RNA Remodeling Proteins: Methods and Protocols [2nd ed.] 9781071609347, 9781071609354

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Methods in Molecular Biology 2209

Marc Boudvillain Editor

RNA Remodeling Proteins Methods and Protocols Second Edition

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 Remodeling Proteins Methods and Protocols Second Edition

Edited by

Marc Boudvillain Centre de Biophysique Moléculaire, CNRS UPR4301, Orleans Cedex 2, France

Editor Marc Boudvillain Centre de Biophysique Mole´culaire CNRS UPR4301 Orleans Cedex 2, France

ISSN 1064-3745 ISSN 1940-6029 (electronic) Methods in Molecular Biology ISBN 978-1-0716-0934-7 ISBN 978-1-0716-0935-4 (eBook) https://doi.org/10.1007/978-1-0716-0935-4 © Springer Science+Business Media, LLC, part of Springer Nature 2015, 2021 This work is subject to copyright. All rights are reserved by the Publisher, whether the whole or part of the material is concerned, specifically the rights of translation, reprinting, reuse of illustrations, recitation, broadcasting, reproduction on microfilms or in any other physical way, and transmission or information storage and retrieval, electronic adaptation, computer software, or by similar or dissimilar methodology now known or hereafter developed. The use of general descriptive names, registered names, trademarks, service marks, etc. in this publication does not imply, even in the absence of a specific statement, that such names are exempt from the relevant protective laws and regulations and therefore free for general use. The publisher, the authors, and the editors are safe to assume that the advice and information in this book are believed to be true and accurate at the date of publication. Neither the publisher nor the authors or the editors give a warranty, expressed or implied, with respect to the material contained herein or for any errors or omissions that may have been made. The publisher remains neutral with regard to jurisdictional claims in published maps and institutional affiliations. This 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 Five years after the first edition of the Methods in Molecular Biology volume on RNA remodeling proteins, we have the pleasure to provide the scientific community with a new collection of cutting-edge approaches devoted to the study of this seminal class of proteins. The tremendous progress of RNA biology has confirmed the obvious: yes, RNA functions rest on specific structures and partnerships with proteins. The often complex and highly dynamic ballet of RNA–protein interactions usually starts as soon as the nascent transcript emerges from the transcription machinery. Then, throughout the RNA life cycle, proteins bind and release the RNA chain, remodeling RNA structures and interactions to drive the maturation, quality control, transport, storage, utilization, or destruction of RNA as a function of cellular needs. Different classes of proteins contribute to remodel RNA and RNA–protein complexes. The case of NTP-independent “RNA chaperones” has been addressed in the 2015 edition of this book and in a new, separate volume of the series (RNA chaperones, volume 2106; Heise, Tilman, Ed.). Here, we more specifically focus on proteins that use the energy derived from NTP cofactor (RNA helicases) or RNA phosphodiester backbone (ribonucleases) hydrolysis to remodel RNA or ribonucleoprotein structures. RNA helicases trigger conformational rearrangements but do not modify the RNA chain itself, whereas ribonucleases irreversibly alter the structures of their RNA substrates in maturation or decay pathways. Both RNA helicases and ribonucleases are present in all kingdoms of life where they play key physiological roles and often represent valuable pharmacological targets. In this book, experts of the field share their most recent protocols and bench tricks to dissect the mechanisms of these important factors in vitro and in vivo. Following a general trend of the field, the book also contains protocols to better identify the physiological targets, cofactors, and biological functions of RNA remodeling proteins and describes current attempts to discover drugs able to control the functions of RNA helicases in various pathological contexts. This book is the result of a collective effort and I would like to thank all authors for their enthusiastic contributions and suggestions. I also would like to thank John Walker—the Series editor—for his guidance throughout the process of assembling and editing the book manuscript. Orleans Cedex 2, France

Marc Boudvillain

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

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1 Key Points to Consider When Studying RNA Remodeling by Proteins . . . . . . . . W. Luke Ward and Rick Russell 2 Happy Birthday: 30 Years of RNA Helicases. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Martina Valentini and Patrick Linder 3 Known Inhibitors of RNA Helicases and Their Therapeutic Potential . . . . . . . . . Yosser Zina Abdelkrim, Josette Banroques, and N. Kyle Tanner 4 Identifying RNA Helicase Inhibitors Using Duplex Unwinding Assays . . . . . . . . John C. Marecki, Alicia K. Byrd, and Kevin D. Raney 5 Thermal Shift Assay for Characterizing the Stability of RNA Helicases and Their Interaction with Ligands . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Emmanuel Saridakis and Franck Coste 6 SRCD and FTIR Spectroscopies to Monitor Protein-Induced Nucleic Acid Remodeling . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Frank Wien, Fre´de´ric Geinguenaud, Wilfried Grange, and Ve´ronique Arluison 7 High-Throughput Protein–Nucleic Acid Interaction Assay Based on Protein-Induced Fluorescence Enhancement . . . . . . . . . . . . . . . . . . . . . . . . . . . . ˇ ı´ha Jaroslav Fulnecˇek and Karel R 8 Probing RNA Helicase Conformational Changes by Single-Molecule FRET Microscopy . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Linda Krause and Dagmar Klostermeier 9 A Fluorescent Assay to Monitor Ligand-Dependent Closure of the Hexameric Rho Helicase Ring . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Michael R. Lawson and James M. Berger 10 A Simple Fluorescence Microplate Assay to Monitor RNA-DNA Hybrid Unwinding by the Bacterial Transcription Termination Factor Rho . . . . . . . . . . . Isabelle Simon and Marc Boudvillain 11 Monitoring Enzymatic RNA G-Quadruplex Unwinding Activities by Nuclease Sensitivity and Reverse Transcription Stop Assays . . . . . . . . . . . . . . . . Ewan K. S. McRae, Steven J. Dupas, Negar Atefi, and Sean A. McKenna 12 Using Magnetic Tweezers to Unravel the Mechanism of the G-quadruplex Binding and Unwinding Activities of DHX36 Helicase . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Huijuan You, Yu Zhou, and Jie Yan

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Contents

Characterization of the Brr2 RNA Helicase and Its Regulation by Other Spliceosomal Proteins Using Gel-Based U4/U6 Di-snRNA Binding and Unwinding Assays . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Eva Absmeier and Markus C. Wahl Monitoring Acetylation of the RNA Helicase DDX3X, a Protein Critical for Formation of Stress Granules. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Makoto Saito, Vytautas Iestamantavicius, Daniel Hess, and Patrick Matthias In Vivo Cross-Linking Followed by polyA Enrichment to Identify Yeast mRNA Binding Proteins. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Sarah F. Mitchell Deciphering the Dynamic Landscape of Transcription-Associated mRNP Quality Control Components Over the Whole Yeast Genome . . . . . . . . . . . . . . . . Ke´vin Moreau, Aure´lia Le Dantec, and A. Rachid Rahmouni Probing the Conformational State of mRNPs Using smFISH and SIM . . . . . . . . Srivathsan Adivarahan and Daniel Zenklusen Probing Transcriptome-Wide RNA Structural Changes Dependent on the DEAD-box Helicase Dbp2 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Yu-Hsuan Lai and Elizabeth J. Tran In Situ Hybridization-Proximity Ligation Assay (ISH-PLA) to Study the Interaction of HIV-1 RNA and Remodeling Proteins. . . . . . . . . . . . Daniela Toro-Ascuy, Aracelly Gaete-Argel, Victoria Rojas-Celis, and Fernando Valiente-Echeverria RNAi Screening to Identify Factors That Control Circular RNA Localization. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Deirdre C. Tatomer, Dongming Liang, and Jeremy E. Wilusz A Cell-Free System for Investigating Human MARF1 Endonuclease Activity . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Hana Fakim and Marc R. Fabian Monitoring eIF4E-Dependent Nuclear 30 End mRNA Cleavage . . . . . . . . . . . . . . Mildred Delaleau An In Vitro Approach To Study RNase III Activities of Plant RTL Proteins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Cyril Charbonnel, Anne de Bures, and Julio Sa´ez-Va´squez Analysis of Bacillus subtilis Ribonuclease Activity In Vivo . . . . . . . . . . . . . . . . . . . . Laetitia Gilet, Olivier Pellegrini, Aude Trinquier, Anastasia Tolcan, Delphine Allouche, Fre´de´rique Braun, Sylvain Durand, and Ciara´n Condon

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Assay of Bacillus subtilis Ribonuclease Activity In Vitro . . . . . . . . . . . . . . . . . . . . . . 403 Olivier Pellegrini, Laetitia Gilet, Aude Trinquier, Anastasia Tolcan, Delphine Allouche, Sylvain Durand, Fre´de´rique Braun, and Ciara´n Condon Protein Pulldown Assays to Monitor the Composition of the Bacterial RNA Degradosome . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 425 Steven W. Hardwick, Ben F. Luisi, and Marilis V. Marques

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

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Contributors YOSSER ZINA ABDELKRIM • Expression Ge´ne´tique Microbienne, UMR8261 CNRS, Institut de Biologie Physico-Chimique, Universite´ de Paris, Paris, France; Molecular Epidemiology and Experimental Pathology (LR16IPT04), Institut Pasteur de Tunis/Universite´ de Tunis el Manar, Tunis-Belve´de`re, Tunisia EVA ABSMEIER • Laboratory of Structural Biochemistry, Institute of Chemistry and Biochemistry, Freie Universit€ at Berlin, Berlin, Germany; MRC Laboratory of Molecular Biology, Structural Studies Division, Cambridge, UK SRIVATHSAN ADIVARAHAN • De´partement de biochimie et me´decine mole´culaire, Universite´ de Montre´al, Montre´al, QC, Canada DELPHINE ALLOUCHE • Expression Ge´ne´tique Microbienne, UMR8261, CNRS, Institut de Biologie Physico-Chimique, Universite´ de Paris, Paris, France VE´RONIQUE ARLUISON • Universite´ de Paris, Paris, France; Laboratoire Le´on Brillouin LLB, CEA, CNRS UMR 12, Universite´ Paris Saclay, CEA Saclay, Gif-sur-Yvette, France NEGAR ATEFI • Department of Chemistry, University of Manitoba, Winnipeg, MB, Canada JOSETTE BANROQUES • Expression Ge´ne´tique Microbienne, UMR8261 CNRS, Institut de Biologie Physico-Chimique, Universite´ de Paris, Paris, France; PSL Research University, Paris, France JAMES M. BERGER • Department of Biophysics and Biophysical Chemistry, Johns Hopkins University School of Medicine, Baltimore, MD, USA MARC BOUDVILLAIN • Centre de Biophysique Mole´culaire (UPR 4301), CNRS, Orle´ans, France FRE´DE´RIQUE BRAUN • Expression Ge´ne´tique Microbienne, UMR8261, CNRS, Institut de Biologie Physico-Chimique, Universite´ de Paris, Paris, France ALICIA K. BYRD • Department of Biochemistry and Molecular Biology, University of Arkansas for Medical Sciences, Little Rock, AR, USA CYRIL CHARBONNEL • Laboratoire Ge´nome et De´veloppement des Plantes, UMR 5096, CNRS, Universite´ Perpignan Via Domitia, Perpignan, France CIARA´N CONDON • Expression Ge´ne´tique Microbienne, UMR8261, CNRS, Institut de Biologie Physico-Chimique, Universite´ de Paris, Paris, France FRANCK COSTE • Centre de Biophysique Mole´culaire, CNRS UPR4301, Orle´ans Cedex 2, France ANNE DE BURES • Laboratoire Ge´nome et De´veloppement des Plantes, UMR 5096, CNRS, Universite´ Perpignan Via Domitia, Perpignan, France MILDRED DELALEAU • Centre de Biophysique Mole´culaire (UPR4301), CNRS, Orle´ans, France STEVEN J. DUPAS • Department of Chemistry, University of Manitoba, Winnipeg, MB, Canada SYLVAIN DURAND • Expression Ge´ne´tique Microbienne, UMR8261, CNRS, Institut de Biologie Physico-Chimique, Universite´ de Paris, Paris, France MARC R. FABIAN • Lady Davis Institute for Medical Research, Jewish General Hospital, Montreal, QC, Canada HANA FAKIM • Lady Davis Institute for Medical Research, Jewish General Hospital, Montreal, QC, Canada

xi

xii

Contributors

JAROSLAV FULNECˇEK • Central European Institute of Technology (CEITEC), Masaryk University, Brno, Czech Republic ARACELLY GAETE-ARGEL • Laboratory of Molecular and Cellular Virology, Virology Program, Faculty of Medicine, Institute of Biomedical Sciences, Universidad de Chile, Santiago, Chile; HIV/AIDS Workgroup, Faculty of Medicine, Universidad de Chile, Santiago, Chile FRE´DE´RIC GEINGUENAUD • Plateforme CNanoMat, UFR SMBH, Universite´ Paris 13, Bobigny, France; Inserm, U1148, Laboratory for Vascular Translational Science, UFR SMBH, Universite´ Paris 13, Bobigny, France LAETITIA GILET • Expression Ge´ne´tique Microbienne, UMR8261, CNRS, Institut de Biologie Physico-Chimique, Universite´ de Paris, Paris, France WILFRIED GRANGE • Institut de Physique et Chimie des Mate´riaux de Strasbourg, CNRS UMR 7504,, Strasbourg, France; Universite´ de Paris, Paris, France STEVEN W. HARDWICK • Department of Biochemistry, University of Cambridge, Cambridge, UK DANIEL HESS • Friedrich Miescher Institute for Biomedical Research, Basel, Switzerland VYTAUTAS IESTAMANTAVICIUS • Friedrich Miescher Institute for Biomedical Research, Basel, Switzerland DAGMAR KLOSTERMEIER • Institute for Physical Chemistry, University of Muenster, Muenster, Germany LINDA KRAUSE • Institute for Physical Chemistry, University of Muenster, Muenster, Germany N. KYLE TANNER • Expression Ge´ne´tique Microbienne, UMR8261 CNRS, Institut de Biologie Physico-Chimique, Universite´ de Paris, Paris, France YU-HSUAN LAI • Department of Biochemistry, Purdue University, West Lafayette, IN, USA; Department of Developmental Neurobiology, St. Jude Children’s Research Hospital, Memphis, TN, USA MICHAEL R. LAWSON • Department of Structural Biology, Stanford University School of Medicine, Stanford, CA, USA AURE´LIA LE DANTEC • Centre de Biophysique Mole´culaire, UPR 4301 du CNRS, Orle´ans, France DONGMING LIANG • Department of Biochemistry & Biophysics, University of Pennsylvania Perelman School of Medicine, Philadelphia, PA, USA PATRICK LINDER • Faculty of Medicine, Department of Microbiology and Molecular Medicine, University of Geneva, Gene`ve, Switzerland BEN F. LUISI • Department of Biochemistry, University of Cambridge, Cambridge, UK W. LUKE WARD • Department of Molecular Biosciences, Institute for Cellular and Molecular Biology, University of Texas at Austin, Austin, TX, USA JOHN C. MARECKI • Department of Biochemistry and Molecular Biology, University of Arkansas for Medical Sciences, Little Rock, AR, USA MARILIS V. MARQUES • Departamento de Microbiologia, Instituto de Cieˆncias Biome´dicas, Universidade de Sa˜o Paulo, Sa˜o Paulo, SP, Brazil PATRICK MATTHIAS • Friedrich Miescher Institute for Biomedical Research, Basel, Switzerland SEAN A. MCKENNA • Department of Chemistry, University of Manitoba, Winnipeg, MB, Canada; Department of Biochemistry and Medical Genetics, University of Manitoba, Winnipeg, MB, Canada; Manitoba Institute for Materials, University of Manitoba, Winnipeg, MB, Canada EWAN K. S. MCRAE • Department of Chemistry, University of Manitoba, Winnipeg, MB, Canada

Contributors

xiii

SARAH F. MITCHELL • Department of Chemistry and Biochemistry, Loyola Marymount University, Los Angeles, CA, USA KE´VIN MOREAU • Centre de Biophysique Mole´culaire, UPR 4301 du CNRS, Orle´ans, France OLIVIER PELLEGRINI • Expression Ge´ne´tique Microbienne, UMR8261, CNRS, Institut de Biologie Physico-Chimique, Universite´ de Paris, Paris, France A. RACHID RAHMOUNI • Centre de Biophysique Mole´culaire, UPR 4301 du CNRS, Orle´ans, France KEVIN D. RANEY • Department of Biochemistry and Molecular Biology, University of Arkansas for Medical Sciences, Little Rock, AR, USA ˇ I´HA • Central European Institute of Technology (CEITEC), Masaryk University, KAREL R Brno, Czech Republic VICTORIA ROJAS-CELIS • Instituto de Ciencias Biomedicas, Facultad de Ciencias de la Salud, Universidad Autonoma de Chile, Santiago, Chile RICK RUSSELL • Department of Molecular Biosciences, Institute for Cellular and Molecular Biology, University of Texas at Austin, Austin, TX, USA JULIO SA´EZ-VA´SQUEZ • Laboratoire Ge´nome et De´veloppement des Plantes, UMR 5096, CNRS, Universite´ Perpignan Via Domitia, Perpignan, France MAKOTO SAITO • Friedrich Miescher Institute for Biomedical Research, Basel, Switzerland; Faculty of Sciences, University of Basel, Basel, Switzerland; Broad Institute of MIT and Harvard, Cambridge, MA, USA EMMANUEL SARIDAKIS • Laboratory of Structural and Supramolecular Chemistry, Institute of Nanoscience and Nanotechnology, National Centre for Scientific Research “DEMOKRITOS”, Athens, Greece ISABELLE SIMON • Mole´culaire (UPR 4301), CNRS, Orle´ans, France; Ecole doctorale Sante´, Sciences Biologiques et Chimie du Vivant (ED 549), Universite´ d’Orle´ans, Orle´ans Cedex 2, France DEIRDRE C. TATOMER • Department of Biochemistry & Biophysics, University of Pennsylvania Perelman School of Medicine, Philadelphia, PA, USA ANASTASIA TOLCAN • Expression Ge´ne´tique Microbienne, UMR8261, CNRS, Institut de Biologie Physico-Chimique, Universite´ de Paris, Paris, France DANIELA TORO-ASCUY • Instituto de Ciencias Biomedicas, Facultad de Ciencias de la Salud, Universidad Autonoma de Chile, Santiago, Chile ELIZABETH J. TRAN • Department of Biochemistry, Purdue University, West Lafayette, IN, USA; Purdue University Center for Cancer Research, Purdue University, Hansen Life Sciences Research Building, West Lafayette, IN, USA AUDE TRINQUIER • Expression Ge´ne´tique Microbienne, UMR8261, CNRS, Institut de Biologie Physico-Chimique, Universite´ de Paris, Paris, France MARTINA VALENTINI • Faculty of Medicine, Department of Microbiology and Molecular Medicine, University of Geneva, Gene`ve, Switzerland FERNANDO VALIENTE-ECHEVERRIA • Laboratory of Molecular and Cellular Virology, Virology Program, Faculty of Medicine, Institute of Biomedical Sciences, Universidad de Chile, Santiago, Chile; HIV/AIDS Workgroup, Faculty of Medicine, Universidad de Chile, Santiago, Chile MARKUS C. WAHL • Freie Universit€ at Berlin, Institute of Chemistry and Biochemistry, Laboratory of Structural Biochemistry, Berlin, Germany FRANK WIEN • Synchrotron SOLEIL, L’Orme des Merisiers Saint Aubin, Gif-sur-Yvette, France

xiv

Contributors

JEREMY E. WILUSZ • Department of Biochemistry & Biophysics, University of Pennsylvania Perelman School of Medicine, Philadelphia, PA, USA JIE YAN • Mechanobiology Institute, National University of Singapore, Singapore, Singapore HUIJUAN YOU • School of Pharmacy, Tongji Medical College, Huazhong University of Science and Technology, Wuhan, China DANIEL ZENKLUSEN • De´partement de biochimie et me´decine mole´culaire, Universite´ de Montre´al, Montre´al, QC, Canada YU ZHOU • Mechanobiology Institute, National University of Singapore, Singapore, Singapore

Chapter 1 Key Points to Consider When Studying RNA Remodeling by Proteins W. Luke Ward and Rick Russell Abstract Cellular RNAs depend on proteins for efficient folding to specific functional structures and for transitions between functional structures. This dependence arises from intrinsic properties of RNA structure. Specifically, RNAs possess stable local structure, largely in the form of helices, and there are abundant opportunities for RNAs to form alternative helices and tertiary contacts and therefore to populate alternative structures. Proteins with RNA chaperone activity, either ATP-dependent or ATP-independent, can promote structural transitions by interacting with single-stranded RNA (ssRNA) to compete away partner interactions and then release ssRNA so that it can form new interactions. In this chapter we review the basic properties of RNA and the proteins that function as chaperones and remodelers. We then use these properties as a foundation to explore key points for the design and interpretation of experiments that probe RNA rearrangements and their acceleration by proteins. Key words RNA remodeling, Helicases, RNA folding, ATP analogs, AMP-PNP, ADP-BeF3

1

Introduction Cellular RNAs interact with proteins for most or all of their functional lifetimes. From the time they are produced by transcription to their destruction by ribonucleases, proteins play critical roles at essentially all stages. Some proteins function as stable partners with RNAs, generating RNA-protein complexes (RNPs), while others interact transiently with RNAs and play such roles as directing the RNA to a subcellular location, packaging or protecting the RNA, or marking the RNA for processes like translation or decay [1]. In addition to functional partners, RNAs interact with broad groups of proteins that accelerate structural transitions of RNA, promoting folding or conformational rearrangements or remodeling RNA-protein complexes by removing protein components. These proteins fall into two groups, as defined functionally: those that use energy from nucleoside triphosphates, typically ATP, and those that do not use an energy source [2–4]. The ATP-dependent

Marc Boudvillain (ed.), RNA Remodeling Proteins: Methods and Protocols, Methods in Molecular Biology, vol. 2209, https://doi.org/10.1007/978-1-0716-0935-4_1, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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proteins are classified as helicase proteins because they are related in sequence and structure to proteins that unwind DNA helices with high processivity in processes like DNA replication [5]. Most of these RNA-interacting proteins are classified within the helicase superfamily 2 (SF2), and indeed many display at least limited helicase activity when measured in vitro [4, 6] (see Notes 1 and 2). The energy-independent proteins represent many broad groups and have no phylogenetic linkage [2, 7], with their most general feature being that they are positively charged, engendering strong interactions with RNA. In the sections below, we summarize the properties of RNA that result in a general requirement for proteins to promote structural transitions, and in this summary we also outline briefly the mechanistic differences between the broad groups of proteins. To the fullest extent possible, we keep the discussion at a very general level with respect to the proteins, without focusing on specific proteins or families. For more detailed descriptions of the specific protein families and their mechanisms, we refer interested readers to recent reviews on RNA helicases [4, 8, 9] and ATP-independent RNA chaperones [10, 11]. We then turn to the design of experiments that probe these protein-mediated RNA structural transitions, highlighting key issues that arise regardless of the particular protein or RNA being studied or the specific experimental method used to probe the process.

2 RNA Structural Features Lead to Its Dependence on Proteins for Structural Rearrangements From both physical and conceptual perspectives, RNA structure begins locally. The most basic unit of RNA structure is short, double-stranded helices, which are defined as RNA secondary structure. Most of these helices are formed by sequences that are local in primary structure and are separated by a few nucleotides that form a loop to close the helix. These local helices can form fast during folding, and they are locally stable even in isolation [12, 13]. Thus, an RNA helix of just a few base pairs can form in an energetically favorable manner in the absence of any other stabilizing structure, and RNA helices of as few as five or six base pairs can persist on timescales of minutes, making structural transitions that require disruption of the helices incompatible with biological timescales [14]. The same basic features also apply to RNA tertiary structures, which are typically base-pairing interactions of various types and can also be stable in isolation. For simplicity, we focus below on RNA secondary structures and direct interested readers to reviews for discussions of protein disruption of RNA tertiary structure [3, 15].

Key Points for RNA Remodeling by Proteins

3

As a consequence of the great stability of local helical structure, RNA folding transitions and conformational changes that involve significant changes in base pairing typically require assistance from proteins (see Note 3). By interacting strongly with single-stranded RNA, either in an ATP-dependent or an ATP-independent manner, a protein can interrupt the interactions between the two strands of a helix, separating the strands and replacing RNA–RNA contacts with RNA–protein contacts (Fig. 1a). For short helices of the lengths that are common in functional RNAs ( kcincorrect in Fig. 2b).

4

General Experimental Considerations for Studying RNA Remodeling by Proteins In the sections below, we describe some key points that arise when monitoring RNA remodeling by proteins and the interactions of these proteins with RNA. Some of these points are very general and indeed apply to measurements of any kinetic or equilibrium process, whereas others are derived from the properties of RNA and remodeling proteins described above. For a simple illustration, we use the example of monitoring unwinding of a helix throughout the discussion below, and we include other types of remodeling reactions as appropriate to illustrate the key points.

Key Points for RNA Remodeling by Proteins

4.1 Conferring Net Directionality to the Remodeling Process

7

Most experimental approaches are only able to monitor a reaction when there is a net directionality to the process. That is, one species (or an ensemble of related species) must be increasing in relative population, while another is decreasing. An exception to this point is in experiments using single-molecule approaches, not covered here, in which a reaction can be monitored as it occurs reversibly at equilibrium [38]. Thus, for ensemble approaches it is necessary to set up the experiment in such a way that one species is initially dominant, and then the activity of the protein accelerates a transition to another species or ensemble that gives a distinct experimental signal. As shown in Fig. 3a, this transition may be unwinding of a helix to give two separate products, or it may be an intramolecular transition that results in a change in structure within RNA. It is important to remember that the reverse reaction can also occur and may be accelerated by the protein. A key first step in design of the experiments is to develop a working knowledge of the system sufficient to identify conditions under which there is a net transition from A to B. It is important to note that A does not have to transition completely to B. It is only necessary for the transition to proceed far enough to generate a reliable experimental signal (see Note 4). With these conditions established, the next general challenge is to design an experimental strategy in which species A will be populated at the start of the experiment, either because it is stable at equilibrium or because it is kinetically trapped. Then the protein will be added or activated, resulting in or accelerating a net transition to species B. The key point is that the reaction can be set up in two stages. The purpose of the first stage is to generate as much species A as possible, and the purpose of the second stage is to monitor the activity of the protein as it promotes the net conversion of species A to species B. Figure 3b shows a simple experimental design for monitoring helix unwinding by a helicase protein using dilution. In this experiment, a small RNA helix is formed in stage 1 and then unwound in stage 2. The key difference in conditions between the stages is the concentration of the RNA oligonucleotides that form the helix. In the first stage, they are present at high concentrations, well above the equilibrium dissociation constant for helix formation, so that nearly all of the RNA will be present as a helix. If one of the strands is more convenient to monitor, that strand should be present at a lower concentration than the complementary strand to maximize its incorporation into the helix. Note that the ionic strength can also be increased in this stage, if desired, to further stabilize the helix [39–41]. Empirically, a monovalent salt concentration of 0.5–1 M is strongly stabilizing for helix formation (Na+ or K+, typically as salts with Cl
, acetate, or glutamate anion). In the second stage, the concentration of the helix is reduced dramatically by diluting it with reaction buffer. If the ionic strength was high in the first stage, it may also be reduced to achieve the

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W. Luke Ward and Rick Russell

A Species B

Species A

B

+

C

Stage 1 Form Species A

Protein + Dilution +/- ATP

Mg2+

Stage 1 Form Species A

Protein + Dilution +/- ATP

Stage 2 Transition to Species B

Stage 2 Transition to Species B Measure fraction of labeled strand in helix

+

Measure fraction of labeled strand in helix 1

1

Fraction 0.8 of 0.6 species 0.4 A

Low [protein]

Fraction 0.8 of 0.6 species 0.4 A

0.2

High [protein]

0.2

No protein

No protein Low [protein] High [protein]

0

0

Time in Stage 2

Time in Stage 2

Fig. 3 General experimental design to monitor an RNA unwinding or remodeling process. (a) General scheme in which there is a net directionality, with state A transitioning to an equilibrium distribution that favors state B (left panel). Middle panel, An RNA helix dissociates into its two strands spontaneously. Right panel, a helical RNA transitions from a kinetically trapped intermediate to a more stable form. (b) Experimental protocol for monitoring RNA unwinding by dilution. In the first stage, the helix is formed at high concentration, with the labeled strand shown in black, and then in the second stage, it is diluted to a much lower concentration in the presence of the protein. The plot shows hypothetical progress curves for the helix in the absence of protein (black, which reaches an endpoint of 0.5 to indicate that the concentration of the unlabeled strand after dilution is in the range of the equilibrium constant for dissociation) and in the presence of increasing concentrations of protein (red and cyan curves). The protein may decrease the endpoint, as shown, if a significant fraction of the ssRNA products remain bound to the protein, as shown schematically above. (c) Experimental protocol for monitoring an RNA folding transition. In stage 1, the RNA is transferred from a denatured state to one that supports folding. It rapidly forms species A even though species B is more stable, hypothetically because the smaller loop of species A allows its formation to be preferred kinetically. The protein is then added in the second stage, and the transition to a population of predominantly species B is monitored. The plot shows hypothetical progress curves in the absence of protein (black) and in the presence of increasing concentrations of protein (red and cyan)

Key Points for RNA Remodeling by Proteins

9

desired conditions for monitoring protein-mediated helix unwinding. At this time, the protein is also added, along with ATP if desired. In practice, achieving a helix concentration that is low enough to observe unwinding will typically require using excess protein over RNA helix and monitoring a single turnover reaction. The protein concentration should be varied, while maintained in excess of the helix, to identify the concentration range over which there is a dependence. It may be convenient to make the transition from stage 1 to stage 2 with two additions, diluting the helix in the first addition and subsequently adding the protein, which is appropriate provided that the time between the two additions is short enough that the helix does not dissociate significantly in the intervening time. The fraction of the oligonucleotide(s) that remains present as a helix is then monitored as a function of time after the transition to stage 2, as shown in the plot in Fig. 3b, to determine an observed rate constant for the transition. It is a good idea to monitor the reaction in the absence of the protein to have a baseline for interpreting the effects of the protein. Depending on the stability of the helix (i.e., the Kd value for helix formation) and its concentration in stage 2, the helix may or may not dissociate fully in the absence of the protein. The protein will be expected to increase the rate constant, and it may also decrease the endpoint (although it is important to remember that proteins can accelerate helix formation as well as dissociation). The reaction should be monitored until it reaches an endpoint, if possible, because the observed rate constant will be equal to the sum of the rate constants for unwinding and formation of the helix. Thus, the observed rate constant is equal to the unwinding rate constant only under conditions that give complete dissociation of the helix to a population only of ssRNAs (see Notes 5 and 6). The same general strategy can be used to follow an intramolecular RNA rearrangement, with a protein promoting interconversion between two alternative secondary structures within an RNA (Fig. 3c). Species A is populated in stage 1 by adding heatdenatured RNA into buffer containing appropriate concentrations of Mg2+ and/or monovalent ion to allow structure formation. Species A accumulates because it is preferred kinetically even though it is less stable than species B, in this case hypothetically because the smaller loop of species A forms more rapidly. By allowing species A to first accumulate in stage 1, the experiment monitors only the effect of the protein on the interconversion of species A and B, and it is not sensitive to any effect the protein may have on the formation of A vs. B from the ensemble of unfolded conformations. In stage 2, the two species can interconvert, and in the absence of the protein, this interconversion favors accumulation of the more stable species B (Fig. 3c). The presence of the protein accelerates this process. For simplicity here, the endpoint is shown

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as being the same in the presence and absence of the protein. Thus, the protein accelerates both the forward and reverse reactions by the same factor, leaving the distribution unchanged at equilibrium. For an ATP-independent protein, this condition must hold, as it is not possible for an enzyme to change the equilibrium distribution of the reactants and products of any reaction. However, for an ATP-dependent protein, the energy of ATP binding and hydrolysis may be used to generate a steady-state distribution of species that is different from the equilibrium distribution (see above) [24, 42]. 4.2 Using a Chase to Selectively Observe the Remodeling Process in One Direction

For some reactions, it is not practical to achieve conditions such that the net reaction proceeds in the desired direction. A good example of this is unwinding of a longer RNA helix, which typically has a Kd value that is sufficiently low that it is not possible to work with concentrations below the Kd value while still having a robust experimental signal (i.e., 20%), the method can also be optimized for 384-well format, although PIFE variability among replicas may be higher. 3. We tested the Em580-10 and Em570-10 filters from BMG Labtech for Cy3 measurement. Although the Em570-10 filter gives higher signal because bandpass window is closer to the Cy3 emission maximum, it is also closer to the excitation window, and we observed higher signal variability caused by light reflections at the edge of well bottom. Therefore, we prefer using the EM580-10 filter. 4. mwPIFE can also be performed without immobilization of oligonucleotide probes. However, we empirically established that in solution measurements give significantly higher PIFE variation between technical replicas than measurements on immobilized probes, which reduces sensitivity of the assay. PIFE as low as 5% above background can be reproducibly detected using immobilized probes. 5. Wash only desired number of wells necessary for the experiment. For example, calculation of a single protein-nucleic acid binding curve with nine different concentrations of a protein (including no protein) in triplicates requires 27 wells. The plate containing unused wells can be stored sealed in the original bag with desiccant, and the remaining wells can be used in a subsequent experiment. 6. Because NeutrAvidin in wells has limited capacity to bind biotin, it is recommended to perform a binding-saturation experiment with each new oligonucleotide probe. We empirically established that immobilization of the oligonucleotide probe in amount that gives approximately threefold increase in fluorescence signal (usually in the range of 0.5–2.5 pmol in 50 μL) above the background is ideal for mwPIFE. More densely immobilized probes may decrease PIFE, likely because probe crowding may limit efficient protein binding. Furthermore, the lower amount of oligonucleotide probe decreases the amount of the protein needed for PIFE analysis. 7. The time needed to measure entire 96-well plate is approximately 27 min. 8. You may first optimize the protein/oligonucleotide probe concentration ratio by measuring protein concentration range. It is also possible to completely replace the buffer with protein solution in situations where higher protein concentration is desired.

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Acknowledgments Our work is supported by the Ministry of Education, Youth and Sports of the Czech Republic from the European Regional Development Fund—Project “REMAP” (No. CZ.02.1.01/0.0/0.0/ 15_003/0000479) and by the Czech Science Foundation (19-21961S). References 1. Webster MW, Stowell JA, Passmore LA (2019) RNA-binding proteins distinguish between similar sequence motifs to promote targeted deadenylation by Ccr4-Not. Elife 8:e40670 2. Jolma A, Yan J, Whitington T et al (2013) DNA-binding specificities of human transcription factors. Cell 152:327–339 3. Re A, Joshi T, Kulberkyte E et al (2014) RNA-protein interactions: an overview. Methods Mol Biol 1097:491–521 4. Dey B, Thukral S, Krishnan S et al (2012) DNA-protein interactions: methods for detection and analysis. Mol Cell Biochem 365:279–299 5. Aramendia PF, Negri RM, Sanroman E (1994) Temperature-dependence of fluorescence and photoisomerization in symmetrical carbocyanines—influence of medium viscosity and molecular-structure. J Phys Chem 98:3165–3173 6. Sorokina M, Koh HR, Patel SS et al (2009) Fluorescent lifetime trajectories of a single fluorophore reveal reaction intermediates during transcription initiation. J Am Chem Soc 131:9630–9631

7. Hwang H, Kim H, Myong S (2011) Protein induced fluorescence enhancement as a single molecule assay with short distance sensitivity. Proc Natl Acad Sci U S A 108:7414–7418 8. Hwang H, Myong S (2014) Protein induced fluorescence enhancement (PIFE) for probing protein-nucleic acid interactions. Chem Soc Rev 43:1221–1229 9. Nadiras C, Delaleau M, Schwartz A et al (2019) A fluorogenic assay to monitor rho-dependent termination of transcription. Biochemistry 58:865–874 10. Rashid F, Raducanu VS, Zaher MS et al (2019) Initial state of DNA-dye complex sets the stage for protein induced fluorescence modulation. Nat Commun 10(1):2104 11. Valuchova S, Fulnecek J, Petrov AP et al (2016) A rapid method for detecting proteinnucleic acid interactions by protein induced fluorescence enhancement. Sci Rep 6:39653 12. Valuchova S, Fulnecek J, Prokop Z et al (2017) Protection of Arabidopsis blunt-ended telomeres is mediated by a physical association with the Ku heterodimer. Plant Cell 29:1533–1545

Chapter 8 Probing RNA Helicase Conformational Changes by Single-Molecule FRET Microscopy Linda Krause and Dagmar Klostermeier Abstract Fo¨rster resonance energy transfer (FRET) is a versatile tool to study the conformational dynamics of proteins. Here, we describe the use of confocal and total internal reflection fluorescence (TIRF) microscopy to follow the conformational cycling of DEAD-box helicases on the single molecule level, using the eukaryotic translation initiation factor eIF4A as an illustrative example. Confocal microscopy enables the study of donor-acceptor-labeled molecules in solution, revealing the population of different conformational states present. With TIRF microscopy, surface-immobilized molecules can be imaged as a function of time, revealing sequences of conformational states and the kinetics of conformational changes. Key word Conformational dynamics, Single-molecule FRET, TIRF, DEAD-box helicase, eIF4A

1

Introduction RNA unwinding by DEAD-box RNA helicases is mediated by a helicase core, formed by two flexibly linked domains [1]. During ATP-dependent RNA unwinding, these helicases undergo a nucleotide-dependent conformational cycle. Closing of the helicase core in response to binding of double-stranded RNA substrates and ATP is the first step in duplex destabilization and accelerates dissociation of one strand of the RNA duplex [2]. The release of phosphate after ATP has been hydrolyzed is coupled to reopening of the helicase core and to release of the second strand of the RNA duplex [3]. Flanking domains and interaction partners regulate helicase activity by modulating the conformational cycle [4, 5]. Conformational changes are thus directly linked to the nucleotide cycle, the helicase activity, and to the regulation of unwinding. Fo¨rster (or fluorescence) resonance energy transfer (FRET) is based on a distance-dependent interaction between two dyes, termed donor and acceptor [6]. When donor and acceptor fluorophores are introduced at suitable positions, conformational changes can be followed by changes in the efficiency of FRET. FRET experiments

Marc Boudvillain (ed.), RNA Remodeling Proteins: Methods and Protocols, Methods in Molecular Biology, vol. 2209, https://doi.org/10.1007/978-1-0716-0935-4_8, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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on the single-molecule level (smFRET) avoid ensemble averaging and enable the determination of the distributions of conformers and their interconversion. FRET can also be used to determine intra- or intermolecular distances, which then serve as constraints to model global conformations of multi-domain proteins or multisubunit complexes. As an example, the RNA-binding domain of YxiN has been positioned relative to the helicase core to generate a structural model of the full-length protein in solution [7, 8]. smFRET has also been applied to study RNA binding, translocation, and unwinding by helicases [9]. Similar approaches to the ones we describe here for RNA-binding proteins have been used extensively to investigate protein–DNA interactions, e.g., in the transcription initiation complex [10]. FRET occurs via the through-space interaction of the transition dipoles of donor and acceptor [6]. The rate constant kt of the energy transfer depends on the distance of the fluorophores rDA:  6 R0 1 kt ¼ ð1Þ r DA τD R0 is the Fo¨rster distance, the donor-acceptor distance where the FRET efficiency is 50 %, and τD is the fluorescence lifetime of the donor fluorophore. Energy transfer to the acceptor competes with the depopulation of the excited state of the donor via radiative and non-radiative processes. The transfer efficiency E thus depends on the ratio of the rate constant for energy transfer kt and the sum of the rate constants of energy transfer and of radiative and non-radiative processes, kr and knr: E¼

kt kt þ kr þ knr

ð2Þ

By combining Eqs. (1) and (2) and substituting τ1 D for knr + kr, the distance dependence of the FRET efficiency is obtained as E¼

R60 R60 þ r 6DA

ð3Þ

Experimentally, the FRET efficiency is determined from the measured donor intensity ID and acceptor intensity IA, corrected for background fluorescence, according to Eq. (4): E¼

IA IA þ ID

ð4Þ

To convert FRET efficiencies into inter-dye distances, the Fo¨rster distance and parameters correcting for instrument non-ideality are required. By converting FRET efficiencies into distances for each individual molecule, distance histograms can then be calculated. For detailed information on the determination of Fo¨rster distances and correction parameters, see [11].

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Conformational changes of single molecules are studied in solution by using a confocal microscope with laser excitation (Fig. 1f–h). The small size of the confocal volume, in combination with concentrations of the donor-acceptor-labeled protein in the picomolar range, ensures that not more than one molecule resides in the confocal volume at any given time. Each single molecule is detected as a burst of fluorescence intensity. The events from several thousand single molecules are then compiled into a FRET efficiency histogram. The observation time is limited by the diffusion time of the molecule through the confocal volume, which is in the range of a few milliseconds. TIRF microscopy enables monitoring of surface-immobilized single molecules on a timescale from seconds to minutes (Fig. 1a–e). In addition to the distribution of conformers, this method also provides information on their interconversion and reveals the sequence in which they occur as well as rate constants for these conformational changes (see [4, 12] for kinetic studies on the conformational cycle of eIF4A). Here, we describe how to analyze the conformation and conformational changes of helicases, using our work on the DEADbox RNA helicase eIF4A as an illustrative example [4, 12]. eIF4A is a minimal DEAD-box helicase consisting of two RecA-like domains [13, 14], whereas the helicase core is flanked by N- and C-terminal domains in many other representatives of the DEAD-box family [14]. eIF4A associates with the cap-binding protein eIF4E and the scaffold protein eIF4G into the eIF4F complex, which is a central player in translation initiation. eIF4G binds to both RecA-like domains of eIF4A and stabilizes a half-open conformation [15, 16].

2

Materials All solutions are prepared with ultrapure water and are sterilefiltered through a 0.2 μm membrane prior to use. Solutions for smFRET measurements are treated with active charcoal overnight to remove fluorescent impurities. The charcoal is removed by filtration through a 0.45 μm membrane, and the solutions are degassed. Buffers are prepared from stock solutions by dilution prior to the measurement. For fluorescent labeling, two cysteine residues within a suitable distance for FRET are required in the protein of interest; these need to be introduced by site-directed mutagenesis in surface-exposed positions outside conserved motifs. Intrinsic solvent-accessible cysteine residues need to be replaced by other amino acids, typically serine, alanine, leucine, isoleucine, or valine, by site-directed mutagenesis (see [11] for the general work flow). Immobilization of the protein of interest on streptavidin-functionalized surfaces requires biotinylation, but alternative immobilization schemes have been used [17]. It is crucial to confirm that the donor-acceptor-labeled protein variant retains wild-type-like enzymatic activity.

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Fig. 1 Principle of confocal and TIRF smFRET measurements. (a) Assembly of TIRF slides; sample channels are delimited by adhesive tape between the PEGylated cover slip (top) and microscope slide (bottom). (b) Fluorescently labeled molecules are immobilized on the surface via a biotin-streptavidin interaction. Several

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2.1 Material Needed for All Procedures

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1. 5 activity buffer: 250 mM Tris/HCl pH 7.5, 400 mM KCl, 2.5 mM MgCl2, 5 mM DTT, 5 % (v/v) glycerol. 2. 1.5 mL protein low-binding reaction microtubes. 3. Ultrapure water.

2.2 Donor-Acceptor Labeling

1. Double-cysteine variants of eIF4A [16, 18] or of your own protein of interest. 2. MicroSpin30 columns (BioRad; one per labeling reaction). 3. Microcentrifuge. 4. Alexa555- and Alexa647-maleimide. 5. Buffer A: 50 mM Tris/HCl pH 7.5, 200 mM NaCl, 1 mM tris (2-carboxyethyl)phosphine (TCEP). 6. Buffer B: 50 mM Tris/HCl pH 7.5, 200 mM NaCl, 2 mM β-mercaptoethanol. 7. Spectrophotometer.

2.3 Determination of Correction Parameters

1. Single-cysteine variants of eIF4A [16, 18] or of your own protein of interest.

2.4 Confocal Microscopy

1. Cuvettes: Lab-Tek® Chambered #1.0 Borosilicate Coverglass System with six chambers (200 μL chambers) or Hamamatsu Microwell Slide (20 μL). 2. Microscope: homebuilt confocal microscope based on an inverted Olympus IX70 microscope with laser excitation (frequency-doubled and pulse-picked output of a pulsed titanium-sapphire laser, 475 nm) and avalanche photodiode detectors, Becker&Hickl SPC600 counting card; alternative commercially available microscope: PicoQuant Microtime200, with LDH-DC 485, LDH-DC 530, and LDH-DC 640 laser

ä Fig. 1 (continued) measurements on the same sample are performed by moving the focus along the sample channel. (c) The fluorescence intensity of labeled proteins within the field of view is imaged as a function of time (top). Images from the donor and acceptor channel are aligned to assign donor and acceptor signals from the same molecule (bottom). (d) From fluorescence intensity time traces for each single molecule (top), FRET time traces (bottom) are calculated. (e) FRET efficiencies from all molecules are compiled into histograms, which reveal the different conformations populated (top). Rate constants for conformational changes are derived from the observed dwell times in the different FRET states (bottom). (f) smFRET with confocal microscopy uses cuvettes (top) to study molecules in solution. Bursts of fluorescence intensity are detected when a labeled molecule diffuses through the confocal volume (bottom). (g) The photons emitted by donor (green) and acceptor (red) are detected as a function of time. Note that the photon counts from the acceptor are displayed on an inverted scale for clarity. (h) The FRET histogram is compiled from FRET values calculated from donor and acceptor intensities for each molecule detected one after the other

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diodes for excitation; and single-photon avalanche diodes, Picoquant TimeHarp 260 counting card (Picoquant, Berlin [18, 19]). 3. 10 mg/mL bovine serum albumin (BSA). 2.5 Preparation of Slides for TIRF Microscopy

1. Microscope slides (Thermo Scientific, 76  26 mm). 2. Microscope cover glasses (Marienfeld, 24  40 mm). 3. Cover slip racks and washing container. 4. Shandon™ Vertical Staining Jars for 16 slides (Thermo Scientific). 5. Grace Bio-Labs SecureSeal™ Adhesive Sheet 0.12 mm (SigmaAldrich). 6. Plasma cleaner (oxygen or argon). 7. Sonicator. 8. Tweezers. 9. Empty pipette tip boxes with insert. 10. 3 mL glass syringe with canule. 11. 10 % Alconox in ultrapure water. 12. 1 M KOH. 13. Methanol. 14. N-[3-(Trimethoxysilyl)propyl]ethylenediamine (aminosilane). 15. Glacial acetic acid. 16. 50 mM MOPS pH 7.5. 17. mPEG succinimidyl valerate (MW 5000, Laysan Bio) and biotin PEG succinimidyl valerate (MW 5000, Laysan Bio). 18. 50 mL Falcon tubes. 19. Nitrogen flow. 20. 500 mL glass measuring cylinder. 21. 500 mL beaker. 22. Crystallizing dish.

2.6

TIRF Microscopy

1. Slides: homebuilt slides with sample channels and PEGylated glass surfaces. 2. Microscope: inverted Olympus IX81 TIRF microscope (Olympus, Mu¨nster, Germany) with a 532 nm laser diode for excitation, a high numerical aperture temperature-controlled objective (UAPON 100, oil immersion; NA ¼ 1.49), a dual-view beam splitter, and an ANDOR iXon3 EM-CCD camera for detection [4].

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3. 3 imaging buffer: 50 nM protocatechuate 3,4-dioxygenase (PCD), 2.5 mM protocatechuic acid (PCA), 1 mM Trolox, 0.1 mg/mL BSA in 1 activity buffer. 4. 1 mg/mL streptavidin in 10 mM Tris/HCl pH 7.5. 5. Empty pipette tip container. 6. Whatman™ gel-blotting-paper GB005. 7. Biotinylated protein of interest.

3

Methods

3.1 Protein Labeling with Donor and Acceptor Fluorophores

1. The storage buffer of the spin column is removed, and the column is washed three times with 500 μL buffer A using centrifugations at 1000 g for 1 min at room temperature (RT) (see Note 1). 2. Dilute the protein (e.g., eIF4A) to 70 μM in 70 μL volume in buffer A. 3. Exchange the buffer to buffer A using the spin column (1000 g, 4 min, RT). 4. Determine the concentration of the protein photometrically. 5. Dissolve the Alexa555- and Alexa647-maleimide dyes in DMSO to a concentration of 20 μg/mL (see Note 2). 6. Use 60 μL of the protein in buffer A (from step 3) for labeling. Perform the labeling reaction in a low-binding microtube. Use a twofold molar excess of Alexa555 maleimide (donor) and a fivefold molar excess of Alexa647 maleimide (acceptor). Mix the dyes in the lid of the tube and then spin down to mix rapidly with the protein. Carry out the labeling reaction in the dark for 1 h at RT (see Note 3). 7. During incubation of the protein with the dyes, wash the spin column from step 1 three times with 500 μL buffer B (1000 g, 1 min, RT). 8. Stop the labeling reaction and remove free dye by buffer exchange to buffer B using the spin column (1000 g, 4 min, RT). Collect the labeled protein in a low-binding microtube. 9. Wash the spin column with buffer B as described in step 8 and apply the protein solution to the column a second time. Collect the protein in a low-binding tube (see Note 4). 10. To determine the labeling efficiency, record an absorption spectrum of the labeled protein from 230 to 720 nm. This typically requires a five- to sixfold dilution with buffer B (see Note 5).

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11. Determine the labeling efficiency by calculating the concentration of the acceptor dye from the absorption at the maximum absorption wavelength of the acceptor (650 nm for Alexa647). The concentration of the donor dye can be calculated from the total absorption at the absorption wavelength of the donor (554 nm for Alexa555) after subtracting the absorption caused by the acceptor dye. The protein concentration is determined from the absorption at 280 nm, corrected for contributions of the donor and acceptor to absorption at this wavelength (see Note 6). 3.2 Determination of Correction Parameters

1. Label the two single-cysteine variants separately with the donor dye or with the acceptor dye (four labeling reactions, following the procedure described in Subheading 3.1 (see Note 7). 2. Incubate a cuvette filled with 0.75 mg/mL BSA in 1 activity buffer for at least 30 min prior to the measurement. 3. Use one of the donor-labeled protein solutions (from step 1) to adjust the laser intensity. Dilute the protein solution to 100 nM dye concentration in 1 activity buffer and adjust the laser intensity so that you obtain a fluorescence intensity of 10.000–30.000 cps. Use this laser intensity for all measurements. 4. Measure the fluorescence (in cps) of the donor-labeled protein for 60 s and the fluorescence of the acceptor-labeled protein for 300 s. Also, measure the fluorescence background of 1 activity buffer for 600 s (see Note 8). 5. If you measure smFRET with your protein in the presence of ligands, you have to repeat the measurements described in the presence and absence of each ligand (see Note 9).

3.3 Confocal Microscopy

1. To prevent unspecific protein binding to the cuvette, incubate the cuvette (Fig. 1f) with 0.75 mg/mL BSA in 1 activity buffer for at least 30 min prior to the measurement. Use 20 μL or 200 μL total volume, depending on the cuvette used (see Notes 10 and 11). 2. Dilute the donor-acceptor-labeled protein to a donor concentration of approximately 1–2 nM in 1 activity buffer. The final donor concentration in the cuvette is 150 pM (see Note 12). 3. Remove the BSA from the cuvette and wash three times with 20 μL (200 μL) 1 activity buffer. Remove any residual buffer (see Note 13). 4. Start by pipetting 5 buffer and the required volume of ultrapure water into the cuvette and then add the labeled protein and cofactors or nucleotide (see Note 14). 5. Measure the fluorescence from the confocal volume for 20–30 min (see Note 15).

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1. Wash tweezers, cover slip racks, washing containers, and microscope slide jars with 10 % Alconox solution (see Notes 16 and 17). 2. Discard the Alconox solution and rinse with ultrapure water several times. 3. Clean 8 microscope slides and 12 cover slips thoroughly with 10 % Alconox (see Note 18). 4. Rinse three times with ultrapure water (see Note 19). 5. Fill the containers with 1 M KOH and sonicate microscope slides and cover slips in KOH for 20 min. 6. Rinse slides and cover slips once with 1 M KOH and three times with ultrapure water. 7. Dry the microscope slides and cover slips under nitrogen flow (see Notes 20 and 21). 8. Clean the microscope slides and cover slips in the plasma cleaner for 10 min. Keep track which side is the upward-facing side of the microscope slides (see Note 22). 9. Wash the microscope slides and cover slips with water several times and then once with methanol (work with methanol under the hood). 10. Fill the containers with fresh methanol and sonicate the microscope slides and cover slips for 20 min. 11. Wash the microscope slides and cover slips once with methanol. 12. Incubate microscope slides and cover slips in a solution containing 1 % (v/v) aminosilane and 5 % (v/v) glacial acetic acid in methanol twice for 10 min in the dark and sonicate for 1 min in between (see Note 23). 13. Wash the microscope slides and cover glasses once with methanol, then several times with water, and dry in nitrogen flow. 14. Prepare a solution of 22 % mPEG and 2 % biotin PEG in 800 μL 50 mM MOPS/NaOH pH 7.5. 15. Fill the empty pipette tip boxes with ultrapure water (2 cm filling height). Place the microscope slides on the insert with the plasma-cleaned site facing upwards (see Note 24). 16. Pipette 60 μL of the mPEG solution from step 14 in the middle of the microscope slide surface and place a cover slip on top (see Note 25). 17. Incubate the slides overnight in the dark (see Note 24). 18. Separate cover slips and microscope slide carefully (see Note 26). 19. Rinse microscope slides and cover slips extensively with water and dry in nitrogen flow (see Note 27).

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20. Cut the adhesive tape into stripes of approx. 5 cm  0.3 cm. 21. Place five strips on the PEG-coated side of the microscope slide; these are the boundaries of the sample channels (Fig. 1a; see Note 28). 22. Place a cover slip on the microscope slide, with the PEG-coated surface facing the channels (Fig. 1a,b; see Note 29). 23. Store two slides back to back in one 50 mL Falcon tube and fill with nitrogen. Place in a vacuum-sealed bag and store at 20  C (see Note 30). 3.5

TIRF Microscopy

1. Wash the channel to be used with 200 μL 1 activity buffer (see Notes 31 and 32). 2. Dilute the streptavidin stock solution fivefold to 0.2 mg/mL in 1 activity buffer and incubate channel for 10 min with 50 μL of this streptavidin dilution in the dark (see Note 33). 3. Dilute the donor-acceptor-labeled protein to a final concentration of 30 pM in 50 μL 1 imaging buffer. Perform a serial dilution in four steps: one ten-fold, two 100-fold, and a final step to 30 pM. Use low-binding microtubes for all dilution steps (see Note 34). 4. Wash the channel with 100 μL 1 activity buffer. 5. Incubate the channel with the donor-acceptor-labeled protein in the dark for 10 min (see Note 33). 6. Wash the channel with 100 μL 1 imaging buffer. 7. Put a drop of mineral oil on the objective of the microscope and place the slide upside down onto the microscope stage. Fluorescence from the donor-acceptor-labeled protein on the cover slip is then recorded. Check if the density of immobilized protein on the channel surface is sufficiently high (see Note 35). Fluorescence is typically recorded from ten different sections of the channel (see Note 36). 8. Add cofactors and/or ligands to the immobilized protein in 1 imaging buffer and repeat imaging. 9. The general procedure of data analysis is described in [4].

4

Notes 1. Labeling of one protein requires 2 mL of buffer A and 5 mL of buffer B; adjust volumes when performing more than one labeling reaction. 2. To prepare dye stocks, dissolve the maleimides in DMSO to a concentration of 10 mM. Split into 5 μL aliquots (50 nmol per aliquot). Remove the DMSO in a speed-vac apparatus and store the dried maleimides at 20  C until needed. Dissolved dyes can be stored at 20  C for approximately 3 months.

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3. The molar excess of dyes and the incubation time have to be optimized for each protein of interest. As a starting point, a two-fold molar excess of Alexa555 and a five-fold molar excess of Alexa647 or a 2.5-fold molar excess of Alexa488 and a 3.5fold molar excess of Alexa546 can be used. Labeling times may vary between 5 min and 1 h, depending on the accessibility of the cysteine residues. If labeling efficiencies are low after 1 h, the solvent accessibility is most likely not sufficient. 4. Solutions of labeled protein can be stored overnight in the fridge on ice. For longer periods, the protein solution may be shock-frozen in liquid nitrogen and stored at 80  C. This depends on protein stability and needs to be tested for the protein of interest. 5. The diluted protein solution used to record the spectrum can be used for smFRET measurements on the same day. 6. The sum of donor and acceptor labeling efficiencies must be equal to or less than 200% per protein with two cysteine residues. If a higher labeling efficiency is determined, this may be due to free dye, which can be removed in an additional washing step using the spin column, or due to unspecific binding. For more detailed information about the labeling process, see [11]. 7. If you label a monomeric protein with two cysteines, you obtain a mixture of six different species of labeled protein in the end (donor/donor, donor/acceptor, acceptor/donor, acceptor/acceptor, plus the two singly labeled species). 8. Since the laser is emitting at the maximum absorption wavelength of the donor, the signal of the acceptor fluorophore is low. To increase the signal-to-noise ratio, the measuring times of acceptor-labeled protein are higher than for donor-labeled proteins. 9. For a detailed description of the analysis, see [11]. 10. For smFRET measurements at room temperature, the small cuvettes can be used. At higher temperatures, the larger cuvettes should be used to minimize effects due to evaporation. In both cases, the cuvettes should be covered during the measurement, either by a lid or by a microscope cover glass to minimize evaporation. 11. In general, 50 mL of 5 activity buffer are prepared and stored at 4  C. Buffers can be used for several weeks when stored at 4  C. It may be useful to prepare buffer in 2 mL aliquots for daily use. 1 buffer is also prepared in a volume of 2 mL for measurements on the same day. 12. Typically, the protein solution is diluted in two to three steps. Dilutions of labeled protein are prepared in low-binding tubes and stored on ice protected from light. Dilute solutions of labeled protein should be prepared freshly after 2 h.

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13. When filling the chamber, make sure you touch the bottom of the slide with the pipette tip only on the side of the chamber to prevent scratching the surface. 14. You can perform sequential smFRET measurements by adding more cofactors or nucleotides to the same sample. After three consecutive measurements, a new sample with fresh labeled protein should be used. 15. It is reasonable to record fluorescence from the protein in buffer first to test for sufficient signal-to-noise ratio. 16. Prepare 500 mL Alconox solution in a beaker. To clean the containers, fill some of the solution inside and rub with your fingers (wear gloves!). Smaller parts such as racks can be incubated in the beaker and cleaned by shaking or rubbing. 17. It is useful to have two staining jars and several coverslip racks. For the plasma cleaner, you need coverslip racks made of plastic, for other purposes metal racks may also be used. 18. Cover slips break easily, so prepare more than needed as backup. 19. Place slides and cover slips into the respective container and racks. To wash slides and slips, fill the containers with water. 20. When handling microscope slides and cover slips, you can hold them in the corner either with tweezers or with your fingers. Touch only the sides with your fingers, not the surfaces. If you have a flexible tube attached to the nitrogen flow, place a 1 mL Eppendorf pipette tip in the front to direct the flow onto slides and cover slips. Place dried microscope slides and cover slips into dry staining jar and plastic racks. 21. When drying the microscope slides or cover slips, visually inspect them and pay attention to any possible residue on the glass surfaces. If the surface is not clean, use a wash bottle filled with ultrapure water to remove any residue. 22. Microscope slides can be placed into the plasma cleaner without a rack; cover slips are placed in the plastic rack. Leave one spot free between cover slips in the rack. It is important to remember which side of the microscope slides was exposed to the plasma. Mark the staining jar on one side and place the microscope slides with the activated side facing the mark. Cover slips standing in the rack are clean from both sides. 23. Prepare the aminosilane solution in a glass cylinder. Remove aminosilane from the storage bottle with the glass syringe and a metal canule. 24. To prevent drying of the slides, this reaction needs to be carried out in a humid environment. Close the lid of the box when incubating overnight.

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25. Avoid air bubbles between the microscope slide and the cover slip. Remove bubbles by gently pressing onto the cover slip with a pipette tip. 26. Move the cover slip over the edge of the microscope slide until you can hold a corner with the tweezers. Move carefully upwards without breaking off corners. 27. Rinsing works best with a washing bottle over a crystallizing dish. Make sure there is no residue from the PEG solution left. Keep track of the orientation of the microscope slides and cover slips. 28. For removing the foil from the adhesive paper, tweezers with sharp ends are helpful. 29. Once you placed the cover slip onto the slide, press along the stripes with a pipette tip to seal and to avoid leakage between the channels during measurements. 30. Slides can be stored at 20  C for approximately 2–4 months. 31. To wash or fill the channel on the slide with a pipette, hold a piece of Whatman™ paper to the opening on the opposite side of the channel. 32. Channels are filled most easily with medium size pipette tips of 100–200 μL. If the solution forms a drop instead of entering the channel, try to hold the pipette more upright. If a bubble has formed, fill the channel from the other side. Extra-fine pipette tips may also help. 33. To incubate the TIRF slide in the dark, an empty pipette tip container can be used as cover. 34. 500 μL of 3 imaging buffer are prepared freshly every day; the dilution into 1 imaging buffer for smFRET measurements is performed freshly every 2–3 h. Store on ice. 35. If the immobilization reaction was successful, labeled protein is detected with uniform density throughout the field of view. Spots from individual proteins need to be clearly distinguishable. If spots are not separated, reduce the concentration of the labeled protein during immobilization. 36. To obtain well-defined FRET histograms, time traces of 80–100 molecules and a total number of detected events of 8000 are required. Rate constants can be derived from these traces if a majority shows changes in FRET efficiency during the observation time.

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References 1. Sengoku T, Nureki O, Nakamura A et al (2006) Structural basis for RNA unwinding by the DEAD-box protein Drosophila vasa. Cell 125:287–300. https://doi.org/10. 1016/j.cell.2006.01.054 2. Theissen B, Karow AR, Ko¨hler J et al (2008) Cooperative binding of ATP and RNA induces a closed conformation in a DEAD box RNA helicase. PNAS 105:548–553 3. Aregger R, Klostermeier D (2009) The DEAD box helicase YxiN maintains a closed conformation during ATP hydrolysis. Biochemistry 48:10679–10681. https://doi.org/10.1021/ bi901278p 4. Harms U, Andreou AZ, Gubaev A, Klostermeier D (2014) eIF4B, eIF4G and RNA regulate eIF4A activity in translation initiation by modulating the eIF4A conformational cycle. Nucleic Acids Res 42:7911–7922. https:// doi.org/10.1093/nar/gku440 5. Andreou AZ, Klostermeier D (2013) The DEAD-box helicase eIF4A. RNA Biol 10:19–32. https://doi.org/10.4161/rna. 21966 6. Fo¨rster T (1946) Energiewanderung und fluoreszenz. Naturwissenschaften 33:166–175 7. Samatanga B, Andreou AZ, Klostermeier D (2017) Allosteric regulation of helicase core activities of the DEAD-box helicase YxiN by RNA binding to its RNA recognition motif. Nucleic Acids Res 45:1994–2006. https:// doi.org/10.1093/nar/gkx014 8. Karow AR, Klostermeier D (2009) A conformational change in the helicase core is necessary but not sufficient for RNA unwinding by the DEAD box helicase YxiN. Nucleic Acids Res 37:4464–4471. https://doi.org/10. 1093/nar/gkp397 9. Rothenberg E, Ha T (2005) Single-molecule FRET analysis of helicase functions. In: Abdelhaleem MM (ed) Helicases methods and protocols. Humana Press, pp 29–43 10. Heilemann M, Hwang LC, Lymperopoulos K, Kapanidis AN (2009) Single-molecule FRET analysis of protein-DNA complexes. In: Moss T, Leblanc B (eds) DNA-protein

interactions principles and protocols, 3rd edn, pp 503–521 11. Andreou AZ, Klostermeier D (2012) Conformational changes of DEAD-box helicases monitored by single molecule fluorescence resonance energy transfer. In: Methods in enzymology, 511th edn, pp 75–109 12. Andreou AZ, Harms U, Klostermeier D (2019) Single-stranded regions modulate conformational dynamics and ATPase activity of eIF4A to optimize 50 -UTR unwinding. Nucleic Acids Res 47:5260–5275. https:// doi.org/10.1093/nar/gkz254 13. Linder P, Lasko PF, Ashburner M et al (1989) Birth of the D-E-A-D box. Nature 337:121–122 14. Schmid SR, Linder P (1992) D-E-A-D protein family of putative RNA helicases. Mol Microbiol 6:283–291 15. Schu¨tz P, Bumann M, Oberholzer AE et al (2008) Crystal structure of the yeast eIF4AeIF4G complex: an RNA-helicase controlled by protein-protein interactions. Proc Natl Acad Sci U S A 105:9564–9569. https://doi. org/10.1073/pnas.0800418105 16. Hilbert M, Kebbel F, Gubaev A, Klostermeier D (2011) eIF4G stimulates the activity of the DEAD box protein eIF4A by a conformational guidance mechanism. Nucleic Acids Res 39:2260–2270. https://doi.org/10.1093/ nar/gkq1127 17. Roy R, Hohng S, Ha T (2008) A practical guide to single-molecule FRET. Nat Methods 5:507–516. https://doi.org/10.1038/nmeth. 1208 18. Andreou AZ, Klostermeier D (2014) eIF4B and eIF4G jointly stimulate eIF4A ATPase and unwinding activities by modulation of the eIF4A conformational cycle. J Mol Biol 426:51–61. https://doi.org/10.1016/j.jmb. 2013.09.027 19. PicoQuant Microtime200. https://www. picoquant.com/products/category/fluores cence-microscopes/microtime-200-timeresolved-confocal-fluorescence-microscopewith-unique-single-molecule-sensitivity. Accessed 28 Oct 2019

Chapter 9 A Fluorescent Assay to Monitor Ligand-Dependent Closure of the Hexameric Rho Helicase Ring Michael R. Lawson and James M. Berger Abstract The bacterial Rho protein is an exemplar RecA-family hexameric helicase that assists with the termination of RNA polymerase activity on a variety of transcripts. During its catalytic cycle, Rho both loads onto and translocates along RNA through a series of tightly regulated, ligand-dependent conformational changes. Here we describe an assay to track Rho as it switches from an open-ring (RNA-loading) to a closed-ring (RNA-translocation) configuration by monitoring the association of a fluorescein-labeled RNA to Rho’s central pore as a change in fluorescence anisotropy. The assay, which is in principle adaptable to the study of ligand-dependent isomerization events in other ring-shaped translocases, is readily amenable to 384-well format plates and small-molecule screening efforts. Key words Rho, RNA, ATPase, Transcription termination, Hexameric helicase, High-throughput screening, Fluorescence anisotropy, Ring translocase

1

Introduction Processive, ring-shaped translocases harness the energy provided by ATP hydrolysis to move either protein or nucleic acid strands through a central pore in the assembly. The bacterial transcription termination factor Rho, a RecA-family hexameric RNA helicase, uses its translocation activity to halt the synthesis of cytosine-rich mRNAs and anti-sense transcripts [1]. Prior to RNA binding, Rho initially adopts an open-ring configuration competent for loading onto client nucleic acid segments [2]. Following RNA binding, and in the presence of an ATP cofactor, Rho transitions into a closedring and translocation-competent state that encircles the RNA strand [3, 4]. Given the widespread distribution of ring translocases throughout nature, there is substantial interest in developing methods to monitor their action on substrates. Although not all ring translocases transition between fully open and closed states as part of their catalytic cycle, many will bind tightly to substrate polymers only

Marc Boudvillain (ed.), RNA Remodeling Proteins: Methods and Protocols, Methods in Molecular Biology, vol. 2209, https://doi.org/10.1007/978-1-0716-0935-4_9, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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Fig. 1 Overview of the assay. On their own, short RNA oligos tumble quickly and display low fluorescence anisotropy. In the presence of nucleotide, Rho will bind to the RNA, slowing its tumbling rate and increasing the fluorescence anisotropy signal

when nucleotide is present [4–6]. This switch-like behavior affords a straightforward approach to distinguish between key functional intermediates using fluorescence-based methods predicated on substrate binding. For Rho, we devised an assay involving a dye-labeled RNA and a non-hydrolyzable ATP analog (ADPlBeF3) to track this rearrangement using fluorescence anisotropy (FA) (Fig. 1). Using this approach, we discovered that numerous known agonists and antagonists of Rho activity (including the antibiotic bicyclomycin, cytosine-rich nucleic acids, and the transcription factor NusG) are potent modulators of Rho ring closure [7, 8]. The simplicity and scalability of this assay render it useful not only for looking at how natural ligands and Rho-binding partners modulate ring status [7, 8] but also for conducting high-throughput screens aimed at identifying new small-molecule agonists and antagonists of Rho function.

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Materials 1. Milli-Q purified water. 2. Rho protein from Escherichia coli, purified as described previously (see ref. 3), stored at a concentration of
30 mg/mL in size exclusion buffer (500 mM KCl, 50 mM Tris, 10% (v:v) glycerol, 1 mM dithiothreitol, pH 7.5) in 100 μL aliquots at 80  C. 3. UV-vis spectrophotometer with microvolume measurement capacity (e.g., NanoDrop from Thermo Fisher Scientific). 4. 25 mM ADPlBeF3 (see Notes 1–3).

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5. Fluorescence microplate reader. The protocol below is for a Synergy Neo2 plate reader (BioTek) but should be easily adapted to any commercial reader. 6. Tabletop centrifuge equipped with rotor and adapters for microplates. 7. 10 mg/mL BSA in Milli-Q water. Store as 500 μL aliquots at 80  C. 8. 100 μM 50 6-fluorescein-labeled RNA oligonucleotide in MilliQ water. Store as 10 μL aliquots at 80  C. 9. 1 M dithiothreitol stock solution in Milli-Q water. Store as 1 mL aliquots at 80  C. 10. Multichannel pipette with 2–20 μL range capability (e.g., 12-channel Rainin Pipet-Lite Multi Pipette L12-20XLS+). 11. 1 assay buffer: 150 mM KCl, 5% (v:v) glycerol, 5 mM MgCl2, 0.5 mM (tris(2-carboxyethyl)phosphine) [TCEP], 20 mM HEPES, pH 7.5. Prepare 2 L and store at 4  C. 12. 2 assay buffer: 300 mM KCl, 10% (v:v) glycerol, 10 mM MgCl2, 1 mM TCEP, 40 mM HEPES, pH 7.5. Prepare 100 mL and store as 1 mL aliquots at 20  C. 13. Black, 384-well low-volume microplates treated for low non-specific binding (e.g., Corning NBS plates #3544). 14. 96-well round bottom plate (e.g., Greiner #650101). 15. Stir plate, small stir bar, and 1 L glass beaker. 16. 0.1 mL disposable dialysis floating chambers with 10 kDa molecular weight cutoff (e.g., 10 k MWCO Slide-A-Lyzer MINI dialysis devices from Thermo Fisher Scientific). 17. Benchtop microcentrifuge. 18. FAconvert.py” script from https://github.com/jaglawson/ platereader_code/ 19. Prism (GraphPad) or equivalent data analysis software.

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3.1 Buffer Exchange of Rho into Assay Buffer

1. Thaw Rho aliquots on ice. Two Rho aliquots (100 μL each) are generally sufficient for a full 384-well plate. 2. Pool and transfer the Rho solution into a 10 k MWCO Slide-ALyzer MINI dialysis float. Multiple floats should be used if more than 200 μL of Rho is required. 3. Place the dialysis float into a beaker containing a stir bar and 1 L of 1 assay buffer, and allow to dialyze overnight on a stir plate at 4  C (low speed).

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4. Empty buffer from beaker, and replace it with 1 L of fresh 1 assay buffer. Dialyze for at least two additional hours at 4  C. 5. Save the buffer from beaker (hereafter referred to as “dialysis buffer”), and transfer the Rho sample from the dialysis float to a microtube. Store sample on ice. 3.2 Measuring RNA Binding to Rho by Fluorescence Anisotropy

1. Quantify Rho concentration in dialyzed sample based on A280 reading (ε ¼ 95,580 M1 cm1 for E. coli Rho hexamer). 2. Thaw frozen aliquots of 10 mg/mL BSA, 1 M dithiothreitol, 2 assay buffer, and 100 μM fluorescein-labeled RNA on ice. 3. Spin down fluorescein-labeled RNA aliquot at 13,000  g in a benchtop microcentrifuge, and dilute to 1 mL with 990 μL of dialysis buffer (1 μM final concentration). 4. Assemble a “3 RNA master mix” with the following composition per 1 mL of 3 mix required: 60 μL of 100 μM fluorescein-labeled RNA, 15 μL of 1 M dithiothreitol, 150 μL of 10 mg/mL BSA, 165 μL of 2 assay buffer, and 610 μL of dialysis buffer. 5. Dilute the Rho sample from step 1 to threefold above the highest concentration desired for assays with dialysis buffer. Place dilution in first column of a 96-well plate. 6. Perform ten serial twofold dilutions of Rho with dialysis buffer (so columns 1–1 contain the dilution series of Rho), and omit Rho entirely from column 12 as a no-protein control (Fig. 2; see Notes 4–5). 7. Add 16 μL of the 3 RNA master mix to the requisite number of wells in a 96-well plate (Fig. 2). 8. Transfer 16 μL of 3 Rho stocks from dilution series to 3 RNA samples with a multichannel pipette. Mix thoroughly and set a timer for 30 min (Fig. 2; see Note 6). 9. Dilute 25 mM ADPlBeF3 with an equal volume of 2 assay buffer, and then dilute further to 3 the final desired concentration of nucleotide with dialysis buffer (herein described as 3 nucleotide). Transfer 16 μL of 3 nucleotide to wells 1–12 of a 96-well plate (Fig. 2). 10. Once the time from step 8 has elapsed, add 16 μL of 3 nucleotide to the pre-mixed 3 Rho/3 RNA with a multichannel pipette, and mix thoroughly. Set multichannel pipette to 20 μL, mix thoroughly again, and set another timer for 30 min (Fig. 2; see Note 6). 11. Once the time from step 10 has elapsed, use the multichannel pipette to transfer two 20 μL replicates of every Rho/RNA/ nucleotide mix to a 384-well plate (“assay plate”), with replicates in adjacent wells such that each 12-well row from the

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Fig. 2 Schematic of sample preparation

96-well plate fills a 24-well row on the assay plate (Fig. 2; see Note 7). Be sure to only depress the pipette down to the first stop to avoid introducing bubbles at this stage. Set a timer for 10 min (see Note 6). 12. Spin the assay plate down in a tabletop centrifuge at 4000  g for 2 min—this step is necessary to remove any bubbles introduced in pipetting and to flatten out the meniscuses of every well. 13. Pre-warm the BioTek Neo2 plate reader to 30  C, and follow on-screen prompts to switch in the proper filters for fluorescein FA reading after loading the appropriate program. Select the appropriate well range and set the PMTs to 65/61. Set to read once per minute (or minimal interval allowed by instrument, generally ~1:15 if reading a full plate) for 30 min (see Note 8). 14. Press start run, define file name, and place plate on carriage. Press OK when timer from step 11 has elapsed, and allow reading for the full 30 min.

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3.3 Data Analysis (See Note 9)

1. Inspect plots of individual parallel and perpendicular values with respect to time. These values generally display a period of rapid change (likely indicative of temperature equilibration) that runs the first 5–10 min and then a period of stability that is maintained through the end of the run. Any of the sets of readings in the stable period are suitable for further analysis; we generally use the reading from 10 to 15 min. 2. Click on Data Reduction and toggle display of FA values in mA. Set time to the set of readings that correspond to a stable period of parallel and perpendicular fluorescence intensity values, and click the excel button. Save the spreadsheet that pops up with an appropriate file name. 3. Open the Excel spreadsheet with FA values, and copy/paste all the data (with row name, but without column names) into a plain text file using a text editor program such as Komodo Edit. For example, if readings had been taken from rows A and B, the file would appear as:

A 171 174 164 165 138 131 93 89 68 65 57 59 52 54 52 53 51 52 53 51 52 48 48 47 B 167 172 167 166 135 133 89 88 66 66 56 57 53 54 52 51 50 51 52 51 52 49 49 45

4. Save this data as a plain.txt file (e.g., assay_01.txt), and move it to the same directory as the “FAconvert.py” script (available at https://github.com/jaglawson/platereader_code/). 5. Replace the text defined as “inputfilename” with the name of your .txt file. 6. Open a terminal window, and navigate to the directory with the data and the FAconvert.py script. 7. Type the following command: “python FAconvert.py.” 8. Open the output files (defined by the variables “outputfilename” and “outputfilename2”; default is to append “averaged_subtracted_” and “averaged_subtracted_tabbed_” to the input file name) generated by the script. Each pair of replicates (e.g., wells 1 and 2) has now been averaged, and the baseline (defined as the average of 23 and 24) has now been subtracted from every one of these pairs of replicates (see Note 10). Thus, in the above example, the “averaged_subtracted_” output file from the above example would now look like this: A

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The “averaged_subtracted_tabbed_” output file would appear as: A

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9. Open Prism, and click “enter __ replicate values side-by-side subcolumns,” and set __ to read the number of replicates you expect to have for this particular experiment (note each pair of duplicates from a particular master mix above will only count as one replicate and has already been averaged at this point). 10. Copy/paste the data into Prism as Y values, replace the former row titles (e.g., “A” or “B”) with a meaningful descriptor, and input the final concentrations of Rho hexamer in the dilution series as X values. 11. Press the “Analyze” button, select nonlinear curve fit, and select all data series you want to fit. Select “One site—specific binding with Hill Slope.” 12. Save the output Kd values from the fits (Fig. 3).

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Notes 1. We have found it best to make ADPlBeF3 fresh every time. It is prepared by adding 93.75 μL of 1 M NaF to 18.75 μL of 1 M BeCl2 (Sigma). It is important to pre-pipette the BeCl2 into a tube first, and then add the NaF to this while mixing rapidly with a pipette; adding BeCl2 to pre-measured NaF can cause the solution to turn white and cloudy. We then add 62.5 μL of 100 mM ADP (purchased as a powder, resuspended in Milli-Q water, and pH adjusted to ~7 with NaOH) and 75 μL of MilliQ water, which yields 250 μL of 25 mM ADPlBeF3.

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Fig. 3 Example of processed, fit data from the assay (reproduced with permission from ref. 7)

2. Beryllium is extremely toxic. As such, great care should be taken when handling these solutions, and proper waste streams maintained for all wet and dry waste. Gram quantities of BeCl2 salts release toxic chlorine gas when resuspended in water, so dissolution should only be done in a fume hood with extreme caution. 3. Recently, it has become difficult to procure BeCl2 from commercial sources; BeSO4 is a viable alternative (Irina Artsimovitch, personal communication). 4. If one hopes to determine the impact of a ligand other than RNA or ADPlBeF3 on Rho ring closure (e.g., protein cofactors, primary site nucleic acids, small molecules), the ideal way to do this in our experience is to include these at 3 the desired final concentration in the Rho master mixes. If the ligand is a lyophilized small molecule or RNA, it should be brought into solution using the dialysis buffer; if it is a protein, then it should be co-dialyzed with Rho throughout both dialysis steps (in the latter case, it also must be accounted for in the A280 reading obtained in Subheading 3.2, step 1). A Rho stock concentration higher than 30 mg/mL may be needed at the start to achieve saturation of ring closure at the high protein concentrations. Alternatively, one can assemble samples for FA using four successive, 13 μL pipetting steps (using 4 Rho, RNA, and nucleotide master mixes) rather than three 16 μL steps, but the same rules apply with respect to using the dialysis buffer to resuspend and/or co-dialyze the ligand with Rho. 5. FA readings can be sensitive to mismatches in buffer conditions, especially glycerol. Be mindful of this fact when adapting this protocol to fit your needs. 6. The ring closure assay is effectively an endpoint assay, rather than an equilibrium assay (because ADPlBeF3 is a tight inhibitor). The final Kd,app values from this assay are somewhat

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sensitive to the incubation times from the addition of Rho to RNA until reading and also sensitive to the time from nucleotide addition to the Rho/RNA mixture until starting the read. We find that results are repeatable on a day-to-day basis if incubation times are strictly limited to 30 min of Rho with RNA, 30 min of the Rho/RNA mixture with ADPlBeF3, and then 10 min for transfer and starting the read. 7. The scripts that will be used to process the data generated assume that the Rho-free control samples are in columns 23 and 24. 8. While it is possible to collect data from a full 384-well plate, we have found that collection of data from half of a plate at a time (corresponding to one 96-well plate of master mixes, with each sample measured in duplicate) is a more plausible goal given the timelines required for repeatability described in Note 6. 9. While analysis steps described here are specific to the file formats generated by a BioTek Neo2 plate reader, the same general principles could be applied to analyze data generated by other instruments. 10. The “averaged_subtracted_tabbed_” format is compatible with most data processing programs (e.g., Prism); however the “averaged_subtracted_” file is also generated since it would be more conducive to further scripting of this analysis.

Acknowledgments This work was supported by G. Harold and Leila Y. Mathers Foundation and the National Institute of General Medical Sciences (R37-071747), to J.M.B. M.R.L. gratefully acknowledges support from the A.P. Giannini Foundation. References 1. Mitra P, Ghosh G, Hafeezunnisa M, Sen R (2017) Rho protein: roles and mechanisms. Annu Rev Microbiol 71:687–709. https://doi. org/10.1146/annurev-micro-030117-020432 2. Skordalakes E, Berger JM (2003) Structure of the rho transcription terminator: mechanism of mRNA recognition and helicase loading. Cell 114(1):135–146 3. Thomsen ND, Berger JM (2009) Running in reverse: the structural basis for translocation polarity in hexameric helicases. Cell 139 (3):523–534. https://doi.org/10.1016/j.cell. 2009.08.043 4. Thomsen ND, Lawson MR, Witkowsky LB, Qu S, Berger JM (2016) Molecular mechanisms

of substrate-controlled ring dynamics and substepping in a nucleic acid-dependent hexameric motor. Proc Natl Acad Sci U S A 113(48): E7691–E7700. https://doi.org/10.1073/ pnas.1616745113 5. Bashore C, Dambacher CM, Goodall EA, Matyskiela ME, Lander GC, Martin A (2015) Ubp6 deubiquitinase controls conformational dynamics and substrate degradation of the 26S proteasome. Nat Struct Mol Biol 22 (9):712–719. https://doi.org/10.1038/nsmb. 3075 6. Strycharska MS, Arias-Palomo E, Lyubimov AY, Erzberger JP, O’Shea VL, Bustamante CJ, Berger JM (2013) Nucleotide and partner-protein

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control of bacterial replicative helicase structure and function. Mol Cell 52(6):844–854. https:// doi.org/10.1016/j.molcel.2013.11.016 7. Lawson MR, Dyer K, Berger JM (2016) Ligandinduced and small-molecule control of substrate loading in a hexameric helicase. Proc Natl Acad Sci U S A 113(48):13714–13719. https://doi. org/10.1073/pnas.1616749113

8. Lawson MR, Ma W, Bellecourt MJ, Artsimovitch I, Martin A, Landick R, Schulten K, Berger JM (2018) Mechanism for the regulated control of bacterial transcription termination by a universal adaptor protein. Mol Cell 71(6):911–922. e914. https://doi.org/10. 1016/j.molcel.2018.07.014

Chapter 10 A Simple Fluorescence Microplate Assay to Monitor RNA-DNA Hybrid Unwinding by the Bacterial Transcription Termination Factor Rho Isabelle Simon and Marc Boudvillain Abstract Transcription termination factor Rho contributes to shape the transcriptomes of many bacteria and is essential in a large subset of them. Although the transcription termination function of Rho is not always easy to reconstitute and to study in vitro, assays based on the ATP-dependent RNA-DNA hybrid unwinding activity of the factor can prove useful to dissect Rho mechanisms or to seek new antibiotics targeting Rho. However, current in vitro assays of Rho helicase activity are time-consuming, as they usually require radiolabeling of the hybrid substrates and analysis of reaction products by gel electrophoresis. Here, we describe a fluorescence-based microplate assay that informs on Rho helicase activity in a matter of minutes and allows the multiplexed analysis of conditions required for primary biochemical characterization or for drug screening. Key words Rho, Transcription, Termination, Helicase, RNA, Fluorescence

1

Introduction Transcription termination factor Rho is a homo-hexameric, ringshaped protein factor that has major regulatory functions in bacteria (reviewed in [1–3]). The molecular mechanisms underlying the biological functions of Rho are complex and involve multiple rearrangements of the Rho ring structure and of its network of interactions with the RNA substrate (see Chap. 9 in this volume and [4, 5]), as well as intricate interplays with the transcription and translation machineries and a variety of cofactors [1–3]. Several biochemical assays have been developed over the years to characterize the Rho factor, from basic RNA binding and ATPase assays [6, 7] to more sophisticated transcription termination assays [8– 11]. Monitoring of the ATP-dependent RNA-DNA hybrid unwinding activity of Rho [12] is also a method of choice to characterize the factor in vitro [13, 14]. This helicase activity derives from oligonucleotide “roadblock” displacement upon

Marc Boudvillain (ed.), RNA Remodeling Proteins: Methods and Protocols, Methods in Molecular Biology, vol. 2209, https://doi.org/10.1007/978-1-0716-0935-4_10, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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directional translocation of the Rho factor along single-stranded RNA substrates [15–17]. It thus represents a good compromise to probe the molecular motor features of Rho (as in termination assays) under conditions where the nucleic acid substrate and reaction setup or ingredients can be easily manipulated and varied (as in RNA binding and ATPase assays). For instance, RNA-DNA substrates having complex, multipartite architectures or bearing specific chemical modifications can be used in the helicase assay to probe Rho stepping mechanism [18–20], a task that is not possible with the other biochemical assays. In most Rho helicase experiments, a RNA-DNA hybrid substrate is first modified with a 32P label (most often on the “tracking” RNA strand) and then purified by polyacrylamide gel electrophoresis (PAGE). Then, the substrate is mixed with Rho and ATP in a suitable reaction buffer, and reaction aliquots are withdrawn and quenched at various times. The reaction aliquots are then analyzed by PAGE to reveal the fractions of single-stranded RNA products formed upon Rho-directed hybrid unwinding as a function of time [14]. In a variant of this protocol, the 32P label is replaced by a fluorochrome (usually on the DNA oligonucleotide strand) in order to analyze the PAGE gel with a fluorescence imager [13]. Although the method is very efficient (see Note 1), it is burdensome and time-consuming and does not allow frequent record of the helicase activity over time nor sample multiplexing. To address these limitations, we have developed a new assay of Rho helicase activity that is based on the development of a fluorescence signal upon unwinding of a tripartite RNA-DNA hybrid labeled with a fluorophore-quencher pair (Fig. 1a). The modality of detection and assay conditions were designed for use with a multiplate fluorescence reader, thereby allowing either continuous or endpoint monitoring of Rho helicase activity as well as sample multiplexing suitable for quick biochemical investigations and, potentially, for drug screening (see Note 1).

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Materials Only RNase-free, milliQ-grade deionized water should be used to prepare buffers and to make dilutions. All solutions (and water stocks used for dilutions) should be sterilized by filtration with single-use 0.2 μm pore size filter units. We also recommend using high-quality microtubes, preferably ones intended for low non-specific binding of nucleic acids and proteins.

2.1 Preparation of the RNA Strand by Transcription

1. Benchtop centrifuge and vortex mixer. 2. Dry bath incubator with shaking capability (e.g., Eppendorf ThermoMixer).

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5’- GGACTTCTCCTCTGTCTCCTTCTTCCTTCTCCTCTGTCTCCTTCTTCC-3’

+1 DNA template:

Rut site

T7 prom.

reporter cassette Transcription

RNA transcript:

5’ Hybridization

Tripartite RNA-DNA hybrid:

5’

5’ Alexa488

72 nt

b

c Control

Xylene cyanol

RNA sample

30 bp

- Dabcyl

Dabcyl

27 bp

+ Dabcyl

Correct 3-piece substrate 2-piece hybrid

Pen mark

RNA shadow

Alexa488-labelled Oligo1

Pen mark

Fig. 1 Preparation of the RNA-DNA hybrid substrates for the fluorogenic Rho helicase assay. (a) Schematic of the two-step protocol described in text. (b) Representative image of the UV shadows observed upon UV illumination of the RNA preparative PAGE gel. (c) Representative image obtained after Typhoon FLA9500 fluorescence imaging of a PAGE gel used for RNA-DNA hybrid substrate preparation

3. Saran sheets and blade. 4. 5 mL syringe equipped with a glass wool plug and a 0.2 μm syringe filter. 5. Vertical PAGE system and power supply. 6. N,N,N,N0 -tetramethylethylenediamine (TEMED). 7. 25% (w/v) ammonium persulfate (APS) in water; fresh solution stored at 4  C.

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8. 10 TBE buffer: 0.89 M Tris-base, 0.89 M boric acid, and 20 mM EDTA. 9. 1 TBE buffer obtained by diluting the 10 TBE buffer stock. 10. Phenol:chloroform:isoamyl alcohol (25:24:1) mix, pH 6.7. 11. Diethyl ether. 12. Ethanol absolute (99.8%) stored at 20  C. 13. 0.5 M EDTA solution adjusted to pH 8.0 with NaOH. 14. 1 M10E1 buffer: 10 mM MOPS, 1 mM EDTA, pH 6.0. 15. 3 M sodium acetate solution adjusted to pH 6.5 with acetic acid. 16. Elution buffer: 0.3 M sodium acetate in 1 M10E1 buffer. 17. UV spectrophotometer (preferably one able to measure absorbance from 1 μL samples). 18. Formamide. 19. Control formamide buffer: formamide supplemented with 0.01% (w/v) xylene cyanol and 0.01% (w/v) bromophenol blue. 20. 100 nM DNA template stock in 1 M10E1 buffer (see Note 2). 21. 5 transcription buffer: 400 mM HEPES pH 7.5, 112 mM MgCl2, 100 mM DTT, 5 mM spermidine, 0.05% (v/v) Triton X-100 (see Note 3). 22. 20 μ/μL Superase-IN (Thermo Fisher Scientific). 23. 100 mM rATP, rCTP, rGTP, and rUTP stock solutions. 24. 12.5 μM T7 RNA polymerase (if using a commercial preparation, follow manufacturer’s instructions to adjust enzyme amounts in transcription reactions). 25. Denaturing 6% acrylamide solution. Mix 4.5 mL of a commercial 40% acrylamide:bis-acrylamide [29:1 ratio] solution with 12.6 g of urea and 3 mL 10 TBE, and adjust the volume to ~25 mL with deionized water. Heat in a microwave oven to melt urea (see Note 4). Adjust the volume to 30 mL with deionized water. 26. X-ray intensifying screen or a fluor-coated TLC plate. 27. Handheld 254 nm UV lamp. 2.2 Preparation of the RNA-DNA Hybrid Substrate

1. Items 1–17 from Subheading 2.1. 2. 100 μM stock solutions of oligodesoxyribonucleotides Oligo1 (bearing an Alexa488 fluorochrome at its’ 50 end), Oligo2 (bearing a Dabcyl quencher at its 30 end), and Oligo4 (Table 1) (see Note 5). 3. Purified RNA transcript from Subheading 3.1.

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Table 1 Sequences of the DNA oligonucleotides Name

Sequence

Oligo1

50 Alexa488-CTT-CTC-GAG-GAT-CCA-GAT-CTG-ATA-CCA-TCG-30

Oligo2

50 -CGA-TGA-ATT-CGA-GCT-CGG-TAC-CCG-CAG-Dabcyl-30

Oligo3

50 -CTT-CTC-GAG-GAT-CCA-GAT-CTG-ATA-CCA-TCG-30

Oligo4

50 -CGA-TGA-ATT-CGA-GCT-CGG-TAC-CCG-CAG-30

4. 5 helicase buffer: 750 mM potassium acetate, 100 mM HEPES pH 7.5, 0.5 mM EDTA. 5. 1 helicase buffer obtained by dilution of the 5 buffer stock in sterile deionized water. 6. Native 6% acrylamide solution. Mix 4.5 mL of a commercial 40% acrylamide:bis-acrylamide [29:1 ratio] solution with 3 mL of 10 TBE buffer, and adjust the volume to 30 mL with deionized water (see Note 4). 7. 25% (w/v) Ficoll 400 in sterile deionized water. 8. Control Ficoll buffer: 25% Ficoll 400 solution supplemented with 0.01% (w/v) xylene cyanol and 0.01% (w/v) bromophenol blue. 9. Fluorescent gel imager (e.g., Typhoon FLA 9500 from GE Healthcare) and associated software for gel image analysis (e.g., ImageQuant TL software from GE Healthcare). 2.3 Helicase Microplate Assay

1. Items 1 and 2 from Subheading 2.1. 2. Purified RNA-DNA hybrid from Subheading 3.2. 3. 100 μM stock solution of oligodesoxyribonucleotide Oligo3 (Table 1) (see Note 6). 4. 2.8 μM Rho hexamer stock solution in Rho storage buffer (50% glycerol, 100 mM KCl, 0.1 mM EDTA, 0.1 mM DTT, 10 mM Tris-HCl, pH 7.9). Preparation of the Rho protein from Escherichia coli is detailed in volume 587 of the series [14]. 5. 5 helicase buffer: 750 mM potassium acetate, 100 mM HEPES pH 7.5, 0.5 mM EDTA, 0.5 mg/mL BSA (see Notes 7 and 8). 6. 1 helicase buffer obtained by diluting the 5 helicase stock buffer with water. 7. 8 initiation mix: 8 mM rATP and 8 mM MgCl2 in 1 helicase buffer. 8. 8 control mix: 8 mM rADP and 8 mM MgCl2 in 1 helicase buffer.

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9. Low adsorption, black 384-well microplates (e.g., NBS microplates from Corning) (see Note 9). 10. 16.5 mM bicyclomycin inhibitor solution in 1 helicase buffer. 11. Fluorescence microplate reader equipped with micro-injectors (see Note 10) and associated software. 2.4 Gel-Shift Helicase Assay

1. Items 5–9 from Subheading 2.1. 2. Item 9 from Subheading 2.2. 3. 10% (w/v) sodium dodecyl sulfate (SDS) in water. Store at room temperature. 4. 2 quench buffer: 8% (v/v) Ficoll 400, 1% (w/v) SDS, 40 mM EDTA, 0.3 M sodium acetate. 5. Control helicase buffer: 4% (v/v) Ficoll 400, 0.5% (w/v) SDS, 20 mM EDTA, 0.15 M sodium acetate, 0.01% (w/v) xylene cyanol, and 0.01% (w/v) bromophenol blue. 6. Helicase PAGE solution: 8% (v/v) acrylamide and 0.5% (w/v) SDS in 1 TBE buffer. Mix 6 mL of a commercial 40% acrylamide:bis-acrylamide [19:1 ratio] solution with 1.5 mL of 10% SDS solution and 3 mL of 10 TBE buffer, and adjust the volume to 30 mL with deionized water (see Note 4). 7. SDS run buffer: 1 TBE buffer containing 0.5% (w/v) SDS.

3

Methods

3.1 Preparation of the RNA Strand by Transcription

Our standard RNA strand used for assembly of the RNA-DNA hybrid substrates is 129 nucleotide (nt) long (see substrate C in ref. 19) and is prepared by in vitro transcription of a linear DNA template obtained by PCR amplification (see Note 2). The DNA template contains a pT7 promoter for the bacteriophage T7 RNA polymerase followed by sequences encoding a strong Rut (Rho utilization site) site [21, 22] and the reporter region recognized by the fluorophore- and quencher-labeled oligonucleotides (Fig. 1a). 1. Slowly melt on ice the 5 transcription buffer, the DNA template solution, and the stocks of rNTPs. Homogenize each solution by gentle vortexing before use. 2. In a 1.5 mL microtube, mix 111 μL of water; 50 μL of 5 transcription buffer; 12.5 μL of each the rATP, rCTP, rGTP, and rUTP stock solutions; 1 μL of SUPERase-IN; and 25 μL of the 100 nM DNA template stock. Incubate the mixture for 5 min at 37  C. 3. Add 10 μL of T7 RNA polymerase, and incubate the mixture for 2 h at 37  C under shaking (300–400 rpm).

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4. After 2 h, add another 10 μL of T7 RNA polymerase, and incubate the mixture for 1 h at 37  C (see Note 11). 5. To stop the reaction, add 12 μL of 0.5 M EDTA and 28 μL of 3 M sodium acetate. 6. Add 150 μL of phenol:chloroform:isoamyl alcohol extraction mix. Vortex and centrifuge for 5 min at 18,000 g to separate the organic and aqueous phases. Transfer the top phase (aqueous) to a new microtube. 7. Add 300 μL of diethyl ether, vortex quickly, and centrifuge for 1 min at 18,000 g. Remove and discard the top ether phase. Repeat this procedure once (see Note 12). 8. Add 900 μL of ethanol, vortex, and incubate overnight at 20  C to precipitate RNA. 9. Centrifuge for 30 min at 20,000 g, and then remove and discard supernatant. Add 150 μL of ice-cold ethanol, vortex vigorously, and centrifuge for 15 min at 20,000 g. 10. Remove ethanol, carefully avoiding the RNA pellet (which should be visible at this stage). Leave the microtube open in a dry bath (in a fume hood) for at least 10 min at 30  C to dry the pellet. 11. Dissolve the RNA pellet in 20 μL of M10E1 buffer, and incubate for 10 min at 30  C under shaking (300–400 rpm). The pellet should not be longer visible. Otherwise, incubate for a few minutes longer. 12. Assemble PAGE gel plates and spacers according to the manufacturer’s instructions. We use custom-made 20  20 cm gel plates with 0.8 mm spacers and a 10-teeth comb. 13. Use tape or any other suitable mean to seal hermetically the bottom of the gel plates. 14. Mix 25 mL of the denaturing 6% acrylamide solution with 60 μL of 25% APS and 30 μL of TEMED. Pour the mixture between the plates, carefully avoiding formation of air bubbles, and insert the comb. 15. Once the gel has polymerized (15–20 min at room temperature), remove the comb, and wash the wells with 1 TBE buffer using a 10 mL syringe. Install the gel into a vertical electrophoresis unit, and fill the top and bottom tanks with 1 TBE buffer. Set the power supply at 25 W, and perform a pre-electrophoresis run for 30 min (see Note 13). Turn power off. 16. Add 20 μL of formamide to the RNA sample from step 11, and heat the mixture at 90  C for 1 min. Prepare a control sample by mixing 5 μL of control formamide buffer with 5 μL of

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M10E1 buffer. Flush diffusing urea from gel wells using a syringe filled with 1 TBE. Load the RNA and control samples in well-separated wells. 17. Run the gel at 25 W until the bromophenol blue in the control sample reaches the bottom of the gel (see Note 14). 18. Remove the plates, and carefully wrap the gel in Saran sheets (see Note 15), limiting formation of bubbles and wrinkles as much as possible. 19. Place the gel on a fluor-coated TLC plate, and visualize the transcript band with a 254 nm lamp by UV shadowing (Fig. 1b). 20. Cut the gel piece corresponding to the correct-size RNA shadow with a clean scalpel blade, and transfer the piece to a 2 mL microtube. Use a pipette tip or tissue grinder to crush the gel piece in small fragments, and add 1 mL of elution buffer. Incubate for 1 h at 30  C under shaking (500 rpm). 21. Briefly centrifuge at 2000 g before transferring the supernatant into a 5 mL microtube kept on ice (see Note 16). 22. Add 1 mL of elution buffer to the tube containing the gel fragments, and incubate for 1 h at 30  C under shaking (500 rpm) (see Note 17). 23. Briefly centrifuge at 2000 g, and transfer supernatant to the tube from step 21. To remove gel debris, filter the solution with a 5 mL syringe equipped with a glass wool plug and a 0.2 μm filter. 24. Mix with three volume equivalents of cold ethanol, and incubate overnight at 20  C. 25. Centrifuge for 30 min at 20,000 g. Discard supernatant (watch for pellet, which may be barely visible and stick poorly to the tube), and add 500 μL of ice-cold ethanol. Briefly vortex and centrifuge for 15 min at 20,000 g. Discard supernatant. 26. Leave the microtube open in a dry bath (in a fume hood) for at least 10 min at 30  C to dry the pellet. Dissolve pellet in 50 μL of M10E1 buffer. 27. Use a UV spectrophotometer to determine RNA concentration from the absorbance at 260 nm. We assume that ε260nm ¼ number of nucleotides  104 L/mol/cm. Typical yields range between 200 and 500 pmol of purified RNA from a 250 μL transcription. 28. Store the RNA solution at 20  C. 3.2 Preparation of the RNA-DNA Hybrid Substrate

Oligonucleotides and hybrid substrates containing the Alexa488 or Dabcyl moieties are fragile and should be handled with care (see Note 5). In order to detect unambiguously the position of

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RNA-DNA hybrid species in PAGE gels, we also prepare a variant of the tripartite RNA-DNA substrate (Fig. 1a) devoid of Dabcyl quencher (Fig. 1c). 1. In a 1.5 mL microtube, mix 50 pmoles of RNA (from Subheading 3.1, step 28), 3 μL of 5 helicase buffer, 0.8 μL of 100 μM Oligo1, and 0.8 μL of either Dabcyl-labeled Oligo2 or unlabeled Oligo4 (100 μM each). Adjust final volume to 12.5 μL with water. 2. Heat the mixture for 2 min at 90  C in a dry bath. Remove the tube from the dry bath, and incubate for 15 min at room temperature. 3. Assemble PAGE gel plates using 0.4 mm spacers, and comb (see Note 18) as described in steps 12 and 13 of Subheading 3.1. 4. Mix 15 mL of native 6% acrylamide solution with 35 μL of 25% APS and 35 μL of TEMED. Pour the mixture between the plates and insert the comb. 5. Once the gel has polymerized, remove the comb, and flush the wells with a 10 mL syringe containing 1 TBE. Install the gel into an electrophoresis unit, and fill the top and bottom tanks with 1 TBE. Set the power supply at 140 V, and perform a pre-electrophoresis run for 30 min. Turn off the power. 6. Add 2.5 μL of 25% Ficoll 400 to each sample, and load samples into well-separated gel wells (see Note 19). In a separate well, load a control sample made by mixing 5 μL of M10E1 buffer with 1 μL of control Ficoll buffer. The gel wells should be flushed with 1 TBE right before loading of the samples. 7. Run the gel at 140 V for approximately 4 h, until the bromophenol blue in the control sample is at the very bottom of the gel. 8. Remove and replace one of the plates by a Saran sheet, and create marks with pieces of tape or a permanent marker pen on the Saran sheet (see Note 20). 9. Scan the gel with a fluorescence imager using settings adequate for detection of Alexa488 fluorescence (see Note 21). Print the gel image at real size. 10. Use the tape or pen marks (Fig. 1c) to position the gel on the printed image. Use a clean scalpel blade to cut the gel band corresponding to the correct hybrid substrate, carefully avoiding the band right below (corresponding to a hybrid substrate lacking Oligo2; Fig. 1c). Transfer the gel piece in a 2 mL microtube. 11. Add 1 mL of elution buffer, and incubate for 1 h at 18  C (see Note 22) under shaking (500 rpm).

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12. Briefly centrifuge, and transfer the supernatant into a 5 mL microtube kept on ice. 13. Add 1 mL of elution buffer to tube containing the gel piece. Incubate for 1 h at 18  C (see Notes 17 and 22) under shaking (500 rpm). 14. Briefly centrifuge and transfer the supernatant into the tube from step 12. Mix and dispatch the resulting supernatant pool into 2 mL microtubes (in ~500 μL aliquots), and add 1.5 mL of ethanol per tube (see Note 16). Vortex and incubate overnight at 20  C. 15. Centrifuge for 30 min at 20,000 g and discard supernatant. Add 500 μL of ice-cold ethanol, vortex, and centrifuge for 15 min at 20,000 g. Discard supernatant. 16. Leave the microtubes open at room temperature (in a fume hood) until the pellets are dry. Resuspend each pellet in 10 μL of 1 helicase buffer (see Note 23). 17. Pool sample aliquots, and determine hybrid concentration from absorbance at 260 nm, neglecting the effect of oligonucleotide hybridization on the extinction coefficient: ε260nm ¼ number of nucleotides (all strands)  104 L/mol/ cm. 18. Store hybrid solution at 20  C (see Note 24). 3.3 Helicase Microplate Assay

The following protocol has been developed with a Synergy H1FD instrument (BioTek) equipped with a temperature controller, two microinjectors, and 485/20 nm (excitation) and 530/25 nm (emission) filters for Alexa488 detection but should be easily adapted to any similar fluorescent microplate reader. 1. Switch on the microplate reader and set the temperature chamber to 37  C. Program the instrument method for the experiment at hand (see step 7 or step 16, below). Clean the microinjector circuits with 1 mL of ethanol and then 1 mL of deionized RNase-free water. 2. Warm the 8 initiation mix and the 8 control mix in a dry bath set to 37  C before loading each mix in separate injector reservoirs (see Note 25). 3. In a 1.5 mL microtube, prepare a 241 μL reaction mixture containing 5.8 nM fluorescent hybrid (see Note 26) and 464 nM unlabeled trap oligonucleotide in final 1 helicase buffer. Transfer 80 and 160 μL aliquots to two new microtubes (Samples 1 and 2) (see Note 27). Add 1.3 μL of 1 helicase buffer in Sample 1. 4. Prepare a 1400 nM dilution of the Rho enzyme stock in 1 helicase buffer (see Note 28). Add 2.6 μL of the Rho dilution to Sample 2.

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5. Load 35 μL sample aliquots in a defined order in the wells of a 384-well microplate. For instance, load Sample 1 aliquots in wells 1 and 2 and Sample 2 aliquots in wells 3–6. In this way, each condition will be tested in duplicate in a single experiment. 6. Add 5 μL of 1 helicase buffer in wells 1 and 2 (“no Rho” control replicates). 7. Install the microplate in the instrument, and run the following method (temperature set to 37  C throughout): shake (orbital) the microplate plate for 5 min at 180 rpm. Then, use microinjectors to dispense 5 μL of 8 control mix in wells 3 and 4 (ADP control replicates) and 5 μL of 8 initiation mix in wells 5 and 6 (unwinding reaction replicates). Record Alexa488 fluorescence from each well every 20 s for at least 30 min (Fig. 2a). Set the instrument to normalize fluorescence output I(t) according to: I ðt Þ ¼ I t II0 0 where It and I0 are, respectively, the fluorescence signals recorded at times t and 0. Also set the instrument to average the replicate signals obtained for each condition (Fig. 2b). 8. Repeat steps 1–7 several times, preferably on different days, to appreciate the experimental variation (Fig. 2c) (see Note 29). The above protocol can also be used to assess the effect of drugs or cofactors on Rho helicase activity. The additional reagent can be added directly into the master mix (step 3 above) or in individual reaction samples. In the latter case, the protocol can be modified, as follows (example provided for testing the effect of the bicyclomycin [BCM] inhibitor): 9. Perform steps 1 and 2 above. 10. In a 1.5 mL microtube, prepare a 426 μL master mix containing 6.6 nM fluorescent hybrid and 525 nM unlabeled trap oligonucleotide in 1 reaction buffer. Transfer one 142 μL aliquot and four 71 μL aliquots to five 1.5 mL microtubes (Samples 3–7). 11. Add 18.6 μL of 1 helicase buffer to Sample 3. 12. From the BCM inhibitor stock, prepare two serial BCM dilutions in 1 helicase buffer. Adjust dilutions to the concentration range to test (e.g., 1000 and 100 μM stock dilutions to obtain final BCM concentrations in samples of 100 and 10 μM, respectively). 13. Add 9.3 μL of one of the BCM serial dilutions to Samples 4 and 5 and 9.3 μL of the other serial dilution to Samples 6 and 7. 14. Prepare a 1400 nM dilution of the Rho enzyme stock in 1 helicase buffer (see Note 28). Then, add 2.7 μL of Rho protein in Sample 3 and 1.3 μL in Samples 4–7.

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a Fluorescence (au)

8000

6000

4000

2000

0

0

10

20

30

Time (min)

c

4

(I-I0)/I0 signal

(I-I0)/I0 signal

b

3 2 1 0

0

10

20

Time (min)

30

4 3 2 1 0

ATP -Rho

ADP ATP

ATP + BCM

+Rho

Fig. 2 Detection of Rho-directed RNA-DNA hybrid unwinding with the fluorescent microplate assay. Data in grey, black, blue, and red correspond to samples without Rho, with Rho and ADP, with Rho and ATP, and with Rho, ATP, and 100 μM BCM, respectively. (a) Representative graph showing raw fluorescence traces obtained with sample replicates from the same experiment. (b) Graph showing the evolution of the averaged normalized fluorescence signals as a function of time (see Subheading 3.3, step 7). Data points were fitted to an equation describing biphasic kinetics (pseudo-first-order burst followed by linear steady-state phase), as described
previously [22, 24]: F ðt Þ ¼ A 1  ek exp t þ k lin t , where F(t) is the fraction of unwound product, A is the amplitude of the exponential phase, and kexp and klin are the rate constants of the exponential and linear phases, respectively. (c) Box plot showing the distribution of normalized fluorescence intensities values recorded at t ¼ 30 min (5–7 independent experiments per tested condition)

15. Load 35 μL sample aliquots in a defined order in the wells of a 384-well microplate. For instance, load Sample 3 aliquots in wells 1–4 and Samples 4–7 in wells 5–12 (two wells per sample).

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16. Install the microplate in the instrument, and run the following method (temperature set to 37  C throughout): shake (orbital) the microplate plate for 5 min at 180 rpm. Then, use microinjectors to dispense 5 μL of 8 control mix in wells 1 and 2 (ADP control replicates) and 5 μL of 8 initiation mix in wells 3–12 (helicase reaction replicates). Record and report Alexa488 fluorescence for all wells as described in step 7, above. 17. Repeat steps 9–16 several times with distinct serial dilutions of BCM to cover the inhibitory concentration range adequately (Fig. 3a).

a Relative fluorescence intensity

1 0.8 0.6 0.4 0.2 0 1

10

100

1000

[BCM] (µM)

b 90°C

Dabcyl-less substrate Dabcyl-plus substrate

ADP 0

ATP

30 0

30

ATP +BCM ADP 0

30

0

ATP ATP +BCM

30 0

30 0

30

(min)

RNA-DNA hybrid Oligo1

Fig. 3 Inhibition of Rho helicase activity by BCM. (a) Graph showing BCM inhibition “endpoint” data (taken after 30 min of helicase reaction) obtained with the fluorescence multiplate assay (Subheading  3.3). The data n points were fitted to a standard sigmoid inhibition equation: I BCM ¼ F 0  F max  ½BCM½BCM , where IBCM n n  þ½IC50  is the normalized fluorescence intensity (at 30 min of reaction) as a function of BCM concentration, F0 is the fitted value of IBCM at 0 μM BCM, Fmax is the maximal fraction of the signal that is sensitive to BCM, n is an empirical parameter that defines the shape of sigmoid curve, and IC50 is the half-maximal inhibitory concentration [23]. (b) Representative 8% PAGE gel showing Rho helicase reaction products obtained under different reaction conditions with a tripartite RNA-DNA hybrid substrate (Fig. 1a) containing the unlabeled Oligo4 (Dabcyl-less, left) or Dabcyl-labeled oligo2 (Dabcyl-plus, right) strand. Note that the presence of the Dabcyl quencher prevents fluorescence detection of the Dabcyl-plus RNA-DNA hybrid species in the gel.

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18. Repeat the procedure over the full inhibitory concentration range to appreciate the experimental variation. With this procedure, we estimated an apparent IC50 of ~15 μM (Fig. 3a), which is in reasonable agreement with previous measurements performed with a more classical helicase assay (IC50 ¼ 49 μM) [23]. 3.4

Gel-Shift Assay

To verify that the development of the Alexa488 fluorescence signal observed with the multiplate assay truly derives from Rho helicase activity (the assay cannot distinguish Rho-directed RNA-DNA hybrid unwinding from unwanted side reactions such as RNA degradation), one may control endpoint reaction products by PAGE (see Note 1). PAGE analysis may be performed occasionally (when problems are suspected; see Note 29) or on a more regular basis (as part of a quality control procedure). To facilitate data analysis, experiments with a control RNA-DNA hybrid substrate devoid of Dabcyl (prepared by using Oligo4 instead of Oligo2 from Table 1; see Subheading 3.2, step 1) may be performed in parallel (Fig. 3b). 1. Prepare 0.5 mL microtubes (one per sample to be analyzed) containing 6 μL of 2 quench buffer. To each tube, add 6 μL of the sample to be analyzed (see Note 30). For instance, a control “t0” sample may come from the master mix of Subheading 3.3, step 3 (slightly increasing the overall master mix volume to account for this aliquot withdrawal), while other samples may correspond to endpoint reaction aliquots taken from microplate wells after Subheading 3.3, step 7. Store on ice. 2. Assemble PAGE gel plates as described in steps 12 and 13 of Subheading 3.1 using 0.8 mm spacers and a 15-teeth comb. 3. Mix 25 mL of 8% helicase PAGE solution with 60 μL of 25% APS and 60 μL of TEMED. Pour the mixture between the plates and insert the comb. 4. Once the gel has polymerized, remove the comb, and flush the wells with a 10 mL syringe containing SDS run buffer. Install the gel into an electrophoresis unit, and fill the top and bottom tanks with SDS run buffer. Set the power supply at 140 V, and perform a pre-electrophoresis run for 30 min. Turn off the power. 5. Flush gel wells with SDS run buffer and load samples from step 1. Also load 5 μL of control helicase buffer in a well-separated well (positions of the xylene cyanol and bromophenol blue dyes are used to assess gel migration) (see Note 31). 6. Run the gel at 140 V until the xylene cyanol in the control sample is at ~5 cm from the bottom of the wells (it takes ~1 h).

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7. Replace one of the glass plates with Saran sheet. Scan the gel with a fluorescence imager using settings adequate for detection of Alexa488 fluorescence (see Note 21).

4

Notes 1. Only PAGE analysis of helicase reaction products allows the precise calculation of substrate unwound fractions as function of time and the verification of the integrity of the substrate (s) and reaction products (an important point with fragile RNA components and the risk of degradation by chemicals, RNases, etc.). The fluorogenic assay presented herein should thus be considered as a complementary tool allowing much quicker, early-stage exploration of samples or conditions but not being necessarily suitable for the thorough, late-stage characterization of “hits.” 2. The DNA template is prepared by standard PCR amplification procedures [9] and can be purified with a commercial silicabased PCR purification kit. 3. DTT is used to prevent formation of disulfide bonds inhibiting T7 RNA polymerase but is not stable over time. We recommend using fresh transcription buffer or to supplement old batches with additional DTT. 4. The PAGE solutions can be prepared in advance or in larger volumes and stored for a few days at room temperature protected from light. 5. Oligonucleotides and hybrids bearing the Alexa488 moiety should be protected from light to limit risks of photobleaching. Dabcyl is a collisional quencher requiring close molecular interaction (aromatic stacking) with the fluorophore; its quenching efficiency is not strongly wavelength-dependent. 6. Oligo3 is used as a “trap” competitor in helicase reactions and should have the same sequence than the Alexa488-labeled Oligo1 oligonucleotide. 7. The composition of the helicase buffer may be varied, but ionic strength should be sufficient (>50 mM monovalent salt) to stabilize the RNA-DNA helices within the hybrid substrate. 8. BSA is used as a non-specific protein competitor and contributes to limit the potential adsorption of reactants to microtube walls. 9. We prefer black microplates for their higher signal-to-noise ratio. We like Corning 384-well microplates #3575 for their consistent, low protein and nucleic acid adsorption properties and for their suitable 20–80 μL working volume range.

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10. Make sure that the microplate reader injectors and the microplate holding chamber are RNase-free. Use standard procedures or commercial RNase decontamination reagents to clean the instrument parts as well as surrounding working area if needed. 11. This step is optional but usually improves the yield of RNA transcripts. 12. The tube can be left open in a fume hood for a few minutes before proceeding to the next step (potential traces of ether will thus be eliminated by evaporation). 13. Make sure that the gel plates are warm before ending the pre-electrophoresis step. 14. In a 6% denaturing gel, bromophenol blue migrates as a ~25nt-long RNA strand, while xylene cyanol dye migrates slightly slower than our standard 129-nt-long RNA transcript (Fig. 1b). 15. Use only UV-transparent plastic wrap (e.g., Saran™ wrap). Some commercial plastic wraps contain UV absorbers that can perturb detection of RNA bands by UV shadowing. 16. Precipitation can also be performed in 5 mL microtubes (dispatching 1.25 mL sample and 3.75 mL ethanol per tube) if a high-speed bench centrifuge with an adequate rotor is available (e.g., Eppendorf 5425 centrifuge and FA-10X5 rotor). The hybrid substrate pellets are then more easily detected from their larger sizes and label-dependent colors (dark orange with the Alexa488-Dabcyl pair or yellowish with only Alexa488). 17. Incubation overnight at 4  C is an alternative option. 18. The hybrid substrates can also be purified on a 0.8 mm polyacrylamide gel using a larger volume (25 mL) of acrylamide solution for gel preparation. However, elution from the gel band is less efficient and requires crushing of the gel band into small pieces as well as syringe filtration as in Subheading 3.1, steps 20–23. Final yields are usually lower. 19. We use a comb with 1.5-cm-wide teeth for our 0.4-mm-wide gels, and load sample volumes of 20 μL or less per well. The number of wells used per sample or the width of comb teeth (or gel width, but see Note 18) should be adjusted according to the case at hand. 20. For samples containing Alexa488, we prefer red indelible markers. To facilitate batch-to-batch comparisons, we mark the positions of the xylene cyanol and bromophenol blue dyes from the control sample whenever possible.

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21. Detection settings will depend on the imaging system. In any case, make sure to use high-grade acrylamide and chemicals and to clean thoroughly the imaging system between scans (see manufacturer’s instructions) to limit background fluorescence. 22. Although the tripartite RNA-DNA hybrids are reasonably stable and do not spontaneously dissociate under standard Rho helicase conditions (see Subheadings 3.3 and 3.4), we prefer to use low temperatures (10 pN). To directly measure the average lifetime of a G4 at a given force, one can implement a force-jump procedure involving two force levels: (1) a G4-folding force, a sufficiently low force at which the folded G4 conformation is energetically favored, and (2) G4-unfolding force, a higher force at which G4 unfolds. After the force jump to a target higher force, the lifetime until the G4 unfolding is recorded. At forces greater than 10 pN, the extension difference between the unfolded and folded states of a typical G4

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structure is more than 5 nm (see Note 1). The unfolded G4 can be refolded by switching force back to the G4-folding force. Repeating the procedure many cycles from the tether, one can obtain the average lifetime τ(F) of the G4 in the tether. To eliminate the potential tether-to-tether and experiment-to-experiment variations, data to calculate the average lifetime should be acquired from multiple independent tethers and from multiple independent experiments. For the G4 refolding rate kr(F) (the subscript “r” indicates refolding), at low forces of a few pN, the extension difference between the unfolded and folded states is too small to be unambiguously resolved. Instead, the refolding rate of an unfolded G4 can be quantified by implementing a revised force-jump cycle (see Note 2). With the knowledge of the force-dependent unfolding and refolding rates, the folding energy of the G4 can also be determined (see Note 3). 1.3 Detecting the Binding and Unwinding Activities of DHX36 Based on the Altered Lifetime of G4

2

The DHX36 helicase binds G4 with high affinity, characterized by a sub-nanomolar dissociation constant [36, 37]. While the highaffinity-binding might potentially distort the G4 structure from its most stable conformation, the binding may provide additional energy to stabilize the overall folded conformation of the G4. Thus, it is expected that the binding of DHX36 may alter the average lifetime of the G4 under force, which can potentially be detected using magnetic tweezers. In addition, as DHX36 may bind to a G4 with different affinities at different stages in one ATPase cycle, it may result in different levels of alteration to the average lifetime of the G4. Finally, the ATP-dependent G4-unwinding activity of DHX36 should lead to a faster G4 unfolding. All these could also be potentially detected in experiments using magnetic tweezers.

Materials

2.1 Magnetic Tweezers Components (Fig. 1a)

1. An optical microscope (e.g., Olympus IX71 inverted microscope used in our study). 2. A high-NA oil-immersion objective lens (e.g., 100 oil-immersion UPlanFLN NA1.3, Olympus used in our study). 3. A light-emitting diode (LED) light source (e.g., Thorlabs, MCWHL2-C1 used in our study) used to image the bead using transmission illumination [38, 39] or backscattered illumination [25, 40]. 4. A charge-coupled device (CCD) or a complementary metaloxide-semiconductor (CMOS) camera to image the bead (e.g., the Pike F-032B CCD camera, Adept Turnkey, used in our study).

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5. An objective piezo actuator (e.g., Physik Instrumente P-725. CDD used in our study) to scan the bead images at different imaging planes, for building a library of the diffraction patterns of the same bead at the different heights from the surface [25, 41]. 6. A pair of permanent magnets to generate force to the bead attached to the DNA construct. 7. A single-axis linear step motor (e.g., Moco manipulator from Micos used in our study) to change the vertical position of the pair of magnets, through which the force applied to the bead can be controlled. 8. A program to control the position of the piezo, linear motor, capture the bead images, and determine the 3-D positions of the bead in real time (an in-house written Labview program was used in our lab). 9. A workstation to run the Labview program (e.g., Windows 10 running Labview 2015 in our study). 2.2 Microfluidic Channel Preparation

1. Cover glasses thickness no. 1 (superior Marienfeld Laboratory Glassware). 2. Cover glasses thickness no. 0 (superior Marienfeld Laboratory Glassware). 3. Parafilm or double-sided tape (Grace Bio-Labs, Inc.) 4. Decon 90™ liquid detergent (Decon Laboratories™) or equivalent. 5. 40% (v/v) of Decon 90™ detergent in deionized water. 6. 1% (v/v) 3-aminopropyl triethoxysilane (APTES) solution in methanol. 7. Ultrasonic cleaning bath (e.g., Elmasonic S 30 H, Elma Schmidbauer GmbH). 8. Basic plasma cleaner (e.g. PDC-32G, Harrick Plasma). 9. High-grade methanol (99.6%).

2.3 Reagents, DNA, and Protein Preparation

1. 2 U/μL Q5 hot-start DNA polymerase with 5 Q5 reaction buffer (New England Biolabs). 2. 100 mM dNTP stock. 3. DreamTaq DNA polymerase with 10 DreamTaq reaction buffer (Thermofisher Scientific). 4. Thermocycler. 5. 500 μg/mL solution of lambda phage DNA. 6. 100 μM stock solutions of the following DNA oligonucleotides in water:

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FWD1

50 ATCACCAAGTGCATGGTGCTTGAA CCCGCCTATG

REV1

50 Thiol-CGACTGAGCTGGCAAGCAACTGACCTG

FWD2

50 ATCACCAACGACATGGCAGGAGGGCGAATG

REV2

50 Biotin-CGAA GCAGATCCCACGCAACCAGCTTACGG

G4-forming 50 CTTGTGCACAGACTCGTATGAGGGTGG oligonucleotide GGAGGGTGGGGAATGCAGCCAGGTC AGTAGCGAC-30 Flank 1

50 Phosphate-CTACTGACCTGGCTGC

Flank 2

50 CGAGTCTGTGCACAAGGTGC

7. 10 U/μL BstXI endonuclease with supplied 10 cleavage buffer. 8. 400 U/μL T4 DNA ligase with supplied 10 ligation buffer. 9. Kit to purify DNA fragments from PCR reactions (Invitrogen). 10. UV-vis spectrophotometer with microvolume capability (e.g., NanoDrop™ 2000c, Thermo Fisher Scientific) to determine DNA concentration. 11. 1 phosphate-buffered saline (PBS), pH 7.4. 12. 1 mg/mL sulfosuccinimidyl 4-(N-maleimidomethyl)cyclohexane-1-carboxylate (Sulfo-SMCC) (Thermo Scientific) in 1 PBS buffer. 13. Streptavidin-coated paramagnetic beads (e.g., Dynal M-280, Thermo Scientific). 14. 3 μm diameter polystyrene beads (e.g., Polysciences beads), which serve as a reference to eliminate the mechanical and thermal drift of the sample channel. 15. BSA passivation solution: 10 mg/mL BSA and 1 mM 2-mercaptoethanol in PBS. 16. 1 unwinding assay buffer: 10 mM Tris–HCl (pH 8.0), 100 mM KCl, and 2 mM MgCl2. 17. ADP removal buffer: 200 μg/mL pyruvate kinase from rabbit muscle and 2.5 mM phosho(enol)pyruvic acid. 18. 4 U/μL hexokinase. 19. 200 mM D-glucose. 20. 10 mM stock solutions of ATP, AMP-PNP, and ADP, buffered to pH 7.5 with NaOH. 21. 10 mM stock solution of ADP·AlF4, prepared by incubating 10 mM ATP-depleted ADP, 50 mM NaF, and 10 mM AlCl3

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22. 1 Assay buffer: 10 mM Tris–HCl, pH 8.0, and 100 mM KCl 23. 30 μM stock solution of Drosophila melanogaster or human DHX36 protein, prepared as described previously [14].

3

Methods

3.1 Preparing a Single-Molecule Tether That Contains a G4 Structure Spanned Between Two dsDNA Handles

1. To prepare DNA fragment PCR1 (Fig. 2a), mix 1 μL of lambda phage DNA, 1 μL of FWD1 oligonucleotide, 1 μL of REV1 oligonucleotide, 20 μL of 10 DreamTaq reaction buffer, 4 μL of the dNTP stock, 172 μL of deionized water, and 1 μL of DreamTaq DNA polymerase. 2. Place sample in a thermocycler and run the following program:

Step

Temperature ( C)

Time

Initial denaturation

95

1 min

30 cycles

95 55 72

30 s 30 s 2 min

Final extension

72

5 min

Hold

4

3. Use a PCR cleanup kit to purify the PCR1 fragment, following manufacturer’s instructions. 4. Repeat steps 1–3 to prepare DNA fragment PCR2 (Fig. 2a), replacing oligonucleotides FWD1 and REV1 by oligonucleotides FWD2 and REV2 (see Note 4). 5. Use a microvolume spectrophotometer to determine concentrations of PCR1 (489 base pairs) and PCR2 (593 base pairs) DNA fragments by absorbance at 260 nm, assuming an average extinction coefficient for double-stranded DNA of 0.020 (μg/ mL)1 cm1. 6. Digest each fragment with BstXI restriction enzyme to create sticky ends for the ligation reaction in the next step. Mix 1 μg of each DNA fragment with 1 μL of BstXI, 5 μL of 10 NEB buffer, and deionized water to 50 μL total volume. Incubate the mixtures for 60 min at 37  C. 7. Use a PCR cleanup kit to purify the PCR1 and PCR2 fragments, following manufacturer’s instructions. 8. Mix 1 μL 100 μM G4-forming oligonucleotide, 1 μL 100 μM Flank 1, and 1 μL 100 μM Flank 2 oligonucleotides (see Fig. 2a), in 97 μL deionized water, which produces ~1 μM

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Fig. 2 Schematic of the G4 DNA construct and the single-molecule stretching experiments. (a) The G4 DNA construct is formed by ligation of an ssDNA containing a G4-forming sequence and two dsDNA handles. (b) A G4 DNA construct is tethered between a superparamagnetic bead and a coverslip surface, stretched by a force generated using a magnetic tweezers setup. (c) An example of force-jump measurement from 1 to 54 pN. The ~8 nm stepwise bead height increase after jumping to the higher force indicates the unfolding of G4 (pink arrows). The much larger stepwise bead height occuring at the time of force jump is from bead rotation

annealed product that contains the G4-forming sequence spanned between two double-stranded DNA annealed by Flank 1 and Flank 2. 9. Ligate the product produced in step 8 with the PCR1 and PCR2 dsDNA handles using a molar ratio of 1:1:1 (Fig. 2a). Mix 1 μL PCR 1 (about 5–7 μg), few μL PCR 2 dsDNA handles based on equal molar concentration ratio to PCR1, few μL G4 DNA based on equal molar concentration ratio to PCR1, 1 μL T4 DNA ligase, and 5 μL 10 T4 ligase buffer and deionized water to a final volume of 50 μL. Incubate the mixture min at 16  C for overnight. 10. Use a PCR cleanup kit to purify the ligated DNA construct, following manufacturer’s instructions. 11. Use a microvolume spectrophotometer to determine concentration of the G4 forming DNA construct by absorbance at 260 nm, assuming an average extinction coefficient for double-stranded DNA of 0.020 (μg/mL)1 cm1.

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3.2 Microfluidic Channel Functionalization and Assembly

1. Place the bottom coverslip (thickness no. 1, 22 mm  32 mm) and top coverslip (thickness no. 0, 20 mm  20 mm) into a glass jar and add 40% (v/v) of Decon 90™ detergent. 2. Submerge the jar into the ultrasonic cleaning bath for 30 min. Rinse the coverslips in the jar with distilled water for >10 times to remove the detergent. 3. Dry the coverslips in the jar in the oven. Use basic plasma cleaner to clean the bottom coverslips for 10 min. 4. Merge the bottom coverslips in 1% APTES solution for 1 h at room temperature to functionalize the bottom coverslip with APTES. Then, wash with pure methanol and dry in oven. 5. Place the cleaned top coverslip on the APTES functionalized bottom coverslip, separated with two parallel parafilm stripes to create a flow cell of 10–20 μL in volume (Fig. 1b).

3.3 Attaching the G4 DNA Tether to the Coverslip Surface and Bead

1. Prepare a 1:100 dilution of the polystyrene bead (1.68  109 particles/mL) suspension in 1 PBS. Flow the dilution into the channel of the flow cell from Subheading 3.2, step 5. Incubate for 10 min at room temperature (27  C). 2. Flush the microfluidic channel with 1 PBS to wash out unbound polystyrene beads. The polystyrene beads stably stuck to the bottom surface of the flow cell serve as a reference to eliminate the mechanical and thermal drift of the sample channel. 3. Fill the channel with 1 mg/mL sulfo-SMCC and incubate at room temperature (27  C) for 30 min, to coat the APTESfunctionalized bottom surface with sulfo-SMCC. Flush 1 PBS through the channel to wash out unbound sulfo-SMCC. 4. Dilute thiol-labeled G4 DNA construct to 5 ng/μL with 1 PBS and then introduce the thiol-labeled G4 DNA construct solution to the channel and incubate for 30 min at room temperature (27  C), to attach the G4 DNA construct to the SMCC-coated surface. Then, rinse the channel with 1 PBS. 5. Fill the channel with the BSA passivation solution and incubate at 4  C for 2.5 h. Then, rinse the channel with 1 PBS. Prepare a 1:50 dilution of the original paramagnetic bead suspension (10 mg/mL) in BSA passivation solution and use it to fill the channel. This will attach the biotin-labeled end of the DNA construct to the bottom coverslip surface and allow forming a tether between surface and the superparamagnetic bead (Fig. 2b). Incubate at room temperature (27  C) for 30 min.

3.4 Search for and Characterize Specific Single G4 Tethers

1. Starting from Subheading 3.3, step 5, flush the channel with 500 μL of assay buffer to remove untethered superparamagnetic beads. 2. Align the center of the magnet pair along with the optical axis and mount the channel onto magnetic tweezers (see Note 5).

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3. At low forces (10 pN) to identify the characteristic G4-unfolding signal (Fig. 2c). A single tether should only give rise to a single G4-unfolding step. Switch force back to below 1 pN (Fig. 2c) to allow the unfolded G4 to refold. For the force calibration, see Note 7. 6. Identify the G4-unfolding force, G4-folding force, and the holding time at each force that are suitable for detecting the interaction between DHX36 and the G4 using the force-jump procedure described in Subheading 1.2. Repeat the force cycles (Fig. 2c) at different forces with different holding times to identify a G4-unfolding force at which the G4 unfolds within seconds and a G4-folding force to ensure >90% G4 refolding within 1 min of holding time. The G4-folding force and G4-unfolding force may vary for different G4-forming sequence. 3.5 Force-Jump Assay for DHX36’s G4-Binding and Unwinding Activities

For the particular c-MYC G4 sequence used in the study [16], the G4-folding force and the G4-unfolding force were chosen to be 1 and 54 pN, respectively (Fig. 3a). The average lifetime of the folded G4 at 54 pN was measured to be τ ¼ 6.4  0.4 s (Fig. 3b). The force-jump assay procedure for the c-MYC G4 sequence is (1) hold the tether at 1 pN for 60 s; (2) switch force to 54 pN for 30 s; and (3) switch force back to 1 pN. Repeat the procedure for hundreds of times to obtain the statistics of G4 lifetime at 54 pN. 1. Dilute DHX36 to 10 nM with assay buffer. Flush 30 μL 10 nM DHX36 to the microfluidic channel. Repeat the same forcejump assay on the same single G4 tether. After jumping to 54 pN, the bead height should remain at the level corresponding to folded state of G4 throughout the 30 s holding time at 54 pN (Fig. 3b). Taking into account the ~6.4 s of lifetime of the naked G4, this indicates that the G4 is bound by a DHX36 helicase, which strongly stabilizes the folded G4 structure against mechanical force. 2. Repeat the same assay to measure the average lifetimes of the G4 in 10 nM DHX36 in the presence of 1 mM of different nucleotides, including nucleotide-free state, AMP-PNP, ADP ·AlF4, and ADP-bound states. The nucleotide-free, AMPPNP and ADP-bound states of DHX36 should result in strong

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Fig. 3 Force-jump experiments to measure the interactions between G4 and DHX36. (a) Illustration of detecting the binding and G4-unwinding activities of DHX36. It is expected that, if a binding stabilizes the G4, the lifetime of the G4 at the higher force should be significantly longer than that of naked G4. In contrast, the G4-unwinding activity of DHX36 in the presence of ATP hydrolysis should significantly decrease the lifetime of the G4. (b) A representative time trace of extension change of a G4 construct tether after jumping to ~54 pN for a naked G4, after introducing 10 nM DHX36, and after introducing 10 nM DHX36 and 1 mM ATP confirmed the expectations in (a). Note: As the data in panel (b) were recorded at a constant force of ~54 pN, the bead height change is equal to the extension change of the molecule

stabilization of the G4, while the ADP·AlF4-bound DHX36, which mimics the ATP hydrolysis intermediate state, should not [16]. 3. Repeat the same assay with 10 nM DHX36 and 1 mM ATP (see Note 8). Flow the mixture into the microfluidic channel. The height of the end-attached bead right after jumping from 1 to 54 pN corresponds to the level of unfolded G4 (Fig. 3b), indicating that the G4 is in an unfolded conformation prior to the force jump to ~54 pN.

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Fig. 4 Model for G4 unwinding by DHX36. (a) DHX36 (represented by its two RecA-like domains, the conserved C-terminal domain and the N-terminal RHAU-specific motif (RSM) domain) binds to a G4 structure in (i) nucleotide-free or (ii) ATP-bound form with high affinity, resulting in stabilization of G4. When DHX36 enters an ATP hydrolysis (iii) and product release (iv) intermediate states, the conformational change of DHX36 likely results in G4 destabilization. The DHX36-ssDNA-binding interface is drawn based on the structure of a closely related DEAH-box helicase. (b) Crystal structure of the bovine DHX36 bound to a G4 DNA (orange) (PDB ID: 5VHE) [12]

Taken together, the data collected in steps 1–3 can be used to demonstrate that the conformational changes of DHX36 are driven by its ATPase cycle and that the direct interactions between DHX36 and the G4 play important roles in DHX36-mediated G4 unfolding [16]. Based on the results, an ATP-dependent G4-unwinding mechanism of DHX36 is proposed (Fig. 4).

4

Notes 1. A G4 typically has 20–30 nucleotides. When it is unfolded into single-stranded DNA (ssDNA) under force, the extension of the ssDNA can be estimated based on the bending stiffness of the ssDNA. At forces greater than 10 pN, the force-extension curve of an ssDNA is known to be well described by the wormlike chain polymer model [42], with a contour length of ~0.7 nm per nucleotide and a bending persistence length of ~0.7 nm in typical solution conditions [43, 44]. Based on this model, upon unfolding, ~20 nucleotides of ssDNA released under forces >10 pN will lead to an extension increase of >5 nm, which can be easily resolved in experiments.

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2. A revised method is needed to determine kr(F). After jumping to a target lower force, the G4 tether can be held for a certain duration Δt, within which the unfolded G4 may or may not refold. Whether the G4 refolds during the holding time can be checked by jumping back to a detection force, a higher force at which the bead height difference between the unfolded and folded states can be resolved. Repeating such force-jump procedure for many cycles for a few tethers, one can obtain the folding probability at the target folding force as a function of the holding time Δt. For a two-state transition, the holding time-dependent
 probability r ðF Þ ðkr ðF Þþku ðF ÞÞΔt follows pr ðΔt Þ ¼ kr ðFkÞþk at a force F. 1  e u ðF Þ Thus, fitting the experimental data to the formula, both the folding rate and unfolding rate can also be obtained. 3. At a given force, if the unfolding rate ku(F) and refolding rate kr(F) of a G4 molecule can be measured, the force-dependent folding-free energy, the free energy of the unfolded state minus that of the folded  state,  of the G4 can be determined by ΔG ðF Þ ¼ kB T ln

k r ðF Þ ku ðF Þ

. A positive value indicates a favored

folded state. By removing the contribution from the forcedependent change in the free energy, the zero-force R F foldingfree energy can be calculated by ΔG 0 ¼ ΔG ðF Þ þ 0 Δx ð f Þdf , where Δx( f ) indicates the force–extension curve difference between the unfolded and folded states of the G4 [35, 45, 46]. Therefore, single-molecule force manipulation not only can reveal the transition kinetics of a G4 molecule but also can be used to measure the thermodynamic stability of the molecule. 4. Both dsDNA handles have high GC content (>60%) to prevent DNA melting when DNA is held at high forces. 5. Align the center of the magnet pair before mounting the channel. Use a 4 objective lens to adjust the x- and y-axis of the pair of magnets to align the center in the optical axis of the microscope. This ensures that the direction of force is perpendicular to the imaging plane and that of the magnetic field in the imaging plane. 6. The z-position of the bead is determined based on the diffraction patterns of the bead images. We use an objective piezo actuator (Physik Instrumente P-725.xDD) to image the bead at different imaging planes interspaced by a fixed distance to build a library of the diffraction patterns of the same bead at a different height from the surface. The autocorrelation of the images is fitted to a polynomial function. The z-position of the bead recorded during the experiment is obtained by maximizing its correlation with the fitted polynomial function. A library of images of a surface-stuck bead is also acquired, which

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functions as a reference to eliminate any thermal-mechanical drift of the sample channel. More details can be found in [25, 40]. 7. For a given pair of magnets, the force applied to a certain bead is solely dependent on bead-magnet distance d. Since the saturated magnetic moment of the beads may be different from one another, at the same bead-magnet distance d, the forces applied to different beads may differ from each other. The ratio of forces experienced by the two beads is the same as the ratio of saturated magnetic moments of the two beads, which is a constant. Thus, if a standard force-distance curve, F0(d), is calibrated for one bead, the force applied to another bead can be directly extrapolated by F(d) ¼ cF0(d), where c is a constant for a given bead [40]. The c value of a bead can be calibrated at a single magnet-bead distance d, based on the transverse fluctuation of the bead or based on a structural transition of DNA that is known to occur at certain force. Details can be found in [25, 40]. 8. In order to minimize contaminant perturbation, it is recommended to purify the ATP with ADP removal buffer before the experiment. Similarly, the ADP solution should be treated with hexokinase and D-glucose at room temperature for 2 h for removing the contaminating ATP. After incubation, the hexokinase can be removed using an Amicon Ultra-0.5 column.

Acknowledgments This work was supported by the Ministry of Education Academic Research Fund Tier 1 (to J. Y.); The National Research Foundation, Prime Minister’s Office, Singapore and the Ministry of Education under the Research Centres of Excellence programme (to J. Y.); The National Natural Science Foundation of China (21708009 to H.Y.), and the Fundamental Research Fund for the Central Universities (2017KFYXJJ153 to H.Y.). References 1. Rhodes D, Lipps HJ (2015) G-quadruplexes and their regulatory roles in biology. Nucleic Acids Res 43:8627–8637 2. Kwok CK, Marsico G, Sahakyan AB et al (2016) rG4-seq reveals widespread formation of G-quadruplex structures in the human transcriptome. Nat Methods 13:841–844 3. Hansel-Hertsch R, Di Antonio M, Balasubramanian S (2017) DNA G-quadruplexes in the human genome: detection, functions and

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Chapter 13 Characterization of the Brr2 RNA Helicase and Its Regulation by Other Spliceosomal Proteins Using Gel-Based U4/U6 Di-snRNA Binding and Unwinding Assays Eva Absmeier and Markus C. Wahl Abstract Functional aspects of nucleic acid helicases can be interrogated by various in vitro methods, using purified components, including nucleic acid binding and unwinding assays. Here we describe detailed protocols for the production and purification of the spliceosomal Ski2-like RNA helicase, Brr2, and one of its regulatory factors, the Jab1 domain of the Prp8 protein from yeast. Furthermore, we include a production protocol for radioactively labeled yeast U4/U6 di-snRNA substrate. We describe polyacrylamide gel-based assays to investigate Brr2’s RNA binding and unwinding activities. The purification protocols and activity assays can be easily adapted for the purification and functional interrogation of other helicases, cofactors, and RNA substrates. Key words Brr2 protein, Electrophoretic mobility shift assay, Helicase cofactor, RNA binding, RNA duplex unwinding, Ski2-like helicase, SF2 helicase, U4/U6 di-snRNA

1

Introduction RNA helicases are molecular motor proteins that were initially named after their ability to unwind double-stranded RNA substrates in vitro in an NTP-dependent manner [1]. However, the in vivo functions of these enzymes can be diverse and include nucleic acid annealing, nucleic acid clamping, and rearrangement of nucleic acid-protein complexes [2–7]. Based on conserved motifs and domains, helicases can be divided into six superfamilies (SFs) [8, 9]. Members of SF1 and SF2 are predominantly monomeric, while members of SF3–6 are multimeric, toroidal helicases. All known eukaryotic RNA helicases belong to SF1 or SF2. SF2 members can be further subdivided into nine families and one group (RecG-like, RecQ-like, Rad3/XPD, Ski2-like, T1R, Swi/Snf, RIG-I-like, DEAD-box, DEAH/RHA family, and NS3/NPH-II group).

Marc Boudvillain (ed.), RNA Remodeling Proteins: Methods and Protocols, Methods in Molecular Biology, vol. 2209, https://doi.org/10.1007/978-1-0716-0935-4_13, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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The large spliceosomal Ski2-like RNA helicase, Brr2, acts during spliceosome activation, when it unwinds the extensively basepaired U4/U6 di-snRNA and displaces U4/U6-bound proteins [7, 10–13]. Brr2 has a distinct architecture (Fig. 1a, b), comprising two helicase units (cassettes) with identical domain arrangement, which together form the helicase region (HR) [14–16]. Only the N-terminal cassette (NC) is active in ATP hydrolysis, RNA binding, and RNA unwinding [12, 16]. The C-terminal cassette (CC) is a catalytically inactive pseudo-helicase but serves as an intramolecular regulator of the NC [16, 17]. Additionally, it acts as a landing pad for other spliceosomal proteins [18–21], which can modulate activities of Brr2 [22, 23]. In addition, Brr2 contains a long N-terminal region (NTR) that comprises two-folded domains connected by disordered regions [24, 25]. In isolated Brr2, the NTR can fold back onto the HR and act as another intramolecular regulator of Brr2, which inhibits Brr2 RNA binding, ATPase, and helicase activities by substrate exclusion and conformational clamping [24]. The Jab1 domain of the large spliceosomal Prp8 protein binds directly to the active NC [26, 27]. Depending on the presence (Jab1) or absence (Jab1ΔC) of a C-terminal tail, Jab1 acts as an inhibitor or activator of Brr2, respectively [26, 28]. To investigate the effect of intra- and intermolecular cofactors on Brr2’s activity, we have used, among others, polyacrylamide gel-based U4/U6 di-snRNA binding and unwinding assays. U4/U6 di-snRNA is most likely also a suitable model substrate for other processive RNA helicases, as it offers 30 - and 50 -single-stranded overhangs for initial helicase binding and as U6 snRNA forms an intramolecular stem loop after unwinding, which prevents U4/U6 reannealing [7], making it possible to perform single-round unwinding assays (Fig. 1c).

2

Materials Prepare all solutions using ultrapure water (18.2 MΩ cm at 25  C) and analytical grade reagents. Adjust the pH of the buffers at 4  C unless otherwise stated. Filter all buffers through a 0.22 μm filter. Use filter pipette tips when working with RNA. Diligently follow all safety and waste disposal regulations, when working with radioactive or otherwise harmful reagents.

2.1 General Equipment, Materials, and Software

1. Bench top centrifuge medium (for spin concentrators and 50 mL tubes). 2. Bench top centrifuge small (for 1.5 and 2 mL microfuge tubes). 3. Large high-speed centrifuge (for cell harvest and lysate clearance).

Functional Characterization of the Brr2 RNA Helicase Sec63

A Plug

PWI

RecA1

RecA2

WH

195

Sec63

HB HLH IG L RecA1

RecA2

WH

HB HLH IG

Brr2

2163

N-terminal cassette

N-terminal region (NTR)

C-terminal cassette

Helicase region (HR)

B

Jab1

Jab1

WHCC

HBNC N

RecA1CC

Plug

NC

NC HLH

PWI

C

WHCC

N HBNC

NC

NC

IG

RecA1CC C

L NC

RecA2 NC

WHNC

HB

CC

180°

IGCC HLH

IGCC CC

RecA2

CC

C U6

RecA1NC

RecA1NC

RecA2

HLH

CC

CC

Isolated U6

U4/U6 di-snRNA

5' Stem II

Brr2

Stem I

3'

5'

ISL

3'-overhang 3' 5'

3'

Brr2, U4

U4 5'-SL

Fig. 1 Brr2 structure and U4/U6 di-snRNA substrate. (a, b) Domain organization of Brr2 (a) and 3D structural overview of full-length yeast Brr2 bound to the Jab1 domain of Prp8 (b). NTR, N-terminal region; HR, helicase region; NC, N-terminal cassette; CC, C-terminal cassette; Sec63, Sec63-homology regions. Protein coloring by domains. Plug, purple; PWI-like (PWI), magenta; RecA1-like (RecA1NC/CC), light gray; RecA2-like (RecA2NC/CC), dark gray; winged helix (WHNC/CC), black; helical bundle (HBNC/CC), slate blue; helix-loop-helix (HLHNC/CC), red; immunoglobulin-like (IGNC/CC), lime green; inter-cassette linker (L), violet; Prp8 Jab1, gold. (c) Scheme illustrating base pairing in the U4/U6 di-snRNA substrate (left) and the presumed structure of isolated U6 snRNA (right) after unwinding. The blue Brr2 symbol and the arrow mark the presumed entry region and direction of translocation of Brr2 on U4 snRNA. 50 -SL 50 -stem loop of U4 snRNA, ISL internal stem loop of U6 snRNA

4. Ultrasonic cell disruption machine (e.g., Sonoplus Ultrasonic Homogenizer HD 3100 [Bandelin], VS 70 T probe). ¨ KTA pure [GE Healthcare]). 5. FPLC system (e.g., A 6. Microvolume UV-vis spectrophotometer (e.g., [NanoDrop]).

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7. Mini gel tank for SDS-PAGE (polyacrylamide gel electrophoresis) gels (e.g., [Invitrogen]). 8. Heating block. 9. Temperature-controlled vertical electrophoresis system (e.g., Owl [Thermo Fisher]). 10. Vortex mixer. 11. Magnetic stirrer. 12. Scintillation counter (e.g., PerkinElmer). 13. Phosphorimager with 32P phosphorimaging (phospho)screens and cassettes (e.g., GE Healthcare). 14. Vacuum polyacrylamide gel dryer. 15. 12-Well precast SDS gradient polyacrylamide gels (e.g., NuPAGE™ Bis-Tris gel 4–12% [Thermo Fisher]). 16. Protein loading buffer (e.g., NuPAGE™ LDS sample buffer [Thermo Fisher]). 17. Broad range protein molecular weight standard. 18. Polyacrylamide gel loading tips. 19. Protein staining solution (e.g., InstantBlue™ Protein stain [Expedeon]). 20. Polyacrylamide gel plates (ca. 20 cm  20 cm [W  L], one plate with a notch of ca. 2.5 cm for comb insertion), spacer (1 mm), and 22-teeth comb. 21. Whatman paper. 22. 10% (w/v) APS solution: Dissolve 1 g ammonium persulfate in 10 mL water. 23. N,N,N0 ,N0 -Tetramethylethylenediamine (TEMED). 24. 10 TBE solution: 1 M boric acid, 1 M TRIS–HCl, 25 mM EDTA. Weight 61.8 g of boric acid, 121.1 g of TRIS, and 7.44 g of EDTA and add water to a final volume of 1 L. 25. 1 PBS (phosphate-buffered saline), pH 7.4. 26. Image analysis software (e.g., ImageQuant [GE Healthcare]). 27. Statistical analysis software (e.g., Prism [GraphPad]). 2.2 Saccharomyces cerevisiae Brr2 Production and Purification

1. Synthetic DNA fragments encoding S. cerevisiae Brr2 variants (codon-optimized for expression in insect cells; full-length Brr2, residues 1–2163, Brr2FL (extinction coefficient at 280 nm: 251,770 M1 cm1); N-terminally truncated Brr2, residues 271–2163, Brr2271–2163 (extinction coefficient at 280 nm: 234,350 M1 cm1)) have been cloned into a modified pFL vector (EMBL, Grenoble) to generate bacmids for the production of Brr2 proteins with a TEV-cleavable, N-terminal His10-tag via recombinant Baculoviruses in insect cell culture [24].

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2. SF9 insect cells and SF9 insect cell medium. 3. Transfection reagent. 4. High-five insect cells and high-five insect cell medium supplemented with 1 L-glutamine. 5. Lysis buffer: 50 mM HEPES-NaOH, pH 7.5, 600 mM NaCl, 10 mM imidazole, 0.05% (v/v) NP40, 10% (v/v) glycerol, 2 mM dithiothreitol (DTT). Weight 11.9 g HEPES, 35 g NaCl, and 0.68 g imidazole and add 5 mL of a 10% (v/v) NP-40 solution and 116 mL of a 86% (v/v) glycerol solution. Add cold water to ca. 900 mL and stir until everything is dissolved. Place buffer in an ice bucket and adjust the pH with 1 M NaOH to a pH of 7.5. Add cold water to 1 L. Filter buffer and store at 4  C. Add 2 mL of a filtered 1 M DTT stock immediately before use (1 M DTT stock stored at 80  C in 1 mL aliquots). For TRIS–HCl buffers (employed below), use 37% (w/w) HCl to adjust pH. 6. Ni2+-NTA buffer A: 50 mM HEPES-NaOH, pH 7.5, 600 mM NaCl, 10 mM imidazole, 10% (v/v) glycerol, and 2 mM DTT. Weight 11.9 g HEPES, 35 g NaCl, and 0.68 g imidazole and add 116 mL of a 86% (v/v) glycerol solution. Prepare 1 L as described for item 5. 7. Ni2+-NTA buffer B: 50 mM HEPES-NaOH, pH 7.5, 600 mM NaCl, 250 mM imidazole, 10% (v/v) glycerol, 2 mM DTT. Weight 6 g HEPES, 17.5 g NaCl, and 8.5 g imidazole and add 58 mL of a 86% (v/v) glycerol solution. Prepare 0.5 L as described for item 5. 8. Dilution buffer: 40 mM HEPES-NaOH, pH 7.5. Weight 4.8 g HEPES and prepare 0.5 L as described for item 5. 9. Heparin buffer A: 40 mM HEPES-NaOH, pH 7.5, 50 mM NaCl, 5% (v/v) glycerol, 2 mM DTT. Weight 9.5 g HEPES and 2.9 g NaCl and add 58 mL of a 86% (v/v) glycerol solution. Prepare 1 L as described for item 5. 10. Heparin buffer B: 40 mM HEPES-NaOH, pH 7.5, 1.5 M NaCl, 5% (v/v) glycerol, 2 mM DTT. Weight 4.8 g HEPES and 43.8 g NaCl and add 29 mL of a 86% (v/v) glycerol solution. Prepare 0.5 L as described for item 5. 11. Gel-filtration buffer: 20 mM TRIS–HCl, pH 7.5, 200 mM NaCl, 20% (v/v) glycerol, 2 mM DTT. Weight 2.4 g TRIS and 11.7 g NaCl and add 233 mL of a 86% (v/v) glycerol solution. Prepare 1 L as described for item 5. 12. 0.8 μm syringe filters. 13. 5 mL His Trap FF column [GE Healthcare]. 14. 5 mL HiTrap heparin HP column [GE Healthcare]. 15. S200 16/60 column [GE Healthcare].

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16. EDTA-free protease inhibitors. 17. Centrifugal filters, 100 kDa molecular mass cutoff. 18. 2 mg/mL RNase A solution. Weight 20 mg RNase A and dissolve in 10 mL water. 19. 2 mg/mL DNase solution. Weight 20 mg DNase and dissolve in 10 mL water. 2.3 S. cerevisiae Jab1 Production and Purification

1. Synthetic DNA fragments encoding S. cerevisiae Prp8 Jab1 domain variants (codon-optimized for expression in insect cells; “full-length” Jab1, residues 2147–2413, Jab1 (extinction coefficient at 280 nm: 36,900 M1 cm1); C-terminally truncated Jab1, residues 2147–2398, Jab1ΔC (extinction coefficient at 280 nm: 36,900 M1 cm1)) have been cloned into the pETM-11 vector (EMBL, Heidelberg) for the production of the target proteins with a TEV-cleavable, N-terminal His6tag in Escherichia coli culture [24, 28]. 2. E. coli Rosetta 2 (DE3). 3. 50 mg/mL kanamycin in water. Sterilize by filtration with a 0.2 μm filter and store at 20  C. 4. LB growth medium. 5. LB-agar plates containing 50 μg/mL kanamycin. 6. Auto-inducing growth medium (e.g., MagicMedia™ Medium from Thermo Fisher Scientific) [29]. 7. Ni2+-NTA buffer A: 100 mM TRIS–HCl, pH 7.5, 200 mM NaCl, 10 mM imidazole, 2 mM DTT. Weight 12.1 g of TRIS, 11.7 g of NaCl, and 0.68 g of imidazole. Prepare 1 L as in Subheading 2.2, item 5. 8. Ni2+-NTA buffer B: 50 mM TRIS–HCl, pH 7.5, 200 mM NaCl, 500 mM imidazole, 2 mM DTT. Weight 3 g TRIS, 5.8 g NaCl, and 17 g imidazole. Prepare 0.5 L as in Subheading 2.2, item 5. 9. Dialysis buffer: 50 mM TRIS–HCl, pH 7.5, 200 mM NaCl, 10 mM imidazole, 2 mM DTT. Weight 6 g TRIS, 23.3 g NaCl, and 1.4 g imidazole. Prepare 2 L as in Subheading 2.2, item 5. 10. Gel-filtration buffer: 10 mM TRIS–HCl, pH 7.5, 200 mM NaCl, 2 mM DTT. Weight 0.7 g TRIS, 11.7 g NaCl. Prepare 1 L as in Subheading 2.2, item 5. 11. 5 mL His Trap FF column [GE Healthcare]. 12. S75 26/60 column [GE Healthcare]. 13. EDTA-free protease inhibitors. 14. Dialysis tubing, 10 kDa molecular mass cutoff. 15. Centrifugal filters, 10 kDa molecular mass cutoff. 16. Tobacco etch virus (TEV) protease. 17. 2 mg/mL DNase solution.

Functional Characterization of the Brr2 RNA Helicase

2.4 S. cerevisiae U4/U6 Di-snRNA Preparation ([P32]-Labeled)

199

1. 10 T7 buffer: 1.2 M HEPES-NaOH, pH 7.5, 160 mM MgCl2, 29 mM spermidine, 400 mM DTT. Weight 28.6 g HEPES, 1.5 g MgCl2, 0.4 g spermidine, and 6.2 g DTT. Prepare 100 mL as in Subheading 2.2, item 5. 2. MonoQ buffer A: 20 mM HEPES-NaOH, pH 7.5, 50 mM NaCl. Weight 2.4 g HEPES and 1.5 g NaCl. Prepare 0.5 L as in Subheading 2.2, item 5. 3. MonoQ buffer B: 20 mM HEPES-NaOH, pH 7.5, 1 M NaCl. Weight 2.4 g HEPES and 29.2 g NaCl. Prepare 0.5 L as in Subheading 2.2, item 5. 4. TNES buffer: 20 mM TRIS–HCl, pH 8.0, 300 mM NaCl, 5 mM EDTA, 0.1% (w/v) SDS. Mix 1 mL of a 1 M TRIS–HCl stock with 15 mL of a 1 M NaCl stock, 250 μL of a 1 M EDTA stock, and 500 μL of a 10% (w/v) SDS solution. Prepare 50 mL as in Subheading 2.2, item 5. 5. Annealing buffer: 40 mM TRIS–HCl, pH 7.5, 100 mM NaCl. Mix 2 mL of a 1 M TRIS–HCl stock with 5 mL of a 1 M NaCl stock. Prepare 50 mL as in Subheading 2.2, item 5. 6. Native-PAGE loading buffer: 1 TBE, 30% (v/v) glycerol, 0.033% (w/v) bromophenol blue, 0.033% (w/v) xylene cyanol. Mix 5 mL of a 10 TBE stock with 17.4 mL of a 86% (v/v) glycerol solution with 15 mg of bromophenol blue and xylene cyanol. Prepare 50 mL as in Subheading 2.2, item 5. 7. Duplex preparation polyacrylamide gel buffer: 6% (w/v) acrylamide/bisacrylamide (19:1), 0.5 TBE. Mix 50 mL of 10 TBE and 150 mL acrylamide/bisacrylamide (40% [w/v], 19:1) and add water to a final volume of 1 L. Right before pouring polyacrylamide gel, add 150 μL of a 10% (w/v) APS solution and 8 μL TEMED to 12 mL duplex preparation polyacrylamide gel buffer for one polyacrylamide mini-gel with a preparative comb. 8. 0.6 μg/μL pUC18/pT7U6 plasmid [30] linearized with BamHI. 9. 0.6 μg/μL pUC18/pT7U4 plasmid [10] linearized with StyI. 10. 10 μg/μL T7 RNA polymerase (we use a P266L variant, prepared in the lab, but commercial preparations of wild-type T7 RNA polymerase should do). 11. 0.1 U/μL pyrophosphatase. 12. 1 M MgCl2. 13. NTP mix (100 mM each NTP). 14. 3 M sodium acetate. 15. Isopropanol. 16. 95% and 70% (v/v) ethanol in water.

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17. RNase-free water. 18. 1 U/μL alkaline phosphatase. 19. 10 U/μL T4 polynucleotide kinase (PNK). 20. 3000 Ci/mmol 10 mCi/mL [γ32P]-ATP. 21. G-25 spin columns (e.g., Microspin™ G-25 [GE Healthcare]). 22. Phenol/chloroform/isoamyl alcohol (PCI) [25:24:1] mix, pH 4.5–5. 23. 20 mg/mL glycogen (RNA grade). 24. 6 M LiCl. 25. MonoQ 5/50 GL1 mL column [GE Healthcare]. 26. Mini polyacrylamide gel plates (short plate and spacer plate (1 mm)) and preparative comb (e.g. Mini-PROTEAN® Tetra Handcast Systems [Bio-Rad]). 27. Mini polyacrylamide gel-casting stands (e.g., Mini-PROTEAN® Tetra Handcast Systems [Bio-Rad]). 28. Mini polyacrylamide gel buffer tank and electrode assembly (e.g., Mini-PROTEAN® Tetra Handcast Systems [Bio-Rad]). 2.5 Electrophoretic Mobility Shift Assay (EMSA)

1. 5 binding buffer: 200 mM HEPES-NaOH, pH 7.9 at room temperature (RT), 75 mM NaCl, 12.5 mM Mg(OAc)2. Mix 10 mL of a 1 M HEPES-NaOH stock, 3.75 mL of a 1 M NaCl stock, and 0.625 mL of a 1 M Mg(OAc)2 stock. Prepare 50 mL as in Subheading 2.2, item 5. 2. EMSA polyacrylamide gel buffer: 4% (w/v) acrylamide/ bisacrylamide (75:1), 0.5 TBE native PAGE. Mix 131.6 mL of gel A (30% [w/v] acrylamide) and 26.6 mL Gel B (2% [w/v] bisacrylamide) solutions with 50 mL of a 10 TBE stock and add water to 1 L. Right before pouring polyacrylamide gel, add 625 μL of 10% (w/v) APS and 31 μL of TEMED to 50 mL of EMSA polyacrylamide gel buffer mix for one gel. 3. Native-PAGE loading buffer, see Subheading 2.4, item 6. 4. 1 μg/MlμL acetylated BSA.

2.6 Polyacrylamide Gel-Based Unwinding Assays

1. 10 unwinding buffer: 400 mM TRIS–HCl, pH 7.5 at RT, 500 mM NaCl, 5 mM MgCl2. Weight 2.4 g TRIS and 1.46 g NaCl and add 250 μL of a 1 M MgCl2 stock solution. Prepare 50 mL as in Subheading 2.2, item 5. 2. Stop buffer: 40 mM TRIS–HCl, pH 7.4 at RT, 50 mM NaCl, 25 mM EDTA, 1% (w/v) SDS, 10% (v/v) glycerol, 0.05% (w/v) xylene cyanol, 0.05% (w/v) bromophenol blue. Mix 2 mL of a 1 M TRIS–HCl stock, 2.5 mL of a 1 M NaCl stock, 1.25 mL of a 1 M EDTA stock, 0.5 g SDS, 5.8 mL of a 86% (v/v) glycerol stock, 10 mg xylene cyanol, and 10 mg bromophenol blue. Prepare 50 mL as in Subheading 2.2, item 5.

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3. Unwinding polyacrylamide gel buffer: 6% (w/v) acrylamide/bisacrylamide (19:1), 0.5 TBE. Mix 50 mL of 10 TBE and 150 mL acrylamide/bisacrylamide (40% [w/v], 19:1) and add water to a final volume of 1 L. Right before pouring polyacrylamide gel, add 625 μL of a 10% (w/v) APS solution and 31 μL TEMED to 50 mL unwinding polyacrylamide gel buffer for one gel. 4. 100 mM ATP/MgCl2. 5. 100 mM DTT.

3

Methods All purification steps are performed at 4  C. Proteins and RNAs are always kept on ice during setup of experiments. To avoid RNase contamination, always use gloves and a lab coat. For setup of experiments containing RNA, use RNase-free barrier tips.

3.1 S. cerevisiae Brr2 Production and Purification

1. For initial virus (V0) production, transfect adhesive Sf9 cells in six-well plates with the isolated recombinant baculoviral DNA and incubate for 60 h at 27  C. For a detailed transfection protocol, please refer to [31]. 2. Collect the supernatant and infect a 25 mL Sf9 cell suspension culture, which has been cultivated in the same flask for 2 days at a density of 0.5  106 cells/mL for virus amplification (V1). Measure cell density every 24 h and add fresh medium to keep the cell density below 1  106 cells/mL if needed. Harvest the cells 60 h after cell proliferation arrest. Spin the suspension culture for 10 min at 800  g at room temperature in a 50 mL tube and collect the supernatant containing V1 (see Note 1). Store V1 at 4  C until further usage. 3. For large-scale protein production, keep 400 mL per flask of high-five cells in suspension at a cell density of 1  106 cells/ mL and infect the cells with approximately 1 mL of V1 virus (see Note 2). For one purification run, use about 1.2 L of expression culture (see Note 3). 4. Harvest cells after approximately 72 h postinfection (viability ca. 92%) by centrifugation at 800  g. 5. Resuspend pellet in 40 mL 1 PBS, spin the suspension for 10 min at 800  g at room temperature, discard supernatant, flash-freeze the pellet in liquid N2, and store at 20  C until further usage. 6. Resuspend pellet in 150 mL cold lysis buffer (see Note 4) and add 3 mg DNase and protease inhibitor cocktail (see Note 5).

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7. Lyse cells by sonication (e.g., Sonoplus Ultrasonic Homogenizer HD 3100 [Bandelin], VS 70T probe; 40% intensity, 1 s on, 2 s off, 25 min, two repetitions, let cool down for 10 min in between repetitions; these values are optimized for the stated sonication device and may vary for another model). 8. Clear cell extract by centrifugation at 50,000  g for 45 min, 4  C. 9. Filter supernatant through 0.8 μm syringe filters (see Note 6). 10. Capture the target on a Ni-NTA column, pre-equilibrated with Ni-NTA buffer A (flow rate 2 mL/min). 11. Wash the column with 75 mL Ni-NTA buffer A (flow rate 2 mL/min). 12. Elute the target with a linear gradient (60 mL) from 10 (Ni-NTA buffer A) to 250 mM imidazole (Ni-NTA buffer B) (0–100% buffer B) and collect in 2 mL fractions (flow rate 1 mL/min). 13. Load 5 μL of each fraction, supplemented with 4 loading dye, together with the molecular weight standard on a SDS-PAGE gradient gel and run the gel following the supplier’s recommendations. After the run, stain gel with Coomassie, following the supplier’s recommendations, and evaluate fractions to pool. 14. Concentrate the pooled fractions in a spin concentrator (100 kDa molecular mass cutoff) to a volume of 1–2 mL according to the supplier’s recommendations. Perform 10 min centrifugation steps and mix the sample in between, to avoid a too high concentration in the bottom of the concentrator. 15. Dilute the sample with dilution buffer to a final salt concentration of 80 mM (see Note 7). 16. To remove contaminating RNA, treat the sample with 15 μL 2 mg/mL RNase A for 20 min at room temperature (see Note 8). 17. Load the sample on a heparin column pre-equilibrated with heparin buffer A (flow rate 2 mL/min). 18. Wash the column with 50 mL of heparin buffer A (flow rate 2 mL/min). 19. Elute the sample with a linear gradient (60 mL) from 50 mM NaCl (heparin buffer A) to 1.5 M NaCl (heparin buffer B) (0–100% buffer B) and collect 1 mL fractions (see Notes 9 and 10) (flow rate 1 mL/min). 20. Successful RNA removal is evaluated by measuring the 260/280 nm ratio of the peak fractions in a UV-vis spectrophotometer, which should be below 0.7. Fractions are additionally evaluated by a SDS-PAGE as in step 3 1/13.

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21. Concentrate peak fractions to a volume of 2 mL as in step 14, load on a gel-filtration column pre-equilibrated with gel-filtration buffer, and collect 2 mL fractions (flow rate 1 mL/min) (see Note 11). Peak fractions are evaluated by SDS-PAGE as mentioned in step 13. 22. Pool fractions of interest and concentrate to 10–30 mg/mL as in step 3 1/14. Determine the concentration from absorbance at 280 nm using a UV–vis spectrophotometer (see Subheading 2.2 for extinction coefficients). Aliquot (20 μL) the sample and flash-freeze in liquid N2 and store at 80  C until further usage (see Notes 12 and 13). 3.2 S. cerevisiae Jab1 Production and Purification

1. Transform E. coli Rosetta2 (DE3) cells with the vector of interest, plate on kanamycin-containing plates, and incubate overnight at 37  C. 2. Use a single colony to inoculate 100 mL of LB medium containing 50 μg/mL kanamycin and incubate overnight in a shaker at 200 rpm and 37  C. 3. Inoculate 3 L of auto-inducing medium [29] containing 50 μg/mL kanamycin with 10 mL of the overnight culture and grow for 72 h at 200 rpm and 18  C (see Note 14). 4. Harvest cells by centrifugation at 9000  g, flash-freeze the pellet in liquid N2, and store at 20  C until further usage. 5. Resuspend the pellet in 150–200 mL cold Ni-NTA buffer A. Add 3 mg DNase and protease inhibitor cocktail. 6. Lyse cells by sonication (70% intensity, 1 s on, 2 s off, 25 min, two repetitions, let cool down for 10 min in between repetitions; these values are optimized for the stated sonication device and may vary for another model). 7. Clear cell extract by centrifugation at 50,000  g for 45 min, 4  C. 8. Capture the target on a Ni-NTA column, pre-equilibrated with Ni-NTA buffer A (flow rate 2 mL/min). 9. Wash with 50 mL Ni-NTA buffer A (flow rate 2 mL/min). 10. Elute the target via a linear gradient (60 mL) from 10 mM (Ni-NTA buffer A) to 250 mM imidazole (0–50% buffer B) and collect 2 mL fractions (flow rate 1 mL/min). 11. Evaluate peak fractions via SDS-PAGE (see Subheading 3.1, step 13) and pool the fractions of interest. 12. Determine the concentration by absorbance at 280 nm using a UV–vis spectrophotometer and calculate the protein amount (extinction coefficient provided in Subheading 2.3). Supplement pooled fractions with 1:20 (w/w with respect to protein content) TEV protease and dialyze overnight in 2 L dialysis buffer at 4  C with constant stirring.

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13. Separate cleaved protein from uncleaved protein, His-tag and His-tagged TEV protease via recycling over a Ni-NTA column. Use the same buffers and purification strategy as for the initial Ni-NTA purification step. The protein will be in the flowthrough, which is collected in 10 mL fractions (flow rate 2 mL/min). 14. Evaluate flow-through factions via SDS-PAGE (see Subheading 3.1, step 13). 15. Flow-through fractions, containing the target, are pooled, concentrated to 5 mL as in Subheading 3.1, step 14 (spin filter with 10 kDa molecular mass cutoff) and loaded on a gel-filtration column, pre-equilibrated with gel-filtration buffer (fraction size 3 mL, flow rate 2 mL/min). Peak fractions are evaluated by SDS-PAGE as mentioned in Subheading 3.1, step 13. 16. Concentrate the target protein to 30 mg/mL as in Subheading 3.1, step 13 (spin filter with 10 kDa molecular mass cutoff). Determine the concentration by absorbance at 280 nm using a UV–vis spectrophotometer. Aliquot (20 μL) the sample and flash-freeze in liquid N2 and store at 80  C until further usage (see Note 12). 3.3 S. cerevisiae U4/U6 Di-snRNA Preparation (U4 [P32] Labeled)

1. Set up in vitro transcription reactions for U4 and U6 by mixing the following components:

Reagent

Volume [mL] (final concentration)

Linearized pUC18/pT7U4 or pUC18/ pT7U6 plasmid

1 (60 ng/μL)

T7 polymerase

0.1 (0.1 μg/μL)

T7 buffer (10)

1 (1)

NTP mix (100 mM each NTP)

0.4 (4 mM each NTP)

MgCl2 (1 M)

0.16 (16 mM)

Pyrophosphatase

0.1 (0.001 U/μL)

H2O

7.24

Total

10

2. Incubate each reaction mixture at 37  C for 12 h (see Note 15). 3. Add 1 vol of PCI solution to reaction, vortex for 1 min, and centrifuge at 20,000  g for 15 min.

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4. Carefully remove the upper, aqueous part and transfer into a new tube. 5. Add 0.1 vol of 3 M sodium acetate. 6. Add 1 vol of isopropanol and vortex briefly. 7. Centrifuge sample for 1 h at 20,000  g and 4  C. 8. Remove isopropanol and wash pellets with 95% (v/v) ethanol. 9. Centrifuge sample for 15 min at 20,000  g and 4  C and remove ethanol. 10. Dry pellet for approximately 15 min and resuspend in 5 mL water (see Note 16). 11. Load RNA on an anion exchange column (e.g., MonoQ 5/50 column) pre-equilibrated in MonoQ buffer A. Elute the RNA with a linear gradient (20 mL) from 50 mM (MonoQ buffer A) to 1 M NaCl (MonoQ buffer B) (0–100% buffer B) (flow rate 0.5 mL/min). 12. Pool the peak fractions, determine the concentration from absorbance at 260 nm with a UV–vis spectrophotometer (extinction coefficient at 260 nm: 0.025 (μg/mL)1 cm1), aliquot (20 μL) and freeze the RNA in liquid N2, and store at 80  C until further usage. 13. Set up dephosphorylation of U4 snRNA by mixing the following reagents: Reagent

Volume [μL] (final concentration)

U4 RNA

Var.a (200 ng/μL; 3.7 μM)

Alkaline phosphatase buffer (10)

30 (1)

Alkaline phosphatase (1 U/μL)

15 (0.05 U/μL)

RNase-free water

Ad 300

Total

300

a

Variable, depending on concentration from preparation or stock concentration

14. Incubate at 37  C for 1 h (see Note 17). 15. Perform PCI extraction as in steps 3–10 and dissolve pellet in 15 μL water. Determine RNA concentration from absorbance at 260 nm (concentration should now be of approximately 4.3 μg/μL; 80 μM). 16. Aliquot the RNA, flash-freeze in liquid N2, and store at 80  C until further usage. 17. Set up U4 snRNA labeling reaction by mixing the following reagents:

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Reagent

Volume [μL] (final concentration)

PNK buffer (10)

4 (1)

T4 PNK (10 U/μL)

3 (0.75 U/μL)

RNase inhibitor (40 U/μL)

1 (1 U/μL)

Dephosphorylated U4

Var.a (70 ng/μL; 1.3 μM)

[γ32P]-ATP (10 mCi/mL)

25 (6 mCi/mL)

RNase-free water

Ad 40

Total

40

a

Variable, depending on concentration from preparation or stock concentration

18. Incubate mixture at 37  C for 2 h. 19. Add 10 μL RNase-free water and load on a spin column pre-equilibrated with water according to the supplier’s recommendation. 20. Elute sample (volume should be 50 μL, concentration 1 μM) and measure the radioactivity of 1 μL sample in a scintillation counter (see Note 18). 21. Add 5 μL U6 snRNA (100 μM) and annealing buffer to 200 μL. 22. Heat to 80  C for 5 min. 23. Switch off heating block and let cool down to 70  C. 24. Add 2.5 μL of 1 M MgCl2 and allow to cool down sample to 30  C (approximately 1.5–2 h). 25. During annealing, prepare the 6% native-PAGE polyacrylamide gel. Add APS and TEMED to the duplex preparation polyacrylamide gel buffer and pour a mini polyacrylamide gel with a preparative comb following the supplier’s recommendations. Pre-equilibrate the polyacrylamide gel in 0.5 TBE at 200 V. Use the electrophoresis system according to the supplier’s recommendation. 26. Spin down sample before opening lid. 27. Add 100 μL native-PAGE loading buffer and load sample on the polyacrylamide gel. 28. Run the polyacrylamide gel at 200 V for 1 h until the bromophenol blue dye runs out of the polyacrylamide gel. 29. Wrap the polyacrylamide gel with saran wrap, put in a cassette with the spacer plate on the bottom and the short plate on top (otherwise the radioactivity is shielded too much by the thick spacer plate), and expose a phosphoscreen for 10 min.

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30. Image screen on a phosphorimager and cut out the corresponding (upper) band on the polyacrylamide gel with a scalpel. 31. Cut the polyacrylamide gel piece in very small pieces and transfer into five 2 mL tubes. 32. Add 1 mL TNES buffer to each tube. Rotate tubes overnight at RT. 33. Spin tubes at 20,000  g for 10 min to pellet polyacrylamide gel pieces. 34. Transfer supernatant to new 1.5 mL tubes, 500 μL per tube (see Note 19). 35. Add 1 vol PCI mix to each tube and vortex for 10 s. 36. Spin down sample at 20,000  g for 10 min. 37. Take the aqueous, upper phase and put it into a new 1.5 mL tube, which contains 50 μL of 6 M LiCl and 2 μL 20 mg/mL glycogen (to visualize pellet, see Note 20). 38. Add 700 μL isopropanol and vortex briefly. 39. Flash-freeze the sample in liquid N2. 40. Centrifuge sample at 20,000  g for 1 h at RT (see Note 21). 41. Remove isopropanol and wash pellet with 300 μL 70% (v/v) ethanol by inverting the tubes. 42. Centrifuge for 5 min at 20,000  g. 43. Remove ethanol and let dry pellet in a hood for 15 min. 44. Add 15 μL annealing buffer per tube and let RNA dissolve for 10 min. 45. Pool the RNA and measure the radioactivity of a 1 μL aliquot in a scintillation counter. Knowing the starting and the end radioactivity of the sample allows to estimate the final concentration (assuming no RNA loss and no residual [γ32P] ATP during the initial labeling process, i.e., steps 17–20). 46. Aliquot sample (10 μL), flash-freeze in liquid N2, and store at 20  C until further usage. 3.4 Electrophoretic Mobility Shift Assay (EMSA) (See Note 22)

1. Pre-cool the vertical electrophoresis system to 4  C. 2. Assemble the glass plate with the notch and the large glass plate with spacers on either side and fixate them with clamps (see Note 23). 3. Cast the EMSA polyacrylamide gel by mixing the EMSA polyacrylamide gel buffer with APS and TEMED and pour it without air bubbles between the large glass plate and the short plate from top to bottom.

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4. Insert the comb and store the polyacrylamide gel horizontally until it is completely polymerized. 5. Once the polyacrylamide gel is polymerized, carefully remove the comb and wash the wells with a syringe and 0.5 TBE. 6. Pre-run polyacrylamide gel for 30 min (100 V) in 0.5 TBE in the vertical electrophoresis system. 7. Prepare the binding reaction by mixing the following reagents (see Note 24): Reagent

Volume [μL] (final concentration)

5 binding buffer

3 (1)

RNase inhibitor (40 U/μL)

0.5 (1.3 U/μL)

U4*/U6 di-snRNA

Var.a (1 nM)

Brr2

Var. (titrate desired Brr2 concentrations)

Jab1

Var. (500 nM)

Acetylated BSA (1 μg/μL)

1.5 (0.1 μg/μL)

NaCl stock

Var. (adjust NaCl concentration in each sample, accounting for increasing NaCl brought into the binding reaction with increasing amounts of Brr2)

RNase-free water

Ad 15

Total

15

a

Variable; depending on concentration from preparation or stock concentration

8. Incubate samples for 5 min at RT. 9. Load samples with polyacrylamide gel-loading tips at the very bottom of the polyacrylamide gel wells (see Note 25). 10. Run polyacrylamide gel for 1.5 h at 100 V. 11. Open the polyacrylamide gel plates carefully and put a Whatman paper on top of the gel. The polyacrylamide gel should stick to the paper. 12. Cover with saran wrap and dry in a gel dryer for at least 1 h. 13. Expose a phosphorimager screen overnight. 14. Scan the polyacrylamide gel on a phosphorimager and quantify the band intensities of free U4/U6 and Brr2-bound U4/U6 (migrates higher in the gel) and perform a background subtraction for each band using an image analysis software.

Functional Characterization of the Brr2 RNA Helicase

Brr2271-2163U4*/U6

Unbound Bound

A

209

U4*/U6 0

5

10 15 20 25 30 40 50 60

Brr2

271-2163

concentration [nM]

B Fraction U4*/U6 bound

1.0 0.8 0.6 0.4 Kd = 16.7 nM ± 0.5 nM

0.2 0.0 0

10

Brr2

20 271-2163

30

40

50

60

concentration [nM]

Fig. 2 Analysis of U4/U6 di-snRNA binding by Brr2. (a) Typical EMSA gel showing increasing fractions of U4/U6 di-snRNA shifted by increasing concentrations of isolated, N-terminally truncated Brr2 (Brr2271–2163). Lines labeled “[bound” and “unbound” indicate approximate regions of lanes quantified for assessing Brr2271–2163-bound and unbound RNA in a lane, respectively. (b) Quantification and fitting of data as shown in (a). Data points represent means  SEM of six independent experiments using the same biological samples. The binding isotherm, yielding the Kd, was fit according to the equation: fraction bound ¼ AcBrr2n/(cBrr2n + Kdn), in which A is the maximum of bound RNA, cBrr2 is the Brr2271–2163 concentration, Kd is the dissociation constant, and n is the Hill coefficient. According to data reported in [24], Fig. 5a.

15. Calculate the ratio of free U4/U6 and Brr2-bound U4/U6 for each Brr2 concentration and calculate affinity by fitting data to a Hill equation; fraction bound ¼ AcBrr2n/ (cBrr2n + Kdn), in which A is the maximum of bound RNA, cBrr2 is the Brr2 concentration, Kd is the dissociation constant, and n is the Hill coefficient; Fig. 2) using a statistical analysis software. 3.5 Gel-Based Unwinding Assays (See Note 22)

1. Prepare the unwinding gel the same way as described for the EMSA polyacrylamide gel (Subheading 3.4, steps 1–5), using the unwinding polyacrylamide gel buffer. It is not necessary to pre-run the unwinding polyacrylamide gel. 2. Prepare the unwinding reaction by mixing the following reagents (see Note 26):

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Reagent

Volume (μL) (final concentration)

10 unwinding buffer

8 (1)

50% (v/v) glycerol

12.8 (8% [v/v])

RNase inhibitor (40 U/μL)

1 (0.5 U/μL)

U4*/U6 di-snRNA

Var.a (2 nM)

Brr2

Var. (100 nM)

Jab1

Var. (500 nM)

DTT (100 mM)

1.2 (1.5 mM)

Acetylated BSA (1 μg/μL)

8 (100 ng/μL)

RNase-free water

Ad 80

Total

80

a

Variable, depending on concentration from preparation or stock concentration

3. Carefully flick the tubes. Spin 2 min at 700  g. 4. Incubate 3 min at 30  C (protein binding). 5. Withdraw a 10 μL aliquot (zero time point), transfer it to a new tube, and add 10 μL stop buffer. Keep aliquot on ice. 6. Add 0.7 μL ATP/MgCl2 (1 mM final) to the reaction mixture and mix by stirring with pipette tip. 7. Withdraw 10 μL reaction aliquots at desired time points. Add 10 μL stop buffer to each aliquot and keep on ice. 8. Mix the final 10 μL of the reaction (see Note 26) with 10 μL stop buffer and boil it for 5 min at 80  C for a 100% unwound control. 9. Load 18 μL of each sample in separate wells of the unwinding gel. 10. Run the gel for 1 h at 200 V and 4  C. 11. Transfer, dry, and expose polyacrylamide gel and quantify bands as in Subheading 3.4, steps 11–13. 12. Scan the polyacrylamide gel on a phosphorimager and quantify the band intensities of free U4 and U4/U6 (migrates higher in the gel) and perform a background subtraction for each band using an image analysis software. 13. Calculate the ratio of free U4 and U4/U6 for each time point and fit the data to a first-order reaction; fraction unwound ¼ A (1 – exp.[kut]), in which A is the amplitude of the reaction, ku is the apparent first-order rate constant of unwinding, and t is time (Fig. 3) using a statistical analysis software.

Functional Characterization of the Brr2 RNA Helicase

211

Boil Brr2271-263

U4*/U6

Brr2 Jab1ΔC ss ds

A

U4*/U6

271-263

U4*

U4*

B

U4/U6 snRNA unwound [%]

Time[sec] 0 15 30 60 90 120 150 180 240 300

100 80 60 40 271-2163

Brr2 Brr2271-2163-Jab1ΔC

20 0 0

60

180 120 Time [s]

Protein Brr2 Brr2

A (%)

271-2163

271-2163

-Jab1

240

300

ku [s1]

100.0 ± 4.0 0.721 ± 0.072 ΔC

98.7 ± 1.3 2.304 ± 0.150

Fig. 3 Analysis of Brr2- and Brr2-Jab1ΔC-mediated U4/U6 di-snRNA unwinding. (a) Typical unwinding gels showing increasing fractions of U4/U6 di-snRNA unwound at increasing time points by isolated N-terminally truncated Brr2 (Brr2271–2163; top) or N-terminally truncated Brr2 in complex with C-terminally truncated Jab1 (Brr2271–2163-Jab1ΔC; bottom). Boil—100% unwound control (boiled RNA sample). Lines labeled “ds” and “ss” indicate approximate regions of lanes quantified for assessing the amount of double-stranded U4*/U6 and the amount of single-stranded U4*, respectively, released by Brr2271–2163 or Brr2271–2163-Jab1ΔC. (b) Quantification of data as shown in (a). Data points represent means  SEM of three independent experiments using the same biological samples. The data were fit to first-order reactions according to the equation: fraction unwound ¼ A(1 – exp[kut]), in which A is the amplitude of the reaction, ku is the apparent first-order rate constant of unwinding, and t is time. According to data reported in [7], Fig. S1

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Notes 1. You can use the pellet of the V1 production for a small-scale pull-down experiment to evaluate protein production. 2. The amount of added V1 virus depends on the V1 virus titer and has to be evaluated for each virus anew in a titration experiment. Adjust the virus concentration in such a way that the cells double one more time. 3. In general, full-length Brr2 is produced at lower levels compared to truncated Brr2. 4. Freeze pellets in 50 mL tubes. Add some lysis buffer and let the pellet thaw. Put it into a beaker and add an additional 150–200 mL of lysis buffer on a magnetic stirring device. 5. Use EDTA-free protease inhibitor mix to avoid stripping of Ni2 + from the Ni-NTA column. 6. Filtering the supernatant through a 0.8 μm filter helps to avoid clogging of the column. 7. It is necessary to have a final salt concentration lower than 100 mM to ensure Brr2 binding to the column. 8. RNase is added to the low salt protein dilution, as it works more efficiently in low salt. During the RNase treatment of Brr2, the protein solution will precipitate somewhat, which indicates that the RNA digestion worked. 9. The heparin column removes residual nucleic acids and separates partially degraded Brr2. 10. Brr2 constructs elute at around 30% heparin buffer B, depending on the target construct (e.g., containing or lacking different parts of the NTR). 11. Purifying and freezing Brr2 in buffer containing glycerol are important to preserve activity. 12. Check the purified protein samples for RNase contamination, either using a commercially available kit or by incubating the samples with some test RNA for 30 min at 37  C and inspecting by urea PAGE. 13. Brr2 loses its activity when stored on ice for longer than 1 day. Thus, use a new aliquot for every day of experiments. 14. Growing the cells at 37  C to an OD600 of 0.6 followed by overnight incubation at 18  C gives similar results. 15. Usually U4 snRNA gives (approximately twofold) higher yields than U6 snRNA. 16. If the suspension is too viscous, increase the resuspension volume.

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17. This reaction setup should yield enough material for approximately 15 labeling reactions. 18. This step is important, as this value is used to calculate the concentration of the resulting duplex. 19. Press pipette tip to the bottom of the tube, to avoid sucking in gel pieces. 20. Put glycogen drop on the wall of the tube to avoid premature mixing with the LiCl solution. 21. Place each tube in the same orientation in the centrifuge, so it is easier to find the pellet. 22. Test intrinsic RNA-binding activity of any cofactor prior to EMSA and unwinding assays with Brr2. 23. Silanization of the gel plates prior to use makes it easier to pour the polyacrylamide gels without bubbles. Covering the lower and side corners of the gel plates with sticky tape helps in casting the polyacrylamide gels. Pre-running the polyacrylamide gel is crucial for the EMSA experiments, as it will lead to sharper bands. 24. RNase inhibitor is not necessary when proteins have been tested negative for RNase contamination. 25. It is crucial to load the samples as concentrated as possible to the bottom of the polyacrylamide gel pockets to obtain sharp bands. 26. For each time point and control, 10 μL are withdrawn. Adjust the reaction volume according to the required number of time points to monitor. References 1. Jankowsky E (2011) RNA helicases at work: binding and rearranging. Trends Biochem Sci 36(1):19–29. https://doi.org/10.1016/j. tibs.2010.07.008 2. Halls C, Mohr S, Del Campo M, Yang Q, Jankowsky E, Lambowitz AM (2007) Involvement of DEAD-box proteins in group I and group II intron splicing. Biochemical characterization of Mss116p, ATP hydrolysisdependent and -independent mechanisms, and general RNA chaperone activity. J Mol Biol 365(3):835–855. https://doi.org/10. 1016/j.jmb.2006.09.083 3. Ballut L, Marchadier B, Baguet A, Tomasetto C, Seraphin B, Le Hir H (2005) The exon junction core complex is locked onto RNA by inhibition of eIF4AIII ATPase activity. Nat Struct Mol Biol 12(10):861–869. https://doi.org/10.1038/nsmb990 4. Bono F, Ebert J, Lorentzen E, Conti E (2006) The crystal structure of the exon junction

complex reveals how it maintains a stable grip on mRNA. Cell 126(4):713–725. https://doi. org/10.1016/j.cell.2006.08.006 5. Jankowsky E, Gross CH, Shuman S, Pyle AM (2001) Active disruption of an RNA-protein interaction by a DExH/D RNA helicase. Science 291(5501):121–125. https://doi.org/ 10.1126/science.291.5501.121 6. Bowers HA, Maroney PA, Fairman ME, Kastner B, Luhrmann R, Nilsen TW, Jankowsky E (2006) Discriminatory RNP remodeling by the DEAD-box protein DED1. RNA 12(5):903–912. https://doi.org/10.1261/ rna.2323406 7. Theuser M, Hobartner C, Wahl MC, Santos KF (2016) Substrate-assisted mechanism of RNP disruption by the spliceosomal Brr2 RNA helicase. Proc Natl Acad Sci U S A 113 (28):7798–7803. https://doi.org/10.1073/ pnas.1524616113

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8. Singleton MR, Dillingham MS, Wigley DB (2007) Structure and mechanism of helicases and nucleic acid translocases. Annu Rev Biochem 76:23–50. https://doi.org/10.1146/ annurev.biochem.76.052305.115300 9. Fairman-Williams ME, Guenther UP, Jankowsky E (2010) SF1 and SF2 helicases: family matters. Curr Opin Struct Biol 20 (3):313–324. https://doi.org/10.1016/j.sbi. 2010.03.011 10. Laggerbauer B, Achsel T, Lu¨hrmann R (1998) The human U5-200kD DEXH-box protein unwinds U4/U6 RNA duplices in vitro. Proc Natl Acad Sci U S A 95(8):4188–4192. https://doi.org/10.1073/pnas.95.8.4188 11. Raghunathan PL, Guthrie C (1998) RNA unwinding in U4/U6 snRNPs requires ATP hydrolysis and the DEIH-box splicing factor Brr2. Curr Biol 8(15):847–855. https://doi. org/10.1016/s0960-9822(07)00345-4 12. Kim DH, Rossi JJ (1999) The first ATPase domain of the yeast 246-kDa protein is required for in vivo unwinding of the U4/U6 duplex. RNA 5(7):959–971. https://doi.org/ 10.1017/S135583829999012x 13. Agafonov DE, Deckert J, Wolf E, Odenwalder P, Bessonov S, Will CL, Urlaub H, Luhrmann R (2011) Semiquantitative proteomic analysis of the human spliceosome via a novel two-dimensional gel electrophoresis method. Mol Cell Biol 31 (13):2667–2682. https://doi.org/10.1128/ MCB.05266-11 14. Pena V, Mozaffari-Jovin S, Fabrizio P, Orlowski J, Bujnicki JM, Lu¨hrmann R, Wahl MC (2009) Common design principles in the spliceosomal RNA helicase Brr2 and in the Hel308 DNA helicase. Mol Cell 35 (4):454–466. https://doi.org/10.1016/j. molcel.2009.08.006 15. Zhang L, Xu T, Maeder C, Bud LO, Shanks J, Nix J, Guthrie C, Pleiss JA, Zhao R (2009) Structural evidence for consecutive Hel308like modules in the spliceosomal ATPase Brr2. Nat Struct Mol Biol 16(7):731–739. https:// doi.org/10.1038/nsmb.1625 16. Santos KF, Mozaffari-Jovin S, Weber G, Pena V, Lu¨hrmann R, Wahl MC (2012) Structural basis for functional cooperation between tandem helicase cassettes in Brr2-mediated remodeling of the spliceosome. Proc Natl Acad Sci U S A 109(43):17418–17423. https://doi.org/10.1073/pnas.1208098109 17. Vester K, Santos KF, Kuropka B, Weise C, Wahl MC (2019) The inactive C-terminal cassette of the dual-cassette RNA helicase BRR2 both stimulates and inhibits the activity of the

N-terminal helicase unit. J Biol Chem 295:2097. https://doi.org/10.1074/jbc. RA119.010964 18. van Nues RW, Beggs JD (2001) Functional contacts with a range of splicing proteins suggest a central role for Brr2p in the dynamic control of the order of events in spliceosomes of Saccharomyces cerevisiae. Genetics 157 (4):1451–1467 19. Cordin O, Hahn D, Alexander R, Gautam A, Saveanu C, Barrass JD, Beggs JD (2014) Brr2p carboxy-terminal Sec63 domain modulates Prp16 splicing RNA helicase. Nucleic Acids Res 42(22):13897–13910. https://doi.org/ 10.1093/nar/gku1238 20. Chen HC, Tseng CK, Tsai RT, Chung CS, Cheng SC (2013) Link of NTR-mediated spliceosome disassembly with DEAH-box ATPases Prp2, Prp16, and Prp22. Mol Cell Biol 33(3):514–525. https://doi.org/10. 1128/MCB.01093-12 21. Nguyen TH, Galej WP, Bai XC, Oubridge C, Newman AJ, Scheres SH, Nagai K (2016) Cryo-EM structure of the yeast U4/U6.U5 tri-snRNP at 3.7 A resolution. Nature 530 (7590):298–302. https://doi.org/10.1038/ nature16940 22. Wollenhaupt J, Henning LM, Sticht J, Becke C, Freund C, Santos KF, Wahl MC (2018) Intrinsically disordered protein Ntr2 modulates the spliceosomal RNA helicase Brr2. Biophys J 114(4):788–799. https://doi. org/10.1016/j.bpj.2017.12.033 23. Henning LM, Santos KF, Sticht J, Jehle S, Lee CT, Wittwer M, Urlaub H, Stelzl U, Wahl MC, Freund C (2017) A new role for FBP21 as regulator of Brr2 helicase activity. Nucleic Acids Res 45(13):7922–7937. https://doi. org/10.1093/nar/gkx535 24. Absmeier E, Wollenhaupt J, Mozaffari-Jovin S, Becke C, Lee CT, Preussner M, Heyd F, Urlaub H, Luhrmann R, Santos KF, Wahl MC (2015) The large N-terminal region of the Brr2 RNA helicase guides productive spliceosome activation. Genes Dev 29 (24):2576–2587. https://doi.org/10.1101/ gad.271528.115 25. Absmeier E, Rosenberger L, Apelt L, Becke C, Santos KF, Stelzl U, Wahl MC (2015) A noncanonical PWI domain in the N-terminal helicase-associated region of the spliceosomal Brr2 protein. Acta Crystallogr D Biol Crystallogr 71:762–771. https://doi.org/10.1107/ S1399004715001005 26. Mozaffari-Jovin S, Wandersleben T, Santos KF, Will CL, Lu¨hrmann R, Wahl MC (2013) Inhibition of RNA helicase Brr2 by the C-terminal

Functional Characterization of the Brr2 RNA Helicase tail of the spliceosomal protein Prp8. Science 341(6141):80–84. https://doi.org/10.1126/ science.1237515 27. Nguyen TH, Li J, Galej WP, Oshikane H, Newman AJ, Nagai K (2013) Structural basis of Brr2-Prp8 interactions and implications for U5 snRNP biogenesis and the spliceosome active site. Structure 21(6):910–919. https:// doi.org/10.1016/j.str.2013.04.017 28. Absmeier E, Becke C, Wollenhaupt J, Santos KF, Wahl MC (2017) Interplay of cis- and trans-regulatory mechanisms in the spliceosomal RNA helicase Brr2. Cell Cycle 16 (1):100–112. https://doi.org/10.1080/ 15384101.2016.1255384

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29. Studier FW (2005) Protein production by auto-induction in high-density shaking cultures. Protein Expr Purif 41(1):207–234. https://doi.org/10.1016/j.pep.2005.01.016 30. Fabrizio P, McPheeters DS, Abelson J (1989) In vitro assembly of yeast U6 snRNP: a functional assay. Genes Dev 3(12B):2137–2150. https://doi.org/10.1101/gad.3.12b.2137 31. Bieniossek C, Richmond TJ, Berger I (2008) MultiBac: multigene baculovirus-based eukaryotic protein complex production. Curr Protoc Protein Sci. Chapter 5:Unit 5 20. https://doi.org/10.1002/0471140864. ps0520s51

Chapter 14 Monitoring Acetylation of the RNA Helicase DDX3X, a Protein Critical for Formation of Stress Granules Makoto Saito, Vytautas Iestamantavicius, Daniel Hess, and Patrick Matthias Abstract Stress granules are dynamic structures that assemble in response to various forms of stress, such as heat shock or oxidative stress, among others. We had previously shown that the lysine deacetylase HDAC6 is required for the formation of stress granules, but the substrate important for this function was not known. We recently found that the RNA helicase DDX3X is a novel HDAC6 substrate, which is critical for the formation of stress granules. Through a series of detailed experiments, we showed that, upon stress, DDX3X becomes acetylated in an intrinsically disordered region; this alters its propensity to undergo phase separation and inhibits growth of the stress granules. HDAC6, by deacetylating DDX3X, allows maturation of the stress granules. This work identified acetylation of an RNA helicase as a critical regulator of the stress response. Here, we present methods to analyze the acetylation of DDX3X; these methods can be easily adapted to study the acetylation of other helicases, or other proteins. Key words Protein acetylation, HDAC6, RNA helicases, DDX3X, Mass spectrometry, Immunoblotting, Immunoprecipitation, Stress granules

1

Introduction Protein lysine acetylation was identified more than 50 years ago in histones, and these early experiments showed that acetylation of histones correlates with increased RNA synthesis activity in isolated calf thymus cells [1]. At that time, no transcription factor had been purified or cloned, and the nucleosome—the basic unit of chromatin organization—had not been described yet [2]. In a seminal paper, the authors showed that acetylated histones were not inhibitory to RNA synthesis, unlike non-acetylated histones. This suggested that acetylation modifies how histones interact with the DNA template and that this leads to facilitated RNA synthesis; this therefore established the link between acetylation and gene expression. Following these insightful findings, it took another 30 years to identify and isolate acetylation-modifying enzymes.

Marc Boudvillain (ed.), RNA Remodeling Proteins: Methods and Protocols, Methods in Molecular Biology, vol. 2209, https://doi.org/10.1007/978-1-0716-0935-4_14, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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Because acetylation was thought to be confined to histones, the enzymes were dubbed histone deacetylases (HDACs) or histone acetyltransferases (HATs), and this nomenclature has largely remained in use, although today the generic terms KDACs or KATs (K meaning lysine) would be preferred [2]. Since the initial description of acetylation on histones, acetylation has been evidenced on multiple other proteins, such as tubulin [3], the transcription factor p53, the chaperone HSP90, or the retinoblastoma protein pRb, to name a few; in each case, acetylation led to a modification of the protein functional properties. In addition, the recent development of large-scale proteomics methods based on mass spectrometry (MS) has allowed to define in various cells or organisms the “acetylome,” i.e., the collection of all acetylated sites present in the sample [4]. This led to the discovery of several thousands of acetylation sites on a large fraction of the cellular proteins, so that acetylation is now considered to be possibly the second most prevalent post-translational protein modification (PTM) after phosphorylation [5]. HDAC6 is a mostly cytoplasmic deacetylase which has a unique structure with two catalytic domains and a zinc finger domain binding ubiquitin [6, 7]. It is involved in various cellular processes related to the stress response, such as the formation of aggresomes in response to misfolded proteins accumulation [8, 9] or the formation of stress granules (SGs) following diverse forms of stress [10]. It is also involved in viral infection [11, 12]. The major substrate of HDAC6 is K40 in alpha-tubulin; because tubulin is highly abundant and a suitable antibody is available, detection of tubulin acetylation is easy to do by immunoblotting (IB). Since HDAC6 is the main tubulin deacetylase in most cells, monitoring the level of tubulin acetylation can be used as a proxy for HDAC6 activity. In a recently published study [13], we found that the RNA helicase DDX3X, which has been linked to various processes such as cancer and function of the inflammasome [14–16], is a novel substrate of HDAC6. For this, we examined the acetylome of cells treated with two HDAC6-specific inhibitors (bufexamac or tubacin), as well as that of HDAC6 knockout mouse embryo fibroblasts (KO MEFs) [17]. Comparison with the acetylome of wild type (WT) cells allowed to identify acetylated peptides that were enriched specifically in the inhibitor-treated cells and/or in the KO cells and could be possible novel HDAC6 substrates. Figure 1a shows the primary structure of DDX3X and the location of the acetylated residues; the acetylated lysines all map to a so-called intrinsically disordered region (IDR), a domain that according to computer prediction algorithms does not assume any particular three-dimensional structure. Through an extensive series of experiments, we could show (1) that acetylation of the DDX3X IDR impairs its propensity to undergo phase separation in vitro and (2) that this blocks the maturation of SGs in vivo. This was the first

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Fig. 1 Acetylation sites on DDX3X-IDR1. (a) The structure prediction program VL3-BA defines DDX3X amino acids 1–167 as an N-terminal IDR1 (bottom, Predictor Of Natural Disordered Regions [PONDR] score graph). Amino acid sequence of IDR1 is shown (upper). All ten lysine residues are written in red. (b) Acetylation sites identified by each method: acetylome analysis, DDX3X immunoprecipitation followed by immunoblot (IP-IB) with DDX3X K118Ac-specific antibody, FLAG-DDX3X IP followed by mass spectrometry (MS) analysis. In the case of the experiments done in 293 T cells, the level of protein acetylation was boosted by transfection of the acetyltransferase CBP. Data is from [13, 18]

demonstration that acetylation can modulate phase separation and SG dynamics, and this may have implications in various settings [13]. As is visible in Fig. 1b, different acetylation sites were identified in different cell lines (here, HeLa, MEF, or 293 T) or by the

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different methods used: acetylome, immunoprecipitation (IP)-MS, and IP-IB. Generally, for biochemical analysis of protein modifications such as phosphorylation or acetylation, immunoblotting is the most convenient (in the sense of easy and fast) method, but mass spectrometry is more general and specific. Yet, each method will have its own limitations and sites that are identified by one method will not automatically be identified by the other. Fig. 2 illustrates schematically a number of factors that can influence the detection of protein acetylation: cell type (i.e., the site is only modified in some cell types), methodology (e.g., immunoblotting vs mass spectrometry), reactivity of the antigen, quality of the antibodies, etc. The detection of protein acetylation by immunoblotting can be very challenging, due to, for example, low stoichiometry of the modification or unreliable quality of the anti-acetyl-lysine antibodies; Fig. 3 illustrates the latter point. For this experiment, WT or HDAC6 KO cells were control-treated or treated with the pan-HDAC inhibitor trichostatin A (TSA) in order to increase the acetylation level of cellular proteins. Protein lysates were prepared and analyzed by SDS-PAGE followed by immunoblotting. The upper part of panel B shows the result obtained when using antibodies specific for acetylated tubulin or acetylated histone H3: in each case, the increase in acetylation following inhibitor treatment, or genetic ablation, is easy to detect and unambiguous. The situation is very different when six different anti-acetylated lysine antibodies were used to examine the same extracts. As is visible, the pattern of bands is very different for the different antibodies, and even the most abundant signals, tubulin and histone H3, are only detected by some of the antibodies: for example, Cell Signaling Technologies antibody #9681 detects acetylated tubulin (MW ca. 50 kDa) but fails to detect acetylated H3 (MW ca. 17 kDa). From this experiment, it is immediately evident that detecting acetylation of a low abundance protein by immunoblotting may be very challenging, as the anti-acetyl lysine antibodies vary widely in their sensitivity and specificity. The solution is often to develop an antibody that is specific for the protein of interest, as we have done for the antibody recognizing DDX3X-K118Ac. This is however associated with a significant time investment and cost, and there is no guarantee that a suitable antibody will be obtained. The flowchart in Fig. 4 describes how we have proceeded to examine DDX3X acetylation and to define its role in phase separation and stress granule formation. The relevant protocols to interrogate DDX3X acetylation are described in the methods section. This workflow and the corresponding methods are generally applicable for other helicases, or any protein of interest. Mass spectrometry-based acetylome determination allows detecting the collection of acetylated peptides that are present within the

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Fig. 2 Comparison between acetylome, immunoprecipitation (IP) followed by immunoblotting (IB), and mass spectrometry (MS) analysis. Six possible reasons to explain discrepancy between these three methodologies are presented. The acetylation status of the target protein may depend on cell type (1. Cell specificity). Acetylated species of the target protein may represent only a fraction of the protein and be below the detection threshold (2. Stoichiometry). Several commercial acetylated lysine antibodies (AcKAb), and also site-specific antibodies (SpAb), are available, but there is no consensus in the field about their quality (3. Antibody specificity). The reaction conditions for antibody-antigen interaction are different; digested peptides are immuno-enriched with acetylated lysine antibodies in the acetylome analysis, while undigested denatured proteins are reacted with antibodies on the immunoblot membrane (4. Condition of epitope). Chromatographic behavior and mass spectrometric response vary greatly for different peptides (5. Bias from MS and chromatography). Acetylome analysis measures the acetylation status of each lysine residue individually, whereas immunoblotting detects the acetylation status of the whole protein unless site-specific antibodies are used. Target-specific MS following IP can identify acetylated peptides from the target protein even at the low stoichiometry of acetylation due to the high dynamic range and sensitivity of modern mass spectrometers (6. Output)

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A) Name and supplier ImmuneChem, ICP0380, Anti Acetyl Lysine Cell Signaling Technology #9441, Acetylated-Lysine Cell Signaling Technology #9681, Acetylated-Lysine (Ac-K-103) Cell Signaling Technology #9814, Acetylated-Lysine (Ac-K2-100) Millipore 05-515, Anti-Acetyl-Lysine Antibody (4G12) Thermo Scientific, MA1-2021 Acetyl Lysine Antibody (1C6)

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proteome of a given sample. Alternatively, one can focus on a particular protein. Acetylation of the protein can be induced/ enhanced by treatment with HDAC inhibitors or by transfection of an acetyltransferase such as CBP (CREB-binding protein). In addition, it may be advantageous to develop an acetylation sitespecific antibody, which will be useful for all downstream analysis methods, such as mass spectrometry, immunoprecipitation, immunoblotting, and immunofluorescence.

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Fig. 4 Flowchart showing methodology covered in this article. Acetylome analysis is a promising strategy to obtain a comprehensive view of acetylation at proteome scale. In order to analyze the acetylation status of a specific protein, this article introduces three strategies: mass spectrometry analysis, immunoblotting, and immunofluorescence. As acetylation often occurs at low stoichiometry, the transient expression of a plasmid encoding an acetyltransferase is usually recommended (Subheading 3.2). If one already knows the acetylation site on a target protein, establishment of acetylation (Ac) site-specific antibody for the site (Subheading 3.1) should be considered. One can directly test the status of target acetylation site by immunoblotting without immunoprecipitation (Subheadings 3.3. and 3.5). Moreover, the established acetylation site-specific antibody may also work in immunofluorescence to assess the localization of the target protein (Subheading 3.7). If one assesses the total acetylation level of a target protein with an acetylated lysine antibody, immunoprecipitation would be required to isolate the target protein from other acetylated proteins in cells (Subheadings 3.3 and 3.4). As mass spectrometry analysis works better with enrichment of the target proteins, this can be achieved by overexpression of epitope-tagged target protein and immunoprecipitation with the tag-specific antibody or a protein-specific antibody. Immunoprecipitated target protein can be subjected to immunoblotting (Subheading 3.5) and mass spectrometry analysis (Subheading 3.6)

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Materials

2.1 Establishment and Characterization of DDX3X-K118AcSpecific Antibody

1. Synthetic DDX3X-K118 acetylated peptide, CDRSGFGK (Ac)FERG. 2. Synthetic DDX3X-K118 peptide, CDRSGFGKFERG. 3. Rabbits immunization service.

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4. Imject Maleimide Activated Carrier Protein Spin Kits (Thermo Fisher Scientific). 5. SulfoLink Immobilization Kit for Peptides (Thermo Fisher Scientific). 6. 0.2 M glycine-HCl, pH 2.0. 7. 2 M Tris–HCl, pH 8.5. 8. Nanodrop (Thermo Fisher Scientific) or equivalent UV–vis spectrophotometer with microvolume capability. 9. 3500 Da molecular weight cutoff (MWCO) Spectrum™ Spectra/Por™ or equivalent dialysis membrane. 10. TBS buffer: 50 mM Tris–HCl pH 7.6, 150 mM NaCl. 11. Pierce BCA protein assay kit (Thermo Fischer Scientific). 12. Peptide Coating Kit (TaKaRa). 13. 1-Step™ Slow TMB-ELISA Substrate Solution (Thermo Fisher Scientific). 14. TBST: 0.05% (w/v) Tween 20, 50 mM Tris–HCl pH 7.6, 150 mM NaCl. 15. Blocking buffer: 5% (w/v) bovine serum albumin in TBST. 16. Amersham ECL Rabbit IgG, HRP-linked whole Ab from donkey (GE Healthcare). Dilute 1:2000 in blocking buffer. 17. 2 M sulfuric acid. 2.2 Induction of DDX3X Acetylation

1. HEK293T cells. 2. Dulbecco’s Modified Eagle Medium (DMEM) supplemented with 10% (v/v) fetal bovine serum (FBS). 3. FLAG-DDX3X expression vector plasmid, prepared as described previously [13]. 4. Acetyltransferase CBP expression vector plasmid (kind gift from Dr. Renate Voit, German Cancer Research Center). 5. Lipofectamine 2000 (Invitrogen).

2.3 Preparation of Cell Lysate for Immunoprecipitation and Immunoblotting

1. Tabletop centrifuge. 2. Tube rotator. 3. Phosphate buffered saline (PBS), pH 7.4. 4. Triton lysis buffer: 50 mM Tris–HCl pH 8.0, 150 mM NaCl, 1 mM EDTA, 0.1% (v/v). TritonX-100, 10 μM Trichostatin A, 10 mM nicotinamide, 50 mM sodium butyrate. 5. cOMPLETE™ EDTA-free protease inhibitor tablets (Roche). 6. Pierce BCA protein assay kit (Thermo Fischer Scientific).

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1. DDX3 (D19B4) Rabbit mAb (Cell Signaling Technology). 2. Anti-FLAG M2 Magnetic Beads (Millipore Sigma). 3. Dynabeads Protein G (Thermo Fischer Scientific). 4. Triton lysis buffer: 50 mM Tris, pH 8.0, 150 mM NaCl, 1 mM EDTA, 0.1% Triton X-100, 10 μM Trichostatin A, 10 mM nicotinamide, 50 mM sodium butyrate.

2.5 Immunoblotting to Detect ac-DDX3X

1. 5 SDS-PAGE sample buffer: 0.25% (w/v) bromophenol blue, 500 mM dithiothreitol. (DTT), 50% (w/v) glycerol, 10% (w/v) sodium dodecyl sulfate (SDS), 250 mM Tris–HCl pH 6.8. 2. 1 SDS-PAGE sample buffer: 1:5 dilution of 5 SDS-PAGE sample buffer with water. 3. NuPAGE 4–12% Bis-Tris Protein Gels, 1.0 mm, 12-well (Invitrogen). 4. 20 NuPAGE MES SDS Running Buffer (Invitrogen). 5. 1 NuPAGE MES SDS Running Buffer: 1:20 dilution of 20 NuPAGE MES SDS Running Buffer with water. 6. XCell SureLock Mini-Cell Electrophoresis System. 7. Semi-dry transblot apparatus. 8. Immobilon-P PVDF Membrane (Millipore). 9. TBS buffer: 50 mM Tris–HCl pH 7.6, 150 mM NaCl. 10. Blocking buffer: 5% (w/v) skim milk in TBS buffer. 11. DDX3X-K118Ac-specific antibody (see Subheading 3.1). Dilute to 1 μg/mL in blocking buffer. 12. Acetylated lysine antibody (Cell Signaling Technology). Dilute 1:1000 in blocking buffer. 13. DDX3 (D19B4) Rabbit monoclonal antibody (mAb) (Cell Signaling Technology). Dilute 1:1000 in blocking buffer. 14. TBST: 0.05% (w/v) Tween 20, 50 mM Tris–HCl pH 7.6, 150 mM NaCl. 15. Amersham ECL Rabbit IgG, HRP-linked whole Ab from donkey (GE Healthcare). Dilute 1:2000 in blocking buffer. 16. Amersham ECL western blotting reagents (GE Healthcare). 17. X-ray film. 18. Film developing machine in a dark room.

2.6 Identification of the DDX3X Acetylation Sites

1. Beads wash buffer: 50 mM Tris–HCl pH 8.0, 150 mM NaCl. 2. Digestion buffer: 3 M guanidine hydrochloride, 20 mM 4-(2-hydroxyethyl)-1-piperazinepropanesulfonic acid (HEPPS/EPPS) pH 8.5, 10 mM chloroacetamide, 5 mM tris (2-carboxyethyl)phosphine (TCEP).

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3. 0.2 μg/μL porcine sequencing grade trypsin (Promega) in 0.2 mM HCl. 4. 0.2 μg/μL lysyl endopeptidase Lys-C in 50 mM HEPES pH 8.5. 5. 50 mM HEPES pH 8.5. 6. 20% (v/v) trifluoroacetic acid in water. 7. Ultrasound bath. 8. MS autosampler tube. 9. Dual-column capillary liquid chromatography system coupled to tandem mass spectrometer. 2.7 Immunofluorescenceto Detect ac-DDX3X

1. MEF cells. 2. Cover glasses, circles, 18 mm, thickness 0.09–0.12 mm (Carolina Biological Supply). 3. 0.05 M sodium arsenite solution. 4. PBS. 5. Fixation buffer: 4% (w/v) paraformaldehyde in PBS. 6. 0.005% (w/v) digitonin in PBS. 7. Wash buffer: 0.1% (w/v) TritonX-100 in PBS. 8. Blocking buffer: 10% (v/v) goat serum in wash buffer. 9. DDX3X-K118Ac-specific antibody (see Subheading 3.1). Dilute to 20 μg/mL in blocking buffer. 10. Anti-DDX3X (15D1B11) antibody (BioLegend). Dilute 1:50 in blocking buffer. 11. Alexa Fluor 488 Goat anti-Mouse IgG (H + L) Secondary Antibody (Invitrogen). Dilute 1:500 in blocking buffer. 12. Alexa Fluor 568 Goat anti-Rabbit IgG (H + L) Secondary Antibody (Invitrogen). Dilute 1:500 in blocking buffer. 13. Kimwipe paper. 14. ProLong Gold Antifade Reagent with DAPI (Cell Signaling Technology). 15. Glass slides.

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Methods

3.1 Establishment and Characterization of DDX3X-K118AcSpecific Antibody

1. Conjugate the DDX3X-K118 acetylated peptide with the mcKLH carrier protein using the Imject Maleimide Activated Carrier Protein Spin Kits. Follow the manufacturer’s instructions.

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2. Immunize rabbits with the KLH-conjugated peptide. Typically, this is done through an immunization service such as that of Pocono Rabbit Farm & Laboratory. 3. Following the manufacturer’s instructions, prepare two types of agarose column with the SulfoLink Immobilization Kit for Peptides: K118 acetylated peptide CDRSGFGK(Ac)FERGconjugated agarose column and K118 unacetylated peptide CDRSGFGKFERG-conjugated agarose column. 4. Pass collected serum (from step 2; usually between 50 and 100 mL) over K118 acetylated peptide conjugated agarose column. Elute with 8 mL of 0.2 M glycine-HCl, pH 2.0. 5. Collect 1 mL fractions into tubes, each containing 150 μL of 2 M Tris–HCl, pH 8.5 for neutralization. 6. Read OD280 of the fractions by Nanodrop. Pool all fractions whose OD280 > 0.5. 7. Pass the pooled elution over K118 unacetylated peptide conjugated agarose column, and collect flow through which should represent the desired antibodies (the antibodies that recognize the non-acetylated peptide will be retained on the column). 8. Dialyze the flow through against 1 L TBS (for 12–15 h in the cold room). 9. Measure the concentration of the antibody with the BCA protein assay kit (see Note 1). 10. Prepare K118 acetylated and unacetylated peptide coated plates with Peptide Coating Kit (TaKaRa), following the manufacturer’s instructions. 11. Dilute the antibody-containing fraction (from step 8) to the appropriate concentrations (0.125, 0.25, 0.5, and 1 μg/mL) in blocking buffer to perform an enzyme-linked immunosorbent assay (ELISA) (steps 12–19). 12. Add 100 μL of diluted antibody to the plates, and incubate for 1 h at RT. 13. Remove the solution, and wash the plate with 200 μL TBST for 10 min on a rocker. Repeat the process two more times (three times in total). 14. Add 100 μL of diluted anti-rabbit HRP secondary antibody to the plates, and incubate for 1 h at RT. 15. Remove the solution and wash the plate with 200 μL TBST for 10 min on a rocker. Repeat the process two more times. 16. Add 100 μL of 1-Step Slow TMB-ELISA to each well. 17. Incubate the plate at RT until the desired color develops (5–30 min).

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18. Stop reaction by adding 100 μL of 2 M sulfuric acid to each well. 19. Measure the absorbance at 450 nm and confirm that the prepared antibody recognizes the K118 acetylated peptide. Use the resulting fraction (from step 9) as the DDX3XK118Ac-specific antibody. 3.2 Induction of DDX3X Acetylation

Day 1 1. Seed 2  106 HEK293T cells into a 35 mm dish 1 day before transfection. Day 2 2. To analyze endogenous DDX3X acetylation, transfect acetyltransferase CBP expression vector alone (proceed to step 2a). For mass spectrometry identification of acetylation sites, DDX3X should be enriched by immunoprecipitation. In this case, FLAG-DDX3X transient expression would be helpful (proceed to step 2b) as commercial FLAG M2 antibody efficiently works in immunoprecipitation. a. Transfect the CBP expression vector plasmid into HEK293T cells by Lipofectamine 2000. Follow the manufacturer’s instructions. b. Transfect the CBP expression vector plasmid into HEK293T cells together with the FLAG-DDX3X expression vector plasmid (use a 4:1 CBP:DDX3X DNA amount ratio to account for the large size of the CBP expression vector) by Lipofectamine 2000. Follow the manufacturer’s instructions. 3. Incubate Overnight at 37  C Day 3 4. Passage transfected cells into a 10 cm dish to avoid their overcrowding in a 35 mm dish. 5. To test the effect of HDAC inhibitors, treat transfected cells after their attachment to the dish at this point (see Note 2). 6. Incubate overnight at 37  C. Day 4 (2 days After Transfection) 7. Proceed to Subheading 3.3.

3.3 Preparation of Cell Lysate for Immunoprecipitation and Immunoblotting

1. Remove culture supernatant, and gently rinse transfected HEK293T cells with 10 mL of ice-cold PBS. 2. Scrape cells and transfer them to a 1.5 mL microtube. Collect the cell pellet by centrifugation at 16,100 g for 10 min at 4  C.

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3. Add 300 μL of ice-cold Triton lysis buffer supplemented with a cOMPLETE protease inhibitor tablet to the cell pellet (see Note 3). Resuspend gently by pipetting up and down several times. 4. Incubate the tube on ice for 15 min, and rotate the tube at 10 rpm in cold room for another 15 min to lyse cells completely. 5. Centrifugation at 16,100 g for 10 min at 4  C. Transfer the supernatant to a new 1.5 mL microtube (see Note 4). 6. Measure the protein concentration of lysate by BCA protein assay kit. Adjust the concentration to 1 μg/μL (see Note 5). 7. Proceed to Subheading 3.4 (immunoprecipitation) OR Subheading 3.5 (immunoblotting) directly. 3.4 Immunoprecipitation of DDX3X

3.4.1 Endogenous DDX3X Precipitation by DDX3 Antibody

If one uses acetylated lysine antibody, one first needs to immunoprecipitate DDX3X from the cell lysate so that one can distinguish its acetylation signal from those of other proteins. This immunoprecipitation process should be performed in a cold room. 1. Add 5 μL of DDX3 antibody into 500 μL protein lysate (containing 500 μg protein) from Subheading 3.3, step 6. 2. Rotate tube overnight at 10 rpm in cold room. 3. Prepare ProteinG Dynabeads: Transfer 50 μL of beads into a 1.5 mL microtube and remove supernatant. Add 1 mL of lysis buffer and remove supernatant. Repeat the process five times in total. The beads can be resuspended in 50 μL of lysis buffer for further handling. 4. Add 50 μL of pre-washed Dynabeads ProteinG (from step 3 above) into the tube containing the protein lysate and antibody. 5. Rotate tube at 10 rpm for 1 h in cold room. 6. Remove supernatant. Wash the beads with 1 mL of Triton lysis buffer. Repeat the process five times in total. 7. Proceed to Subheading 3.5 (immunoblotting).

3.4.2 Immunoprecipitation of Transiently Expressed FLAG-DDX3X by FLAG Antibody

1. Add 10 μL of FLAG M2 magnetic beads into 500 μL protein lysate (containing 500 μg protein) from Subheading 3.3, step 6. 2. Rotate tube at 10 rpm for 1 h in cold room. 3. Remove supernatant. Wash the beads with 1 mL of Triton lysis buffer. Repeat the process three times in total. 4. Proceed to Subheading 3.5 (immunoblotting) OR Subheading 3.6 (mass spectrometry).

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3.5 Immunoblotting to Detect Ac-DDX3X

1. Boil samples in SDS-PAGE sample buffer for 10 min at 95  C. We routinely add 4 μL of 5 sample buffer into 16 μL of cell lysate or 20 μL of 1 sample buffer directly to the immunoprecipitated beads. 2. Install 4–12% NuPAGE gel(s) in the electrophoresis apparatus, and fill apparatus with 1 NuPAGE MES SDS Running Buffer. If one detects the acetylation of immunoprecipitated DDX3X, load 18 μL out of 20 μL on a gel (for immunoblotting with acetylated lysine antibody) and the remaining 2 μL on a second gel (for immunoblotting with DDX3X antibody) in order to control the amount of immunoprecipitated DDX3X. 3. Run gel for 35 min at 200 V. 4. Transfer proteins from the gel onto the membrane using semidry transblot apparatus, following manufacturer’s instructions. 5. Place membrane in a tray filled with blocking buffer. Incubate the membrane in blocking buffer on a rocker (set at 10 rpm) at room temperature for 1 h. 6. Place the blocked membrane in a 50 mL Falcon tube with 2 mL of the diluted DDX3X-K118Ac-specific antibody (OR acetylated lysine antibody and DDX3X antibody for immunoprecipitated product; see Note 6). 7. Rotate on a rocker (set at 10 rpm) in the cold room for overnight. 8. Wash the membrane with TBST for 10 min on a rocker (set at 10 rpm). Repeat the process three times in total. 9. Place the blocked membrane in the 50 mL falcon tube with 2 mL of the diluted anti-rabbit HRP secondary antibody. 10. Rotate on a rocker (set at 10 rpm) at room temperature for 1 h. 11. Wash the membrane with TBST for 10 min on rocker. Repeat the process three times in total. 12. React with Amersham ECL western blotting reagents following the manufacturer’s instructions. 13. Detect the signal with an X-ray film using a film developing machine following the manufacturer’s instructions.

3.6 Identification of the DDX3X Acetylation Sites

1. Add 1 mL wash buffer WITHOUT DETERGENTS and without protease inhibitors (see Note 7). Mix by inversion, separate beads magnetically, and remove the supernatant. Repeat the process four times in total. 2. Take 10 μL of Lys-C stock and add 50 μl of digestion buffer (enough for ten reactions).

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3. Remove the wash buffer completely from the beads and add 6 μL of Lys-C in digestion buffer, spin down quickly ( * > 0.01 > ** > 0.001 > ***

In this chapter, we provide a full description of our Rho-based experimental approach and outline its use to investigate the mRNP QC activities over the whole yeast genome. High-throughput RNA sequencing (RNA-seq) was used to survey a broad collection of mRNPs whose biogenesis is affected by Rho action (Fig. 1). This genome-wide perspective of affected mRNPs was extended by generating high-resolution maps (ChIP-seq) of the distribution of the QC components along the yeast chromosomes before and after perturbation of mRNP biogenesis by Rho. As illustrated in Fig. 2, the perturbation of mRNP biogenesis redistributes the binding of QC components over the genome with a large shift of Nrd1 and Nab3 from genomic loci producing ncRNAs prone to processing

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Fig. 2 Rho activity mediates the stimulation of massive recruitment of the QC components (Nrd1, Nab3, Rrp6, and Trf4) to rrp6-sensitive mRNA genomic loci at the expense of their locations within ncRNA genomic features. (a) IGV snapshots of ChIP-seq peaks mapped over two rescued mRNA genomic loci. The ChIP signals for each protein are shown both for Rho and + Rho conditions. (b) Circos plots summarizing the observed dynamic landscape of the four QC components over the 16 yeast chromosomes analyzed under Rho (left plots) and + Rho (right plots) conditions. The two top plots show the ChIP signals detected for the ncRNA genomic loci (CUTs, SUTs, and snoRNAs). The two plots at the bottom show the ChIP signals detected for the genomic features of the Rho-affected mRNAs that were rescued by Rrp6 depletion. The red arrow symbolizes the hijacking of Nrd1 and Nab3 from ncRNA genomic loci to the Rho-affected mRNA genes upon perturbation of mRNP biogenesis by Rho. Student’s t-test (bidirectional) was used to compare Log2 average ChIP-seq signals

and decay such as CUTs (cryptic unstable transcripts), SUTs (stable unannotated transcripts), and snoRNAs (small nucleolar RNAs) to protein-coding genes. Nrd1 and Nab3 are apparently titrated out from ncRNA genomic features by a large recruitment to Rho-affected mRNA gene loci, leading to transcriptional termination defects of ncRNAs [13]. This approach can be adapted to any protein potentially involved at the chromatin level in mRNA surveillance and/or processing. It can also be extended to the study of ncRNAs whose processing relies on the same machineries as mRNA.

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Materials

2.1 Yeast strains, Plasmids, and Growth Media

1. All Saccharomyces cerevisiae strains used for the present work are listed below. They are derived from the wild-type strain BMA41 (MATa or MATα ade2-1 ura3-1 leu2-3112 his3-11,15 trp1Δcan1-100) [14] and were constructed by homologous recombination (see Note 1): – rrp47Δ, like BMA41 with rrp47Δ::KAN, – NAB3-MYC, like BMA41 with Nab3-MYC::HIS3. – NRD1-MYC, like BMA41 with Nrd1-MYC::HIS3. – RRP6-MYC, like BMA41 with Rrp6-MYC::HIS3. – TRF4-MYC, like BMA41 with Trf4-MYC::HIS3. 2. pCM185-Rho-NLS plasmid, a derivative of the pCM185 vector (CEN, TRP) from “EUROSCARF (http://www. euroscarf.de) expressing bacterial Rho factor in yeast with a C-terminal NLS (nuclear localization signal) and under the control of a doxycycline-regulated (Tet-Off) promoter [8]. 3. Liquid media for the growth of yeast cells, prepared from a mix of three different powders. For 900 mL of media, weigh 18 g of glucose, 6.03 g of YNB (Yeast Nitrogen Base) containing ammonium sulfate, and 0.67 g of CSM-Trp (complete synthetic medium without tryptophan) dissolved in 900 mL of distilled water (see Note 2). After dissolution, dispatch into three portions of 300 mL in 500 mL bottles and autoclave. Store the autoclaved media for up to 6 months at room temperature. 4. 1000 stock solution of doxycycline at 5 mg/mL. Dissolve 125 mg of doxycycline in 25 mL of 50% ethanol/water (v/v); store at 20  C. The Tet promoter is repressed by addition of doxycycline to the growth medium at a final concentration of 5 μg/mL just before use. 5. Specific instruments and labware: – Precision balance for weighting the powder for media preparation. – Vortex for homogenization of solution. – Magnetic plate for agitation (with heating function). – Centrifuge for pelleting the yeast cells when needed. – Glass or plastic flasks for cell growth. – Optical density measurement device to monitor cell growth. – Rotary shaker (250–300 rpm) for cell growth at controlled temperature.

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2.2 RNA Extraction Solutions

1. DEPC-treated water. 2. AE buffer: 50 mM sodium acetate, pH 5.2, 10 mM EDTA. 3. 10% (w/v) sodium dodecyl sulfate (SDS) solution in water. 4. Buffer-saturated, phenol-chloroform solutions, at pH 4.6 and at pH 6.7. 5. 3 M sodium acetate solution, pH 5.2. 6. Ethanol absolute (>99%). 7. 75% ethanol in water. 8. Dry ice. 9. 2 U/μL RNase-free DNase I with 10 supplied buffer. 10. Specific instruments and labware: – Dedicated set of pipettes and special RNase-free tips. – Fume hood to avoid inhaling phenol-chloroform vapes. – NanoDrop spectrophotometer or equivalent.

2.3 Chromatin Immunoprecipitation (ChIP)

1. TBS buffer: 20 mM Tris–HCl, pH 7.5, 150 mM NaCl. 2. 1 phosphate-buffered saline (PBS), pH 7.4. 3. PBS/BSA mixture: dissolve 125 mg bovine serum albumin (BSA) in 25 mL 1 PBS. 4. 0.1 M PMSF solution. Dissolve 430 mg of protease inhibitor phenylmethylsulfonyl fluoride (PMSF) in 25 mL of isopropanol, and store as 1 mL aliquots at room temperature until use (see Note 3). 5. 37% formaldehyde solution in water. We use a commercial solution from Sigma, which also contains 10–15% methanol as stabilizer. 6. 2.5 M glycine in water. Sterilize with a 0.22 μm filter, and store as aliquots at 20  C. 7. 1 TE buffer: 10 mM Tris–HCl and 1 mM EDTA, pH 8. 8. Lysis buffer 140: 50 mM HEPES pH 7.5, 140 mM NaCl, 1 mM EDTA, 1% (v/v) Triton X-100, 0.1% (w/v) sodium deoxycholate. Right before use, add 1:100 volume of 0.1 M PMSF (see Note 4). 9. Lysis buffer 360: 50 mM HEPES pH 7.5, 360 mM NaCl, 1 mM EDTA, 1% (v/v) Triton X-100, 0.1% (w/v) sodium deoxycholate. Right before use, add 1:100 volume of PMSF 0.1 M (see Note 4). 10. Washing buffer: 10 mM Tris–HCl pH 8, 0.25 M LiCl, 0.96% (v/v) NP-40, 0.5% (w/v) SDS, 1 mM EDTA. 11. Elution buffer: 50 mM Tris–HCl pH 8, 10 mM EDTA, 1% (w/v) SDS.

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12. 10 mg/mL proteinase K in water. 13. Anti-c-MYC sc-40 antibody at 200 μg/mL (Santa Cruz Biotechnology). 14. Protein G PLUS agarose beads (Sigma). 15. Specific instruments and labware: – Centrifuge with temperature control (must be maintained at 4  C). – Heating bloc. – RNase-/DNase-free tips with barrier filter. – FastPrep device (MB Biomedicals) for cell breakage or an equivalent apparatus. – Quick PCR Purification Kit (see Note 5). – Sonication apparatus (see Note 6). 2.4 Libraries Preparations and NGS Data Treatment

For the preparation of sequencing libraries from immunoprecipitated DNA fragments and inputs, different kits and specific tools are needed: 1. NEBNext Ultra II DNA Library Prep for Illumina from New England Biolabs or equivalent from other suppliers. 2. Qubit assay kit (Thermo Fisher Scientific). 3. Magnetic rack device for washing steps of the NEBNext Ultra II kit. 4. Magnetic beads (specific brands are suggested in the NEBNext Ultra II kit). 5. Spectrophotometer for Qubit assays measurement. 6. Academic core facility or commercial service for highthroughput nucleic acid sequencing, e.g., on an Illumina NextSeq 500 (see Note 7). 7. 64 GB RAM, 12 cores computer, equipped with Linux Ubuntu operating system and R software. 8. Software for NGS data treatment, e.g., as provided by the web-based Galaxy platform (https://usegalaxy.org/).

3

Methods The first parts of each subsection are purely experimental and explain how we used Rho expression in yeast, how we extract RNA to perform high-throughput sequencing, and how we perform chromatin immunoprecipitation to reveal the recruitment of QC components over the genome. The subsequent parts are dedicated to the use of bioinformatics to analyze the genome-wide sequencing results.

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3.1 Cell Growth and Rho-Induction for Perturbation of mRNP Biogenesis

The expression of Rho in yeast is repressed by growing the cells in the presence of the tetracycline analogue, doxycycline (Doxy), at a final concentration of 5 μg/mL, whereas the omission of Doxy in the growth medium allows maximum expression. The cell growth is monitored by measuring the optical density at 600 nm (OD600). 1. Inoculate the S. cerevisiae strain of interest transformed by the plasmid pCM185-Rho-NLS (TRP1 marker) from the 80  C stock or from a streaking on a fresh plate into 5 mL of liquid medium minus tryptophan and with Doxy at a final concentration of 5 μg/mL. Then, let the cells grow in a rotating incubator at 25  C overnight. 2. The following morning, use 800 μL of the culture to inoculate 10 mL of fresh medium containing 0.5 μg/mL of Doxy, and grow the cells at 25  C for approximately 6 h until an OD600 of 0.5. 3. Wash the cells three to four times with fresh medium lacking Doxy by sequential centrifugations at 3500  g for 5 min at 5  C and removal of the supernatant (see Note 8). 4. Dilute the cells and allow them to grow overnight in 200 mL flasks containing 40–45 mL of the medium lacking Doxy to achieve maximum Rho expression. In parallel, grow similar cultures with medium containing 5 μg/mL Doxy for the non-induced samples. After an overnight growth (14–16 h), the Rho-induced cells are typically at an OD600 of 0.25–0.3, whereas the non-induced samples are at an OD600 between 0.8 and 1 (see Note 9). 5. To analyze the samples, dilute the non-induced cultures with fresh medium to adjust the OD600 to the induced cultures before harvesting the cells (RNA extraction) or cross-linking (ChIP).

3.2

RNA Extraction

Total RNA extraction is performed using the acidic hot phenol/ chloroform method according to [15] with some modifications. 1. After adjusting the non-induced and induced cultures to the same lowest OD600, put back the yeast liquid cultures in the agitating incubator at 25  C for 20 min. 2. During this 20 min meantime, prepare all the necessary buffers and stand ready to perform all the following manipulations on ice. 3. Transfer the yeast cultures to 50 mL tubes, and then centrifuge at 4  C, 10 min, 3500  g. Remove the supernatant, and wash the pellet with 1 mL of DEPC-treated water. 4. Resuspend the cell pellet in 1 mL of DEPC-treated water by vortexing, and transfer the suspension to 1.5 mL microtubes. Centrifuge the tubes in a bench centrifuge at 4000  g for

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approximately 1 min. Remove the supernatant, and add 400 μL of AE buffer and 40 μL of 10% SDS, and then vortex to suspend the cells. 5. Transfer the suspensions into 1.5 mL microtubes previously filled with micro glass beads (~1/5 of the tube). The next steps have to be performed under a fume hood. 6. Add an equivalent volume (440 μL) of hot phenol/chloroform pH 4.5 (previously warmed up to 65  C). 7. Vortex the tubes for 1 min, and then incubate them at 65  C for 1 min. This is repeated three times to optimize the mechanical breakage of the cells. 8. Put the samples in ethanol/dry-ice bath for 2 min (alternatively, they can be stored 2 h at 80  C). Put back the samples to the 65  C incubator for 3 min, vortex, and repeat the process twice. 9. Centrifuge the tubes at 4  C, 12 min, 15,000  g in a bench centrifuge, and then carefully pipette the aqueous phase and transfer it in a new tube. After treatment with DNase I (as recommended by the supplier) to remove possible DNA contaminants, add an equivalent volume of phenol/chloroform pH 6.7, vortex, and centrifuge at 4  C for 10 min at 15,000  g. 10. Carefully pipette the aqueous phase, and transfer it in a new tube, and then add sodium acetate to a final concentration of 0.3 M. Add 3 volumes of 100% ethanol, and precipitate at 80  C for 2 h followed by centrifugation at 4  C for 20 min at 15,000  g. Remove the supernatant, and wash the pellet with 75% ethanol. Dry the pellet briefly, and resuspend it in 30 μL of RNase-free water. Then, the RNA can be quantified from a ~1 μL aliquot with a NanoDrop (or equivalent) spectrophotometer (assuming an extinction coefficient at 260 nm for RNA of 0.025 (μg/mL)1 cm1). 3.3 RNA-seq Libraries Preparation and Sequencing

cDNA libraries preparations from isolated RNA were made by I2BC high-throughput sequencing platform (Gif-sur-Yvette) using Illumina ScriptSeq protocol after Ribo-Zero treatment and followed with sequencing on an Illumina NextSeq 500 instrument. We always outsource the preparation of RNA-seq libraries to NGS platforms because they usually have high expertise and are better equipped for the task, especially regarding the quality checks of the samples. For each condition (Rho-repressing or Rho-inducing conditions), two biologically independent samples are used for libraries preparation and sequencing.

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3.4 RNA-seq Data Processing and Computational Analysis

In this section, we provide an overview of the bioinformatics packages and manipulations used for the sequencing analysis and data processing pipeline (see Note 10): 1. Run a quality control analysis of the reads using FastQC v 0.11.5 package. 2. Remove the adaptor’s sequences from the reads using Cutadapt 1.15. 3. Use the bwa 0.7.12 software package to map the reads on the reference genome (Saccharomyces cerevisiae genome V64.1.1). The genome annotations can be downloaded from SGD (https://www.yeastgenome.org/), and specific ncRNAs annotations are taken from [16] for SUTs and from [17] for CUTs. 4. Quality control the alignment to the genome with Samtools 0.1.19. 5. Process the data by counting the number of reads per genomic feature using the package featureCounts V 1.5.0-p1, with -s 1 option. 6. Isolate a working pool of mRNAs from the Wt strain that are affected by Rho action by comparing the mRNA levels between the Rho and +Rho conditions. This determines which mRNAs from the data sets of the Wt strain are becoming downregulated under Rho action. For these differential expression analyses, use DESeq2 package under R environment (V 3.4.4). 7. Perform the same differential analysis on the isolated mRNA pool using the data sets collected from the rrp47Δ strain, which lacks the exonuclease Rrp6. 8. Using the two sets of results obtained above (Wt and rrp47Δ strain), sort the mRNAs by their sensitivity to Rho action and which can be rescued under Rrp6 depletion (Fig. 1a). 9. To visualize the effect of Rho as reads coverage, use the same subsets of the reads results obtained for the selected pools of mRNAs to generate Meta-transcript profiles variations of mRNAs between Wt and the rrp47Δ strain in the absence or presence of Rho (Fig. 1b). To perform these analyses, generate BigWig files using bedtools V2.25.0 (genomecov and bigwigCompare) and UCSC tool BedGraphToBigWig. After computing genome coverage with genomecov, convert the BedGraph outputs to BigWig files using BedGraphToBigWig. Meta-profiles plots are then made using bedtools V2.25.0 with computeMatrix (scale-regions mode, with a bin size of 1 base) on BigWig files and plotProfile for graphical output.

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Each protein of interest within the cross-linked chromatin complex is immunoprecipitated via the MYC tag present at its C-terminus using an antibody recognizing the MYC epitope (anti c-MYC sc-40 antibody). 1. At the end of the overnight growth, as for RNA extraction, adjust the OD600 of the non-induced cells by dilution with medium in order to obtain the same number of cells in each culture/condition. 2. To perform protein-nucleic acid cross-linking within the chromatin, add 1.2 mL of 37% formaldehyde solution to the ~45 mL of yeast culture (from Subheading 3.1) directly into the culture flask, and incubate for 20 min at room temperature on a rotating platform within a fume hood. 3. Stop the cross-linking reaction by adding 2.25 mL of 2.5 M glycine, and incubate 5 min at room temperature, and then put the samples on ice (see Note 11). 4. Transfer the culture into a 50 mL conical tube, and wash the cells three times with cold TBS by applying centrifugation steps at 3500  g for 5 min at 4  C and carefully removing all supernatant (see Note 12). 5. Resuspend the washed cell pellet in 1 mL of lysis buffer 140 by pipetting up and down, and add the cell suspension directly to a 2 mL screw cap microtube, containing glass beads (same volume of beads than the cell suspension). 6. Break the cells with a FastPrep device in a cold room (4 cycles of 30 s at 6000  g with a 1 min pause between each cycle). 7. To recover the cell extract without the glass beads, drill a hole in the bottom of the microtube with an incandescent needle, and then introduce it into a 15 mL conical tube. After removing the screw cap, centrifuge 1 min at 1500  g to recover the extract. 8. After recovering the extract and transferring it to a 2 mL microtube, adjust the volume to 1.2 mL with lysis buffer 140, and then shear the cross-linked chromatin by sonication on ice to reduce average DNA fragments to an approximate size of ~400–500 base pairs (see Note 13). 9. Centrifuge the samples for 5 min at 2500  g and 4  C, and then recover the supernatant (approximately 1 mL), and put it in a new microtube. 10. Take 20 μL of supernatant genomic DNA which will be used as input sample, and mix the remaining part with 4 μL anti c-MYC antibody solution for the IP sample. “Fix the tubes on a rotating plate, and incubate overnight at 4  C.

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11. The next morning, add 37 μL of the protein G PLUS agarose beads slurry, and let the samples rotate for 2 h at 4  C. 12. Perform successive washing steps by 2–3 min centrifugation at 4000  g and supernatant elimination at 4  C (the agarose beads are pulled at the bottom of the tube): 2 times with 1 mL lysis buffer 140, 2 times with 1 mL lysis buffer 360, 1 time with 1 mL washing buffer, and 1 time with 1 mL TE. 13. Elute the DNA from the agarose beads twice with 75 μL of elution buffer during 10 min incubations at 65  C, each time recovering supernatant. 14. The samples (150 μL of IP and the 20 μL of input adjusted to 150 μL by addition of elution buffer) are reverse cross-linked by incubation at 65  C overnight. 15. Add to each sample 15 μL of 10 mg/mL proteinase K, and incubate for digestion 1.5 h at 37  C. Recover the DNA samples using a PCR Purification Kit (see Note 5), following manufacturer’s instructions. 3.6 DNA Libraries Preparations and Sequencing for ChIP-seq

ChIP-seq libraries preparations in duplicates are done following the NEBNext Ultra II DNA Library Prep Kit for Illumina from New England Biolabs as outlined by the supplier. Briefly, the kit workflow can be divided in three major steps (see Note 14). 1. The first step is applied directly on the DNA fragments from the ChIP samples and is based on the enzymatic preparation of both ends of the DNA fragments. 2. The second step consists of adaptors ligation at each end. 3. The third step uses the ligated adaptors as a platform for hybridization of primers (possessing unique barcodes used for multiplexed sequencing) and subsequent amplification of the DNA fragments by PCR. 4. A Qubit assay is then performed following the manufacturer’s instructions to evaluate the amount of DNA obtained at the end of the PCR amplification (see Note 14). 5. The libraries are sent to the preselected high-throughput sequencing platform that runs an Illumina NextSeq 500.

3.7 ChIP-seq Data Analysis

Upon completion of the high-throughput sequencing, download the results and perform the data analyses according to the following steps (see Note 15): 1. Run a quality control of the reads using FastQC v 0.11.5 package. 2. Process the trimming of the adaptor’s sequences from the reads using Cutadapt 1.15 package.

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3. Map the whole reads on the reference Saccharomyces cerevisiae genome (version V64.1.1) with the bwa 0.7.12 software package. 4. Quality control the alignment of the reads to the reference genome using Samtools 0.1.19 package. 5. Proceed to a specific peak-calling step using the PePr V1.1.21 package by calculating reads ratios between IP and input in a comparative mode where only peak signals detected simultaneously in the two replicates are considered. The piled-up peak signals obtained for the perturbed condition (+Rho) and non-perturbed condition (Rho) are subsequently subjected to a step of differential analysis where only peaks appearing for one condition but not the other are scored (see Note 16).

4

Notes 1. Each strain carrying a C-terminally MYC-tagged protein for the ChIP experiments was constructed by integration of appropriate DNA fragments encoding the Tag at the protein gene locus following homologous recombination. The DNA fragments were obtained by PCR amplification with appropriate primers on plasmid template pYM19 [18]. The strains can be obtained from us upon request. Alternatively, similar strains derived from the wild-type background strain BY4741 are available from the EUROSCARF yeast collection (http:// www.euroscarf.de/). 2. The synthetic complete medium is without tryptophan to maintain within the yeast cells the centromeric plasmid pCM185-Rho-NLS that has the TRP1 marker. 3. Stock solutions of PMSF in 100% isopropanol do not need any sterilization, and they are stable for at least 9 months at 15–25  C. 4. Upon addition of 1:100 volume of 0.1 M PMSF, the final working concentration of PMSF is 1 mM. 5. We use the GeneJET PCR Purification Kit (Thermo Fischer), but other brands such as QIAquick PCR Purification Kit (Qiagen) work as well. 6. Depending on what is available, use either a probe sonicator device while maintaining the sample on ice or use a water bath sonicator such as the “Bioruptor (Diagenode), which has the advantage of keeping the samples at 4  C during sonication. 7. We recommend to have an early contact with the NGS outsourcing platform to discuss the specific needs (single end reads or paired reads, number of reads per library), the experimental design, and the schedule for the sequencing operations.

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8. Cell washing is an important step to remove all the residual Doxy and thus to allow full expression of Rho during growth for the induced samples. 9. The difference of the growth density between the cells grown under repressing conditions and those grown under inducing conditions results from the growth defect induced by Rho expression. 10. All the bioinformatics data processing packages are freely available and can be downloaded from respective web sites and run with Linux Ubuntu operating system. Alternatively, the Galaxy webserver (https://usegalaxy.org/) which is under a graphical user interface can also be used for data processing. But this alternative shows less flexibility in terms of option and data manipulations. 11. From this step, try to work on ice so the samples are kept as close as possible to 4  C. 12. If necessary for timing purpose, the processing of the samples can be stopped at this step by storing the cell pellets at 80  C. 13. As specified in Note 6, the shearing of the chromatin can be done with a Bioruptor or a usual sonicator with a slim probe that fits into a 2 mL tube. With our sonicator device (Autotune Model 750 watts), we use the following parameters: 20% amplitude, 7 s of pulse ON, and 3 s of pulse OFF for a total time of 4 min with the probe diving at 2/3 inside the microtube which is maintained on ice. These parameters should be calibrated for each specific sonication device by analyzing the size of the DNA fragments on agarose gel. 14. It is good to know that this kit can be used for a broad range of initial DNA quantity (from 500 pg to 1 μg). Another advice when using this kit is to avoid numerous PCR amplification cycles as far as possible in order to circumvent PCR duplicates that can make analysis bias during bioinformatics processing of the sequencing data. 15. All the bioinformatics data processing packages are freely available and can be downloaded from respective web sites. Note that the data processing of the ChIP-seq results can also be run on the Galaxy webserver (https://usegalaxy.org/) as for RNAseq. 16. Peak calling using the PePr V1.1.21 package allows the analysis of duplicates directly and in a comparative mode when needed. This differential analysis reveals clearly Rho-mediated shifts of recruitment, and, thus, it gives a snapshot of the dynamic landscape of the QC components. Manipulations on peak caller outputs are done mostly within R environment (V 3.4.4). Figures are built with ggplot2 package, and Circos plots are

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created using OmicCircos package. All snapshots used for direct visualization of reads coverage or ChIP peaks distributions were taken with the IGV (Integrative Genomics Viewer) browser. References 1. Bentley DL (2014) Coupling mRNA processing with transcription in time and space. Nat Rev Genet 15:163–175 2. Luna R, Gaillard H, Gonzalez-Aguilera C et al (2008) Biogenesis of mRNPs: integrating different processes in the eukaryotic nucleus. Chromosoma 117:319–331 3. Doma MK, Parker R (2007) RNA quality control in eukaryotes. Cell 131:660–668 4. Fasken MB, Corbett AH (2005) Process or perish: quality control in mRNA biogenesis. Nat Struct Mol Biol 12:482–488 5. Schmid M, Jensen TH (2013) Transcriptionassociated quality control of mRNP. Biochim Biophys Acta 1829:158–168 6. Rougemaille M, Gudipati RK, Olesen JR et al (2007) Dissecting mechanisms of nuclear mRNA surveillance in THO/sub2 complex mutants. EMBO J 26:2317–2326 7. Villa T, Rougemaille M, Libri D (2008) Nuclear quality control of RNA polymerase II ribonucleoproteins in yeast: tilting the balance to shape the transcriptome. Biochim Biophys Acta 1779:524–531 8. Mosrin-Huaman C, Honorine R, Rahmouni AR (2009) Expression of bacterial Rho factor in yeast identifies new factors involved in the functional interplay between transcription and mRNP biogenesis. Mol Cell Biol 29:4033–4044 9. Mosrin-Huaman C, Hervouet-Coste N, Le Dantec A, Stuparevic I, Rahmouni AR (2014) Bacterial Rho helicase: a new tool to dissect mRNP biogenesis and quality control in yeast. Trends Cell Mol Biol 9:79–93 10. Vasiljeva L, Kim M, Mutschler H et al (2008) The Nrd1-Nab3-Sen1 termination complex interacts with the Ser5-phosphorylated RNA polymerase II C-terminal domain. Nat Struct Mol Biol 15:795–804 11. Honorine R, Mosrin-Huaman C, HervouetCoste N et al (2011) Nuclear mRNA quality

control in yeast is mediated by Nrd1 co-transcriptional recruitment, as revealed by the targeting of Rho-induced aberrant transcripts. Nucleic Acids Res 39:2809–2820 12. Stuparevic I, Mosrin-Huaman C, HervouetCoste N et al (2013) Cotranscriptional recruitment of RNA exosome cofactors Rrp47p and Mpp6p and two distinct Trf-Air-Mtr4 polyadenylation (TRAMP) complexes assists the exonuclease Rrp6p in the targeting and degradation of an aberrant messenger ribonucleoprotein particle (mRNP) in yeast. J Biol Chem 288:31816–31829 13. Moreau K, Le Dantec A, Mosrin-Huaman C et al (2019) Perturbation of mRNP biogenesis reveals a dynamic landscape of the Rrp6dependent surveillance machinery trafficking along the yeast genome. RNA Biol 16:879–889 14. Baudin-Baillieu A, Guillemet E, Cullin C et al (1997) Construction of a yeast strain deleted for the TRP1 promoter and coding region that enhances the efficiency of the polymerase chain reaction-disruption method. Yeast 13:353–356 15. 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 16. Xu Z, Wei W, Gagneur J et al (2009) Bidirectional promoters generate pervasive transcription in yeast. Nature 457:1033–1037 17. Neil H, Malabat C, d’Aubenton-Carafa Y et al (2009) Widespread bidirectional promoters are the major source of cryptic transcripts in yeast. Nature 457:1038 18. Janke C, Magiera MM, Rathfelder N et al (2004) A versatile toolbox for PCR-based tagging of yeast genes: new fluorescent proteins, more markers and promoter substitution cassettes. Yeast 21:947–962

Chapter 17 Probing the Conformational State of mRNPs Using smFISH and SIM Srivathsan Adivarahan and Daniel Zenklusen Abstract mRNAs and lncRNAs assemble with RNA-binding proteins (RBPs) to form ribonucleoprotein complexes (RNPs). The assembly of RNPs initiates co-transcriptionally, and their composition and organization is thought to change during the different steps of an RNP life cycle. Modulation of RNP structural organization has been implicated in the regulation of different aspects of RNA metabolism, including establishing interactions between the 50 and 30 ends in regulating mRNA translation and turnover. In this chapter, we describe a single-molecule microscopy approach that combines fluorescent RNA in situ hybridization (smFISH) and structured illumination microscopy (SIM) and allows to measure different aspects of RNP organization in cells, including distances between different regions within individual mRNAs, as well as the overall compaction state of RNAs in different subcellular compartments and environmental conditions. Moreover, we describe a detailed workflow required for image registration and analysis that allows determining distances at sub-diffraction resolution. Key words smFISH, mRNA, mRNPs, lncRNAs, RNA organization, RNA structure, Single-molecule resolution microscopy, Structured illumination microscopy

1

Introduction The regulation of gene expression involves many steps and requires the interplay of RNA and RNA-binding proteins (RBPs). RNP organization and composition changes during the RNA lifecycle as RBPs associate with mRNAs at different steps of the gene regulation pathway [1, 2]. RNAs itself can form extensive local secondary structures as well as long-distance intramolecular interactions through Watson-Crick base pairing and many different nonWatson-Crick interactions that, in turn, can provide additional domains for the binding of RBPs [3, 4]. RBPs, aside from playing a crucial role in regulating many catalytic steps during RNA metabolism such as splicing, 30 processing or degradation, act as important structural components of the RNP contributing to their organization as 3D assemblies. RBP binding can influence RNP

Marc Boudvillain (ed.), RNA Remodeling Proteins: Methods and Protocols, Methods in Molecular Biology, vol. 2209, https://doi.org/10.1007/978-1-0716-0935-4_17, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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organization either by stabilizing or destabilizing local secondary structures or by allowing additional short and long-distance intramolecular interactions with other RBPs through their homo- and hetero-dimerization domains [5]. Progression through the different steps of RNA metabolism involves frequent RNP reorganization. Among the best-studied long-distance rearrangement is establishing the communication between the 50 and 30 ends within mRNAs. During translation, the 50 and 30 ends of the mRNA are thought to be brought together through interactions between the cytoplasmic cap-binding protein eIF4E, the adapter protein eIF4G, and the poly(A) binding protein PABC1. In addition to stimulating translation, these interactions could potentially play a role in regulating RNA decay by bringing the poly(A) tail and cap in close proximity [6, 7]. In contrast to the detailed view of the factors participating in 50 –30 communications that were largely obtained through in vitro and in vivo biochemical and genetic approaches, few current methods exist that allow to directly interrogate such RNP (re-) organization in cells. RNP organization has been studied using different approaches, with X-ray crystallography, NMR and cryo-electron microscopy playing crucial roles in constructing high-resolution structures of RNPs at different scales, from tRNAs to the ribosome. However, aside from ribosomes that have a well-defined 3D structure, neither high-resolution structures nor principles that define the 3D organization of large RNPs have been investigated [5, 8, 9]. In particular, much less is known on the overall structural organization of mRNAs and lncRNAs, thought to be more difficult to study using these structural approaches due to the presumption that these molecules are either less well defined as 3D assemblies and/or get frequently rearranged during and while transiting between different steps of the gene regulation pathway [9]. Despite these difficulties, numerous in vivo and in vitro approaches have been developed to interrogate RNP organization, some focused on determining local secondary structures using various chemical probing approaches, and others on mapping local as well as longrange contacts mediated by protein-RNA, protein-protein, or RNA-RNA interactions [10–12]. Microscopy approaches provide another powerful tool to study RNP organization in cells [13–23]. Here, we describe a single-molecule imaging approach that combines single-molecule resolution in situ hybridization (smFISH) with structured illumination microscopy (SIM) to study RNA organization in fixed cells. The method makes use of the ability to localize signals emitted from single molecules with sub-diffraction resolution using 2D or 3D Gaussian fitting, allowing for a localization precision of around 20 nm using the setup described here. RNAs are detected using 20 nucleotide (nt) long DNA probes, each labeled with a single fluorescent dye and hybridized to RNA in paraformaldehyde fixed

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Fig. 1 Schematic illustrating the imaging and image analysis workflow used to measure conformational states and compaction of single mRNPs using smFISH and structured illumination microscopy

tissue culture cells. Probes hybridizing to different regions labeled with fluorophores that allow spectral separation are then used to measure intramolecular distances within induvidual RNAs. Moreover, using probes targeting multiple regions within an mRNA further allows determining the overall compaction state of RNAs [20]. However, accurately measuring intramolecular distances requires careful image registration and a stable microscopy setup. We describe different aspects of the imaging and image analysis workflow required to achieve such measurements (Fig. 1).

2

Materials Prepare all reagents and solutions using ultrapure water and analytical grade reagents. Store all reagents at room temperature unless stated otherwise.

2.1

Chemicals

1. Ethanol. 2. Bovine Serum Albumin (BSA). 3. Formamide. 4. Puromycin dihydrochloride. 5. Cycloheximide. 6. Sodium arsenite. 7. Homoharringtonine. 8. 50% (w/v) dextran sulfate solution. 9. 32% (w/v) paraformaldehyde (PFA) solution in H2O, methanol free and RNase free.

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10. 200 mM of ribonuclease inhibitor ribonucleoside vanadyl complexes (RVC). 11. Triton X-100. 12. Sodium chloride (NaCl). 13. Potassium chloride (KCl). 14. Sodium phosphate (Na2HPO4·7H2O). 15. Potassium dihydrogen phosphate (KH2PO4). 16. Sodium citrate dihydrate (HOC(COONa)(CH2COONa)2 · 2H2O). 17. 40 ,6-Diamidino-2-phenylindole dihydrochloride (DAPI). 18. 0.1% (w/v) Poly-L-Lysine in H2O. 19. Hydrochloric acid (HCl). 20. Sodium hydroxide (NaOH). 21. Sodium bicarbonate (NaHCO3). 22. E. coli tRNA (Roche). 23. 10 mg/mL salmon sperm DNA (ssDNA) solution. 24. Sterile (e.g., autoclaved) deionized water. 2.2 Buffers and Solutions

1. 10 PBS—Dissolve 80 g NaCl, 2 g KCl, 26.8 g Na2HPO4·7H2O, and 2.4 g KH2PO4 in 800 mL water. Adjust pH to 6.8 before making up the volume to 1 L. pH when diluted to 1 will be 7.4. 2. 20 SSC (3 M NaCl and 0.3 M sodium citrate)—Dissolve 175.3 g of NaCl and 88.2 g of sodium citrate dihydrate in 800 mL of H2O. Adjust the pH to 7.0 with HCl and make up the volume to 1 L. 3. Hybridization solution (10% (v/v) formamide, 2 SSC, 10% (w/v) dextran sulfate)—For 10 mL hybridization solution, add 1 mL of 100% Formamide, 1 mL of 20 SSC, and 2 mL of 50% dextran sulfate. Make up the volume to 1 mL with deionized water. Aliquot and store at 20  C. 4. Washing solution (10% Formamide, 2 SSC)—For 10 mL, add 1 mL of 100% formamide and 1 mL of 20 SSC. Make up the volume to 10 mL with deionized water. Prepare fresh every time. 5. Labeling solution (0.1 M sodium bicarbonate, pH ¼ 8.3)—To 84.01 mg of sodium bicarbonate, add 9 mL of deionized water. Adjust pH with NaOH/HCl if necessary, to 8.3. Make up the volume to 10 mL. Aliquot and store at 20  C. 6. 70% ethanol—diluted in deionized water. 7. 1 PBS—10 PBS diluted in deionized water. 8. 5% (w/v) BSA stock solution.

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9. Permeabilization solution (0.1% (v/v) Triton X-100, 0.5% (w/v) BSA and 1 PBS)—For 50 mL, add 50 μL of Triton X-100, 5 mL of 5% BSA stock solution, and 5 mL of 10 PBS to a 50 mL tube, and make up the volume to 50 mL with deionized water. 2.3 General Laboratory Equipment

1. Fine-tip forceps. 2. Aluminum foil. 3. Parafilm. 4. 37  C incubator. 5. Fume hood. 6. Heating blocks for tubes or a thermocycler. 7. Sterile 0.2 μm syringe filters. 8. Syringes. 9. Speed vacuum centrifuge. 10. UV–vis spectrophotometer. 11. Kimwipes.

2.4

Cell Culture

1. Culture medium: Dulbecco’s modified Eagle’s medium (DMEM) with 10% (v/v) fetal bovine serum. 2. Cultured adherent cell lines grown and passaged regularly (two to three times a week) using trypsin. 3. 12 well plates. 4. Circular 18 mm diameter #1.5 thickness cover glass (Electron Microscopy Sciences). 5. 0.01% (w/v) Poly-L-Lysine in deionized water. Prepare fresh and sterilize with a 0.2 μm syringe filter. 6. Fixation Solution (4% (v/v) PFA and 1 PBS)—For 10 mL, add 1.25 mL of 32% PFA solution and 1 mL of 10 PBS, and make up the volume to 10 mL using deionized water. 7. (Optional) Concentrated drug solution and drug solvent. The concentration of the drug stock will depend on the solubility and working concentration of the tested drug e.g., 50 mg/mL in water for puromycin to inhibit translation by ribosomal removal (working concentration 100 μg/mL), 5 mg/mL in ethanol for cycloheximide to inhibit translation by ribosome stalling (working concentration 100 μg/mL), 10 mg/mL in DMSO for Homoharringtonine to inhibit translation initiation (working concentration 100 μg/mL), and 50 mM in water for sodium arsenite to induce oxidative stress (working concentration 2 mM)).

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2.5 Sample Preparation

1. smFISH probes: From 30 to 48 distinct 20mer DNA oligonucleotides with a 30 amine modification designed using Stellaris RNA FISH probes designer. 2. Succinimidyl (NHS) ester form of dye of interest. For far-red channel, Cy5 or Alexa647 dye; red/orange channel, Cy3 or Dylight 550; green channel, Dylight 488 or Atto488 dye. 3. Qiagen Nucleotide removal kit (Qiagen). 4. 100 nm TetraSpeck™ beads suspension (Thermo Fisher Scientific). 5. 5 mg/mL DAPI stock solution in deionized H2O. Store at 20  C. 6. DAPI working solution: 2.5 μg/mL DAPI in 1 PBS, freshly prepared for each experiment and stored in dark. 7. Clean glass microscopy slides. 8. Antifade mountant (e.g., Prolong Gold, Prolong Diamond, Prolong Glass). 9. Transparent nail polish. 10. 10 mg/mL E. coli tRNA solution. Store at 20  C. 11. ssDNA/tRNA mix—5 mg/mL ssDNA and 5 mg/mL tRNA—mix 500 μL of ssDNA solution with 500 μL of tRNA solution. Store at 20  C.

2.6 Microscope Setup

2.7 Image Analysis and Data Representation

1. Zeiss Elyra PS1 imaging system used for image acquisition in the experiments described below is equipped with 50 mW 405 nm, 100 mW 488 nm, 100 mW 561 nm, and 150 mW 642 nm lasers and four emission filters— (1) BP420–480 + LP750, (2) BP495–590 + LP750, (3) LP570, and (4) LP655, a 63 1.4 NA Plan Apo DIC II oil objective with a working distance of 0.1 mm, EMCCD camera (Andor iXon3 DU-885 K with a chip size of 1004  1002 pixels and pixel size of 8 μm), a piezo stage for high precision focus in Z and step size of 25 nm, and a high performance computer workstation for imaging, image processing, and image analysis. The microscope is stored in a temperature-controlled room to minimize temperature fluctuations. Software required: 1. ZEN black 2012 SP5 (ver14.0.20.201 from Zeiss). 2. FIJI (ImageJ). 3. IDL virtual machine and Localizeapp.sav (ver14.2, see Note 17). 4. AIR Localize (ver1.4, see Note 18).

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5. MATLAB (ver9.3 or newer). 6. R Studio for plotting.

3

Methods

3.1 Probe Designing and Labeling

Depending on the length of the mRNA of interest, probes can be designed against two or more regions within the mRNA. Probes targeting the different regions have to be labeled with dyes that can be spectrally separated using a specific microscope setup (see Note 1). 1. To design probes against multiple regions, choose a ~1000 nt region within the mRNA as template and use the Stellaris probe designer tool to design 30–48 probes against the target. Once designed, probes can either be ordered conjugated with fluorescent dyes (steps 2–7 are not required) or containing a single amine modification at the 30 end that allows post-synthesis labeling with different fluorophores (see Note 2). An alternative protocol for probe labeling as described in Gaspar et al. can also be used [24]. 2. For post-synthesis labeling, the probes corresponding to one target region are pooled in equimolar amounts prior to labeling. Pooled probes can be stored at 20  C for many months. We have efficiently labeled probes stored for several years. 3. The pooled probes can be used for labeling using NHS ester dyes. 4. NHS ester dyes can be purchased as value packs (available as Mono 5-pack for Cyanine dyes, 5  50 μg packs for Dylight dyes), in which case one vial can be used for labeling 20 μg of probes. Alternatively, dyes can be aliquoted to equivalent concentrations as described in [24] (see Note 3). 5. Use one vial of NHS ester dye pack or equivalent aliquoted dye to label 2  10 μg of probes. Aliquot 10 μg (~1–1.2 nmoles) of probes, and dry the probe mix using a speed vacuum evaporator. Resuspend one vial of dye (~50 nmoles) in 30 μL of labeling solution and add 15 μL of the resuspended dye to the probes. Incubate o/n in the dark. 6. Remove unincorporated dye using a nucleotide removing spin column such as Qiagen nucleotide removal kit, following manufacturer’s instructions. Alternatively, probes can also be purified using a size exclusion column as described in [24]. 7. Use a UV-vis spectrophotometer to measure the concentration, and determine the labeling efficiency to ensure a high labeling efficiency for the probes. If the labeling is low ( SingerLab). Save the mask in the same folder as the other images (see Note 22). 6. Repeat steps 3–5 for nuclear mRNAs. 7. Open “LocalizeApp.sav” with IDL Virtual machine, and use it to determine the position of RNA spots in 2D. Adjust the parameters such that only true signals are detected. Compare with the original image to determine the quality of localization. The coordinates of the localized RNAs are saved as “.loc” files. 8. Repeat for all channels. 9. Open MATLAB and navigate to the parent folder containing subfolders, each containing the nuclear and cytoplasmic masks as well as the file containing the coordinates from the localized RNA spots in all channels. 10. To measure colocalization precision, use the script “RNAloc. m.” Distribution of colocalization precision in 2D measurements can be seen in Fig. 2a (right panel). To calculate distance distributions of two color images, run “RNA2.m.” For instance, 50 –30 distance distribution for cytoplasmic MDN1 mRNAs in mock, puromycin, and sodium arsenite treatment is shown in Fig. 2b, c (right panels). For three color images, run “RNA3.m.” Distribution of 2D distances between different regions of cytoplasmic MDN1 mRNAs for mock, homoharringtonine, and puromycin treatment is shown in Fig. 3b.

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11. The scripts will analyze the data and save the distance information in two files—“Nuclear Distances.csv” and “Cytoplasmic Distances.csv” within these subfolders. 12. In addition, “RNA3.m” also saves the coordinates of the spots corresponding to an individual mRNA in two different files— “Cytoplasmic Spots.csv” and “Nuclear Spots.csv.” These coordinates can be used to determine the radius of gyration (see below and Fig. 3c). 3.5.2 mRNP Conformation Analysis in 3D

1. Open 3D images in ImageJ and split the images according to the channels and save them as done for 2D analysis (see Note 21). 2. In ImageJ, make nuclear and cytoplasmic masks as shown previously for 2D image analysis (see Subheading 3.5.1 above). 3. Open MATLAB and run “AIRLOCALIZE.m”. Set the parameters that correpsond to the imaging conditions and microscopy setup. Open 3D stacks for each individual channel with RNA spots, and determine the 3D coordinates of the signals using AIRLOCALIZE. The localized spots are saved in “.loc3” files. Repeat for all other channels. 4. Repeat the process for all images to be quantified. 5. Navigate to the parent folder within MATLAB that contains subfolders, similar to when analyzing 2D images. 6. Run “RNA2_3D.m.” This will calculate distances between RNAs within the masks and save the results in two files – “Cytoplasmic Distances.csv” and “Nuclear Distances.csv.” 3D distance distribution for MDN1 50 –30 distances are shown in Fig. 2b (right panel). To analyze data related to measuring colocalization precision in 3D, use the script ‘RNAloc_3D. m’. Distribution of colocalization precision in 3D measurements can be seen in Fig. 2a (right panel).

3.5.3 Estimating Parameters of mRNP Compaction and Performing Statistical Analysis

Compaction levels of mRNAs can be determined when relative position of three or more regions of an mRNA is known. We use our three-color imaging dataset to determine mRNP compaction using the factor—“mean radius of gyration.” The mean radius of gyration (hRgi) is calculated as follows: vffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffi u 3 X   u Rg ¼ t1=3 ðr k  r mean Þ2 k¼1

where k represents one of the three regions of the mRNP and rk the position of the corresponding region in space determined by Gaussian fitting.

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1. Use the coordinates of different regions of mRNPs saved in the files “Cytoplasmic Spots.csv” and “Nuclear Spots.csv” (see above) and the above formula to calculate the mean radius of gyration. 2. Determine the statistical significance of changes in distance distributions between regions of an mRNA with the Kolmogorov-Smirnov test using the function “ks.test” in R.

4

Notes 1. It is important to take into account the expression levels and isoforms for the investigated transcripts. If the expression level of the mRNA of interest is very high, it becomes challenging to spatially separate individual mRNAs and therefore difficult to determine their individual conformations. Furthermore, if mRNAs have multiple isoforms, it is important to consider which of the isoforms are expressed in a particular cell line and to ensure that probes are specific to and target the isoforms that need to be studied. 2. The number of probes is an important consideration to get a good RNA signal, which in turn is essential to achieve a high localization precision. The number of probes that can be designed to hybridize to a specific region depends on the length and composition of the target sequence. We use a probe design software from LGC Biosearch Technologies called Stellaris Probe designer, which searches for probes of 20 nt in length and a GC content of ~35–55%. Moreover, it excludes repetitive sequences within genomes of different organisms to ensure specificity. Other oligo design tools such as the one described in Tsanov et al. can also be used, but we have predominantly used this software and obtained good results [29]. Stellaris Probe designer recommends the use of a minimum of 25 probes for a single-target sequence, though in our hands we have obtained good RNA signal with as little as 15 probes using our imaging setup for probes labeled in the red and far-red channels. Length and spacing (we use a spacing of 2nt as default) between probes can also be altered if needed. Due to higher autofluorescence in the green channel, we found that we needed at least 35 probes in the green channel to achieve acceptable signal-to-noise and localization precisions. 3. Aliquoting the dyes must be done in a moisture-free environment and using anhydrous DMSO. This is to preserve the reactivity of the dye for longer periods. Aliquoted dyes in DMSO can be stored at 80  C.

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4. We have not noticed an observable loss in FISH signal using aliquots of labeled FISH probes thawed multiple times over a short period of time. However, it is recommended that if the probes are to be used over longer periods, they be aliquoted and stored at 20  C. 5. It is important to be careful while pipetting as some cell lines are prone to detaching from the coverslips when pipetted directly on the face of the coverslip. 6. Ensure that all steps involved in handling of labeled probes or dyes are done with minimal light exposure. 7. Addition of BSA is optional. We have not observed differences in signal when omitting BSA in the buffer. 8. The Z stack interval was optimized to get the minimal interval that resulted in highest localization precision for our imaging setup. Moreover, the number for the z stacks was chosen to cover a considerable volume of the cell while keeping acquisition times low (see also Note 10). 9. By default, the grid sizes allocated to the lasers are different. For the 405 nm, it is 23 μm; for the 488 nm laser, it is 28 nm; for the 561 and 642 nm lasers, it is 34 μm. However, in our hands, we found that the best signal and localization were achieved for the 42 μm grid for all lasers. 10. It is important to ensure that the acquisition time is kept to a minimum. We have noticed that longer acquisitions due to either increased number of z stacks or increased exposure time resulted in a decrease in colocalization precision. This is mostly likely due to mechanical or thermal drift that the sample experiences during acquisition. 11. A high NA objective with a very small point spread function (PSF) is preferable for imaging to get the best results. The PSF of the objective should be measured regularly using 100 nm TetraSpeck beads, and this PSF can be used for reconstruction of the 3D SIM image. Regular measurement of the PSF also helps avoid artifacts during reconstruction due to possible defects in the objective. 12. Registration with beads should be performed every day of imaging. Although the microscope system is relatively stable for short periods of time, in our experience, the parameters for alignment of channels can change over time. 13. Image processing using the 3D SIM option is essential when analyzing the images in 3D. However, images can be processed using the 2D-SIM option, which does not yield a high z resolution but provides similar resolution in xy. Using the experimentally measured PSF did not yield in a higher localization precision when compared to using the option of a

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theoretical PSF when processing images. However, this could vary depending on the quality of the objective. 14. TetraSpeck beads are not very bright when illuminated with a 405 nm laser, and as this channel is essentially used for nuclear marker, it is skipped when performing registration. 15. We chose two different regions of MDN1 (each around 1.2–1.5 kb in length) to test for precision of localization (or “colocalization precision”). We did not observe a difference in colocalization precision between the two regions tested. However, this might not apply to longer sequences which correspond to regions of mRNAs that are no longer diffraction limited. 16. Although the number of mRNAs can vary based on transcript or cell line of choice, for many of the mRNA studied, 50 cells gave us at least 500 RNAs. Quantifying data from more than 500 mRNAs did not significantly alter our observations when measuring mRNA intramolecular distances. However, for low abundant RNAs or the cell type, this number might sometimes be difficult to achieve and require imaging many more cells. 17. “LocalizeApp.sav” is freely available from Dan Larson’s lab website (https://ccr.cancer.gov/Laboratory-of-Receptor-Biol ogy-and-Gene-Expression/daniel-r-larson). IDL Virtual Machine can be downloaded by following instructions from here: https://www.harrisgeospatial.com/Support/Self-HelpTools/Help-Articles/Help-Articles-Detail/ArtMID/10220/ ArticleID/17309/The-IDL-Virtual-Machine. 18. AIRLocalize is available from Timothee Lione’s lab website (http://www.timotheelionnet.net/software/). In addition to AIRLocalize and LocalizeApp, other software packages allow 2D and 3D quantifications, such as FISH-Quant [28]. 19. The plugin was developed in Robert Singer’s laboratory and can be downloaded from [https://github.com/zenklusenlab/ ImageJ_plugins]. The plugin creates a mask using all the ROI in the ROI Manager of ImageJ. 20. It is assumed that the user has a minimal understanding of MATLAB. All the MATLAB scripts required for data quantification and example datasets for data analysis can be found here: https://github.com/zenklusenlab/MolCell_DistanceCalc. 21. The scripts rely on a predefined folder structure and nomenclature of different input files. The scripts are designed to run and analyze data within subfolders and save the output files within the same subfolders. Each subfolder, for instance, contains all the information relevant for data quantification of one field – the localization (“.loc” or “.loc3” files, the nuclear and cytoplasmic masks—also see Note 22). For two-color images,

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the image files corresponding to RNA FISH signal should be saved as “Cy5.tif” and “Cy3.tif” (and the corresponding localization files as “Cy5.loc” and “Cy3.loc”). For three-color images, the image files should be saved as “5p.tif,” “3p.tif,” and “mid.tif,” where 5p, 3p, and mid correspond to RNA signal for the 50 end, 30 end, and middle region, respectively. Example datasets illustrating the folder structure and file names can be found here: https://github.com/zenklusenlab/MolCell_DistanceCalc/tree/master/Example%20Files 22. The mask files should be saved as “Cymask.tif” and “Nucmask. tif” (for cytoplasmic and nuclear masks, respectively) and should be saved in the same subfolder as other data from the same field of view. References 1. Mitchell SF, Parker R (2014) Principles and properties of eukaryotic mRNPs. Mol Cell 54:547–558. https://doi.org/10.1016/j. molcel.2014.04.033 2. Hentze MW, Castello A, Schwarzl T, Preiss T (2018) A brave new world of RNA-binding proteins. Nat Rev Mol Cell Bio 19:327. https://doi.org/10.1038/nrm.2017.130 3. Ganser LR, Kelly ML, Herschlag D, Al-Hashimi HM (2019) The roles of structural dynamics in the cellular functions of RNAs. Nat Rev Mol Cell Biol 20. https://doi.org/ 10.1038/s41580-019-0136-0 4. Lu Z, Chang HY (2016) Decoding the RNA structurome. Curr Opin Struc Biol 36:142–148. https://doi.org/10.1016/j.sbi. 2016.01.007 5. Singh G, Pratt G, Yeo GW, Moore MJ (2015) The clothes make the mRNA: past and present trends in mRNP fashion. Annu Rev Biochem 84:1–30. https://doi.org/10.1146/annurevbiochem-080111-092106 6. Pelletier J, Sonenberg N (2019) The organizing principles of eukaryotic ribosome recruitment. Annu Rev Biochem 88:307–335. https://doi.org/10.1146/annurev-biochem013118-111042 7. Vicens Q, Kieft JS, Rissland OS (2018) Revisiting the closed-loop model and the nature of mRNA 50 –30 communication. Mol Cell 72:805–812. https://doi.org/10.1016/j. molcel.2018.10.047 8. Jones S (2016) Protein–RNA interactions: structural biology and computational modeling techniques. Biophys Rev 8:359–367. https://doi.org/10.1007/ s12551-016-0223-9

9. Dimitrova-Paternoga L, Jagtap P, Chen P-C, Hennig J (2019) Integrative structural biology of protein-RNA complexes. Structure 28:6–28. https://doi.org/10.1016/j.str. 2019.11.017 10. Mitchell D, Assmann SM, Bevilacqua PC (2019) Probing RNA structure in vivo. Curr Opin Struc Biol 59:151–158. https://doi.org/ 10.1016/j.sbi.2019.07.008 11. Wheeler EC, Nostrand EL, Yeo GW (2017) Advances and challenges in the detection of transcriptome-wide protein-RNA interactions. Wiley Interdiscip Rev Rna 9:e1436. https:// doi.org/10.1002/wrna.1436 12. Zhao J, Qian X, Yeung P et al (2019) Mapping in vivo RNA structures and interactions. Trends Biochem Sci 44:555–556. https://doi. org/10.1016/j.tibs.2019.01.012 13. Bjo¨rk P, Wieslander L (2015) The Balbiani ring story: synthesis, assembly, processing, and transport of specific messenger RNA–protein complexes. Annu Rev Biochem 84:65–92. https://doi.org/10.1146/annurev-biochem060614-034150 14. Christensen KA, Kahn LE, Bourne CM (1987) Circular polysomes predominate on the rough endoplasmic reticulum of somatotropes and mammotropes in the rat anterior pituitary. Am J Anat 178:1–10. https://doi.org/10. 1002/aja.1001780102 15. Christensen KA, Bourne CM (1999) Shape of large bound polysomes in cultured fibroblasts and thyroid epithelial cells. Anatomical Rec 255:116–129. https://doi.org/10.1002/( sici)1097-0185(19990601)255:23.0.co;2-o

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16. Brandt F, Carlson L-A, Hartl UF et al (2010) The three-dimensional organization of polyribosomes in intact human cells. Mol Cell 39:560–569. https://doi.org/10.1016/j. molcel.2010.08.003 17. West JA, Mito M, Kurosaka S et al (2016) Structural, super-resolution microscopy analysis of paraspeckle nuclear body organization. J Cell Biol 214:817–830. https://doi.org/10. 1083/jcb.201601071 18. Yamazaki T, Souquere S, Chujo T et al (2018) Functional domains of NEAT1 architectural lncRNA induce paraspeckle assembly through phase separation. Mol Cell 70:1038–1053 . e7. https://doi.org/10.1016/j.molcel.2018.05. 019 19. Mor A, Suliman S, Ben-Yishay R et al (2010) Dynamics of single mRNP nucleocytoplasmic transport and export through the nuclear pore in living cells. Nat Cell Biol 12:543. https:// doi.org/10.1038/ncb2056 20. Adivarahan S, Livingston N, Nicholson B et al (2018) Spatial organization of single mRNPs at different stages of the gene expression pathway. Mol Cell 72:727–738 . e5. https://doi.org/ 10.1016/j.molcel.2018.10.010 21. Adivarahan S, Zenklusen D (2019) Lessons from (pre-)mRNA imaging. Adv Exp Med Biol 1203:247–284. https://doi.org/10. 1007/978-3-030-31434-7_9 22. Khong A, Parker R (2018) mRNP architecture in translating and stress conditions reveals an ordered pathway of mRNP compaction. J Cell Biol 217:201806183. https://doi.org/10. 1083/jcb.201806183 23. Koch A, Aguilera L, Morisaki T et al (2020) Quantifying the spatiotemporal dynamics of

IRES versus Cap translation with singlemolecule resolution in living cells. Biorxiv 2020(01):09.900829. https://doi.org/10. 1101/2020.01.09.900829 24. Ga´spa´r I, Wippich F, Ephrussi A (2018) Terminal deoxynucleotidyl transferase mediated production of labeled probes for single-molecule FISH or RNA capture. Bio-protocol 8. https://doi.org/10.21769/bioprotoc. 2750 25. Zenklusen D, Larson DR, Singer RH (2008) Single-RNA counting reveals alternative modes of gene expression in yeast. Nat Struct Mol Biol 15:1514. https://doi.org/10.1038/nsmb. 1514 26. Thompson RE, Larson DR, Webb WW (2002) Precise nanometer localization analysis for individual fluorescent probes. Biophys J 82:2775–2783. https://doi.org/10.1016/ s0006-3495(02)75618-x 27. Lionnet T, Czaplinski K, Darzacq X et al (2011) A transgenic mouse for in vivo detection of endogenous labeled mRNA. Nat Methods 8:165. https://doi.org/10.1038/nmeth. 1551 28. Mueller F, Senecal A, Tantale K et al (2013) FISH-quant: automatic counting of transcripts in 3D FISH images. Nat Methods 10:277. https://doi.org/10.1038/nmeth.2406 29. Tsanov N, Samacoits A, Chouaib R, et al (2016) smiFISH and FISH-quant - a flexible single RNA detection approach with superresolution capability. Nucleic Acids Res 44: e165. https://doi.org/10.1093/nar/ gkw784

Chapter 18 Probing Transcriptome-Wide RNA Structural Changes Dependent on the DEAD-box Helicase Dbp2 Yu-Hsuan Lai and Elizabeth J. Tran Abstract RNA helicases function in all aspects of RNA biology mainly through remodeling structures of RNA and RNA-protein (RNP) complexes. Among them, DEAD-box proteins form the largest family in eukaryotes and have been shown to remodel RNA/RNP structures and clamping of RNA-binding proteins, both in vitro and in vivo. Nevertheless, for the majority of these enzymes, it is largely unclear what RNAs are targeted and where they modulate RNA/RNP structures to promote RNA metabolism. Several methods have been developed to probe secondary and tertiary structures of specific transcripts or whole transcriptomes in vivo. In this chapter, we describe a protocol for identification of RNA structural changes that are dependent on a Saccharomyces cerevisiae DEAD-box helicase Dbp2. Experiments detailed here can be adapted to the study of other RNA helicases and identification of putative remodeling targets in vivo. Key words Secondary structure, RNA, Helicase, Sequencing, Mapping, Genome-wide

1

Introduction An RNA molecule can often be folded into multiple thermodynamically favored conformations. However, for an RNA to perform its function properly, formation of other non-functional, yet stable, structures has to be prevented or resolved. Potential candidates for solving this RNA folding problem in vivo are RNA chaperones, including RNA helicases, which can bind and remodel RNAs and macromolecular complexes of RNA with proteins (RNPs) [1, 2]. Consistently, one study suggested that ATP-dependent factors, such as RNA helicases, maintain the actively unfolded state of mRNAs in vivo [3]. Therefore, to better understand the biological function of RNA structures and their regulators, identification of the enzymatic targets of RNA helicases becomes the next critical step in the field. DEAD-box proteins are a family of RNA helicases found in all kingdoms of life, with 4 members in E. coli, 26 in Saccharomyces cerevisiae, and 37 in humans [4]. They are characterized by the

Marc Boudvillain (ed.), RNA Remodeling Proteins: Methods and Protocols, Methods in Molecular Biology, vol. 2209, https://doi.org/10.1007/978-1-0716-0935-4_18, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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presence of the 12 conserved motifs in the core [5] and are named by the Asp-Glu-Ala-Asp (D-E-A-D) sequence within motif II. Some members have additional N-terminal or C-terminal domains flanking the core, which provide specificity and/or regulation of these enzymes [6]. Many DEAD-box helicases are involved in steps of gene expression, including transcription, premRNA splicing, export, translation, and RNA decay [7]. Importantly, several human DEAD-box proteins have also been implicated in a variety of diseases and cancer development [8, 9]. Most DEAD-box helicases have an ATP-dependent RNA-unwinding activity in vitro [5]. Some also display ATP-independent RNA annealing [10], an activity proposed to facilitate exchange between RNA conformations [11]. Although non-processive, the duplex unwinding activity has been demonstrated to be functional in several RNA processing steps. For example, Mss116 in S. cerevisiae serves as an RNA chaperone, assisting the folding of functional group I and II introns by disrupting misfolded structures and promoting formation of alternative conformations [12– 14]. DEAD-box helicases, including DDX5 and DDX17, have also been reported to unwind secondary RNA structures in pre-mRNA and thereby regulate alternative splicing and/or miRNA processing [15–17]. Several DEAD-box helicases have also been shown to resolve structures in the 50 UTR of mRNAs to facilitate the initiation of ribosome scanning and translation [18– 20]. These examples suggest that the RNA remodeling activity plays a critical role in gene expression, yet the enzymatic targets of the majority of DEAD-box family remain uncharacterized. Not until 5 years ago was the first genome-wide RNA structure probing method inside cells reported [3, 21, 22]. Since then, several studies have developed methods to map secondary structures in various RNA species and regions in cells from different organisms, including yeast, plant, and humans [23–27]. Methods for mapping intracellular RNA structures can be predominantly categorized into two major groups based on the modification chemistry: chemical modification by dimethyl sulfate (DMS) and selective 20 -hydroxyl acylation analyzed by primer extension (SHAPE) [28]. DMS is a membrane-permeable chemical that methylates accessible adenines and cytosines in an RNA molecule that are not actively engaged in Watson-Crick base pairing [28]. By probing the reactivity of nucleotides to DMS in a given RNA, the level of base pairing can be inferred at single-nucleotide resolution [29, 30]. Because of its ability to rapidly penetrate into cells, DMS probing of RNA structures has been widely applied to cells from various organisms, including bacteria, yeast, plants, and humans [31]. However, DMS-based method can only provide information of As and Cs; therefore, structural mapping can be limited for regions with few or no A/C nucleotide. In SHAPE-based methods, accessible ribonucleotides are acylated by SHAPE reagents at the

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20 hydroxyl position in the ribose, which usually happens in flexible, single-stranded regions of RNAs [32]. Because all four nucleotides can be modified by SHAPE reagents, SHAPE-based methods provide a comprehensive mapping of cellular RNA structures. Nevertheless, they have been applied to only a limited number of cell types and have not been shown to be successful for RNA structural probing in yeast [27]. The modifications produced by DMS or SHAPE reagents can lead to reverse transcription stops or mutations in the cDNAs during library preparation. Consequently, sites of modification can be identified by the location of adaptor ligation or nucleotide conversion in sequences. Thus, the level of modification at each nucleotide reflects the state of RNA folding and structures. To date, only a few genome-wide studies investigated the impact of DEAD-box helicases on cellular RNA structures. One such example is the recent study of the role of Ded1, a DEAD-box protein required for translation initiation, in RNA structure remodeling and translation efficiency in S. cerevisiae [20]. By comparing mRNA structure profiles of the wild type and ded1 mutant, it was found that loss of Ded1 helicase activity leads to a striking change in RNA accessibility in the 50 UTR as compared to other regions and that the regions of structural changes align with the sites of Ded1 binding [20]. These results collectively indicate that Ded1 facilitates translation initiation through unwinding of mRNA structures in 50 UTRs. This paradigm illustrates that combination of structural probing with other genome-wide methods provides rich information on the enzymatic targets and molecular mechanisms of DEAD-box helicases inside cells. Our lab used DMS to map RNA structures in wild type and dbp2Δ to pinpoint DBP2-dependent changes [33]. The regions of DBP2-dependent structural alteration were also well correlated with Dbp2-binding sites identified and aligned well just upstream of annotated polyadenylation sites. This provided a putative model for how a DEAD-box protein may remodel RNA/RNP structures transcriptome wide in vivo. Here, we describe the method for probing Dbp2-dependent RNA structural changes using DMS-seq in S. cerevisiae. This protocol is largely based on the development of this technique [24], with additional optimizations to decrease undesired, self-ligated primers. This method can be further extended to studies of other RNA helicase targets in intact cells.

2

Materials

2.1 DMS Treatment of Yeast Cells

1. YP liquid medium: dissolve 10 g of yeast extract and 20 g of peptone in 900 mL of deionized water, and autoclave.

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2. 20% (w/v) glucose solution: dissolve glucose powder in nuclease-free water, and autoclave the solution. 3. Wild-type (BY4741) strain: MATa his3Δ1 leu2Δ0 met15Δ0 ura3Δ0 (Open BioSystems). 4. dbp2Δ strain: MATa dbp2::KanMx6 his3Δ1 leu2Δ0 met15Δ0 ura3Δ0 [34]. 5. UV–VIS spectrometer. 6. Dimethyl sulfate (DMS). 7. 4.8 M 2-mercaptoethanol (BME). 8. Isoamyl alcohol. 9. Centrifuge. 10. AE buffer: 50 mM sodium acetate (NaOAc, pH 5.2), 10 mM ETDA. 2.2 Purification and DNase Treatment of Total RNAs

1. Nuclease-free water. 2. 10% (w/v) sodium dodecyl sulfate (SDS) solution. 3. [5:1] Acid phenol/chloroform mix, pH 4.5. 4. [24:1] chloroform/isoamyl alcohol solution. 5. Ethanol. 6. 3 M sodium acetate in nuclease-free water, pH 5.2. 7. Refrigerated microcentrifuge. 8. Nanodrop. 9. 2 U/μL TURBO DNase (Thermo Fisher Scientific). 10. 20 mg/mL glycogen, RNA grade (Thermo Fisher Scientific). 11. 70% ethanol: mix 7 mL of ethanol with 3 mL of nuclease-free water.

2.3 Primer Extension and Denaturing Polyacrylamide Gel Electrophoresis (PAGE)

1. Nuclease-free water. 2. Fluorescent 5.8S rRNA primer: 50 -/56FAM/AAATGACGCT CAAACAGGCATG-30 . 3. Deoxynucleotide (dNTP) solution mix (10 mM each). 4. Dideoxynucleotides: a set of ddATP, ddTTP, ddCTP, and ddGTP at 5 mM each. 5. 200 U/μL SuperScript III Reverse Transcriptase with 5 First Strand buffer supplied with SuperScript III (250 mM Tris–HCl [pH 8.3 at room temperature], 375 mM KCl, 15 mM MgCl2) (Thermo Fisher Scientific). 6. Digital dry bath or water bath. 7. 2 N NaOH. 8. 2 N HCl.

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9. Formamide (>99.5%). 10. Urea (powder). 11. 12.5 TTE buffer: dissolve 108 g of Tris base, 36 g of taurine, and 2 g of EDTA in 1 L of nuclease-free water. 12. 40% acrylamide (19:1). 13. 10% (w/v) ammonium persulfate (APS): dissolve 100 mg of ammonium persulfate in 1 mL of nuclease-free water. 14. Tetramethylethylenediamine (TEMED). 15. Denaturing gel loading buffer: 90% (v/v) formamide in 0.5 TTE buffer. 16. Polyacrylamide gel electrophoresis (PAGE) apparatus. 17. Fluorescence gel imager. 2.4 Purification of Polyadenylated RNAs and Generation of cDNAs

1. Nuclease-free water. 2. Poly(A)Purist MAG Kit (Thermo Fisher Scientific). 3. 2 U/μL RNase H. 4. Phenol equilibrated with Tris–HCl (pH 8.0). 5. Random hexamer fused with Illumina TruSeq adapter: 50 -CA GACGTGTGCTCTTCCGATCTNNNNNN-30 . 6. TEN buffer: 10 mM Tris–HCl, pH 7.5, 1 mM EDTA, 250 mM NaCl. 7. ssDNA linker: 50 -/5Phos/ GAGCGTCGTGTAG/3SpC3/-30 .

NNNAGATCGGAA

8. CircLigase ssDNA ligase reaction kit (Epicentre). The kit includes 10 reaction buffer, 100 U/μL CircLigase enzyme, and 1 mM ATP and 50 mM MnCl2 stock solutions. 9. SYBR Gold (Thermo Fisher Scientific). 10. 10 TBE buffer. Use commercial source or standard procedure for preparation. 2.5 PCR Amplification and Validation of cDNA Libraries

1. Thermocycler. 2. Nuclease-free water. 3. Ex Taq DNA polymerase, supplied with a 10 reaction buffer and 2.5 mM dNTP mixture (Takara). 4. Illumina TruSeq primers (bolded letters are barcodes for multiplexed libraries):

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5. TOPO TA cloning kit for sequencing (Thermo Fisher Scientific). 6. Wizard Plus Minipreps DNA Purification System (Promega).

3

Methods

3.1 Optimization of DMS Modification in Yeast Cells 3.1.1 DMS Treatment

1. For each condition being tested, grow 50 mL of wild-type and dbp2Δ yeast cells in YP with 2% glucose at 30  C to an OD 600 nm of 0.5–0.7. Prepare one additional culture for the negative (no DMS) and sequencing controls. 2. Add DMS to the liquid culture to a final concentration of 10 mM (see Note 1). Do not add DMS in the control culture. 3. Incubate cells for 5, 10, or 15 min at 30  C with vigorous shaking (see Note 2). 4. Quench the reaction with 75 mL of 4.8 M BME and 25 mL of isoamyl alcohol (see Note 3). 5. Harvest the cells by centrifugation at 2700  g for 5 min, and remove the supernatant. 6. Wash cell pellets with 10 mL of 4.8 M BME, pipetting up and down to disrupt pellet. Harvest the cells by centrifugation at 2700  g for 5 min, and remove the supernatant. 7. Wash cell pellets again with 20 mL of AE buffer, pipetting up and down to disrupt pellet. Harvest the cells by centrifugation at 2700  g for 5 min, and remove the supernatant.

3.1.2 Purification and DNase Treatment of Total RNAs

1. Re-suspend cells in 400 μL of AE buffer, and add 40 μL of 10% SDS and 400 μL of acid phenol/chloroform (5:1). 2. Vortex the mixture, and place at 65  C for 10 min, vortexing samples once every minute. 3. Incubate samples on ice for 5 min. 4. Centrifuge samples at 17,000  g for 5 min, and move the aqueous (top) layer to new tubes.

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5. Add 300 μL of chloroform/isoamyl alcohol (24:1) and mix thoroughly. Repeat step 4. 6. Precipitate the RNAs by adding 50 μL of 3 M sodium acetate and 1 mL ethanol, and incubate at 80  C for at least 30 min. 7. Centrifuge at 17,000  g for 15 min at 4  C and discard the supernatant. 8. Wash the RNA pellet with 500 μL of 70% ethanol, re-suspending pellet by vortexing to remove excess salt (see Note 4). After suspension place samples in a 80  C freezer for at least 15 min to precipitate. 9. Centrifuge samples at 17,000  g for 15 min at 4  C, and discard the supernatant. Dry samples at room temperature. 10. Re-suspend the purified RNAs in 100–150 μL nuclease-free water, and measure the RNA concentration using a NanoDrop or other UV–VIS spectrometer at 260 nm. At least 30 μg of RNA will be needed for the following step. 11. Set up a 150 μL DNase reaction mixture as follows: Component

Volume (μL)

Final quantity

Purified total RNAs

?

30 μg

10X reaction buffer

15

1X

TURBO DNase

3

6U

Nuclease-free water

?

Up to 150 μL

12. Incubate the samples at 37  C for 30 min. 13. Purify the DNase-treated RNAs by adding 150 μL of acid phenol/chloroform (5:1), and repeat steps 4–9 of this section, with addition of 1 μL glycogen (20 mg/mL) during precipitation. 14. Re-suspend RNAs in 20 μL nuclease-free water, and measure the RNA concentration after DNase digestion with a NanoDrop or other UV–VIS spectrometer at 260 nm. At least 20 μg of RNA will be needed for the following steps. 3.1.3 Primer Extension

1. Set up reverse transcription reactions on ice as follows:

Component

Volume (μL)

Final quantity

DNase-treated RNAs

?

20 μg

2 μM fluorescent 5.8S rRNA primer

1

2 pmol

dNTP mix, 10 mM

1

1 mM

Nuclease-free water

?

Up to 12 μL

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2. Incubate the reaction mixture at 65  C for 5 min, and then place the tube on ice for at least 1 min. 3. For sequencing controls, add the following materials to the reaction mix (see Note 5):

Component

Volume (μL)

Final concentration

5 First Strand buffer

4

1

DTT (0.1 M)

1

5 mM

ddT or ddG (5 mM)

1

0.25 mM

SUPERase-In (20 U/μL)

1

1 U/μL

SuperScript III Reverse Transcriptase (200 U/μL)

1

10 U/μL

4. For other samples, add the following materials (see Note 6):

Component

Volume (μL)

Final concentration

5 First Strand buffer

4

1

DTT (0.1 M)

1

5 mM

Nuclease-free water

1

N/A

SUPERase-In (20 U/μL)

1

1 U/μL

SuperScript III Reverse Transcriptase (200 U/μL)

1

10 U/μL

5. Incubate the samples at 25  C for 10 min and then at 50  C for 50 min. 6. Stop the reactions and degrade RNAs by adding 4 μL of 2 N NaOH, and then incubate samples at 95  C for 3 min. 7. Neutralize the solutions by adding 4 μL of 2 N HCl. 8. Precipitate, wash, and dry the cDNAs as described in steps 6–9 of Subheading 3.1.2, ensuring inclusion of 1 μL of glycogen (20 mg/mL) during precipitation. 3.1.4 Visualization of Primer Extension Products

1. Re-suspend cDNAs in 12 μL of denaturing gel loading buffer (see Note 7). 2. Resolve primer extension products using denaturing PAGE. To prepare 50 mL of 8% gel solution, mix the following materials following instructions below (see Note 8).

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Component

Quantity

Urea

21 g

40% acrylamide (19:1)

10 mL

12.5 TTE buffer

4 mL

Nuclease-free water

Up to 50 mL

10% ammonium persulfate

500 μL

TEMED

50 μL

3. Mix urea, acrylamide, 12.5 TTE, and water, making sure urea is completely dissolved. 4. Prepare large format (20 cm 40 cm) glass plates with 0.35 mm spacers and a 20-well comb. Arrange the spacers and glass plates and clip, with at least three clips on each side, and lay gel horizontally on the bench. It should be supported slightly above the bench surface by the clips. 5. Mix APS and TEMED into the acrylamide solution from step 3, and pipet within gel plates using a 25 mL seropipet at a consistent speed. While pipetting, knock firmly along the leading edge of the acrylamide solution to prevent bubbles. Once gel is full, carefully place the comb, and leave horizontal to polymerize. 6. When gel is polymerized, place in gel running apparatus, and fill with running buffer (1 TTE). Wash the wells thoroughly using a pipet or syringe to remove accumulated urea. 7. Pre-run gel at 30 W for 1 h before loading samples. 8. Load DMS-treated samples and sequencing controls, and run at 30 W until leading dye band (bromophenol blue) is near the bottom of the gel. 9. Remove gel sandwich from running apparatus. Dry the outside of the plates, and visualize cDNAs in the gel using gel imaging system to detect the fluorescence signal from the FAM-labeled primer (Fig. 1). It is not necessary to disassemble the plates for detection. 10. For each lane, quantify the fully extended product at the top of the gel image, and compare the signal to the signal of the whole lane (including DMS modification bands and the fully extended product). If the DMS treatment is optimized for single-hit kinetics, the quantity of the full-length product should be ~75% of all the cDNAs generated [24]. Single-hit kinetics are optimal to minimize chance that a structural

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Fig. 1 Primer extension products of DMS-modified RNAs. The first lane contains only the fluorescent primer, indicating the start point of primer extension. The intense signal at the top in the rest of the image represents the fully extended product. The “A” and “C” lanes are sequencing controls with ddTTP or ddGTP in the primer extension reaction (lanes 2 and 3). The positions of bands in these two lanes correspond to the adenine and cytosine residues in the probed sequence. In the no-DMS control (lane 4), some weak bands are detected in addition to the full-length product, which are likely due to random fall-off of the reverse transcriptase or to stops induced by RNA structures. Unlabeled arrows on the right side indicate positions of DMS-induced stops. Single-hit kinetics (full-length product ~75%) was determined by calculating the % of full-length product/total. Lanes 5, 6, and 7 correspond to 93%, 76%, and 73% full-length product. 10 min was chosen for genome-wide experiments

change is induced by a modification elsewhere in the RNA. Primer extension products should be quantified using ImageJ or ImageQuant, determining the amount of full-length product as a fraction of total products (Fig. 1). 3.2 Purification of Polyadenylated RNAs

1. Use total RNAs purified from the optimal DMS treatment condition (see Fig. 1) to prepare RNA solutions with a final concentration of 600 μg/mL in nuclease-free water. The total

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amount of RNA necessary for the following steps should be at least 300 μg. 2. Perform poly(A) selection using the Poly(A)Purist MAG Kit according to the manufacturer’s manual. The amount of recovered RNAs is expected to be 1–2% of the input total RNAs from exponentially growing S. cerevisiae. 3. After poly(A) selection, measure the RNA concentration using Nanodrop to ensure appropriate recovery. 3.3

DNase Treatment

1. Treat 3 μg of poly(A) RNAs with Turbo DNase. Each reaction includes 1 μL (2 Units) of Turbo DNase, 3 μL of 10 reaction buffer, 3 μg of poly(A) RNAs, and nuclease-free water up to 30 μL. 2. Incubate the samples at 37  C for 30 min. 3. Purify the DNase-treated RNAs by adding 100 μL of acid phenol/chloroform (5:1), and repeat steps 4–7 of Subheading 3.1.2, with addition of 1 μL glycogen (20 mg/mL) during precipitation. 4. Wash the RNA pellet with 70% ethanol. Centrifuge samples at 17,000  g for 15 min at 4  C, and discard the supernatant. Dry samples at room temperature. 5. Re-suspend RNAs in nuclease-free water. Measure the RNA concentration using Nanodrop.

3.4 Generation of cDNA

1. Set up a 12 μL reverse transcription mix with the following materials: Component

Volume (μL)

Final quantity

DNase-treated poly(A) RNAs

?

2 μg

Random hexamer fused with Illumina TruSeq 1 adapter, 100 μM

10 μM

dNTP mix, 10 mM

1

1 mM

Nuclease-free water

?

Up to 12 μL

2. Incubate the reaction mixture at 65  C for 5 min, and place the tube on ice afterward for at least 1 min. 3. Add the following materials to the reaction mix:

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Component

Volume (μL)

Final concentration

5 First Strand buffer

4

1

DTT, 0.1 M

2

0.01 M

SUPERase-In (20 U/μL)

1

1 U/μL

SuperScript III Reverse Transcriptase (200 U/μL)

1

10/μL

4. Incubate the samples at 25  C for 10 min and then at 50  C for 50 min. 5. Stop the reaction by heating the samples at 85  C for 5 min. Chill the tubes on ice. 6. Add 1 μL of RNase H (2 U/μL) to each sample to degrade RNAs, and incubate the reactions at 37  C for 20 min. 7. Purify cDNAs by adding 100 μL of phenol (pH 8.0), and repeat steps 4–9 in Subheading 3.1.2, with addition of 1 μL glycogen (20 mg/mL) during precipitation. 8. Re-suspend cDNAs in 10 μL of denaturing gel loading buffer, and resolve them on a 8% denaturing PAGE gel as described in Subheading 3.1.4, steps 2–7. 9. Visualize cDNAs by staining in a 1X solution of SYBR Gold in 0.5 TBE for 20 min at room temperature with gentle shaking. Excise bands with a razor blade, and collect gel pieces containing cDNA fragments larger than 40 nt (Fig. 2a). 10. Crush the gel pieces by ejecting through a syringe, and soak the gel pieces in 600 μL TEN buffer (enough to cover gel pieces) in a 1.5 mL microtube on a rocker at 4  C overnight to elute cDNAs. Collect the elution solution, carefully avoiding drawing gel debris, and purify cDNAs by ethanol precipitation as described in Subheading 3.3, steps 3–4. 3.5 ssDNA Linker Ligation

1. Re-suspend the gel-purified cDNA pellets from Subheading 3.4, step 10 in 14 μL of nuclease-free water. 2. For each cDNA sample, prepare a ligation reaction as follows: Component

Volume (μL) Final quantity

cDNA

14 μL

?

ssDNA linker, 100 μM

1

5 μM

CircLigase reaction buffer, 10

2

1

ATP, 1 mM

1

0.05 mM

MnCl2, 50 mM

1

2.5 mM

CircLigase ssDNA ligase (100 U/μL) 1

5 U/μL

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Fig. 2 Representative PAGE images of DNA fragments after reverse transcription (a) and ssDNA linker ligation (b). The two samples (lanes 2 and 3) shown are from two biological replicates of wild type after DMS treatment. Bands pointed by arrows indicate positions of undesired, excess primers (a) or the ligation product of RT primer and ssDNA linker (b)

3. Incubate samples at 65  C overnight (for at least 12 h), and then inactivate CircLigase by heating samples at 85  C for 15 min. 4. Purify ligated cDNAs by extractions with phenol (pH 8.0) and then with chloroform/isoamyl alcohol (24:1), followed by ethanol precipitation as described in Subheading 3.3, steps 4 and 5. 5. Re-suspend cDNAs in 10 μL of denaturing gel loading buffer, and resolve them on a 8% denaturing PAGE gel as described in Subheading 3.1.4, steps 2–7.

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6. Visualize cDNAs by staining with SYBR Gold for 20 min at room temperature. Excise bands with a razor blade, and collect gel pieces containing cDNA fragments larger than 60 nt (Fig. 2b). 7. Recover ligated cDNAs from gel pieces as described in step 10 of Subheading 3.4. 3.6 PCR Amplification of cDNA Libraries

1. For each sample, re-suspend the ligated, size-selected cDNA in 10 μL of nuclease-free water. 2. Set up a PCR reaction as below:

Volume (μL)

Final quantity

Ligated cDNA (or water for primer-dimer control)

5 μL

?

dNTP mixture (2.5 mM each)

2

0.2 mM

Ex Taq buffer, 10

2.5

1

Illumina TruSeq forward primer (5 μM)

1

0.2 μM

Illumina TruSeq reverse primer (5 μM)

1

0.2 μM

Ex Taq polymerase (50 U/μL)

0.5

0.1 U/μL

Nuclease-free water

13

13 μL

Component

3. Amplify the cDNAs by PCR using the following conditions (see Note 9). Step Initial denaturation

Temperature

Time

Number of cycles



1 min

1



98 C

Denaturation Annealing Extension Final extension

98 C 55  C 72  C 72  C

10 s 30 s 1 min 10 min

10–35

Incubation

4 C

2 min

1

1

4. Resolve PCR products on a large format 8% PAGE gel. Gel should be assembled as above, but no urea is added to the gel mixture, and 1 TBE is used as the running buffer instead of 1 TTE. 5. Pre-run gel for at least 30 min at 5 V/cm. 6. Load samples, and run at 5 V/cm, keeping an eye on the xylene cyanol dye, which will run at ~160 bp in an 8% gel (see Note 10).

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7. Collect fragments larger than 150 bp as described in Subheading 3.4, steps 8–10. Re-suspend purified libraries in 20 μL of nuclease-free water. 3.7 Validation of Libraries

1. Perform cloning of size-selected PCR products using TOPO TA cloning kit for sequencing according to the manufacturer’s instructions. 2. Extract plasmid DNA from at least five colonies for each library using the Wizard Plus Miniprep DNA Purification kit and manufacturer’s instructions. Use a commercial service (or in-house facility if available) to sequence the inserted fragments. The inserted sequences should correspond to the transcriptome of interest (excluding ribosomal DNA). If that is the case, then proceed to the next steps (which are usually handled by expert deep sequencing facilities or commercial service companies and are thus not highly detailed below). 3. Examine the library size distribution using an Agilent Bioanalyzer. The library should not contain products below 150 bp, such as undesired primer-dimer by-products. 4. Quantify libraries by qPCR to determine the appropriate amount for pooling libraries in the following sequencing runs. 5. Perform sequencing on an Illumina HiSeq 2500 platform for 2  100 bp paired-end cycle run (see Note 11). In our case, we pooled three barcoded libraries to run in a single lane of a HiSeq flow cell.

3.8 Bioinformatics Analysis of the Sequencing Data

4

The bioinformatics processing steps and calculations are detailed elsewhere [35]. Examples of the normalized DMS reactivity and the difference in wild type and dbp2Δ are shown in Fig. 3.

Notes 1. DMS is highly toxic. You must wear gloves and use a fume hood for all steps involving DMS. 2. In addition to the length of treatment time, DMS concentration can also be varied and tested to obtain optimal reactivity conditions. 3. Both BME and isoamyl alcohol have strong smells, and the mixed solution can leak from tube caps during centrifugation. Perform these steps in fume hoods, and seal the lid of centrifuge tubes carefully with parafilm. 4. If the DNA pellet is not fully washed, retained salt can lead to poor resolution and distorted gel images. In addition, excess salt will also reduce the efficiency of poly(A) selection in Subheading 3.2.

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Fig. 3 Normalized DMS reactivity in wild-type and dbp2Δ cells at the 30 ends of selected transcripts (a–c). Decreased reactivity indicates that the nucleotides are more protected by RNA/RNP structures and vice versa. The change dependent on DBP2 is further visualized by subtracting the reactivity value in wild type from the value in dbp2Δ (bottom panel). This is reproduced from [33], with permission from Genetics

5. The ratio between dNTP and ddTTP (or ddGTP) may need to be optimized to detect and clearly visualize most of the adenines (or cytosines) in the sequence [36]. 6. If multiple reactions are performed, a master mix can be made and aliquoted to sample tubes from the last step. This will

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reduce the handling time and chance of RNase contamination in the reagents and samples. 7. Do not add dye in the loading buffer, as it will impact the fluorescence signal for gel imaging. To roughly track the positions of DNA fragments during electrophoresis, load 10 μL of gel loading dye (0.25% bromophenol blue, 0.25% xylene cyanol FF in 90% formamide) in an empty well. Loading of unextended fluorescent primer in an empty lane also helps to identify the start of the sequence. 8. For gel running, TTE is used instead of the typical Tris-borate EDTA (TBE) buffer to increase resolution. TTE gives sharper bands than TBE [37]. 9. The number of amplification cycles may need to be optimized, and products may need to be visualized on a gel. The final concentration of libraries may be insufficient for downstream analysis and sequencing with too few cycles of amplification, whereas excess amplification can lead to production of undesired, nonspecific bands. 10. In addition to amplified libraries, load the non-templated primer-dimer control for PAGE analysis to pinpoint the sizes of potential nonspecific products. 11. Paired-end sequencing is recommended, as it will provide comprehensive coverage information for each nucleotide in the downstream bioinformatics analysis.

Acknowledgments We thank Dr. Yiliang Ding (John Innes Centre) for help in protocol optimization and construction of sequencing libraries. We also thank Dr. Sharon Aviran (UC Davis) for discussion on experimental design that was critical for downstream bioinformatic analysis and for conducting bioinformatics necessary to visualize significant reactivity change genome wide. This work was supported by NIH R01GM097332 to E.J.T., P30CA023168 for core facilities at the Purdue University Center for Cancer Research. References 1. Schroeder R, Barta A, Semrad K (2004) Strategies for RNA folding and assembly. Nat Rev Mol Cell Biol 5:908–919. https://doi.org/10. 1038/nrm1497 2. Doetsch M, Schroeder R, Fu¨rtig B (2011) Transient RNA-protein interactions in RNA folding. FEBS J 278:1634–1642

3. Rouskin S, Zubradt M, Washietl S et al (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 4. Fairman-Williams ME, Guenther UP, Jankowsky E (2010) SF1 and SF2 helicases: family matters. Curr Opin Struct Biol 20:313–324

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16. Kar A, Fushimi K, Zhou X et al (2011) RNA helicase p68 (DDX5) regulates tau exon 10 splicing by modulating a stem-loop structure at the 50 splice site. Mol Cell Biol 31:1812–1821. https://doi.org/10.1128/ MCB.01149-10 17. Moy RH, Cole BS, Yasunaga A et al (2014) Stem-loop recognition by DDX17 facilitates miRNA processing and antiviral defense. Cell 158:764–777. https://doi.org/10.1016/j. cell.2014.06.023 18. Svitkin YV, Pause A, Haghighat A et al (2001) The requirement for eukaryotic initiation factor 4A (elF4A) in translation is in direct proportion to the degree of mRNA 50 secondary structure. RNA 7:382–394. https://doi.org/ 10.1017/S135583820100108X 19. Soto-Rifo R, Rubilar PS, Limousin T et al (2012) DEAD-box protein DDX3 associates with eIF4F to promote translation of selected mRNAs. EMBO J 31:3745–3756. https://doi. org/10.1038/emboj.2012.220 20. Guenther UP, Weinberg DE, Zubradt MM et al (2018) The helicase Ded1p controls use of near-cognate translation initiation codons in 50 UTRs. Nature 559:130–134. https://doi. org/10.1038/s41586-018-0258-0 21. Ding Y, Tang Y, Kwok CK et al (2014) In vivo genome-wide profiling of RNA secondary structure reveals novel regulatory features. Nature 505:696–700. https://doi.org/10. 1038/nature12756 22. Wan Y, Qu K, Zhang QC et al (2014) Landscape and variation of RNA secondary structure across the human transcriptome. Nature 505:706–709. https://doi.org/10.1038/ nature12946 23. Talkish J, May G, Lin Y et al (2014) Mod-seq: high-throughput sequencing for chemical probing of RNA structure. RNA 20:713–720. https://doi.org/10.1261/rna.042218.113 24. Ding Y, Kwok CK, Tang Y et al (2015) Genome-wide profiling of in vivo RNA structure at single-nucleotide resolution using structure-seq. Nat Protoc 10:1050–1066. https://doi.org/10.1038/nprot.2015.064 25. Zubradt M, Gupta P, Persad S et al (2016) DMS-MaPseq for genome-wide or targeted RNA structure probing in vivo. Nat Methods 14:75–82. https://doi.org/10.1038/nmeth. 4057 26. Ritchey LE, Su Z, Tang Y et al (2017) Structure-seq2: sensitive and accurate genome-wide profiling of RNA structure in vivo. Nucleic Acids Res 45:e135. https:// doi.org/10.1093/nar/gkx533

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Chapter 19 In Situ Hybridization-Proximity Ligation Assay (ISH-PLA) to Study the Interaction of HIV-1 RNA and Remodeling Proteins Daniela Toro-Ascuy, Aracelly Gaete-Argel, Victoria Rojas-Celis, and Fernando Valiente-Echeverria Abstract The mechanisms involved in the posttranscriptional control of the replicative cycle of the human immunodeficiency virus (HIV), specifically the molecular events which allow the interaction between the viral genomic RNA (gRNA) and the cellular machinery for the transport, translation, or intracellular packaging, have not been yet elucidated. In this chapter, we describe the in situ hybridization-proximity ligation assay (ISH-PLA) to characterize interactions between the genomic RNA (gRNA) of HIV-1 and viral proteins or host proteins involved in nuclear export and translation initiation. We also present data that validate the ISH-PLA as a simple and useful tool to study HIV-1 gRNA-protein interactions within cells. Key words AIDS, HIV-1, HIV-1 RNA, Proximity ligation assay, ISH-PLA, RNA-protein interaction

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Introduction Human immunodeficiency virus type 1 (HIV-1) is a member of the lentivirus subgroup of the Retroviridae family, responsible for one of the most important human pandemic diseases that still has no cure: the acquired immunodeficiency syndrome (AIDS) [1]. Following the HIV-1 infection, the viral genome is converted into a double-stranded DNA molecule by the reverse transcriptase (RT), transported to the nucleus, and inserted into a cellular chromosome [2]. The proviral insert depends on RNA polymerase II for transcription, where RNA-binding proteins (RBPs) associated with an HIV-1 genomic RNA (gRNA) are a determining factor for the storage, degradation, or packaging into new viral particles [3–5]. Many techniques have been developed to study RNA-protein interactions, which can be classified as RNA-centric methods or protein-centric methods, depending on the molecule they start with [6]. The proximity ligation assay (PLA) is a highly specific

Marc Boudvillain (ed.), RNA Remodeling Proteins: Methods and Protocols, Methods in Molecular Biology, vol. 2209, https://doi.org/10.1007/978-1-0716-0935-4_19, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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and sensitive method that produces countable spots allowing the detection of endogenous protein-protein interactions and also interactions with low-abundance proteins [7]. The PLA uses a pair of oligonucleotide-labeled antibodies binding in proximity (30–40 nm apart) to two proteins in a complex [8]. In brief, two primary antibodies against two target proteins are recognized by secondary antibodies conjugated to short DNA oligonucleotides. If the two proteins are in proximity in the same complex, the oligonucleotides will be ligated, producing circular DNA, which is amplified by rolling circle amplification (RCA) using a DNA polymerase. Then, fluorescently labeled oligonucleotides are hybridized to the RCA product, and the PLA signal is detected using fluorescence microscopy as discrete spots [8]. To study possible interactions between HIV-1 gRNA and host cellular factors, we developed a variant of PLA. The association of RNA fluorescence in situ hybridization (FISH), which allows to identify changes in the localization of viral RNA, with immunofluorescence (IF) analysis can provide information on the co-localization of proteins with the viral RNA. Yet, this information is not sufficient to unambiguously confirm an interaction. To address this limitation, we combined RNA in situ hybridization (ISH) with PLA to evaluate proximity between HIV-1 gRNA and proteins of interest (Fig. 1) [9–11]. To validate the approach, we performed the in situ hybridization – proximity ligation assay (ISH-PLA) between wild-type HIV-1 gRNA (pNL4.3-wt) and Gag (p17) or CRM1 (which is known to function as the partner of Rev. in the nuclear export of HIV-1 gRNA) and between HIV-1 gRNA (pNL4.3-ΔRev) and Rev protein. Our results demonstrate the interaction between HIV-1 gRNA and Rev. or Gag or CRM1 (Fig. 2a, b). We also performed an ISH-PLA coupled to IF to observe the interaction of gRNA with Gag and the expression of both molecules (gRNA and Gag) in the same cell (Fig. 2c). We have also used this technique to study the interaction of HIV-1 gRNA with several cellular proteins involved in nuclear export and translation initiation, such as CBP80, eIF4E, eIF4G, eIF4AI, and eIF3G; we focused on determining whether the Rev./RRE axis is important for the recruitment of these cellular proteins to HIV-1 gRNA. We quantified the interaction between HIV-1 gRNA and the cellular proteins mentioned above, in the presence or absence of Rev, using our ISH-PLA protocol [9]. Our results validate ISH-PLA as a simple and useful tool to study HIV-1 gRNA-protein interactions within cells. Of note, this protocol may also be used to determine the subcellular localization of the gRNA-protein interactions.

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Fig. 1 Schematic of the ISH-PLA coupled to IF protocol. (a) Viral RNA is recognized by a digoxigenin-11-UTPcontaining RNA probe during the hybridization step. Primary antibodies to detect a partner protein (e.g., viral Gag) and digoxigenin are used. (b) Secondary PLA antibodies attached to oligonucleotides are added to recognize the primary protein and anti-digoxigenin antibodies. (c) A connector oligonucleotide is added and ligated, resulting in a circular DNA only if the targeted protein and RNA are in close proximity (811 nt in length) circular RNAs accumulated in the nucleus upon Hel25E depletion. Indeed, reporter plasmids that generate Drosophila circular RNAs of distinct lengths confirmed that the length of the mature circular RNA dictates the nuclear export mechanism [35]. Humans encode two homologs of Hel25E—UAP56 (DDX39B) and URH49 (DDX39A)—that are 90% identical to one another [38, 39], yet these helicases have distinct roles in controlling localization of human circular RNAs [35]. Depletion of UAP56 causes long (>1298 nt in length) circular RNAs to become enriched in the nucleus, while depletion of URH49 causes nuclear retention of short (50% by comparing radioactivity in FT solution (incorporated) versus that retained on column (free) using Geiger counter. 5. Add all of the radioactive probe (~50 μL) to tube containing pre-hybridizing membrane (see Subheading 3.3.2) and hybridize with rotation for at least 4 h at 42  C (see Note 14).

3.3.4 Membrane Washing and RNA Detection

1. Remove and safely discard the radioactive hybridization buffer (see Note 15). 2. Rinse the membrane briefly with at least 10 mL of washing solution 1 by shaking the tube and discard safely the washing buffer (see Note 15). 3. Add 10 mL of the washing solution 1, and place the tube in the hybridization oven for 10 min at 42  C. Then, discard the washing solution safely (see Note 15). 4. Add 10 mL of the washing solution 2 and place the tube in the hybridization oven for 10 min at 42  C. Then, discard the washing solution safely (see Note 15). 5. Repeat step 4 two more times. 6. Air-dry the membrane on a sheet of 3MM Whatman paper, cover both with plastic wrap, and expose it to a PhosphorImager screen overnight (4 h can be sufficient if the probed RNA is abundant). 7. Reveal image on a PhosphorImager. In a rifampicin experiment, typically we will see the band corresponding to the mRNA of interest decreasing steadily over time. Stable RNAs (e.g., ribosomal RNAs) will not decrease and indeed can be used to normalize for loading variations. RNA sizes may also change, e.g., when analyzing mutant strains lacking processing enzymes.

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Notes 1. It is best to work directly in a 37  C room or right next to a 37  C incubator (air shaker or water bath). 2. For data points that are close together, it is advisable to find a lab partner to run the centrifuge, remove supernatants, and freeze samples, while the other adds rifampicin, samples the cultures, and mixes with frozen azide. 3. Rifampicin is light sensitive and stored resuspended in DMSO or DMF in foil covered tubes; wear gloves during manipulation. 4. For minimal medium (e.g., MD or M9), it is important to wash the pellet by resuspending in 1 mL of TE/NaCl buffer to remove excess phosphate and to centrifuge a second time before freezing the pellet on dry ice. 5. When working with RNAs, be careful to avoid any contamination with RNases. Wear gloves at all times, and prior to your experiments, clean your bench and your pipettes with water then ethanol or RNase away solution. We keep a separate set of micropipettes reserved for RNA work. 6. OD260/OD280 ratios are generally poor for RNAsnap™ preps. 7. Samples should be equilibrated at room temperature before measurement of RNA concentration to avoid interference from the SDS present in the solution that precipitates in the cold. 8. Work under a chemical fume hood. 9. Be careful not to dry the pellet too much because it will be difficult to resuspend. 10. After transfer, RNAs can be visualized with a portable UV light at 254 nm. Mark the ribosomal RNAs and the bands of the RNA marker with a pencil, pressing strongly. This will score the membrane and hold the radioactivity during the hybridization, allowing the visualization of the marker without its labeling. 11. Silanization of the plate should be done under a fume hood (only 1 plate should be treated!). 12. Use a plastic tool to avoid damaging the glass plates. 13. Dry the Whatman 3MM paper with paper towel to remove liquid excess and allow the gel to stick to the Whatman 3MM paper. 14. If the RNA of interest is expressed at a very low level, the radiolabeled oligonucleotide can be replaced by an α-32P-UTP-labeled riboprobe complementary to the RNA. This will often improve the signal, but note that it is difficult

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to remove if the membrane is to be reprobed for another RNA. We usually make riboprobes between 200 and 500 nucleotides in length, using a PCR template with a T7 promoter integrated into the lower oligo (see Chapter 25). The pre-hybridization and hybridization temperatures should be increased to 68  C if a riboprobe is used instead of radiolabeled oligonucleotide. 15. All radioactive materials should be handled by properly trained scientists, following safety rules and local regulations.

Acknowledgments This work was supported by funds from the CNRS (UMR8261), Universite´ de Paris, the Agence Nationale de la Recherche (ARNrQC; BaRR) and the Labex (Dynamo) program. References 1. Durand S, Condon C (2018) RNases and helicases in gram-positive bacteria. Microbiol Spect 6(2). https://doi.org/10.1128/microbiolspec. RWR-0003-2017 2. Baumgardt K, Gilet L, Figaro S, Condon C (2018) The essential nature of YqfG, a YbeY homologue required for 30 maturation of Bacillus subtilis 16S ribosomal RNA is suppressed by deletion of RNase R. Nucleic Acids Res 46 (16):8605–8615. https://doi.org/10.1093/ nar/gky488 3. Durand S, Braun F, Lioliou E, Romilly C, Helfer AC, Kuhn L, Quittot N, Nicolas P, Romby P, Condon C (2015) A nitric oxide regulated small RNA controls expression of genes involved in

redox homeostasis in Bacillus subtilis. PLoS Genet 11(2):e1004957. https://doi.org/10. 1371/journal.pgen.1004957 4. Putzer H, Gendron N, Grunberg-Manago M (1992) Co-ordinate expression of the two threonyl-tRNA synthetase genes in Bacillus subtilis: control by transcriptional antitermination involving a conserved regulatory sequence. EMBO J 11:3117–3127 5. Stead MB, Agrawal A, Bowden KE, Nasir R, Mohanty BK, Meagher RB, Kushner SR (2012) RNAsnap: a rapid, quantitative and inexpensive, method for isolating total RNA from bacteria. Nucleic Acids Res 40(20):e156. https://doi.org/10.1093/nar/gks680

Chapter 25 Assay of Bacillus subtilis Ribonuclease Activity In Vitro Olivier Pellegrini, Laetitia Gilet, Aude Trinquier, Anastasia Tolcan, Delphine Allouche, Sylvain Durand, Fre´de´rique Braun, and Ciara´n Condon Abstract Ribonucleases can cleave RNAs internally in endoribonucleolytic mode or remove one nucleotide at a time from either the 50 or 30 end through exoribonuclease action. To show direct implication of an RNase in a specific pathway of RNA maturation or decay requires the setting up of in vitro assays with purified enzymes and substrates. This chapter complements Chapter 24 on assays of ribonuclease action in vivo by providing detailed protocols for the assay of B. subtilis RNases with prepared substrates in vitro. Keywords Ribonuclease, Protein purification, RNA substrates, In vitro transcription, Radioactive labelling

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Introduction The Gram-positive model organism Bacillus subtilis has >20 different ribonucleases (RNases) involved in mRNA and stable RNA maturation and decay [1]. Remarkably, fewer than half of these RNases overlap with the well-studied E. coli model. In B. subtilis, the main endoribonucleases involved in mRNA decay are RNase Y, which cleaves in single-stranded AU-rich regions, and RNase III, which cleaves dsRNA. The main 50 -exoribonuclease is RNase J1, which functions in a stoichiometric complex with its paralog RNase J2, while the main 30 -exoribonucleases are PNPase, RNase PH, RNase R, and YhaM. rRNA processing involves RNase III, RNase J1, YqfG, Mini-III, and RNase M5 in B. subtilis, while tRNA processing requires RNase P and either RNase Z or the four 30 -exoribonucleases, depending nominally on whether the tRNA has an encoded CCA. RNases are routinely assayed in vitro to definitively demonstrate that the effect of an enzyme observed in vivo is direct rather than indirect (see Chapter 24). In vitro assays are also used to establish an enzyme’s biochemical (Km, Kd) or kinetic parameters (Kcat). We generally use purified His-tagged enzymes for these

Marc Boudvillain (ed.), RNA Remodeling Proteins: Methods and Protocols, Methods in Molecular Biology, vol. 2209, https://doi.org/10.1007/978-1-0716-0935-4_25, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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assays. For new uncharacterized enzymes, we typically start with standard buffer at neutral pH, 100 mM NaCl, and 10 mM Mg2+ as the most likely metal ion and a reducing agent (e.g., dithiothreitol [DTT]/β-mercaptoethanol) and then optimize each of these buffer components in turn. Although assays can be performed on generic RNA targets, it is preferable that the true substrates be used. Our standard operating procedure is to synthesize specific RNA substrates by transcribing PCR products containing an integrated phage T7 promoter. These can be labelled with α-32P-UTP directly or can be synthesized as unlabelled substrates and labelled later at their 50 or 30 extremities using γ-32P-ATP or α-32P-pCp. For very short RNA substrates (