chem Bio

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19. TURBO DNase. 20. 8% TBE-urea-polyacrylamide mini-gel. 21. Formamide buffer (2): 95% (v/v) Formamide, 5 mM EDTA, pH 8.0, 0.02 5% (w/v) SDS, 0.025% (w/v) bromophenol blue, 0.025% (w/v) xylene cyanol FF. Store at 20  C. 22. RNA elution buffer: 300 mM Sodium acetate, pH 5.3, 1 mM EDTA, pH 8.0, 100 U/ml RNasin Plus (add the RNasin immediately before use). 2.2 In Vitro Pseudouridylation with Recombinant PUS

1. Recombinant PUS of interest: For purification strategy for several PUS, please see reference [5]. 2. 5 Pseudouridylation buffer: 500 mM Tris pH 8.0, 500 mM ammonium acetate, 25 mM MgCl2, 0.5 mM EDTA. Filter sterilize using a 0.2 μm filter. 3. 100 mM DTT. 4. Deionized distilled H2O. 5. Acid phenol. 6. Chloroform. 7. Isopropanol. 8. 100% Ethanol. 9. 70% Ethanol. 10. 3 M Sodium acetate, pH 5.3.

2.3

CMC Treatment

1. BEU buffer: 7 M Urea, 4 mM EDTA, pH 8.0, 50 mM bicine, pH 8.5. Filter sterilize using a 0.2 μm filter. Adjust pH to ~9.0 using NaOH and HCl. 2. 0.5 M CMC in BEU buffer: Make fresh immediately before CMC treatment, 0.5 M CMC (CMC (Sigma, cat. no. C106402)) in BEU buffer. 3. Sodium carbonate buffer: 50 mM Na2CO3, pH 10.4, and 2 mM EDTA, pH 8.0. Prepare from stocks of 1 M Na2CO3, pH 10.4, and 0.5 M EDTA, pH 8.0, and adjust the final pH to 10.4 using sodium bicarbonate. Filter sterilize using 0.2 μm filter. 4. 40 mM EDTA, pH 8.0. 5. 3 M Sodium acetate, pH 5.3. 6. GlycoBlue. 7. 100% Ethanol. 8. 70% Ethanol. 9. 10 mM Tris–Cl, pH 8.0.

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2.4 Reverse Transcription and Library Construction

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1. RT buffer w/o Mg2+ (10): 500 mM Tris–Cl, pH 8.6, 600 mM NaCl, and 100 mM DTT. Store at 20  C. 2. Gel-purified RT primer /5Phos/NNNNNNNNNGATCGTCGGACTGTA GAACTCTGAACGTGTAGATC/iSp18/CACTC A/iSp18/ CCTTGGCACCCGAGAATTCCAGTC C TT GGTGCCCGAGTG 3. 240 mM MgCl2. 4. AMV RT. 5. 25 mM dNTPs. 6. 1 N NaOH. 7. 1 N HCl. 8. CircLigase ssDNA ligase and accompanying buffer (Epicentre). 9. 1 mM ATP. 10. 50 mM MnCl2. 11. Phusion High-Fidelity (HF) DNA polymerase and HF buffer. 12. RP1 primer—Library PCR F: AATGATACGGCGACCACCGAGATCTACAC GTTCA GAGTTCTACAGTCCGA 13. Barcoded reverse PCR primers (XXXXXX indicates unique barcodes)—Library PCR R: CAAGCAGAAGACGGCATAC GAGATXXXXXXGTGACTGGAGTTCCTTGGCACCCGA GAATTCCA

3

Methods

3.1 DNA Pool Design (Validation of Sites and PUS Assignment)

1. Design a pool of DNA oligo sequences that include the pseudouridine sites of interest and flanking sequences. For each pseudouridine of interest, extract the coordinate and desired amount of flanking sequence centered on the pseudouridine (e.g., for an RNA sequence length of 130 nucleotides in length include the pseudouridine and 65 nucleotides of upstream and 64 nucleotides of downstream sequence, see Note 1). 2. The DNA oligos should include the T7 promoter ( GCTAA TACGACTCACTATAGGG ) upstream of the sequence of interest for in vitro transcription and a common sequence at the 30 end for cDNA synthesis and library construction (CAC TCGGGCACCAAGGAC). 3. Order the commercially synthesized pool (e.g., Twist Biosciences) of DNA oligo sequences (Fig. 1).

3.2 PCR Amplify the DNA Pool

1. Resuspend the lyophilized pool of DNA oligos at 0.5 ng/μL in 10 mM Tris–HCl, pH 8.0.

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2. Prepare a PCR master mix for a 500 μL PCR reaction: PCR master mix

1

11

5 HF buffer

10 μL

110 μL

10 μM Pool PCR F primer

1 μL

11 μL

10 μM Pool PCR R primer

1 μL

11 μL

10 mM dNTPs

1 μL

11 μL

DMSO

1.5 μL

6.5 μL

H2O

34.25 μL

376.75 μL

Oligo Pool

1 μL

11 μL

Phusion polymerase

0.25 μL

2.75 μL

3. Distribute the PCR master mix across PCR tubes. Place in thermocycler for 12 PCR cycles as indicated below (see Note 2). Initial denaturation

98  C for 30 s

Cycle Denature

98  C for 10 s

Anneal

58  C for 30 s

Extend

72  C for 30 s

Final extension

72  C for 5 min

4. Cast two 8% non-denaturing TBE-polyacrylamide mini-gels. 5. Add 100 μL of 6 DNA loading buffer to the 500 μL PCR reactions to 1 final concentration. Include a 10 bp DNA ladder and run the PCR product evenly distributed across all the lanes of two 8% TBE polyacrylamide gels for 50 min at 200 V. Stain each gel in 15 mL of 0.5 TBE with SYBR Gold (1:10,000) rocking for 5 min, visualize with blue light illumination (Fig. 2), and cut out bands (see Note 3). 6. Elute PCR product by placing gel slices in two 1.5 mL tubes containing 750 μL of DNA elution buffer. Rock overnight at room temperature. 7. Transfer solution with eluted DNA into new 1.5 mL tubes and precipitate DNA by adding 750 μL isopropanol and 2 μL GlycoBlue to each tube and vortexing. Incubate at 20  C for at least 30 min. Spin at maximum speed on microfuge for 30 min to pellet DNA. Wash pellets with 750 μL of 70% ethanol and spin down at maximum speed on microfuge for 10 min.

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Fig. 2 Gel image of PCR of pool sequences. Representative gel image of PCR product distributed across multiple wells

8. Remove supernatant and allow pellets to dry for 10 min at room temperature. 9. Resuspend precipitated DNA in 8 μL of water. 3.3 In Vitro Transcribe RNA

1. Set up the in vitro transcription reaction using the MEGAshortscript T7 Transcription kit as follows: To 8 μL of PCR product (Subheading 3.2) add: 2 μL 10 buffer. 2 μL ATP. 2 μL UTP. 2 μL CTP. 2 μL GTP. 2 μL enzyme mix 2. Incubate at 37  C for 2 h, add additional 2 μL of T7 polymerase, and incubate for another 2 h. 3. Add 2 μL TURBO DNase and incubate at 37  C for 15 min to degrade the DNA template. 4. Cast 8% TBE-urea-polyacrylamide mini-gel and pre-run for 20 min at 200 V. 5. Add 24 μL of 2 formamide buffer to the sample and heat denature at 95  C for 2 min. 6. Load 6 μL of sample into each well of the gel, include a 10 bp DNA ladder, and run at 200 V for 60 min. 7. Stain each gel in 15 mL of 0.5 TBE with SYBR Gold (1:10,000) rocking for 5 min, visualize (Fig. 3), and cut out bands (see Note 4).

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Fig. 3 Gel image of in vitro-transcribed RNA pool. Representative image of gel containing RNA in vitro transcribed from the pooled DNA template distributed across multiple wells

8. Elute RNA by placing gel slices in 750 μL of RNA elution buffer and incubate overnight at 4  C on a rocking platform. 9. Transfer the supernatant to a fresh 1.5 mL tube and precipitate RNA by adding 750 μL isopropanol and 2 μL GlycoBlue and vortexing. Incubate at 20  C for at least 30 min. Spin down at maximum speed in microfuge for 30 min to pellet RNA. 10. Wash pellets with 750 μL of 70% ethanol and spin down at maximum speed in microfuge for 10 min. 11. Remove supernatant and allow pellets to dry for 10 min at room temperature. 12. Resuspend dried RNA pellets in 20 μL of 10 mM Tris, pH 8. 3.4 In Vitro Pseudouridylation with Recombinant PUS

Include a –CMC (mock treated) and a +CMC sample for each PUS treatment condition. Include a no-PUS control along with the individual PUS samples. These negative control samples will allow for identification of reverse transcriptase stops that are CMC and PUS dependent indicative of a pseudouridine. 1. Fold the RNA (see Note 5) by diluting the RNA stock solution to 5 μM final concentration. 2. Prepare a master mix to fold RNA by combining 6 μL (30 pmol) of 5 μM RNA stock solution and add 33.5 μL of water for each PUS being tested and a no-PUS control. 3. Incubate the RNA reactions at 75  C for 2 min to denature the RNA. 4. Place the sample on ice for 5 min. 5. Add 10 μL of 5 pseudouridylation buffer to the RNA solution for each PUS being tested and a no-PUS control. 6. Incubate sample at 37  C for 20 min to refold RNA.

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7. Prepare the remaining components of the in vitro pseudouridylation reaction and pre-warm at 30  C prior to addition of folded RNA. Folded RNA and recombinant PUS (see Note 6) should be added last. Final component concentrations are 2 mM DTT, 1 pseudouridylation buffer, and 600 nM PUS. In vitro pseudouridylation reaction: 90 μL 5 Pseudouridylation buffer 10 μL 100 mM DTT 2.4 μL Recombinant PUS (130 μM). 46.5 μL of folded RNA. Bring volume to 500 μL with water. 8. Incubate reactions at 30  C for 45 min to complete pseudouridylation. 45 min is sufficient for nearly complete pseudouridylation by many PUS in our hands. 9. Immediately add 500 μL of acid phenol to samples to stop the reactions, vortex, and spin for 5 min at maximum speed in a microfuge. 10. Transfer top phase to a new tube, add 500 μL of chloroform, vortex, and spin for 5 min at maximum speed in a microfuge. 11. Transfer top phase to a new tube; add 1 volume of isopropanol, 1/9 volume of sodium acetate, and 2 μL of glycogen; and vortex. Incubate the reaction at 20  C for at least 30 min and spin at maximum speed for 30 min in a microfuge to precipitate the RNA. 12. Wash the RNA pellet with 750 μL of 70% ethanol and spin down at maximum speed in a microfuge for 10 min. 13. Remove supernatant and allow pellets to dry for 10 min at room temperature. 14. Resuspend pellet in 30 μL of water. 3.5 CMC Modification and Reversal

1. To begin CMC modification, make a fresh 0.5 M CMC stock in BEU buffer. See Note 7. 2. Split RNA sample into two 0.2 mL tubes by transferring 12 μL of RNA into a tube for the mock-treated (–CMC) and 18 μL of RNA into a separate tube for the CMC-treated sample. Bring the volume of each sample to 20 μL with water (see Note 8). 3. Add 2.9 μL of 40 mM EDTA to each sample and incubate at 80  C for 3 min in a thermocycler (see Note 9). 4. Add 100 μL of 0.5 M CMC in BEU buffer to +CMC samples and 100 μL of BEU buffer to –CMC samples. 5. Incubate RNA for 45 min at 40  C with shaking at 2  g in a thermomixer. 6. Transfer samples to fresh 1.5 mL tubes. 7. Precipitate the RNA to clean up the reaction by adding 1 mL of 100% ethanol, 50 μL of sodium acetate, and 2 μL of GlycoBlue (see Note 10).

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8. Incubate the reaction at 20  C for at least 30 min. 9. Spin at maximum speed for 30 min in a microfuge to precipitate the RNA. 10. Wash pellets 2 by adding 500 μL of 70% ethanol and spin down for 10 min in a microfuge at maximum speed and 4  C. 11. Dry pellets for 10 min at room temperature. 12. To begin the CMC reversal, resuspend each pellet in 30 μL of sodium carbonate buffer. 13. Incubate the RNA for 2 h at 50  C with shaking at 1000 rpm in thermomixer. 14. Precipitate RNA by adding 2 μL GlycoBlue, 1/9 volume of sodium acetate, and 2.5 volumes of ethanol and incubating at 20  C for 30 min. 15. Spin at maximum speed for 30 min in a microfuge to precipitate the RNA. 16. Wash pellets 2 by adding 500 μL of 70% ethanol and spin down for 10 min in a microfuge at maximum speed and 4  C. 17. Allow pellets to dry for 10 min at room temperature. 18. Resuspend in 8 μL of 10 mM Tris–HCl, pH 8. 19. Begin size selection of full-length RNA after CMC reversal (see Note 11) by casting an 8% TBE-urea-polyacrylamide mini-gel and pre-run for 20 min at 200 V. Meanwhile, add 8 μL of 2 formamide buffer to samples, heat at 95  C for 2 min, and then place on ice. 20. Load gel with samples and run for 60 min at 200 V. Include a 10 bp DNA ladder on the gel. 21. Stain gel with 15 mL of 0.5 TBE and SYBR Gold (1:10,000) for 5 min, visualize stained gel (Fig. 4, see Note 12), and cut out full-length product (~151 nt) using the 10 bp DNA ladder as a guide for size. 22. Elute RNA from gel slices in 400 μL of RNA elution buffer by incubating overnight at 4  C on a rocking platform. 23. Transfer eluate into a new tube, precipitate by adding 1 mL of 100% ethanol and 2 μL of GlycoBlue, and vortex. Incubate at 20  C for at least 30 min. 24. Spin at a maximum speed for 30 min in a microfuge to precipitate the RNA. 25. Wash pellets 2 by adding 500 μL of 70% ethanol and spin down for 10 min in a microfuge at maximum speed and 4  C. 26. Allow pellets to dry for 10 min at room temperature. 27. Resuspend in 6.2 μL H2O.

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Fig. 4 Size selection of full-length RNA. Representative image of gel purification of full-length RNA after CMC reversal 3.6 Reverse Transcription

1. Transfer 6.2 μL of RNA into clean PCR tubes. 2. Add 1 μL of gel-purified RT primer (RT 25 μM) and 0.8 μL of 10 RT buffer without magnesium. In parallel prepare a no-RNA reaction control with 6.2 μL of H2O. 3. Anneal the RT primer by incubating as follows: 1. 65  C for 4 min 2. 55  C for 2 min 3. 45  C for 2 min 4. 42  C for 2 min 4. Centrifuge briefly to collect condensation and place on ice. 5. Prepare an extension master mix with the following components per reaction: Extension master mix: 0.6 μL 10 RT buffer w/o Mg2 2.24 μL 25 mM dNTPs 1.16 μL 240 mM MgCl2 1.0 μL RNasin Plus 1.0 μL AMV RT. 6. Add 6 μL of extension mix to the annealing reaction. 7. Incubate for 1 h at 42  C in thermocycler. 8. Cast as many 8% TBE-urea-polyacrylamide mini-gels as needed for the number of samples. Pre-run gels for 20 min at 200 V. 9. Add 1.5 μL 1 N NaOH to each sample and incubate for 15 min at 98  C to hydrolyze RNA templates.

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Fig. 5 Size selection of truncated cDNA. Representative image of gel purification of truncated cDNA

10. Add 1.5 μL 1 N HCl to each reaction to neutralize pH. 11. Add 17 μL of 2 formamide buffer to each reaction. 12. Heat samples for 2 min at 95  C and place on ice. 13. Load samples, include a 10 bp DNA ladder, and run 8% TBEurea-polyacrylamide mini-gels for 68 min at 200 V. 14. Stain gel with 15 mL of 0.5 TBE and SYBR Gold (1:10,000) for 5 min, visualize stained gel, and cut out the truncated cDNA of expected size relative to the position of the expected pseudouridine in the oligo and taking into account the length added by the RT primer. Example: For an RNA pool consisting of 130 nt oligos with the expected pseudouridine at position 66, and using the 30 adapter and RT primer noted above, the truncated cDNAs should run at ~155 nt. Cut a band on the gel /+30 nucleotides the expected size (Fig. 5, see Note 13). 15. Elute cDNA by placing each gel slice in 400 μL DNA elution buffer by incubating at room temperature with rocking overnight. 16. Precipitate by adding 1 mL of ethanol and 2 μL GlycoBlue, vortexing and incubating at 20  C for at least 30 min. 17. Spin down at maximum speed for 30 min. 18. Wash pellets with 750 μL of 70% ethanol and spin down at maximum speed in a microfuge for 10 min. 19. Allow pellets to dry for 10 min at room temperature. 20. Resuspend each pellet in 15 μL 10 mM Tris, pH 8.

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3.7 cDNA Circularization (See Note 14)

391

1. Prepare circularization master mix with the following components per reaction: Circularization master mix components per reaction: 2 μL 10 Circ ligase buffer. 1 μL 1 mM ATP. 1 μL 50 mM MnCl2 2. Add 4 μL circularization master mix to each cDNA sample and mix by pipetting. 3. Dilute circ ligase (1:2) in 2 circ ligase buffer by adding 1 μL of circ ligase and 1 μL of 2 circ ligase buffer. 4. Add 1 μL of 0.5 circ ligase dilution (step 3) to each sample and mix. 5. Incubate for 6 h at 60  C. 6. Heat inactivate circ ligase by incubating for 10 min at 80  C.

3.8

Diagnostic PCR

1. Perform a diagnostic PCR to determine the optimal PCR cycle number for each library. PCR master mix components per reaction: 3.34 μL 5 HF buffer. 0.33 μL 10 mM dNTPs. 0.84 μL 10 μM RP1 primer. 0.84 μL 10 μM barcoding primer. 10.17 μL H2O. 0.17 μL Phusion polymerase 2. Transfer 15.7 μL of master mix into individual PCR tubes and add 1 μL circularized cDNA. 3. Mix well and divide the sample into two reactions for testing two PCR cycles (e.g., 10, 12). 4. Set PCR to the following cycles: Initial denaturation

98  C for 30 s

Cycle Denature

98  C for 10 s

Anneal

60  C for 20 s

Extend

72  C for 40 s

Final extension

72  C for 5 min

6. Add 6 DNA loading dye to each reaction. 7. Run on 8% non-denaturing TBE polyacrylamide mini-gel for 40 min at 200 V.

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8. Evaluate the optimal cycle number which amplifies the desired product but does not produce extra bands from overamplification. 3.9

Final PCR

1. Set up PCR reaction. PCR master mix components per reaction: 10.0 μL 5 HF buffer 1.0 μL 10 mM dNTPs 2.5 μL 10 μM RP1 primer 2.5 μL 10 μM barcoding primer 30.5 μL H2O 0.5 μL Phusion. 2. Add 3 μL circularized cDNA to each tube. Mix and run PCR program described above with the chosen number of cycles for each sample. 3. Add 10 μL of 6 DNA loading dye to each reaction. 4. Run on each PCR reaction split over three lanes in 8% non-denaturing TBE polyacrylamide mini-gels for 40 min at 200 V. 5. Stain gel with 15 mL of 0.5 TBE and SYBR Gold (1:10,000) for 5 min, visualize stained gel, and cut out PCR product (Fig. 6 and see Note 15). 6. Elute PCR product by placing gel slice in 400 μL DNA elution buffer and incubating at room temperature with rocking overnight. 7. Transfer eluate into a new tube and precipitate by adding 1 mL of 100% ethanol and 2 μL of GlycoBlue and vortexing. Incubate at 20  C for at least 30 min. 8. Wash pellets 1 with 750 μL of 70% ethanol and spin down at maximum speed in a microfuge for 10 min. 9. Allow pellets to dry for 10 min at room temperature. 10. Resuspend in 10 μL 10 mM Tris, pH 8.

3.10 Next-Generation Sequencing and Pseudouridine Detection

1. Submit libraries for sequencing on Illumina HiSeq single-end 40–75 bp reads. 2. Primer sequences at the 50 end of the amplicon can be trimmed from the reverse read using cutadapt [6]. 3. PCR duplicates can be collapsed by virtue of the 10 N UMI introduced with the RT primer during library construction using fastx_collapser [7]. 4. Processed reads can then be mapped to a custom bowtie index of pool sequences using tophat2 [8].

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Fig. 6 Final library PCR. Representative image of gel purification of the final library PCR

5. Pseudouridylation signal can be calculated for each position in a 51 nt window centered at the expected Ψ, and the fraction of reads in the window whose 50 ends map to that position is calculated for each paired /+CMC sample. Pseudo-seq signal is the difference in fractional reads between the +CMC and CMC libraries multiplied by the window size. The reported pseudo-seq signal corresponds to the nucleotide 30 to the Ψ. 6. We have assigned pseudouridylated substrates to individual PUS from single replicates by using a Grubbs’ outlier test (significance level alpha set to 0.05) to identify sites that have pseudo-seq signal values that deviate from the normal distribution of peak heights for all the other conditions (all other PUS and no-PUS samples) [5]. A target site was assigned to a PUS if it was called as an outlier exclusively in the corresponding PUS sample and had a pseudo-seq signal >1.0. Other suitable statistical approaches might be applied to assign a PUS or RNA-modifying enzyme to a substrate (see Note 16). 3.11 Kinetic Analysis to Identify Structural and Sequence Motifs that Drive Modification

To determine the RNA sequence and structural features that govern pseudouridylation specificity by a particular PUS, we have included modifications to the base protocol as outlined below. Pseudo-seq signal is an end-point measurement, which is affected by capture biases and is not comparable across different sites. To compare pseudouridylation efficiency of different substrates and to determine the contribution of specific elements to pseudouridylation of those substrates we have developed a kinetic approach. This approach determines the relative initial velocity (v0,rel) for each sequence from a series of time points.

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Define PUS Recognition Features: Identify sequence or structural features in common among targets of a particular PUS. The common features can be determined from sites identified as targets by genetic assignment in cells (pseudo-seq) or from direct targets of a PUS identified by an initial in vitro pseudouridylation assay with a DNA pool of sites of interest as described above. To identify common sequence features among targets of a PUS determine the frequency of nucleotides at each position surrounding all the pseudouridines (e.g., Weblogo [9]) and/or perform a motif enrichment analysis (e.g., MEME [10]). To identify shared structural features among targets, fold the RNA sequences in silico (e.g., RNA Fold) and use structure probing data (e.g., SHAPE-seq [11], DMS-seq [12]) to refine the structural features if available. Design Sequence and Structure Perturbing Mutants: Design mutations to test the sequence and structural requirements for modification. To test the sequence constraints on modification you can include sequences in the pool that test all possible sequence variations of the identified motif. Keep each position of the motif constant while varying the other positions to all possible nucleotides. To interrogate the importance of structural features, design structure disrupting and compensatory mutants to disrupt common features of the identified structures. For example, to determine the importance of a stem in a stem-loop, design mutants that disrupt and restore base pairing of the stem. Design weak stemdisrupting mutants by randomly selecting 25% of base pairs for mutation, and strong stem-disrupting mutants by randomly selecting an additional 25% of base pairs for mutations. Restore base pairing of the mutants to generate compensatory mutants. Append a unique barcode for each sequence variant upstream of the 30 adapter sequence to be able to map and distinguish reads coming from closely related sequences. The length of the barcode can be determined by determining the appropriate hamming distance (number of nucleotide differences between two barcodes) given the total number of sequences to be included in the pool.

3.11.2 In Vitro Pseudouridylation Time Course

Since the fraction of reads in the +CMC samples mapping to the Ψ-dependent RT stop position correlate well with pseudo-seq signal [5], CMC libraries can be omitted from this time course experiment to reduce the total number of samples. Modify the following steps in Subheading 3.4 above (In vitro pseudouridylation with recombinant PUS) to include samples across a 15-min time course: Step 8: Include reactions for multiple time points between 0 and 15 min. This range of time points will need to be determined empirically for each PUS to select suitable time points prior to saturation.

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Step 9: Stop each reaction by snap freezing the tubes in liquid nitrogen to quickly stop each time point. Proceed to addition of acid phenol once samples for all time points have been collected and snap frozen. 3.11.3 Next-Generation Sequencing and Kinetic Analysis

Modify the following steps in Subheading 3.10 above (Nextgeneration sequencing and pseudouridine detection): Step 1: In order to read the 10 N UMI and the stop position at the 50 end and the barcode at the 30 end of the amplicon, the libraries for the kinetic analysis of wild-type and structure-disrupting mutants need to be sequenced using paired-end 40–75 bp reads. Step 2: Primer sequences at the 50 end of the amplicon can be trimmed from the reverse read using cutadapt. Trimmed pairedend reads are then merged with PEAR. Step 3: PCR duplicates can be collapsed by virtue of the 10 N UMI introduced with the RT primer during library construction using fastx_collapser [7]. Collapsed reads need to be trimmed of the nucleotides corresponding to the barcode 30 end and of 30 adapter sequence using cutadapt. Step 5: For the kinetic analysis, the pseudouridine signal is calculated as the fraction of reads whose 50 ends map to the expected stop position. To calculate the relative initial velocities for each substrate, the background signal (fraction of reads) from the average of two 0-min time points is subtracted from each time point and any negative values are set to 0. The values for each time point are normalized by the maximum signal (fraction of reads) obtained for the wild-type sequence. A linear regression analysis is then performed to obtain the initial velocity (slope). Differences in the relative initial velocity (v0,rel) between all wild-type and mutant substrates of the same type (e.g., stem disrupted) are calculated by performing a paired, two-tailed Student’s t-test ( pvalue < 0.05).

4

Notes 1. We have used DNA pools that are 170 nucleotides in length. Oligo synthesis companies can now synthesize oligo pools of up to 300 nucleotides in length. For our experiments, we have included up to tens of thousands of sequences. However, the number of sequences that can currently be synthesized commercially as pools appears to be limitless (e.g., Twist Bioscience). Therefore, the throughput of this experiment can be scaled up or down according to experimental needs. 2. Test various PCR cycles to identify the optimal amplification cycle of the pool to maximize yield and prevent nonspecific

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overamplification products. An appropriate annealing temperature should be selected based on the primer sequences used. 3. Use blue-light illumination instead of UV to avoid damaging the DNA template. 4. Use blue-light illumination instead of UV to avoid damaging the input RNA for in vitro pseudouridylation reactions. 5. PUS proteins and other tRNA-modifying enzymes in many cases recognize structural features of folded RNA. Therefore, it is important to fold the purified in vitro-transcribed RNA to ensure modification. The RNA is denatured in water followed by addition of 5 pseudouridylation buffer which includes magnesium to facilitate folding at 37  C. 6. Purify the recombinant PUS or modifying enzyme of interest. This will need to be optimized for each enzyme; as such we have not included a protocol here. See [5] for the purification strategy for several PUS. 7. It is important to prepare a fresh 0.5 M CMC BEU buffer solution before use. 8. More RNA is distributed to the +CMC sample to account for lower recovery of CMC-modified RNA. 9. This step denatures the RNA to increase accessibility of pseudouridines that might be protected by structure and improve derivatization with CMC. 10. Use of ethanol precipitation is important because it yields better recovery of CMC-modified RNA than isopropanol precipitation. 11. During the incubation at 50  C under alkaline conditions for CMC reversal there is significant RNA degradation. We have found that gel purifying the full-length RNA allows for uniform coverage of the oligo sequence and prevents a 30 read coverage bias that is caused by degradation during this step. 12. Use blue-light illumination since reverse transcription is very inefficient on UV-damaged RNA template. 13. It is likely that only the full-length product will be visible on the gel. Cut the truncated cDNAs (due to CMC-dependent stops one nucleotide 30 to the expected Ψ position) to enrich for the pseudouridine-containing RNAs using a suitable-sized ladder as a reference. 14. A cDNA 50 adapter ligation strategy can be used instead of cDNA circularization using the long RT primer. This strategy overcomes the inefficient circularization of longer RNAs by circ ligase. 15. It is important to gel purify the library PCR product away from the contaminating PCR product that results from amplification

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of circularized RT primer to decrease the proportion of unusable reads. 16. One shortcoming of pseudo-seq signal is that it uses RT stop information exclusively at the position of interest and does not account for noise in the window that might lead to sporadic peaks at positions of interest. Additionally, the Grubbs’ outlier analysis is unable to assign sites to multiple PUS that might exhibit redundant targeting. To overcome these limitations, we have been exploring the use of a Z-score-based calculation of pseudouridylation in order to account for noise in the window.

Acknowledgments This work was supported by NIH (R01GM101316) to W.V.G. and a Jane Coffin Childs Postdoctoral Fellowship 161624T to N.M.M. References 1. Carlile TM, Rojas-Duran MF, Zinshteyn B, Shin H, Bartoli KM, Gilbert WV (2014) Pseudouridine profiling reveals regulated mRNA pseudouridylation in yeast and human cells. Nature 515:143–146. https://doi.org/10. 1038/nature13802 2. Schwartz S, Bernstein DA, Mumbach MR, Jovanovic M, Herbst RH, Leo´n-Ricardo BX, Engreitz JM, Guttman M, Satija R, Lander ES, Fink G, Regev A (2014) Transcriptome-wide mapping reveals widespread dynamic-regulated pseudouridylation of ncRNA and mRNA. Cell 159:148–162. https://doi.org/10.1016/j. cell.2014.08.028 3. Li X, Zhu P, Ma S, Song J, Bai J, Sun F, Yi C (2015) Chemical pulldown reveals dynamic pseudouridylation of the mammalian transcriptome. Nat Chem Biol 11:592–597. https:// doi.org/10.1038/nchembio.1836 4. Gilbert WV, Bell TA, Schaening C (2016) Messenger RNA modifications: form, distribution, and function. Science (80-) 352:1408–1412. https://doi.org/10.1126/science.aad8711 5. Carlile TM, Martinez NM, Schaening C, Su A, Bell TA, Zinshteyn B, Gilbert WV (2019) mRNA structure determines modification by pseudouridine synthase 1. Nat Chem Biol 15:966–974. https://doi.org/10.1038/ s41589-019-0353-z 6. Martin M (2011) Cutadapt removes adapter sequences from high-throughput sequencing reads. EMBnet J. https://doi.org/10.14806/ ej.17.1.200

7. Gordon A, Hannon GJ, Gordon (2014) FASTX-toolkit. [Online] http://hannonlab. cshl.edu/fastx_toolkit 8. Kim D, Pertea G, Trapnell C, Pimentel H, Kelley R, Salzberg SL (2013) TopHat2: accurate alignment of transcriptomes in the presence of insertions, deletions and gene fusions. Genome Biol. https://doi.org/10.1186/gb2013-14-4-r36 9. Crooks GE (2004) WebLogo: a sequence logo generator. Genome Res 14:1188–1190. https://doi.org/10.1101/gr.849004 10. Bailey TL, Johnson J, Grant CE, Noble WS (2015) The MEME suite. Nucleic Acids Res 43:W39–W49. https://doi.org/10.1093/ nar/gkv416 11. Lucks JB, Mortimer SA, Trapnell C, Luo S, Aviran S, Schroth GP, Pachter L, Doudna JA, Arkin AP (2011) Multiplexed RNA structure characterization with selective 20 -hydroxyl acylation analyzed by primer extension sequencing (SHAPE-Seq). Proc Natl Acad Sci 108:11063–11068. https://doi.org/10. 1073/pnas.1106501108 12. Rouskin S, Zubradt M, Washietl S, Kellis M, Weissman JS (2014) Genome-wide probing of RNA structure reveals active unfolding of mRNA structures in vivo. Nature 505:701–705. https://doi.org/10.1038/ nature12894

Chapter 23 Targeted RNA m6A Editing Using Engineered CRISPR-Cas9 Conjugates Xiao-Min Liu and Shu-Bing Qian Abstract N6-methyladenosine (m6A) is a major epitranscriptomic mark exerting crucial diverse roles in RNA metabolisms, including RNA stability, mRNA translation, and RNA structural rearrangement. m6A modifications at different RNA regions may have distinct molecular effects. Here, we describe a CRISPR-Cas9based approach that enables targeted m6A addition or removal on endogenous RNA molecules without altering the nucleotide sequence. By fusing a catalytically inactive Cas9 with engineered m6A modification enzymes, the programmable m6A editors are capable of achieving RNA methylation and demethylation at desired sites, facilitating dissection of regional effects of m6A and diversifying the toolkits for RNA manipulation. Key words N6-methyladenosine, CRISPR-Cas9, RNA targeting, Methylation, Demethylation

1

Introduction N6-methyladenosine (m6A) has emerged as a critical epitranscriptomic code to modulate gene expression through influencing a variety of RNA fates, such as mRNA translation, RNA stability, and RNA structural switch [1]. The biogenesis of m6A is catalyzed co-transcriptionally by a core heterodimeric methyltransferase complex consisting of METTL3 and METTL14 [2]. The discovery of demethylases FTO and ALKBH5 implicates the reversible feature of m6A [3, 4]. The dynamic nature of m6A has also been observed in response to stress [5]. Transcriptome-wide sequencing reveals that m6A preferably occurs in the consensus motif RRACH (R, purine and H, non-guanine base) along the transcripts [6, 7]. Within mRNA, m6A is enriched in the 30 untranslated region (30 UTR) and near stop codons. The asymmetric distribution suggests regional effects of m6A. For instance, m6A residues within the 50 UTR and coding regions enhance translation efficiency [8–10], whereas m6A on 30 UTR regulates RME1 mRNA levels [11]. The functionality of m6A is largely exerted through direct binding to

Mary McMahon (ed.), RNA Modifications: Methods and Protocols, Methods in Molecular Biology, vol. 2298, https://doi.org/10.1007/978-1-0716-1374-0_23, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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different “reader” proteins [12]. Additionally, m6A has been found to affect RNA-protein interactions via local structural switch [13]. Despite the remarkable advances in the functional identification of m6A modifications in cellular and physiological processes, dissecting regional or even site-specific effects of this mark on particular RNAs remains challenging. Recent advances in RNA-targeting approaches enable RNA tracking, stabilizing, and editing in living cells [14]. The discovery of clustered regularly interspaced short palindromic repeats (CRISPR) tremendously contributes to the simplicity and diversity of RNA manipulation. As a typical class 2 type II member, CRISPR-associated protein 9 (Cas9) has been widely used for precision genome editing in diverse biological systems [15]. Together with engineered single guide RNA (sgRNA), Cas9 protein effectively recognizes and cleaves the double-stranded DNA (dsDNA) bearing protospacer adjacent motif (PAM). Given this remarkable site-specific mechanism, the Cas9 system has been devised to target single-strand RNA (ssRNA) molecules in the presence of a short oligonucleotide PAMmer [16]. By fusing with green fluorescent protein (GFP) or PIN RNA endonuclease, catalytically inactive Cas9 (dCas9) enables live-cell tracking of cellular RNA localization or targeted degradation of endogenous toxic RNAs [17, 18]. With the availability of RNA-targeting Cas9 toolbox, programmable editing of m6A modifications on RNA has become feasible. We developed an engineered platform that enables programmable m6A methylation and demethylation via delivery of dCas9-METTL3-METTL14 and dCas9-ALKBH5/FTO, together with designed sgRNA and corresponding PAMmer into living cells (Fig. 1) [19]. Very recently, other groups have also successfully developed similar strategies for engineered m6A editing using alternative RNA-targeting systems [20–25]. Collectively, these powerful methodologies should facilitate a greater mechanistic understanding of the epitranscriptome. In this chapter, we describe detailed protocols for designing RNA-targeting components and three complementary techniques for quantitative m6A detection after dCas9 conjugate-mediated m6A editing. We also discuss both advantages and limitations in conducting these experiments.

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Materials

2.1 Cell Culture and Molecular Biology Reagents

1. Cell line of interest: Mouse embryonic fibroblasts (MEF) and HeLa cell lines. 2. Dulbecco’s modification of Eagle’s medium (DMEM). 3. 10% Fetal bovine serum. 4. Opti-MEM medium.

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Fig. 1 Schematic overview of targeted RNA m6A editing using engineered CRISPR-Cas9 conjugates. (a) Diagram of engineered m6A “writers” by fusing dCas9 with methyltransferase domains of METTL3 and METTL14. The sgRNA and PAMmer are base-paired to nearby regions of the m6A consensus motif on target RNA. PAM sequence NGG is supplied by PAMer that hybridizes to the target RNA. (b) Schematic diagram of engineered m6A “erasers” by fusing dCas9 with full-length ALKBH5 or FTO. The sgRNA and PAMmer are basepaired to nearby regions of the m6A site on target RNA. PAM sequence NGG is supplied by PAMer that hybridizes to the target RNA

5. Phosphate-buffered saline (PBS). 6. Transfection reagent: Lipofectamine 2000 is used for transfection of dCas9 fusion and sgRNA (see Subheading 2.2). Lipofectamine RNAiMax is used for transfection of synthesized PAMmers (see Subheading 2.2). 7. Isopropyl β-d-1-thiogalactopyranoside (IPTG). 8. E. coli competent cells: Subcloning efficiency DH5α competent cells and BL21(DE3) competent cells. 9. Luria-Bertani (LB) medium: 1% w/v Tryptone, 0.5% w/v yeast extract, and 0.5% w/v sodium chloride. 2.2 Preparation of RNA-Targeting Components

1. Backbone plasmids: dCas9 fusion editing plasmids (M3-M14-dCas9, M3D395A-M14-dCas9, ALKBH5-dCas9, ALKBH5H204A-dCas9, FTO-dCas9, and FTO FTOH231AD233A -dCas9) are derived from pCMV-BE2 vector (Addgene #73020). Targeting sgRNA plasmids originate from Cas9 sgRNA vector (Addgene #68463). These and additional backbone plasmids mentioned in the protocol are all available from Addgene (www.addgene.org). 2. Target gene-specific DNA oligos and HPLC-purified PAMer sequences (consisting of mixed DNA and 20 OMe RNA bases) are commercially synthesized by Integrated DNA

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Technologies. As an example, the sgRNA sequence and PAMer sequence for targeting MALAT1 are shown below: MALAT1 sgRNA: CAACUUAAUGUUUUUGCAUU. MALAT1 PAMer: mUTmAAmGTmUGmGGGmAT mUAmCTmCTmUGmATmCTmUG. 3. Q5 Site-Directed Mutagenesis Kit. 2.3 In Vitro Methylation Assay

1. RNA probe (50 -CGAUCCUCGGCCAGGACCAGCCUUCC CCAG -30 ) is derived from Hsp70 50 UTR and commercially synthesized from Thermo Fisher Scientific. 2. Purified M3M14, M14M3, and their inactive mutant proteins. 3. 1 M Tris buffer, pH 7.5. 4. 100 mM ZnCl2 buffer. 5. Triton-X. 6. 100 mM DL-dithiothreitol (DTT). 7. Glycerol. 8. RNaseOUT™ Recombinant Ribonuclease Inhibitor. 9. [Methyl-3H]AdoMet (PerkinElmer, NET155250UC). 10. Trizol reagent. 11. Chloroform. 12. 3 M Sodium acetate, pH 5.2. 13. 100% Ethanol. 14. Centrifuge 5424R. 15. Scintillation counting.

2.4 m6A Immunoprecipitation Coupled with RT-qPCR

1. Trizol reagent. 2. RNA fragmentation buffer: 10 mM Tris–HCl pH 7.0, 10 mM ZnCl2. 3. Anti-m6A antibody (Synaptic Systems, 202,003) and anti-m6A antibody (Millipore Cat#ABE572). 4. 1  m6A immunoprecipitation (IP) buffer: 10 mM Tris–HCl pH 7.4, 150 mM NaCl, and 0.1% Igepal CA-630. 5. RNaseOUT Recombinant Ribonuclease Inhibitor. 6. Ribonucleoside vanadyl complex (RVC). 7. Nuclease-free water. 8. m6A elution buffer: 6.7 mM N6-methyladenosine 50 -monophosphate sodium salt, 10 mM Tris–HCl pH 7.4, 150 mM NaCl, and 0.1% Igepal CA-630. 9. 0.5 M EDTA. 10. Protein A/G beads. 11. 3 M Sodium acetate, pH 5.2.

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12. Ethanol. 13. 20 mg/mL Glycogen. 14. High Capacity cDNA Reverse Transcription Kit. 15. Power SYBR Green PCR Master Mix. 16. Centrifuge 5424 R. 17. LightCycler 480 Real-Time PCR System. 18. Target gene primers: As an example, the primer sequences used for MALAT1 are shown: P1, 50 - CGTAACGGAAGTAATT CAAG-30 ; P2, 50 -GTCAATTAATGCTAGTCCTC-30 . 2.5 Site-Specific m6A Detection Assay

1. Trizol reagent. 2. Anti-m6A antibody (Synaptic Systems). 3. Immunoprecipitation buffer: 50 mM Tris–HCl, pH 7.4, 100 mM NaCl, 0.05% Igepal CA-630. 4. 3 M Sodium acetate, pH 5.2. 5. 100% Ethanol. 6. Tth DNA Polymerase and Tth buffer. 7. 25 mM MnCl2. 8. Fluorescein-12-dUTP (Jena Bioscience). 9. Novex TBE-Urea Sample Buffer (2). 10. 5 Novex TBE Running Buffer. 11. 15% Novex TBE-Urea gels. 12. Heat block. 13. Stratalinker. 14. Typhoon 9400 variable mode imager. 15. Target gene primer: As an example, the primer sequence used for MALAT1 is shown: P3, 50 - CAATTAATGCTAGTCCT CAGGATTTAAAAAATAATCTT AACTCAAAG-30 .

2.6 SELECT Assay for m6A Detection

1. Trizol reagent. 2. 3 M Sodium acetate, pH 5.2. 3. 100% Ethanol. 4. Deoxynucleotide (dNTP) solution mix. 5. CutSmart buffer: 50 mM KAc, 20 mM Tris-HAc, 10 mM MgAc2, 100 μg/mL BSA. 6. Bst 2.0 DNA polymerase. 7. SplintR ligase. 8. 100 mM Adenosine 50 -triphosphate (ATP). 9. Power SYBR Green PCR Master Mix.

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10. Mastercycler® nexus. 11. LightCycler 480 Real-Time PCR System. 12. Target gene primers: As an example, the primer sequences used for MALAT1 are shown: P4, 50 -tagccagtaccgtagtgcgtgGGATT TAAAAAATAATCTTAACTCAAAG-30 ; P5, 50 -5phos/CCAA TGCAAAAACATTAAGT cagaggctgagtcgctgcat-30 ; P6, 50 ATGCAGCGACTCAGCCTCTG -30 ; P7, 50 - TAGCCAG TACCGTAGTGCGTG-30 .

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Methods

3.1 Verification of Enzymatic Activities of Catalytic Domain Fusions of METTL3 and METTL14

Structural studies have revealed that METTL3-METTL14 heterodimer harbors strong catalytic activity of m6A biogenesis. METTL3 primarily functions as the catalytic core through binding of the DPPW motif to the methyl donor AdoMet, while METTL14 serves as an RNA-binding scaffold [26, 27]. Thus, we constructed a single-chain m6A methyltransferase (MTase) by linking two MTase domains derived from human METTL3 and METTL14, respectively. We expressed and purified two fusion proteins with different orientations of the MTase domains (Fig. 2a). As negative controls, their inactive D395A mutant proteins are also purified in parallel for the methylation assay. The methylation assay enables the measurement of methyltransferase activity using tritium (3H)labeled methyl donor S-adenosyl-methionine (SAM), as the radioactive methyl group is transferred onto the nucleoside after the reaction (Fig. 2b).

Fig. 2 Verification of enzymatic activities of catalytic domain fusions of METTL3 and METTL14. (a) Schematic diagram of the core m6A methyltransferase complex comprising methyltransferase domains of METTL3 and METTL14. M3M14 and M14M3 are generated with different orientations of MTase domains. (b) Illustration of the experimental principle for methylation assay. (c) Measurement of methyltransferase activities of fusion proteins M3M14, M14M3, and their inactive mutants using a synthesized RNA substrate (n ¼ 4, p < 0.001)

Targeted RNA m6A Editing 3.1.1 Generation of M3M14, M14M3, and Their Mutant Fusion Proteins

3.1.2 Measurement of Methyltransferase Activity

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Cloning of M3M14, M14M3, and their mutants into pGEX-6P-1 was performed as previously described [19]. Four different constructs were sequence-verified and transformed into BL21 (DE3) competent E. coli. The bacteria are grown in LB medium under 37  C for about 4 h followed by IPTG induction for another 3 h. Cells were collected and lysed for protein purification according to the manufacturer’s instructions. 1. Measure the concentration of the four purified proteins. 2. Add 100 nM purified protein with 50 μL reaction mixture containing 400 nM RNA probe, 20 mM Tris, pH 7.5, 50 μM ZnCl2, 1 mM DTT, 0.01% Triton-X, 0.2 U/μL RNaseOUT, 1% glycerol, and 0.5 μCi [methyl-3H]AdoMet (see Note 1). 3. Incubate the reaction mixture at 30  C for 1 h (see Note 2). 4. Add 75 μL of nuclease-free water and stop the reaction by adding 375 μL of Trizol reagent. 5. Homogenize the mixture by pipetting up and down several times and incubate at room temperature for 5 min. 6. Add 100 μL of chloroform and homogenize the mixture again followed by incubation for 5 min. Centrifuge the sample for 15 min at 12,000  g at 4  C. 7. Transfer the aqueous phase containing the RNA to a new tube. Add 25 μL of sodium acetate, 625 μL of 100% ethanol, and 1 μL of glycogen to the aqueous phase. 8. Incubate at 20  C for at least 2 h and centrifuge for 15 min at 12,000  g at 4  C. 9. Discard the supernatant and resuspend the pellet in 500 μL of 75% ethanol. 10. Centrifuge for 5 min at 7500  g at 4  C. Discard the supernatant and air-dry the RNA pellet for 8 min. 11. Add 20 μL of nuclease-free water to dissolve the pellet. 12. Set up nuclease-free water only as a negative control. Place test tubes containing 8 μL of purified RNA or water only into the test rack. 13. Select the right tag (3H) for scintillation counting. Start automatic counting after placing the rack in the machine (see Note 3). 14. Record the data, and remove the rack and test tubes from the scintillation counter. 15. Repeat the assay and draw the graphic panel. The radioactive levels of 3H-methyl-incorporated RNA (disintegrations per minute, DPM) are used to quantify the methyltransferase activity of different proteins (Fig. 2c).

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3.2 Preparation of RNA-Targeting Components for Methylation

As M3M14 exhibits higher activity than M14M3, M3M14-dCas9 is used for targeted methylation. While dCas9 fuses with catalytically inactive protein, M3D395AM14 serves as a negative editing control (see Note 4). Cloning of M3M14-dCas9 and M3D395AM14-dCas9 is performed as previously described [19].

3.2.1 Design of sgRNA and PAMer

sgRNA bearing λ2 sequence (CAUGGCAUUCCACUUAUCAC) serves as a non-targeting negative control. 27-nt PAMer containing mixed DNA and 20 OMe RNA bases is designed as previously described [16, 17]. It is advisable to make sgRNA-targeting sequence and PAMer-targeting sequence having eight overlapping nucleotides. The supply of PAMmer enables the specific targeting of RNA transcripts while effectively minimizing any targeting of their corresponding dsDNA template (see Note 5).

3.2.2 Transfection of Components for Targeted Methylation into MEF Cells or Cell Line of Interest

1. Grow MEF cells in growth medium in a 5% CO2 cell culture incubator with 95% humidity. Split MEF cells a day before the transfection so that the cells are approximately 80% confluent for experiments. 2. Several minutes before transfection, remove the growth medium and wash cells twice using 5 mL of PBS. Replace it with 4.8 mL of Opti-MEM medium. 3. Prepare transfection mixture containing M3M14-dCas9 and sgRNA at a mass ratio of 5:3. For one 10 cm dish of MEF cells, 7.5 μg of M3M14-dCas9 plasmid and 4.5 μg of sgRNA plasmid are mixed with 600 μL of Opti-MEM medium in a 1.5 mL Eppendorf tube. 24 μL of lipofectamine 2000 is mixed with 600 μL of Opti-MEM medium in another tube. After 5 min, mix the components of two tubes and incubate for 20 min (see Note 6). 4. Add 1.2 mL of DNA-lipid complex to the cells grown in 4.8 mL of Opti-MEM. 5. 20 min after the first transfection, prepare the transfection of PAMer. 150 pmol PAMer is mixed with 600 μL of Opti-MEM medium in an Eppendorf tube. 45 μL of lipofectamine RNAiMAX reagent is mixed with 600 μL of Opti-MEM medium in another tube. After 5 min, mix the components of two tubes and incubate for 20 min (see Note 7). 6. Add 1.2 mL of RNA-lipid complex to the cells grown in 6 mL of Opti-MEM. 7. After 6 h, add 7.2 mL of DMEM medium supplemented with 10% FBS into the cells (see Note 8). 8. Replace the medium with normal DMEM medium containing 10% FBS the next morning. 9. Harvest the transfected MEF cells for m6A detection after at least 24 h.

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3.3 Preparation of RNA-Targeting Components for Demethylation

As the non-AlkB domains are crucial for the enzymatic functionalities of ALKBH5 and FTO, full-length CDSs of both demethylases are fused to the amino terminus of dCas9. To generate inactive controls, introduce a H204A mutation to ALKBH5 and (H231A, D233A) double mutations to FTO (see Note 9).

3.3.1 Cloning of Editing Constructs for Targeted Demethylation

Full-length coding sequences of human ALKBH5 and FTO are cloned to generate ALKBH5-dCas9 and FTO-dCas9 as previously described [19]. The resulting plasmids are used as templates to generate the demethylase inactive mutant plasmids (ALKBH5H204A-dCas9 and FTOH231A/D233A-dCas9) using Q5 Site-Directed Mutagenesis Kit according to the manufacturer’s instructions.

3.3.2 Transfection of Components for Targeted Demethylation into HeLa Cells

1. Prepare transfection mixture containing ALKBH5-dCas9 and sgRNA at a mass ratio of 3:1. For one 10 cm dish of HeLa cells, 7.5 μg of ALKBH5-dCas9 plasmid and 2.5 μg of sgRNA plasmid are mixed with 600 μL of Opti-MEM medium in an Eppendorf tube. 24 μL of lipofectamine 2000 is mixed with 600 μL of Opti-MEM medium in another tube. After 5 min, mix the components of two tubes and incubate for 20 min (see Note 6). 2. Add 1.2 mL of DNA-lipid complex to the cells grown in 4.8 mL of Opti-MEM. 3. 20 min after the first transfection, prepare the transfection of the PAMer. 150 pmol PAMer is mixed with 600 μL of OptiMEM medium in a 1.5 mL of Eppendorf tube. 45 μL of lipofectamine RNAiMAX reagent is mixed with 600 μL of Opti-MEM medium in another tube. After 5 min, mix the components of the two tubes and incubate for 20 min (see Note 7). 4. Add 1.2 mL of RNA-lipid complex to the cells grown in 6 mL of Opti-MEM. 5. After 6 h, add 7.2 mL of DMEM medium supplemented with 10% FBS into the cells (see Note 8). 6. Replace the medium with standard DMEM medium containing 10% FBS the next morning. 7. Harvest the transfected HeLa cells for m6A detection after at least 24 h.

3.4 Detection of Methylation Using m6A Immunoprecipitation Coupled with RT-qPCR

In this approach, isolated total RNA is initially converted to 100–200 nucleotide (nt) fragments after random cleavage by ZnCl2. Methylated fragments are enriched for RT-qPCR analysis by immunoprecipitation using anti-m6A antibodies. It is worth noting that the probes for qPCR should be designed across the

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Fig. 3 Detection of m6A status on target site within MALAT1 transcript using three distinct approaches. (a) RT-qPCR analysis of m6A levels on A2577 of MALAT1 in HeLa cells transfected with active or inactive ALKBH5-dCas9 in the presence of targeting sgRNA (n ¼ 4, p < 0.001). (b) Site-specific detection of m6A on A2577 of MALAT1 in HeLa cells transfected with active or inactive ALKBH5-dCas9 in the presence of targeting sgRNA. The increased probe+1 signal for A2577 is a result of reduced methylation by ALKBH5-dCas9 but not the inactive mutant. ALKBH5-dCas9-mediated targeted demethylation is noneffective on a control non-targeted m6A site. (c) SELECT-guided detection of m6A levels on A2577 of MALAT1 in HeLa cells transfected with active or inactive ALKBH5-dCas9 in the presence of targeting sgRNA (n ¼ 4, p < 0.01). Ctrl indicates non-targeting sgRNA control in a, b, and c

target m6A site (Fig. 3a, upper panel). Here, we illustrated the detection of a m6A site at A2577 on MALAT1 after ALKBH5dCas9-mediated targeted demethylation (see Note 10). 1. 36–48 h after transfection, wash HeLa cells in a 10 cm dish twice with PBS. Isolate total RNA by adding 1 mL Trizol reagent. Procedures for RNA isolation are performed according to the manufacturer’s protocol (see Note 11). 2. Fragment 180 μL of isolated RNA (around 1 μg/μL) in 20 μL of freshly prepared RNA fragmentation buffer at 94  C for 5 min (see Note 12). 3. Immediately add 20 μL of 0.5 M EDTA. Vortex and spin down the tubes and place the mixture on ice. 4. To precipitate fragmented RNA, add 22 μL of 3 M sodium acetate, pH 5.2, 550 μL of 100% ethanol, and 2 μL of glycogen. Mix the contents and incubate at 20  C for at least 2 h. 5. Centrifuge for 15 min at 12,000  g at 4  C. Discard the supernatant and resuspend the pellet in 800 μL of 75% ethanol. 6. Centrifuge for 5 min at 12,000  g at 4  C and air-dry the pellet. Add 400 μL nuclease-free water to dissolve the pellet (see Note 13).

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7. Mix the remaining 375 μL of RNA with 5 μL of RNaseOUT (40 U/μL), 5 μL of RVC (200 mM), 3.5 μg anti-m6A antibody (Synaptic Systems), 3.5 μg anti-m6A antibody (Millipore), and 100 μL 5  m6A IP buffer. Rotate the mixture on a rotator in cold room for 2 h. 8. Prepare protein A/G beads by washing the beads three times using 1  m6A IP buffer. 9. Add the washed protein A/G beads to the m6A-IP mixture and rotate for 3 h at 4  C. 10. Wash the IP mixture with 1 mL of 1  m6A IP buffer four times by gentle centrifugation at 800  g for 5 min. 11. Pellet the beads after the last wash. Incubate the beads with 200 μL m6A elution buffer for 1 h at 4  C (see Note 14). 12. To precipitate the methylated RNA, add 20 μL of 3 M sodium acetate, pH 5.2, 500 μL of 100% ethanol, and 2 μL of glycogen. Mix the contents and incubate at 20  C for at least 2 h. 13. Pellet methylated RNA and wash the pellet with 800 μL of 75% ethanol. 14. Dissolve the RNA pellet in 15 μL nuclease-free water. Measure the concentration of input and immunoprecipitated RNA. 15. Prepare 20 μL of RT reaction mix containing 2 μL of 10 RT buffer, 0.8 μL of dNTP mix (100 mM), 2 μL of 10 RT random primers, 1 μL of MultiScribe™ Reverse Transcriptase, 1 μL of RNase Inhibitor, and 13.2 μL of immunoprecipitated RNA. For input RNA, use 2 μg and bring to 13.2 μL with nuclease-free water (see Note 15). 16. Place the PCR tubes in a thermal cycler using the conditions: 25  C for 10 min, 37  C for 120 min, and 85  C for 5 min. 17. cDNAs for both methylated and input RNAs are used for qPCR. Prepare 10 μL of qPCR mix containing 5 μL of 2 SYBR Green Master Mix, 2 μL of forward primer P1 (1 μM), 2 μL of reverse primer P2 (1 μM), and 1 μL of cDNA. 18. Run qPCR under the following conditions: 95  C, 5 min; 40 cycles of 95  C, 15 s and 60  C, 45 s. Quantify the methylated RNA and evaluate the efficiency of targeted demethylation by comparing the relative methylated RNA between targeting and non-targeting samples (Fig. 3a) (see Note 16). 3.5 Detection of m6A Using Site-Specific Methylation Assay

The site-specific m6A detection assay couples m6A antibody crosslinking with fluorescein-dUTP incorporation during primer elongation. After cross-linking, m6A antibodies fixed to methylated adenosine substantially block the incorporation of labeled dUTP during reverse transcription (Fig. 3b, upper panel). On a scanned gel, the low fluorescein intensity of transcribed products at the

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expected size should reflect high m6A levels at specific adenosine sites on RNA. Here, we illustrated the detection of the m6A site A2577 on MALAT1 after ALKBH5-dCas9-mediated targeted demethylation. 1. 36–48 h after transfection, wash HeLa cells in a 60 mm dish twice with PBS. Isolate total RNA by adding 1 mL of Trizol reagent. Procedures for RNA isolation are performed according to the manufacturer’s instructions. 2. Dilute 20 μg total RNA in 450 μL immunoprecipitation buffer and incubate with 1 μg anti-m6A antibody (Synaptic Systems) by rotating at 4  C for 2 h. 3. The antibody-RNA mixture is cross-linked twice with 0.15 J/ cm2 of 254 nm UV light using a Stratalinker. 4. Precipitate RNA by adding 1 mL of ethanol, 45 μL of sodium acetate, and 2 μL of glycogen. Mix the contents and incubate at 20  C for at least 2 h. 5. Centrifuge for 15 min at 12,000  g at 4  C. Discard the supernatant and resuspend the pellet in 1 mL of 75% ethanol. 6. Centrifuge for 5 min at 12,000  g at 4  C and air-dry the pellet. Add 10 μL of nuclease-free water to dissolve the pellet. 7. Perform reverse transcription in a 7 μL of reaction containing m6A-cross-linked RNA, Tth buffer, and 50 pmole primer P3. 8. Anneal the reaction mixture by heating at 95  C for 10 min and gradually cooling to room temperature. 9. Add 5 U of Tth enzyme and 1 mM MnCl2 to the mixture and heat at 70  C for 3 min (see Note 17). 10. Perform primer extension by adding 1 μM fluorescein-12dUTP and incubate for 15 min at 70  C (see Note 18). 11. Add 10 μL 2 TBE-urea dye and heat for 3 min. Resolve the products immediately on a 15% Novex TBE-urea gel (see Note 19). 12. Monitor the fluorescein signal by placing the gel in a Typhoon 9400 variable mode imager or similar. Evaluate the efficiency of targeted demethylation by comparing the signal intensity between targeting and non-targeting samples (Fig. 3b) (see Note 20). 3.6 Detection of Methylation Using SELECT Assay

SELECT method utilizes the capacity of m6A to impede the Bst DNA polymerase-mediated single-base elongation and SplintR ligase-mediated nick ligation [28]. After two rounds of selection, the amount of final products generated from m6A-containing RNA is much less than products from unmodified RNA. qPCR can be used to quantify the products. A low amount of qPCR products would indicate high m6A levels at specific adenosine on RNA. The

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SELECT products of indicated site should be normalized to the RNA abundance of indicated transcript bearing this site. Here, we illustrated the detection of the m6A site A2577 on MALAT1 after ALKBH5-dCas9-mediated targeted demethylation. 1. 36–48 h after transfection, wash HeLa cells in 60 mm dish twice using PBS. Isolate total RNA by adding 1 mL of Trizol reagent. Procedures for RNA isolation are performed as described in the manufacturer’s manual. 2. Incubate 5 ug total RNA with 40 nM Down Primer P4, 40 nM Up Primer P5, and 5 μM dNTP in 17 μL of 1 CutSmart buffer and anneal at a temperature gradient: 90  C for 1 min, 80  C for 1 min, 70  C for 1 min, 60  C for 1 min, 50  C for 1 min, and then 40  C for 6 min. 3. Incubate 17 μL of the annealing products with a 3 μL of enzyme mixture (0.01 U Bst 2.0 DNA polymerase, 0.5 U SplintR ligase, and 10 nmol ATP) at 40  C for 20 min. Denature the samples by heating to 80  C for 20 min (see Note 21). 4. In parallel, 2 μg of total RNA is synthesized into cDNA in 20 μL of RT reaction mix containing 2 μL of 10 RT buffer, 0.8 μL of dNTP mix (100 mM), 2 μL of 10 RT random primers, 1 μL of MultiScribe™ Reverse Transcriptase, 1 μL of RNase Inhibitor, and nuclease-free water (see Note 22) following the manufacturer’s instructions. 5. Prepare 10 μL of qPCR mix for both ligated product and target transcript. The reaction contains 5 μL of 2 SYBR Green Master Mix, 1 μL of forward primer P6 (2 μM), 1 μL of reverse primer P7 (2 μM), and 3 μL of cDNA. 6. Run qPCR on a LightCycler 480 Real-Time PCR System at the following condition: 95  C, 5 min; (95  C, 10 s; 60  C, 45 s)  40 cycles. 7. Normalize the SELECT products of indicated m6A site to the amount of indicated m6A-containing RNA. Evaluate the efficiency of targeted demethylation by comparing the normalized SELECT product between targeting and non-targeting samples (Fig. 3c).

4

Notes 1. Always wear protective clothing and disposable gloves when working with radioactive [methyl-3H]AdoMet in designated areas. 2. It is necessary to set up a negative control reaction which contains a protein (e.g., BSA) without methyltransferase activity.

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3. Always clean and perform wipe tests of the experimental area and equipment to avoid any radioactive contamination in the laboratory. 4. To examine the effectiveness of targeted m6A addition, it is recommended to select candidate transcripts with low basal methylation at desired sites. 5. M3M14-dCas9-mediated targeted methylation is more effective in a window of around ten nucleotides. The secondary structure and binding status of RNA may affect the functional window and editing efficiency. 6. The mass ratio of dCas9 editing plasmid and sgRNA for transfection may need to be optimized for different cell types based on the expression efficiency of dCas9 fusion proteins and endogenous abundance of target RNA. 7. To exclude the possibility that the binding of the fusion enzyme or targeting sgRNA may influence the methylation status of the target RNA, it is recommended to set up two negative controls (methyltransferase inactive control and non-targeting sgRNA control) when preparing targeting components for transfection. 8. To ensure efficient transfection, the components for targeted m6A editing are retained instead of being removed from the medium. 9. It is recommended to choose candidate RNAs harboring high basal methylation at desired loci when investigating the effectiveness of targeted m6A demethylation. 10. Multiple pairs of primers can be designed to test the qPCR efficiency for probing methylated RNA. Amplicons with a length of 100–200 base pairs are ideal for qPCR analysis. 11. The abundance of endogenous target RNA may affect the efficiency of RT-qPCR analysis. It is recommended to enrich more methylated RNA fragments before RT-qPCR validation by preparing more cells for transfection if transcript level is low. 12. It is recommended to closely follow the fragmentation time and temperature to generate short transcripts with a length of 100–200 nt. 13. Save 25 μL as input for normalization of the m6A IP. 14. To minimize nonspecific binding, it is essential to elute methylated RNA by competition rather than by Trizol extraction of the entire bead-antibody conjugates. 15. It is suggested to use all the eluted m6A-containing RNAs in order to increase the amplification signal of the RT-qPCR.

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16. Quantify m6A levels of a particular RNA by normalizing to the corresponding input RNA amount. In parallel, methylation of other m6A sites can be examined to assess potential off-target effect. 17. Tth enzyme and buffer can be replaced with other reverse transcriptases and reaction buffers (e.g., SuperScript III). Remember to change the optimal temperature for probe extension. 18. Protect the fluorescein-12-dUTP from exposure to light during the procedure. 19. It is necessary to denature the RT product and load singlestrand DNA markers before running the gel. 20. Primer elongation can be conducted for alternative m6A sites to evaluate potential off-target activity of programmable editors. 21. The elongation and nick ligation are conducted in one reaction to simplify the procedure and minimize the degradation of RNA. 22. To reduce detection bias due to varied amounts of target RNA in different samples, the basal levels of target transcript should also be quantified.

Acknowledgments This work was supported by US National Institutes of Health (R21CA227917, R01GM1222814) and HHMI Faculty Scholar (55108556). References 1. Shi H, Wei J, He C (2019) Where, when, and how: context-dependent functions of RNA methylation writers, readers, and erasers. Mol Cell 74(4):640–650 2. Liu J, Yue Y, Han D, Wang X, Fu Y, Zhang L, Jia G, Yu M, Lu Z, Deng X, Dai Q, Chen W, He C (2014) A METTL3-METTL14 complex mediates mammalian nuclear RNA N6-adenosine methylation. Nat Chem Biol 10(2):93–95 3. Jia G, Fu Y, Zhao X, Dai Q, Zheng G, Yang Y, Yi C, Lindahl T, Pan T, Yang YG, He C (2011) N6-methyladenosine in nuclear RNA is a major substrate of the obesity-associated FTO. Nat Chem Biol 7(12):885–887 4. Zheng G, Dahl JA, Niu Y, Fedorcsak P, Huang CM, Li CJ, Va˚gbø CB, Shi Y, Wang WL, Song SH et al (2013) ALKBH5 is a mammalian RNA

demethylase that impacts RNA metabolism and mouse fertility. Mol Cell 49(1):18–29 5. Meyer KD, Jaffrey SR (2014) The dynamic epitranscriptome: N6-methyladenosine and gene expression control. Nat Rev Mol Cell Biol 15(5):313–326 6. Meyer KD, Saletore Y, Zumbo P, Elemento O, Mason CE, Jaffrey SR (2012) Comprehensive analysis of mRNA methylation reveals enrichment in 30 UTRs and near stop codons. Cell 149(7):1635–1646 7. Dominissini D, Moshitch-Moshkovitz S, Schwartz S, Salmon-Divon M, Ungar L, Osenberg S, Cesarkas K, Jacob-Hirsch J, Amariglio N, Kupiec M, Sorek R, Rechavi G (2012) Topology of the human and mouse m6A RNA methylomes revealed by m6A-seq. Nature 485(7397):201–206

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8. Zhou J, Wan J, Gao X, Zhang X, Jaffrey SR, Qian SB (2015) Dynamic m6A mRNA methylation directs translational control of heat shock response. Nature 526(7574):591–594 9. Meyer KD, Patil DP, Zhou J, Zinoviev A, Skabkin MA, Elemento O, Pestova TV, Qian SB, Jaffrey SR (2015) 50 UTR m6A promotes cap-independent translation. Cell 163 (4):999–1010 10. Mao Y, Dong L, Liu XM, Guo J, Ma H, Shen B, Qian SB (2019) m6A in mRNA coding regions promotes translation via the RNA helicase-containing YTHDC2. Nat Commun 10(1):1–11 11. Bushkin GG, Pincus D, Morgan JT, Richardson K, Lewis C, Chan SH, Bartel DP, Fink GR (2019) m6A modification of a 30 UTR site reduces RME1 mRNA levels to promote meiosis. Nat Commun 10(1):1–13 12. Patil DP, Pickering BF, Jaffrey SR (2017) Reading m6A in the transcriptome: m6A-binding proteins. Trends Cell Biol 28 (2):113–127 13. Liu N, Dai Q, Zheng G, He C, Parisien M, Pan T (2015) N6-methyladenosine-dependent RNA structural switches regulate RNA-protein interactions. Nature 518 (7540):560–564 14. Smargon AA, Shi YJ, Yeo GW (2020) RNA-targeting CRISPR systems from metagenomic discovery to transcriptomic engineering. Nat Cell Biol 22(2):143–150 15. Wang H, La Russa M, Qi LS (2016) CRISPR/ Cas9 in genome editing and beyond. Annu Rev Biochem 85(1):227–264 16. O’Connell MR, Oakes BL, Sternberg SH, East-Seletsky A, Kaplan M, Doudna JA (2014) Programmable RNA recognition and cleavage by CRISPR/Cas9. Nature 516 (7530):263–266 17. Nelles DA, Fang MY, O’Connell MR, Xu JL, Markmiller SJ, Doudna JA, Yeo GW (2016) Programmable RNA tracking in live cells with CRISPR/Cas9. Cell 165(2):488–496 18. Batra R, Nelles DA, Pirie E, Blue SM, Marina RJ, Wang H, Chaim IA, Thomas JD, Zhang N, Nguyen V, Aigner S, Markmiller S, Xia G, Corbett KD, Swanson MS, Yeo GW (2017) Elimination of toxic microsatellite repeat expansion

RNA by RNA-targeting Cas9. Cell 170 (5):899–912 19. Liu XM, Zhou J, Mao Y, Ji Q, Qian SB (2019) Programmable RNA N6-methyladenosine editing by CRISPR-Cas9 conjugates. Nat Chem Biol 15(9):865–871 20. Rau K, Ro¨sner L, Rentmeister A (2019) Sequence-specific m6A demethylation in RNA by FTO fused to RCas9. RNA 25 (10):1311–1323 21. Shinoda K, Suda A, Otonari K, Futaki S, Imanishi M (2020) Programmable RNA methylation and demethylation using PUF RNA binding proteins. Chem Commun 56 (9):1365–1368 22. Liu J, Dou X, Chen C, Chen C, Liu C, Xu MM, Zhao S, Shen B, Gao Y, Han D, He C (2020) N6-methyladenosine of chromosomeassociated regulatory RNA regulates chromatin state and transcription. Science (80- ) 367 (6477):580–586 23. Li J, Chen Z, Chen F, Xie G, Ling Y, Peng Y, Lin Y, Luo N, Chiang CM, Wang H (2020) Targeted mRNA demethylation using an engineered dCas13b-ALKBH5 fusion protein. Nucleic Acids Res 48(10):5684–5694 24. Zhao J, Li B, Ma J, Jin W, Ma X (2020) Photoactivatable RNA N6-Methyladenosine editing with CRISPR-Cas13. Small 1907301:1–8 25. Wilson C, Chen PJ, Miao Z, Liu DR (2020) Programmable m6A modification of cellular RNAs with a Cas13-directed methyltransferase. Nat Biotechnol 38:1431–1440 26. Wang X, Feng J, Xue Y, Guan Z, Zhang D, Liu Z, Gong Z, Wang Q, Huang J, Tang C, Zou T, Yin P (2016) Structural basis of N6-adenosine methylation by the METTL3METTL14 complex. Nature 534 (7608):575–578 27. Wang P, Doxtader KA, Nam Y (2016) Structural basis for cooperative function of Mettl3 and Mettl14 methyltransferases. Mol Cell 63 (2):306–317 28. Xiao Y, Wang Y, Tang Q, Wei L, Zhang X, Jia G (2018) An elongation- and ligation-based qPCR amplification method for the radiolabeling-free detection of locus-specific N6-methyladenosine modification. Angew Chem Int Ed 57(49):15995–16000

INDEX A Activated charcoal Norit® ............................................. 363 Adapter Depletion Solution ................................ 161, 162 Adapter/sRNA co-fold structures ............................... 154 ADP-ribosylation ARTs ........................................................................ 231 cellular processes ..................................................... 231 DarTG system ......................................................... 232 human DNA repair PARPs..................................... 232 NAD+ ....................................................................... 231 PARPs ...................................................................... 231 posttranslational modification process ................... 231 RNA ......................................................................... 232 toxin-antitoxin system ............................................ 232 ADP-ribosylhydrolase enzyme ..................................... 231 ADP-ribosyltransferase protein .................................... 233 ADP-ribosyltransferases (ARTs)................................... 231 Affinity gel electrophoresis ........................................... 218 Agencourt RNAClean XP bead purifications ...................................... 59, 65, 66 Agilent 2100 Bioanalyzer .........................................83, 94 Agilent TapeStation ........................................................ 34 Agilent’s “Source Optimizer” software .............. 296, 297 Agilent’s MassHunter Workstation Software .............. 297 AG-rich sequence ............................................................ 16 Alkaline hydrolysis.............................................. 82, 87, 88 AlkAniline-Seq technology alkaline hydrolysis......................................... 82, 87, 88 aniline cleavage ....................................................83, 89 bioinformatic analysis..........................................83, 92 extensive RNA dephosphorylation ....................82, 88 library preparation........................................ 83, 89, 90 library purification..................................................... 90 library quality assessment ............................ 83, 91, 92 library quantification.................................... 83, 90, 91 library sequencing ...............................................83, 92 PCR purification ....................................................... 83 RNA extraction ......................................................... 79 RNA quality assessment......................................86, 87 RNA quantification .............................................82, 86 total RNA extraction TRIzol™................................................. 81, 85, 86 yeast/bacteria ......................................... 79, 84, 85 ALKBH5 ......................................................................... 32

ALKBH5-dCas9-mediated targeted demethylation ..................................... 408, 410 ALKBH8 ....................................................................... 198 Alternative media .......................................................... 290 Amersham Hybond-XL membrane .................... 202, 209 Amino acids ................................................................... 217 Aniline cleavage ............................................................... 83 Anti-reverse cap analogs (ARCAs) ............................... 344 APB affinity gel electrophoresis..........219, 220, 222, 224 APB Northern analysis.................................................. 229 Applications of RNA modifications gene expression regulation ..................................... 342 modified nucleotides............................................... 342 mRNA termini ............................................... 344, 345 RNA-based biosensors ............................................ 342 synthetic mRNA.................................... 343, 345, 346 Application-specific integrated circuit (ASIC) .............. 53 Arabidopsis thaliana ..................................................... 198

B Bacteria ............................................................................ 93 Bacterial sonication buffer ................................... 199, 203 BAM mapping file ........................................................... 22 Barcoded reverse PCR primers..................................... 383 Base-calling...................................................................... 41 dependency................................................................ 48 differential.................................................................. 47 EpiNano-Error .......................................................... 42 modification probabilities ......................................... 42 RNA modifications ................................................... 43 samples....................................................................... 42 systematic.............................................................42, 46 BED format ..................................................................... 22 Beta-mercaptoethanol................................................... 194 Bioanalyzer electrodes .................................................... 93 Biochemical assays......................................................... 348 Bioinformatic analysis ..................................................... 79 Biological regulation ..................................................... 247 Bisulfite conversion ....................................................... 135 Bisulfite sequencing ...................................................... 336 Blue-light illumination.................................................. 396 Brassica napus gel purification ..................................... 164 Brassica napus total RNA ............................................. 160 BstI enzyme.......................................................... 186, 194

Mary McMahon (ed.), RNA Modifications: Methods and Protocols, Methods in Molecular Biology, vol. 2298, https://doi.org/10.1007/978-1-0716-1374-0, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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416 Index

AND

PROTOCOLS

C Calling PTMs ChIP-seq experiments................................................. 9 m1A peaks .................................................................... 9 Ψ sites........................................................................... 9 Canonical nucleoside isotopologues ............................ 305 Canonical nucleotides DNA oligonucleotides .............................................. 61 5S and 5.8S rRNA synthesis..................................... 63 gel purification .......................................................... 61 PCR synthesis ............................................................ 65 RNA Ampure XP bead purifications ........................ 64 Capillary electrophoresis................................................. 79 Carbodiimide (CMC) ................................................... 380 CD3-methionine-labeled culture ................................. 303 cDNA 3’-linker ligation................................................ 104 cDNA 50 adapter ligation strategy ............................... 396 cDNA molecules ............................................................... 6 Cell-based assays .................................................. 347, 348 Cell extracts ................................................................... 319 Cell-free in vitro translation systems............................ 348 Cell metabolism ............................................................ 332 Cellular immune receptors ........................................... 347 Cellular processes .......................................................... 231 Cellular RNAs ............................................................... 135 Cfr enzymes................................................................... 106 CG-/AU-rich sequences .............................................. 304 Charcoal extraction ....................................................... 375 Chemical-assisted RNA fragmentation ........................ 103 Classical phenol:chloroform extraction ....................... 161 CLIP-based methods .................................................... 106 Clustered regularly interspaced short palindromic repeats (CRISPR) ........................................ 400 CMC-modified RNA .................................................... 396 Coding sequence (CDS)............................................... 330 Comparative NAIL-MS experiments ........................... 289 Concatemerization ........................................................ 154 Conventional nanopore sequencing .............................. 71 Coomassie gel................................................................ 212 Coomassie stain protocols ............................................ 204 Coverage analysis ............................................................ 24 CRISPR-associated protein 9 (Cas9)........................... 400 CRISPR-Cas9-based approach cell culture ...................................................... 400, 401 in vitro methylation assay ....................................... 402 m6A detection assay....................................... 403, 404 m6A immunoprecipitation ............................ 402, 403 m6A immunoprecipitation coupled with RT-qPCR ............................................ 407–409 METTL3-METTL14 heterodimer catalytic activity ................................................. 404 methyltransferase activity measurement........... 405 pGEX-6P-1........................................................ 405

molecular biology reagents............................ 400, 401 RNA-targeting components preparation...... 401, 402 RNA-targeting components, demethylation cloning ............................................................... 407 non-AlkB domains ............................................ 407 transfection ........................................................ 407 RNA-targeting components, methylation M3D395AM14 ................................................. 406 MEF cells ........................................................... 406 sgRNA and PAMer design ............................... 406 SELECT assay ................................................ 410, 411 site-specific m6A methylation assay .............. 408, 410 Curlcake sequences ......................................................... 32 Cutadapt .......................................................................... 22 Cy3-labeled RNA oligo ................................................ 239 Cytosine-50 -triphosphate (CTP) .................................. 360 Cytosol........................................................................... 328

D Dark Reader transilluminator..................... 155, 159, 163 DarTG system ............................................................... 232 Deacylation ........................................................... 221–223 Deadenylated mRNAs .................................................. 334 Delta-delta Ct (ddCt) .......................................... 180, 181 Demultiplexing.............................................................. 164 Denaturing urea PAGE gel.................................. 232, 235 Deoxynucleotide (dNTP).................................... 110, 403 DESeq2-adjusted P-values ........................................... 120 Diode array detector (DAD) ........................................ 287 Direct RNA nanopore sequencing (dRNA-seq) .............................................32, 34 Direct RNA sequencing................................................ 265 adapter ligation....................................................38, 39 input RNA ................................................................. 38 RMX adapter ligation .........................................40, 41 RT ........................................................................39, 40 S. cerevisiae................................................................. 35 Direct RNA sequencing datasets base-calling ................................................... 41, 43, 44 current intensity values features extraction.............. 44 distinct strategies ....................................................... 42 mapping ..................................................................... 42 programs/modules ................................................... 42 Disease Ontology ............................................................ 24 DisGeNET disease genes................................................ 24 DMEM D0422 ............................................................. 290 DMEM medium .................................................. 303, 407 DMS-seq........................................................................ 394 DNA LoBind tube .......................................................... 66 DNA oligo sequences ................................................... 361 DNA oligonucleotides .................................................. 208 DNA oligos .......................................................... 210, 373 dNTPs alternative concentrations ................................ 183 Double-stranded DNA (dsDNA) ................................ 400

RNA MODIFICATIONS: METHODS DRACH........................................................................... 16 D-tube Dialyzers ............................................................. 72 Dynamic multiple reaction monitoring (dMRM).............................................. 297, 299

E E. coli BL21 (DE3) ....................................................... 373 Electropherogram representation ................................ 164 Electrophoresis ..................................................... 228, 372 Engineered CRISPR-Cas9 conjugates......................... 401 Ensembl Bacteria Genome Database (EMBL-EBI)................................................ 117 Enzymatic digestion, RNA phosphodiesterase I............................... 249, 252, 253 S1 nuclease digestion .............................................. 252 Enzyme-RNA covalent ................................................. 106 EpiNano 1.2 suite ........................................................... 43 EpiNano algorithm datasets....................................................................... 34 dRNA-seq .................................................................. 34 in vitro-transcribed constructs (see Modified/ unmodified in vitro-transcribed constructs preparation) IVT RNAs ...........................................................32, 33 polyA tailing ........................................................33, 34 RNA base modifications detection........................... 32 softwares .................................................................... 34 EpiNano-Error ................................................................ 42 base-calling error....................................................... 48 Epinano_Predict module .......................................... 47 RNA-modified sites.............................................48, 49 EpiNano script folder...................................................... 43 EpiNano suite.................................................................. 42 EpiNano-SVM ................................................................. 42 predict RNA modifications.................................45, 46 train EpiNano models .........................................44, 45 Epitranscriptome ........................................................... 288 Epitranscriptomic RNA modifications......................... 123 Epitranscriptomics ............................................... 262, 327 Eppendorf DNA LoBind tube ....................................... 65 Ethanol .......................................................................... 119 Ethanol precipitation .................................................... 396 Eukaryotic cells and tissues .......................................... 255 Eukaryotic translation................................................... 349 Eukaryotic translation initiation factor 2 (eIF2α) ......................................................... 345 Extensive RNA dephosphorylation..........................82, 88

F FASTA sequences............................................................ 42 FASTQ files ........................................................ 5, 11, 150 FASTX toolkit ............................................................... 131 Fibrillarin (FBL) ............................................................ 172

AND

PROTOCOLS Index 417

FileZilla .......................................................................... 120 FLAG-tagged enzyme.......................................... 108, 111 Fluidigm Access Array Integrated Fluidic Circuit (IFC)....................................... 137, 139 Fluorescein-12-dUTP................................................... 413 Formamide .................................................................... 159 Forward experiment...................................................... 303

G Galaxy web platform ..................................................... 116 Gamma-purification lanes............................................. 205 Gamma-toxin assay ....................................................... 212 Gamma-toxin expression ..................................... 199, 203 Gamma-toxin in vitro assay .......................................... 201 Gamma-toxin induction ............................................... 203 Gamma-toxin protein ................................................... 211 Gamma-toxin purification .......................... 200, 203, 204 GENCODE database...................................................... 17 Gene expression ............................................................ 331 Gene Ontology (GO) ..................................................... 24 GeneJET PCR Purification ......................................83, 90 Gene-specific primer (GSP)................................. 174–176 Gene-specific sequence (GS) ........................................ 143 GGACA k-mers ............................................................... 45 Glass wool filtration ...................................................... 375 Glucose .......................................................................... 227 GlycoBlue ...................................................................... 119 Glycoblue™ coprecipitant.............................................. 82 Glycosidic bond............................................................. 281 Green fluorescent protein (GFP) ................................. 400 Grubbs’ outlier analysis ................................................ 397 Guanosine-50 -triphosphate (GTP)............................... 360 Guppy base-caller .......................................................41, 48

H High definition (HD) adapters ........................... 154, 156 High-dNTP conditions ................................................ 181 High-performance liquid chromatography (HPLC)...................................... 280, 296, 304 High-resolution LC-MS ............................................... 287 High-throughput in vitro pseudouridylation assay cDNA circularization .............................................. 391 circularization master mix components ........ 391, 392 CMC treatment..................................... 381, 387, 388 DNA pool design .................................................... 383 mutations........................................................... 394 PUS recognition features.................................. 394 DNA pool PCR amplification ....................... 383, 384 DNA sequences pool .............................................. 380 extension master mix .............................................. 389 HF buffer................................................................. 392 IVT RNA ........................................................ 385, 386 kinetic analysis ......................................................... 393

RNA MODIFICATIONS: METHODS

418 Index

AND

PROTOCOLS

High-throughput in vitro pseudouridylation assay (cont.) NGS and kinetic analysis......................................... 395 NGS and pseudouridine detection ............... 392, 393 PCR reaction ........................................................... 392 recombinant PUS.................................. 381, 386, 387 RT and library construction ................. 383, 389, 390 time course ..................................................... 394, 395 μL phusion............................................................... 392 μM RP1 primer ....................................................... 392 High-throughput sequencing ........................................ 77 HiScribe T7 In Vitro Transcription Kit protocol ....... 140 HiScribeTM T7 Quick High Yield RNA Synthesis...................................................61, 65 Homogenization ........................................................... 206 Hot acid phenol .............................................................. 79 Housekeeping (HK) ..................................................... 181 HTSeq-count script ...................................................... 117 Human 18S rRNA .......................................................... 54 Human reference genome sequence (hg19) ................. 17 Hybridization .................... 202, 213, 220, 221, 223–226 Hydrolysis............................................................. 234, 240

I IgG IP negative control................................................ 124 Illumina Experimental Manager software...................... 94 Illumina MiSeq..................................................... 137, 149 Illumina multiplex primer (IDT) ................................. 110 Illumina sequencers ............................................... 94, 164 Immunoprecipitation FLAG-tagged protein binding ............................... 112 FLAG-tagged protein elution ................................ 112 resin preparation...................................................... 111 resin recycling.......................................................... 112 In vitro methylation assay............................................. 402 In vitro MGFP transcript.............................................. 149 In vitro transcription (IVT)............................................ 61 In vitro-transcribed 5’-m7G-capped RNA .................. 254 In vitro-transcribed mRNAs......................................... 340 In vitro-transcribed RNA ............................................. 347 In vitro-transcribed RNAs (IVT RNAs) ..................32, 33 Inorganic pyrophosphatase (IPPase) .................. 361, 373 Inosine ........................................................................... 334 Integrated fluidic circuit (IFC) ........................... 145–147 Integrated Genome Viewer (IGV).......12, 117, 118, 131 Internal m7G in mRNA analytical method validation ................................... 254 biological samples ..................................250–252, 257 chemicals and reagents................................... 249, 250 differential enzymatic digestion phosphodiesterase I..........................249, 252, 253 S1 nuclease digestion ............................... 249, 252 enzymatic digestion ................................................ 254 equipment................................................................ 251 eukaryotic cells and tissues ..................................... 255

5’-m7G-capped RNA synthesis..................... 250, 251 LC-ESI-MS/MS ..................................................... 255 nucleoside standards ............................................... 248 nucleosides analysis, LC-ESI-MS/MS................... 253 oligonucleotides ...................................................... 248 rice sample ............................................................... 256 Internal standard (ISTD) ............................................. 289 Ion-exchange chromatography .................................... 309 IP RC files ......................................................................... 9 Isopropyl ß-d-1-thiogalactopyranoside (IPTG).......... 199 Isotopologues calibration ................................................................ 305 labeled/unlabeled canonicals ................................. 301 modified nucleosides............................................... 281 NAIL-MS samples................................................... 289 nucleobase ............................................................... 281 physicochemical attributes...................................... 280 precursor/product ion............................................ 297 RNA molecules ....................................................... 289 SILIS ............................................................... 297, 305 IVT-derived rRNA strands ............................................. 61 IVT-generated rRNA ...................................................... 71 IVT reactions.............................................................72, 73

J JBrowse integration visualization................................... 25

K KAPA analysis ................................................................ 116 Kapa HiFi Hotstart PCR kit......................................... 163 KAPA library quantification kit .................................... 116 Klen Taq V669L ........................................................... 173 Kluyveromyces lactis ....................................................... 199

L Labeled adenine ............................................................ 303 Labeled isotopologues .................................................. 303 Labeled methionine ...................................................... 303 LC-MS/MS absolute quantification............................ 300 LC-MS/MS analysis ..................................................... 281 LC-MS/MS measurement calibration ................................................................ 295 data analysis ............................................................. 297 dMRM method ....................................................... 297 gradient.................................................................... 296 HPLC ...................................................................... 296 individual nucleosides optimization....................... 297 isotopologues .......................................................... 297 modified nucleosides............................................... 301 MS signals................................................................ 297 nucleoside m7G ....................................................... 301 nucleosides and regression curves ................. 299, 301 quantitative MassHunter software ......................... 301

RNA MODIFICATIONS: METHODS source optimization ................................................ 296 Labeled nucleosides ...................................................... 303 LC-MS buffer................................................................ 304 LC-MS vials ................................................................... 287 Library preparation, miCLIP-MaPseq PCR amplification ................................................... 115 R1R DNA................................................................ 115 RNA 30 end dephosphorylation ............................. 114 TGIRT-III, template-switching reaction............... 114 thermostable ligation .............................................. 115 Library quantification ................................................... 111 Ligation bias .................................................................. 154 Linear polyacrylamide (LPA)............................... 201, 207 Linearized 28S plasmid DNA ........................................ 72 Liquid chromatography-coupled tandem mass spectrometry (LC-MS/MS) ....................... 279 Liquid chromatography-electrospray ionization-tandem mass spectrometry (LC-ESI-MS/MS)......................248, 252–255 Liquid chromatography-mass spectrometry (LC-MS) ............................................. 218, 262 Long noncoding RNA (lncRNA) ...................23, 31, 137 Low dNTP methods engineered enzymes ................................................ 173 primer extension assay ............................................ 172 RT ............................................................................ 173 RTL-P ..................................................................... 174, (see also ReverseTranscription Under Low dNTP Conditions Followed by PCR (RTL-P)) SuperScript III......................................................... 173 Luria-Bertani (LB) ........................................................ 199 Lysing yeast cells ........................................................... 319 Lysis and DNase treatment .......................................... 111

M m1A peaks calling .............................................................. 9 m1A-seq read mapping ..................................................... 7 m1Ψ-containing mRNAs.............................................. 342 M3M14-dCas9-mediated targeted methylation ......... 412 m5C differential chemical reactivity............................. 135 m6A ............................................................................... 349 m6A epitranscriptome .................................................... 16 m6A immunoprecipitation coupled with RT-qPCR ............................................ 402, 403 antibodies ................................................................ 407 isolated RNA ........................................................... 408 methylated RNA ..................................................... 409 PCR tubes ............................................................... 409 protein A/G beads.................................................. 409 RT reaction .............................................................. 409 targeted demethylation ........................................... 409 transfection .............................................................. 408 m6A MeRIP-seq BAM files index ......................................................... 21

AND

PROTOCOLS Index 419

data and software ...................................................... 17 extract FASTQ format files....................................... 19 library preparation and sequencing....................17, 18 modification site/peak calling .................................. 21 quality control ........................................................... 20 RNAmod ................................................................... 17 samples....................................................................... 19 sequencing quality..................................................... 19 short read mapping ................................................... 20 m6A methylation .......................................................... 332 m6A methyltransferase ................................................. 404 m6A modification .................................................. 15, 329 m6A modification BED formats .................................... 23 m6A RNA modifications ................................................ 32 m6A RT-QPCR-based approach biological samples ................................................... 193 Cts difference calculation ....................................... 192 data analysis ............................................................. 192 primer design QPCR ....................................................... 188, 189 RT ............................................................. 188, 189 QPCR analysis ................................................ 188, 191 RNA extraction/quantification .................... 186, 187, 189, 190 RT ................................................................... 187, 190 schematic representation ........................................ 187 m6A-containing RNAs ................................................. 412 m6A-LAIC-seq.............................................................. 186 m6Am-containing mRNAs........................................... 333 m6Am modification ...................................................... 333 m7G per tRNA quantification ...................................... 301 m7G-seq chemical-assisted method cDNA 3’-ligation .................................................... 102 cDNA 3’-linker ....................................................... 100 conversion into abasic site .......................99, 101, 102 data processing and analysis ................................... 102 fragmented mRNA preparation .......................98–100 misincorporation ....................................................... 97 mRNA decapping............................................. 99, 100 PCR amplification .......................................... 100, 102 RT ...............................................................98, 99, 102 3’-adapter ligation.............................................99–101 Mass spectrometry-based analysis ................................ 248 Mass spectrometry (MS)............................. 172, 279, 280 Maxima RT (MRT) enzyme......................................... 187 mcm5s2U modification gamma-toxin endonuclease .................................... 199 gamma-toxin expression ................................ 199, 203 gamma-toxin in vitro assay ..................................... 201 gamma-toxin induction .......................................... 203 gamma-toxin nuclease assay ................................... 208 gamma-toxin purification ..................... 200, 203, 204 multicellular eukaryotes .......................................... 198

RNA MODIFICATIONS: METHODS

420 Index

AND

PROTOCOLS

mcm5s2U modification (cont.) neurological and neurodevelopmental disorders....................................................... 197 Northern blot probe design ................................... 208 Northern blotting protocol........................... 201, 202 Northern Northern blotting analysis............ 208–210 organisms................................................................. 197 purified protein coomassie detection .................................... 201, 204, 205 qRT-PCR analysis .......................................... 202, 211 RNA extraction ....................................................... 201 RNA preparation............................................ 205–207 single-cell organisms ............................................... 198 stress response pathways ......................................... 197 tRNA............................................................... 197–199 mcm5s2U-dependent endonucleolytic cleavage ......... 208 Membrane stripping...................................................... 226 MEME ........................................................................... 394 meRanCall ..................................................................... 148 meRanCompare ............................................................ 148 meRanTK ...................................................................... 148 meRIP/m6A-seq .......................................................... 124 MeRIP-seq.............................................................. 16, 186 Messenger RNA (mRNA) ..................................... 15, 357 chemical modifications............................................ 328 next-generation sequencing technologies ............. 327 Meta-gene coverage plots...........................................9, 10 Methylated RNA immunoprecipitation and sequencing (MeRIP-seq) .................................................. 15 Methylation ................................................................... 335 Methylation-iCLIP (miCLIP) approach...................... 106 Methyl-RNA immunoprecipitation (meRIP).............. 123 Methyltransferase (METTL3) ........................................ 15 Mettl3 knockdown (shMETTL3).................................. 23 MGFP in vitro transcripts ............................................ 140 Michaelis constant (KM) ............................................... 369 Michaelis-Menten equation.......................................... 369 Michaelis-Menten kinetic parameters .......................... 358 miCLIP-MaPseq advantages................................................................ 107 cell lysis ........................................................... 107, 108 FLAG-tagged enzyme............................................. 111 gel purification and RNA extraction ...................... 113 immunoprecipitation .................... 106–108, 111–112 library preparation..................................110, 114–115 library quantification............................................... 111 lysis and DNase treatment ...................................... 111 proteinase K treatment ........................................... 112 qPCR quantification ............................................... 116 RNA fragmentation .............................. 110, 113, 114 RNA isolation.......................................................... 108 sequencing ............................................................... 111 sequencing read mapping and analysis ......... 116–117 Micro Bio-Spin gel........................................................ 150

Microcentrifuge.................................................... 160, 376 MicroRNA (miRNA) ............................................. 24, 153 MidRand adapters ......................................................... 154 Minimum free energy (MFE)......................................... 24 MINLEN parameter ....................................................... 94 MiSeq Reagent Kit v3................................................... 148 MiSeq sequencing ......................................................... 146 Misincorporation........................................................... 103 Modification calling tools ............................................... 22 Modified ribonucleosides ............................................. 343 Modified/unmodified in vitro-transcribed constructs preparation AmpliScribe™ T7 High Yield Transcription ...........................................36, 37 enzymatic digestion ............................................35, 36 plasmid transformation and isolation....................... 35 polyA tailing ........................................................37, 38 RNAClean XP bead stock......................................... 38 RNeasy Qiagen Kit ................................................... 37 MODOMICS database................................................. 119 Molecular feature extraction (MFE)............................ 273 Monoclonal anti-FLAG M2-peroxidase ...................... 119 Monosaccharides ........................................................... 217 Mouse embryonic fibroblasts (MEF)........................... 400 mRNA 3’ termini ................................................. 333, 334 mRNA 5’ cap........................................................ 330, 333 mRNA life cycle .............................................................. 16 chemical modifications.......................... 329, 330, 332 cytosol ...................................................................... 328 eukaryotic pre-mRNAs ........................................... 328 gene expression in eukaryotes ................................ 328 protein–mRNA interaction..................................... 328 RNA–RNA interactions .......................................... 328 mRNA modifications ............................................. 16, 379 gene expression ....................................................... 248 m7G (see Internal m7G in mRNA) qualitative and quantitative analysis ....................... 255 translation (see mRNA translation) mRNA termini modifications .............................. 347, 348 mRNA translation ......................................................... 399 ac4C ......................................................................... 337 cell-based assays....................................................... 346 functional consequences ......................................... 330 hm5C .............................................................. 338, 339 human health and biology...................................... 350 inosine...................................................................... 334 internal modifications .................................... 348, 349 m1A ......................................................................... 335 m3C ................................................................ 336, 337 m5C ......................................................................... 336 m5U................................................................ 337, 338 m7G ......................................................................... 335 molecular mechanisms ............................................ 350 mRNA 3’ termini ........................................... 333, 334

RNA MODIFICATIONS: METHODS mRNA 5’ cap.................................................. 330, 333 mRNA life cycle ...................................................... 346 mRNA stability........................................................ 347 mRNA termini modifications ........................ 347, 348 pseudouridine........................................ 339, 340, 342 RNAmodification applications (see Applications of RNA modifications) structural and biochemical analyses ....................... 350 mRNA-based therapeutics............................................ 343 MS ladder sequencing, RNA acid degradation, RNA ........................................... 270 acid-mediated hydrolysis......................................... 264 anchor-based computational algorithm ................ 273, 274, 276 biotin/streptavidin capture and release ........ 267, 268 CMC conversion ................................... 268, 271, 275 direct sequencing .................................................... 265 epitranscriptomics ................................................... 262 isomeric nucleotides................................................ 263 ladder components/fragments............................... 262 LC-MS analysis............................................... 262, 273 LC-MS elution buffers ........................................... 268 manually reading sequences, RNA sample mixture....................................... 273, 274, 276 mass spectrometer ................................................... 262 mixed RNA sample ............................... 270, 271, 275 1D MS sequencing.................................................. 263 peptides.................................................................... 261 RNA 3´-end labeling and biotin ................... 269, 274 RNA acid hydrolysis................................................ 262 RNA mixtures ......................................................... 263 streptavidin beads, biotinylated RNA ................... 269, 270, 275 synthetic RNA oligonucleotides.................... 266, 267 synthetic single RNA .............................................. 265 3´-end labeling protocol ......................................... 267 2D data analysis....................................................... 263 2D LC-MS-based method...................................... 263 2D mass-tR plots ..................................................... 272 2D-HELS MS Seq ......................................... 265, 266 MS-based transcriptome-wide m5C mapping............. 336 MSigDB functional gene sets ......................................... 24 MTase domains ............................................................. 404 Multiple-turnover pseudouridine formation ............... 368 MultiScribe™ Reverse Transcriptase ........................... 411 Multistep thermal cycling conditions .......................... 147 Mutation of C118 (C118A)......................................... 106

N N1-methyladenosine (m1A) .....................................5, 335 N4-acetylcytidine (ac4C)..................................... 126, 337 N6-methyladenosine (m6A)................... 15, 31, 105, 126 biological processes ................................................. 185 critical epitranscriptomic code................................ 399

AND

PROTOCOLS Index 421

dynamic nature ........................................................ 399 internal RNA modification ..................................... 185 miCLIP/IP.............................................................. 186 modifications ........................................................... 185 RNA-protein interactions ....................................... 400 3’ UTR .................................................................... 399 transcriptome-wide sequencing.............................. 399 translation efficiency................................................ 185 N7-methylguanosine (m7G) .......................................... 16 5’-m7G-cap structure ............................................. 248 mammals.................................................................. 248 mass spectrometry-based analysis........................... 248 methylation....................................................... 97, 335 METTL1 ................................................................. 335 mRNA 5’ cap........................................................... 335 mRNAs .................................................................... 335 NaBH4-mediated reduction .................................... 97 RNA modification ..................................................... 97 rRNA ....................................................................... 248 tRNA........................................................................ 248 N-acryloyl-3-aminophenylboronic acid (APB) ........... 218 NAIL-MS experiments calibration ................................................................ 295 categories ................................................................. 288 cell culture ............................................. 280, 281, 302 comparative .................................................... 288, 289 digestion ......................................................... 286, 287 epitranscriptome...................................................... 288 HEK293 cells labelling .................................. 290, 292 isotopologues ................................................. 280, 281 LC-MS/MS (see LC-MS/MS measurement) modified nucleosides............................................... 281 pulse-chase............................................................... 289 RNA digestion/filtration............................... 294, 295 RNA isolation.......................................................... 292 RNA isolation and purification .............................. 286 RNA purification oligonucleotide hybridization assay ........ 293, 294 SEC ........................................................... 292, 293 sensitive quantification............................................ 281 SILIS preparation.......................... 286, 289–291, 303 Nanodrop ...................................................................... 190 Nanopolish ....................................................................... 44 Nanopore-based human nuclear-encoded rRNA sequencing adapter ligation and cleanup ..............................59, 60 anticipated throughput ............................................. 61 base-calling ................................................................ 61 biological human rRNA............................................ 55 canonical nucleotides .......................................... 61–65 canonical rRNAs.................................................. 55–57 data analysis and visualization .................................. 61 linearized DNA plasmid bearing .............................. 66 oligomer splint preparation ...................................... 58

RNA MODIFICATIONS: METHODS

422 Index

AND

PROTOCOLS

Nanopore-based human nuclear-encoded rRNA sequencing (cont.) PCR protocol ............................................................ 65 reaction assemble, PCR tube..............................65, 66 RNA isolation......................................................56, 58 splint annealing and ligation ..............................58, 59 total RNA isolation ................................................... 55 XhoI restriction digest (see XhoI restriction digest, plasmids) Nanopore genomic DNA ............................................... 54 Nanopore sequencing ..................................................... 53 Nanopore strand ............................................................. 63 Native RNA ..................................................................... 32 Natural-occurring RNA modification density ............. 305 NCBI Sequence Read Archive ......................................... 5 NCBI/Primer-BLAST.................................................. 189 Ncs2-Ncs6 thiolase ....................................................... 198 N-cyclohexyl-N’-(2-morpholinoethyl)carbodiimide metho-p-toluenesulfonate (CMC) ................................... 5, 268, 271, 275 NEBNext® Multiplex Small RNA Library Prep Set for Illumina® ........................................................ 93 Network of Cancer Gene................................................ 24 Neural networks .............................................................. 53 NEXTflex V3 kit ........................................................... 162 Next-generation sequencing (NGS) .............16, 124, 153 Nicotinamide adenine dinucleotide phosphate (NADPH) ........................................... 316, 319 NMR spectroscopy........................................................ 319 Nm-seq .......................................................................... 172 Noncoding RNAs ................................................ 171, 327 Non-phosphorylated RNA oligo (noP ssRNA)....................................... 235, 237 Nonradioactive cyanine-tagged RNA oligos .......................................... 234, 239, 240 Nonradioactive UTP..................................................... 373 Normalization control .................................................. 202 Northern blot......................................220, 221, 223–226 Northern blot probe design ......................................... 208 Northern blotting ......................................................... 218 Northern blotting analysis................................... 208–210 Northern blotting DNA oligos .................................... 212 Northern blotting protocol................................. 201, 202 NSun RNA methyltransferase ...................................... 106 Nucleic acid isotope labeling-coupled mass spectrometry (NAIL-MS) .................................................. 280 Nucleoside modifications ............................................. 135 Nucleoside standards .................................................... 248 Nucleosides.................................................................... 282

O Oligo synthesis .............................................................. 395 Oligonucleotide hybridization ................... 209, 293, 294 Oligonucleotides ............................................70, 157, 248

One-step thermal cycling conditions ........................... 145 OPCR melting curve analysis .............................. 191, 192 Optimized QQQ parameters ....................................... 298 Oxford Nanopore Technologies ..............................41, 60

P PAGE purification ......................................................... 372 PAMer-targeting sequence ........................................... 406 PA-mod-seq................................................. 125, 126, 132 Parallel amplification ..................................................... 136 PARalyzer ...................................................................... 131 PARP-like orthologous proteins .................................. 232 PATH environment ........................................................ 22 PCR amplicons..................................................... 137, 150 PCR amplification ..........................................94, 143, 154 PCR cycles ............................................................ 166, 395 PCR optimization ........................................................... 71 PCR primer ............................................................ 71, 150 PCR-amplified library ................................................... 163 PCR-based template ....................................................... 73 PEG8000 concentration............................................... 104 Periodate oxidation .............................................. 222, 223 pET28a plasmids........................................................... 203 Pharmacodynamic (PD) ............................................... 343 Pharmacokinetic (PK)................................................... 343 Phenol:chloroform extraction ...................................... 161 Phenol:choroform:isoamyl alcohol (PCA) .................... 93 PhiX library.................................................................... 150 Phosphodiesterase I digestion ............................. 252, 253 Photoactivatable ribonucleoside-enhanced cross-linking and immunoprecipitation (PAR-CLIP)................................................. 124 Photo-crosslinking-assisted m6A sequencing (PA-m6A-seq) antibody-RNA complexes....................................... 124 IP..................................................................... 126–129 poly(A) purification................................................. 128 RNA elution ................................................... 127, 130 RNA end repair ..................................... 127, 129, 130 RNA modifications ................................................. 124 RNA preparation..................................................... 126 RNase footprinting step ......................................... 125 RNA-seq read peaks................................................ 126 sequencing data analysis ......................................... 131 sequencing libraries preparation.................... 127, 130 tissue culture................................................... 126, 127 total RNA extraction............................................... 128 Pico RNA chip ................................................................ 93 Piwi-interacting RNAs (piRNAs)................................. 153 Plant miRNAs ............................................................... 154 Plasmid DNA isolation ................................................... 66 p-nitrophenylphosphate (pNPP).................................. 132 Poly ADP-ribose polymerase (PARPs) ........................ 231 Poly(A)-binding proteins (PABPs) .............................. 333

RNA MODIFICATIONS: METHODS Poly(A)-specific ribonuclease (PARN)......................... 344 PolyA tailing .................................................................... 39 Polyethylene glycol (PEG) ........................................... 155 Posttranscriptional modifications, tRNAs biosynthesis ............................................................. 309 cellular extract preparation ................... 311, 312, 315 cellular stress............................................................ 309 EDTA....................................................................... 320 elution fractions ...................................................... 320 in extract NMR ....................................................... 310 NADPH................................................................... 319 NMR monitoring..........................313, 316, 318, 321 NMR spectroscopy.................................308–310, 320 reagents.................................................................... 310 RNA transcription................................................... 320 RNAs ....................................................................... 307 T7 in vitro transcription ......................................... 318 time-resolved NMR ....................................... 309, 317 transcription and purification .......310, 311, 313, 314 ultrapure deionized water....................................... 310 unlabeled cellular extract ........................................ 309 yeast cellular extracts............................................... 321 yeast extracts............................................................ 309 yeast tRNAPhe NMR sample .................................. 321 Preadenylated 3’ adapter .............................................. 154 Pre-trained SVM models ................................................ 46 Primer sequences........................................................... 187 Primestar GXL polymerase ............................................. 73 Protein/non-RNA analysis........................................... 304 Proteinase K treatment ................................................. 109 Protein–mRNA interactions ......................................... 349 Protein-RNA cross-link formation............................... 107 Protein-RNA interaction assay ..................................... 124 Protein-RNA species..................................................... 107 Proteomics..................................................................... 261 Protospacer adjacent motif (PAM) .............................. 400 Pseudo-Seq analysis......................................................... 11 Pseudo-Seq data mapping ................................................ 7 Pseudo-seq signal ................................................. 393, 397 Pseudouridine (Ψ)............................................5, 105, 135 C–C glycosidic isomerization, uridine base........... 339 discovery .................................................................. 379 eIF2α........................................................................ 345 formation ................................................................. 374 gene expression ....................................................... 340 human diseases ........................................................ 342 in vitro transcription ............................................... 349 intermolecular RNA–RNA ..................................... 340 m1Ψ ......................................................................... 345 m6A modification ................................................... 329 mammalian mRNA ................................................. 339 mRNA secondary structures .................................. 342 mRNA stability............................................... 340, 342 mRNA............................................................. 341, 342

AND

PROTOCOLS Index 423

NAP57/DKC1 ....................................................... 339 nucleobase ............................................................... 358 posttranscriptional RNA modifications ................. 339 protein expression ................................................... 345 RNA–protein interactions ...................................... 340 stand-alone enzymes/H/ACA............................... 357 structural isomer ..................................................... 357 translation ................................................................ 345 tritium release assay................................................. 358 uridine modification................................................ 357 Pseudouridine-containing RNAs ................................. 396 Pseudouridine synthases (PUS) attacking C20 position............................................. 358 bacterial.................................................................... 376 characterizing steps ................................................. 359 concentrations ................................................ 372, 375 dilution .................................................................... 369 [5-3H]-labeled substrate RNAs.............................. 365 [5-3H]-substrate RNA............................................ 376 H/ACA snoRNP .................................................... 363 high-throughput ..................................................... 380 isomerization ........................................................... 358 RNA modification ................................................... 377 Pseudouridylation ......................................................... 339 PTM-containing RNA fragments .................................... 3 PTM mapping techniques ................................................ 6 Pulse-chase NAIL-MS experiments ............................. 289

Q QPCR primer design .................................................... 188 QPCR primers...................................................... 183, 193 Q-tRNA modifications APB affinity gel electrophoresis.................... 219, 220, 222, 224 biological materials.................................................. 219 deacylation ...................................................... 221–223 detection ................................................ 218, 225, 226 5’-End Labeling ...................................................... 225 human and mouse cells........................................... 219 hybridization ................................. 220, 221, 223–226 membrane stripping ................................................ 226 in mouse cells ................................................. 222, 223 northern blot................................. 220, 221, 223–226 nutrition-dependent levels...................................... 218 periodate oxidation ........................................ 222, 223 quantification........................................................... 226 semidry transfer ....................................................... 223 tissues ....................................................................... 219 TRIzol reagent ........................................................ 221 Quality trimming .............................................................. 7 Quantitative PCR (qPCR)................................... 153, 186 Qubit dsDNA Broad Range Assay Kit......................... 146 Qubit fluorometer RNA BR assay ................................. 68 Qubit fluorometer RNA HS assay ................................. 60

RNA MODIFICATIONS: METHODS

424 Index

AND

PROTOCOLS

QubitTM BR RNA Assay kit.......................................... 69 QubitTM dsDNA BR assay kit ...................................... 65 Quench-flow apparatus................................................. 359 Quenching medium ...................................................... 303 Queuosine (Q) RNA modifications ................................................. 218 7-deazaguanine ....................................................... 217 synthesize................................................................. 218

R RA1 buffer..................................................................... 194 Radical SAM RNA-methylating enzymes radical-based mechanism ........................................ 106 RNA m5C methyltransferases ................................ 106 substrates and modification sites ............................ 106 Radioactive [methyl-3H]AdoMet................................ 411 Radioactive enzyme assay ............................................. 360 Radioactive isotope tritium .......................................... 358 Radioactive NAD+ ............................................... 235–237 Radioactively labeled RNA oligos ....................... 237, 238 Radioactivity ......................................................... 233, 234 Randomization .............................................................. 154 RC index file (RCI)........................................................... 9 Reactive oxygen species ................................................ 338 Read 1 sequencing primer (R1R DNA) ...................... 115 Read mapping FastQC tool................................................................. 7 m1A-seq ....................................................................... 7 Pseudo-Seq data .......................................................... 7 Real-time quantitative PCR (RT-qPCR) ..................... 175 Receiver operating characteristic (ROC) ....................... 45 Recombinant PUS ..............................381, 386, 387, 396 Retention time window (ΔRT) .................................... 297 Retrotranscription (RT)................................................ 186 Reverse transcriptase (RT).............................5, 6, 98, 154 Reverse Transcription Under Low dNTP Conditions Followed by PCR (RTL-P) cell cultivation and collection ................................. 177 cell culture ............................................................... 176 GSP .......................................................................... 175 high-and low-dNTP conditions .................... 178, 179 mRNA...................................................................... 175 Nm site .................................................................... 175 qPCR analysis ................................................. 179, 180 quantification.................................................. 180, 181 RNA extraction .............................................. 177, 178 RNA isolation.......................................................... 176 RT and qPCR .......................................................... 176 RT placement .......................................................... 175 RT-qPCR................................................................. 175 sensitivity ................................................................. 175 snoRNA-guided Nm...................................... 175, 177 rf-rctools .......................................................................... 12 RiboMeth-seq ............................................................... 172

Ribonucleoside modifications ...................................... 217 Ribo-seq analyses .......................................................... 337 Ribosomal RNA (rRNA) ................................... 5, 24, 357 Ribosome......................................................................... 54 RibOxi-seq..................................................................... 172 RIP experiments.............................................................. 11 RlmN enzymes ............................................ 106, 108, 111 RlmN-mediated methylation........................................ 107 RNA 3’-ligation ............................................................ 103 RNA 30 end dephosphorylation ................................... 114 RNA ADP-ribosylation................................................. 232 denaturing urea PAGE gel............................. 232, 235 hydrolysis ........................................................ 234, 240 nonradioactive cyanine-tagged RNA oligos.......................................... 234, 239, 240 pre-run gel ............................................................... 235 radioactive NAD+ ........................................... 235–237 radioactively labeled RNA oligos .................. 237–239 radioactivity .................................................... 233, 234 RNA oligo preparation ........................................... 235 RNA Ampure XP bead purifications .............................. 64 RNA-based biosensors .................................................. 342 RNA-based therapeutics ............................................... 343 RNA-binding proteins (RBPs) ....................................... 16 RNA bisulfite conversion method bioinformatics analysis ............................................ 148 bisulfite oligonucleotide primer design ........ 141, 143 cDNA synthesis ..................................... 138, 142, 144 IFC.................................................................. 145–147 individual PCR amplification......................... 144, 145 IVT.................................................................. 137, 140 library sequencing components ............................. 139 MiSeq sequencing .......................................... 146, 148 multiNA microelectrophoresis system ................... 139 PCR amplicon amplification................................... 139 PCR amplicon purification/quantification............ 139 RNase-free/DNase-free H2O................................ 137 RT ............................................................................ 142 second-generation sequencing ............................... 137 sodium bisulfite solution preparation................................. 138, 140, 141 total RNA extraction...................................... 137, 140 RNA degradation ........................................ 193, 396, 413 RNA digestion............................................................... 287 RNA Framework calling PTMs ......................................................... 9–10 data retrieval ................................................................ 5 GitHub repository ...................................................... 4 output files and interpretation ...........................10, 11 PATH........................................................................... 4 read counting .............................................................. 8 reads mapping ............................................................. 7 reference index ............................................................ 6 refGene ........................................................................ 6

RNA MODIFICATIONS: METHODS SAM/BAM format ..................................................... 5 UNIX-based OS.......................................................... 4 XML files ................................................................... 11 RNA immunoprecipitation (IP)................................... 124 RNA integrity.................................................................. 79 RNA ligase 1 (RNL1)................................................... 154 RNA metabolism........................................................... 123 RNA methylation .......................................................... 105 RNA methyltransferases................................................ 106 RNA modification mapping ......................................... 124 RNA modifications ....................................................... 105 commercial antibodies .............................................. 32 discovery .................................................................... 15 dynamic modifications .............................................. 16 epitranscriptome..................................................15, 16 learning algorithms ................................................... 16 MeRIP-seq data ........................................................ 16 MS technique .......................................................... 279 NGS ........................................................................... 32 pseudouridine............................................................ 31 sequencing techniques ............................................ 279 single-molecule resolution........................................ 32 throughout evolution ............................................... 77 transcriptome............................................................. 15 RNA modification tritium release assay data analysis .................................................... 368, 369 [5-3H]-substrate RNA crush and soak purification..............362, 364–367 IVT...........................................360, 361, 363, 364 limitations ................................................................ 360 multiple turnover conditions.................................. 374 pseudouridine................................................. 358, 359 pseudouridylation ................................................... 360 quench-flow........................................... 363, 370, 372 reaction buffer ......................................................... 363 RNA pseudouridylation .......................................... 358 S. cerevisiae H/ACA snoRNP ....................... 365, 368 single-turnover reactions ........................................ 359 RNA pellet............................................................ 133, 374 RNA posttranscriptional modification (PTMs) discovery ...................................................................... 3 high-throughput sequencing methods ...................... 3 RIP approaches ........................................................... 3 RNA structure ............................................................. 3 RNA precipitation ......................................................... 304 RNA pseudouridylation liquid scintillation analyzer ..................................... 358 Michaelis-Menten kinetic parameters .................... 374 PUS .......................................................................... 358 quench-flow apparatus ............................................ 369 RNA concentration................................................. 375 rRNA fragment ....................................................... 365 tritium release assay (see RNA modification tritium release assay)

AND

PROTOCOLS Index 425

yeast PUS1 .............................................................. 380 RNA re-precipitations................................................... 132 RNA sequence/structure effects.................................. 154 RNA structure mapping ................................................... 6 RNA transcription......................................................... 320 RNAClean XP bead stock............................................... 38 RNA-methylating enzymes .......................................... 105 RNAmod ......................................................................... 16 analysis outputs ................................................... 23–25 data input................................................................... 22 default parameters ..................................................... 23 flexible parameters..................................................... 22 functional modules.................................................... 18 GitHub ...................................................................... 22 JBrowse tool .............................................................. 22 workflow .................................................................... 18 RNA-modified sites...................................................48, 49 RNA-modifying enzymes .................................... 106, 379 RNase contamination ................................................... 205 RNase inhibitor ............................................................... 56 RNase-free water .................................................. 207, 363 RNA-seq-based Nm mapping methods....................... 172 RNAsnap protocol ........................................................ 205 RNA-targeting approaches ............................................................... 400 Cas9 toolbox ........................................................... 400 components ............................................................. 400 systems ..................................................................... 400 RRACH k-mers.........................................................46, 48 rRNA subunits .............................................................. 304 RT enzymes ................................................................... 103 RT primer extension ....................................................... 79 RT primer sequences .................................................... 188 RT reaction .................................................................... 191 RT SuperScript III ........................................................ 173 RTL-P quantification .................................................... 181 RT-qPCR analysis ................................................ 408, 412

S S. cerevisiae rRNAs ............................................................ 6 S1 nuclease digestion ........................................... 249, 252 S-adenosyl-L-methionine (SAM) ........................ 316, 404 SAM/BAM files ..........................................................8, 17 SAMtools ................................................................ 61, 131 SCARLET...................................................................... 186 SDS solution.................................................................. 227 SDS-PAGE electrophoresis ................................. 204, 205 Second-generation sequencing .................................... 135 SEC-purified tRNA....................................................... 294 SELECT-guided detection ........................................... 408 SELECT products................................................ 410, 411 Semidry transfer of RNAs............................................. 223 SeqLogo plots ................................................................. 24 Sequencing .................................................................... 280

RNA MODIFICATIONS: METHODS

426 Index

AND

PROTOCOLS

Sequencing read mapping and analysis enrichment analysis ................................................. 117 sequence processing/alignment.................... 116, 117 stop sites/mismatches............................................. 117 sgRNA-targeting sequence........................................... 406 Shearing ......................................................................... 150 Shimadzu Microchip Electrophoresis System MCE®-202 MultiNA .................................. 144 Shrimp Alkaline Phosphatase (rSAP) ............................. 99 SILAC proteomics experiments ................................... 288 SILISGen1 labelling........................................................ 302 SILISGen2 labelling........................................................ 289 Simian virus 40 (SV40)................................................. 125 Single guide RNA (sgRNA) ......................................... 400 Single-chain m6A methyltransferase (MTase)............. 404 Single-nucleotide primer extension ............................. 186 Single-strand RNA (ssRNA) ............................... 233, 400 Single-turnover pseudouridine formation ................... 371 Site-specific m6A detection assay ................................. 408 Size-adjusted concentration ......................................... 116 Size-exclusion chromatography (SEC) ............... 292, 293 Small interfering RNAs (siRNAs) ................................ 153 Small nuclear RNA (snRNA) ....................................... 357 Small nucleolar ribonucleoprotein (snoRNP)............. 357 Small nucleolar RNAs (snoRNAs) ............................... 339 Small RNAs (sRNAs) eukaryotic regulatory .............................................. 153 expression levels ...................................................... 154 insects and plants..................................................... 154 library preparation process ..................................... 154 miRNA profiles ....................................................... 154 molecules ................................................................. 154 NGS ......................................................................... 153 quantitative research tools ...................................... 153 sRNA library preparation protocol (see TS5 protocol) snoRNA KO ......................................................... 182, 183 snoRNA-guided 20 -O-methylation .............................. 174 snoRNA-guided methylations...................................... 175 Sodium bisulfite solution.............................................. 148 Sodium borohydride (NaBH4)...................................... 79 SRA accession IDs............................................................. 5 sRNA-adapter ligation .................................................. 154 SRA database ................................................................... 50 SRA toolkit ...................................................................... 17 sRNA circularization ..................................................... 154 Stable isotope labeling compounds .................... 280, 291 Standard protocol (TS)................................................. 155 Static trimming strategy.................................................. 12 Streptavidin-coated Dynabeads® .................................. 304 Subsequent alkaline phosphatase (rSAP) ..................... 103 Substrate RNA concentration ...................................... 375 Sulfuric acid (H2SO4) ................................................... 319 SUPERaselIn™ RNase Inhibitor ................................ 103

SuperScript III Reverse Transcriptase (SSIII RT) .....................................40, 173, 202 Support vector machine (SVM) ..................................... 47 SYBR Green Real-Time PCR Master Mix................... 211 Synthetic mRNA vaccines............................................. 343 Synthetic mRNAs................................................. 343–348 Synthetic RNA oligonucleotides ......................... 266, 267

T Tandem mass spectrometry (MS/MS) ........................ 281 TapeStation................................................................39, 48 Targeted m6A editing................................................... 412 TBE-urea denaturing polyacrylamide gel .................... 160 Temperature-adjustable shaker..................................... 211 TGIRT reverse transcriptase ......................................... 107 Therapeutic modality .................................................... 343 Thermo Fisher Trizol LS protocol............................... 205 ThermoPol buffer ......................................................... 193 Thermostable 50 AppDNA/RNA Ligase .................... 115 Thermostable group II intron reverse transcriptase (TGIRT) .....................................................107, 110, 120 Thermostable ligation................................................... 115 Thiolated nucleosides ................................................... 304 Time-resolved NMR ..................................................... 317 tiRNA purification ........................................................ 304 Toll-like receptors (TLRs) ............................................ 346 Total RNA extraction ..................................................... 54 Traditional fragmentation buffer ................................. 103 Train EpiNano models..............................................44, 45 Transcriptome-wide analyses ............................... 332, 335 Transcriptome-wide sequencing methods ................... 379 Transfer RNA (tRNA) .................................................. 357 anticodon stem-loop ............................................... 217 posttranscriptional modifications (see Posttranscriptional modifications, tRNAs) Translation end sits (TES) .............................................. 24 Translation start sites (TSS) ........................................... 24 Trim Galore ..................................................................... 17 Tris-EDTA (TE).............................................................. 93 Tritium release reaction buffer ..................................... 362 TRIzol reagent .............................................................. 221 TRIzol™ extraction ........................................................ 93 Trm9p-Trm112p methyltransferase............................. 198 TRMT9B genes............................................................. 198 tRNA modification analysis ..................................................................... 218 enzymatic digestion and LC-MS/MS ................... 218 tRNA-modifying enzymes ............................................ 396 tRNA sequence ............................................................. 208 TRPT1 .................................................................. 232, 233 TS5 protocol data analysis

RNA MODIFICATIONS: METHODS raw sequence files treatment.................... 164, 165 trimmed sequences mapping ................... 165, 166 library preparation gel purification .................................158, 163, 164 5’ adapter ligation .................................... 158, 162 PCR amplification ............................................. 158 PCR amplifications............................................ 163 RT ............................................................. 158, 162 unligated 3’ adapter elimination ..................................158, 161, 162 3’ adapter ligation .................................... 158, 161 preadenylated 3’ HD adapter preparation................................. 156, 160, 161 small RNAs isolation............................................... 155 sRNA isolation ...................................... 155, 159, 160 TUG1 transcript............................................................ 188 Two-dimensional (2D) ................................................. 262 Two-dimensional hydrophobic end-labeling strategy to MS-based sequencing (2D-HELS MS Seq)........................... 265, 266 2’-O-methylation (Nm)................................................ 338 chemistry ................................................................. 172 dNTPs...................................................................... 174 mRNA and pre-mRNA........................................... 171 noncoding RNAs .................................................... 171 nucleoside ribose ..................................................... 172 posttranscriptional ribose modification ................. 171 RT ................................................................... 172, 173 snoRNAs.................................................................. 172 Two-step thermal cycling conditions........................... 145

AND

PROTOCOLS Index 427

U UCSC genome database................................................... 6 Ultrapure water ............................................................. 218 UNIX/Linux operating system ..................................... 17 Unlabeled nucleosides .................................................. 303 Untranslated regions (UTRs)....................................... 330 Uridine-50 -triphosphate (UTP) ................................... 361 Uridine-containing mRNAs ......................................... 345

W Wash buffer (WSB) ......................................................... 60 Watson-Crick base pairing ............................................ 213 Western blot analysis ..................................................... 119 WIGGLE tracks............................................................... 12

X XhoI restriction digest, plasmids Agencourt RNAClean XP bead purification ........... 67 canonical transcripts purification........................68, 69 DNA concentration .................................................. 67 electrophoresis and gel purification ......................... 67 library preparation, polyadenylated canonical 28S rRNA.............................................................. 70 library preparation, splint adapters.....................69, 70

Y Yeast cellular extracts .................................................... 321 Yeast H/ACA snoRNPs ............................................... 372 Yeast Saccharomyces cerevisiae ....................................... 198 Yeast tRNAPhe ............................310, 314, 316, 318, 321