RNA-Chromatin Interactions: Methods and Protocols [1st ed.] 9781071606797, 9781071606803

Table of contents :
Front Matter ....Pages i-x
Tuning the Expression of Long Noncoding RNA Loci with CRISPR Interference (Lovorka Stojic)....Pages 1-16
Identification of Chromatin Binding Sites for Long Noncoding RNAs by Chromatin Oligo-Affinity Precipitation (ChOP) (Marianna Nicoletta Rossi, Rossella Maione)....Pages 17-28
Knockdown of Nuclear lncRNAs by Locked Nucleic Acid (LNA) Gapmers in Nephron Progenitor Cells (Masaki Nishikawa, Norimoto Yanagawa)....Pages 29-36
Design and Application of a Short (16-mer) Locked Nucleic Acid Splice-Switching Oligonucleotide for Dystrophin Production in Duchenne Muscular Dystrophy Myotubes (Célia Carvalho, Maria Carmo-Fonseca)....Pages 37-50
Targeting Polyadenylation for Retention of RNA at Chromatin (Evgenia Ntini, Ulf Andersson Vang Ørom)....Pages 51-58
Simultaneous Detection of RNAs and Proteins with Subcellular Resolution (Sunjong Kwon, Koei Chin, Michel Nederlof)....Pages 59-73
In Vivo Crosslinking of Histone and RNA-Binding Proteins (Yong-Eun Kim, Kyoon Eon Kim, Kee K. Kim)....Pages 75-88
Proteomics-Based Systematic Identification of Nuclear Proteins Anchored to Chromatin via RNA (Kyoko Hiragami-Hamada, Naoki Tani, Jun-ichi Nakayama)....Pages 89-99
2D Saturation Transfer Difference NMR for Determination of Protein Binding Sites on RNA Guanine Quadruplexes (Ewan K. S. McRae, David E. Davidson, Sean A. McKenna)....Pages 101-113
Mapping Transcriptome-Wide and Genome-Wide RNA–DNA Contacts with Chromatin-Associated RNA Sequencing (ChAR-seq) (Charles Limouse, David Jukam, Owen K. Smith, Kelsey A. Fryer, Aaron F. Straight)....Pages 115-142
Identification of cis-Elements for RNA Subcellular Localization Through REL-seq (Yafei Yin, Xiaohua Shen)....Pages 143-160
Transcriptome-Wide Mapping of Protein–RNA Interactions (Xianju Bi, Xiaohua Shen)....Pages 161-173
Proximity RNA-seq: A Sequencing Method to Identify Co-localization of RNA (Jörg Morf, Steven W. Wingett)....Pages 175-194
Immunoprecipitation of DNA:RNA Hybrids Using the S9.6 Antibody (Hunter R. Gibbons, Thomas M. Aune)....Pages 195-207
Characterization of R-Loop Structures Using Single-Molecule R-Loop Footprinting and Sequencing (Maika Malig, Frederic Chedin)....Pages 209-228
Analysis of RNA–DNA Triplex Structures In Vitro and In Vivo (Anna Postepska-Igielska, Alena Blank-Giwojna, Ingrid Grummt)....Pages 229-246
Analyzing RNA–DNA Triplex Formation in Chromatin (Rodrigo Maldonado, Gernot Längst)....Pages 247-254
Mapping RNA–Chromatin Interactions In Vivo with RNA-DamID (Seth W. Cheetham, Andrea H. Brand)....Pages 255-264
Back Matter ....Pages 265-266

Citation preview

Methods in Molecular Biology 2161

Ulf Andersson Vang Ørom Editor

RNA-Chromatin Interactions Methods and Protocols

METHODS

IN

MOLECULAR BIOLOGY

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

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

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

RNA-Chromatin Interactions Methods and Protocols

Edited by

Ulf Andersson Vang Ørom Department for Molecular Biology and Genetics, Aarhus University, Aarhus C, Denmark

Editor Ulf Andersson Vang Ørom Department for Molecular Biology and Genetics Aarhus University Aarhus C, Denmark

ISSN 1064-3745 ISSN 1940-6029 (electronic) Methods in Molecular Biology ISBN 978-1-0716-0679-7 ISBN 978-1-0716-0680-3 (eBook) https://doi.org/10.1007/978-1-0716-0680-3 © Springer Science+Business Media, LLC, part of Springer Nature 2020 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 RNA is an essential molecule for numerous biological processes. Research in recent years has taught us that RNA–protein interactions play an important role for both RNA and protein function. With the focus on noncoding RNA and their function, the nuclear and chromatinassociated function of RNA has received increasing attention. It is my hope that researchers interested in RNA-chromatin interactions and functionality can use this book as a starting point to expand their experimental approaches toward the numerous outstanding questions in this new and expanding field. Aarhus C, Denmark

Ulf Andersson Vang Ørom

v

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

1 Tuning the Expression of Long Noncoding RNA Loci with CRISPR Interference . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Lovorka Stojic 2 Identification of Chromatin Binding Sites for Long Noncoding RNAs by Chromatin Oligo-Affinity Precipitation (ChOP) . . . . . . . . . . . . . . . . . . . Marianna Nicoletta Rossi and Rossella Maione 3 Knockdown of Nuclear lncRNAs by Locked Nucleic Acid (LNA) Gapmers in Nephron Progenitor Cells. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Masaki Nishikawa and Norimoto Yanagawa 4 Design and Application of a Short (16-mer) Locked Nucleic Acid Splice-Switching Oligonucleotide for Dystrophin Production in Duchenne Muscular Dystrophy Myotubes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Ce´lia Carvalho and Maria Carmo-Fonseca 5 Targeting Polyadenylation for Retention of RNA at Chromatin. . . . . . . . . . . . . . . Evgenia Ntini and Ulf Andersson Vang Ørom 6 Simultaneous Detection of RNAs and Proteins with Subcellular Resolution . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Sunjong Kwon, Koei Chin, and Michel Nederlof 7 In Vivo Crosslinking of Histone and RNA-Binding Proteins . . . . . . . . . . . . . . . . . Yong-Eun Kim, Kyoon Eon Kim, and Kee K. Kim 8 Proteomics-Based Systematic Identification of Nuclear Proteins Anchored to Chromatin via RNA . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Kyoko Hiragami-Hamada, Naoki Tani, and Jun-ichi Nakayama 9 2D Saturation Transfer Difference NMR for Determination of Protein Binding Sites on RNA Guanine Quadruplexes. . . . . . . . . . . . . . . . . . . . . . . Ewan K. S. McRae, David E. Davidson, and Sean A. McKenna 10 Mapping Transcriptome-Wide and Genome-Wide RNA–DNA Contacts with Chromatin-Associated RNA Sequencing (ChAR-seq) . . . . . . . . . . Charles Limouse, David Jukam, Owen K. Smith, Kelsey A. Fryer, and Aaron F. Straight 11 Identification of cis-Elements for RNA Subcellular Localization Through REL-seq . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Yafei Yin and Xiaohua Shen 12 Transcriptome-Wide Mapping of Protein–RNA Interactions . . . . . . . . . . . . . . . . . Xianju Bi and Xiaohua Shen

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29

37 51

59 75

89

101

115

143 161

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14

15

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17 18

Contents

Proximity RNA-seq: A Sequencing Method to Identify Co-localization of RNA . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Jo¨rg Morf and Steven W. Wingett Immunoprecipitation of DNA:RNA Hybrids Using the S9.6 Antibody . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Hunter R. Gibbons and Thomas M. Aune Characterization of R-Loop Structures Using Single-Molecule R-Loop Footprinting and Sequencing . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Maika Malig and Frederic Chedin Analysis of RNA–DNA Triplex Structures In Vitro and In Vivo. . . . . . . . . . . . . . . Anna Postepska-Igielska, Alena Blank-Giwojna and Ingrid Grummt Analyzing RNA–DNA Triplex Formation in Chromatin . . . . . . . . . . . . . . . . . . . . . Rodrigo Maldonado and Gernot L€ a ngst Mapping RNA–Chromatin Interactions In Vivo with RNA-DamID. . . . . . . . . . . Seth W. Cheetham and Andrea H. Brand

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

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247 255 265

Contributors THOMAS M. AUNE • Vanderbilt University Medical Center, Nashville, TN, USA XIANJU BI • Department of Basic Medical Sciences, School of Medicine, Tsinghua University, Beijing, China ALENA BLANK-GIWOJNA • Division of Molecular Biology of the Cell II, German Cancer Research Center, DKFZ-ZMBH Alliance, Heidelberg, Germany ANDREA H. BRAND • The Gurdon Institute and Department of Physiology, Development and Neuroscience, University of Cambridge, Cambridge, UK MARIA CARMO-FONSECA • Instituto de Medicina Molecular, Faculdade de Medicina, Universidade de Lisboa, Lisbon, Portugal CE´LIA CARVALHO • Instituto de Medicina Molecular, Faculdade de Medicina, Universidade de Lisboa, Lisbon, Portugal FREDERIC CHEDIN • Department of Molecular and Cellular Biology and Genome Center, University of California Davis, Davis, CA, USA SETH W. CHEETHAM • Mater Research Institute-University of Queensland, Woolloongabba, QLD, Australia KOEI CHIN • OHSU Center for Spatial Systems Biomedicine, Oregon Health and Sciences University, Portland, OR, USA; Department of Biomedical Engineering, Oregon Health and Sciences University, Portland, OR, USA DAVID E. DAVIDSON • Department of Chemistry, University of Manitoba, Winnipeg, MB, Canada KELSEY A. FRYER • Department of Biochemistry, Stanford University School of Medicine, Stanford, CA, USA; Department of Genetics, Stanford University School of Medicine, Stanford, CA, USA HUNTER R. GIBBONS • Vanderbilt University Medical Center, Nashville, TN, USA INGRID GRUMMT • Division of Molecular Biology of the Cell II, German Cancer Research Center, DKFZ-ZMBH Alliance, Heidelberg, Germany KYOKO HIRAGAMI-HAMADA • Division of Chromatin Regulation, National Institute for Basic Biology, Okazaki, Aichi, Japan; Department of Basic Biology, School of Life Science, The Graduate University for Advanced Studies (SOKENDAI), Okazaki, Aichi, Japan DAVID JUKAM • Department of Biology, Stanford University, Stanford, CA, USA KEE K. KIM • Department of Biochemistry, Chungnam National University, Daejeon, Republic of Korea KYOON EON KIM • Department of Biochemistry, Chungnam National University, Daejeon, Republic of Korea YONG-EUN KIM • Department of Biochemistry, Chungnam National University, Daejeon, Republic of Korea SUNJONG KWON • OHSU Center for Spatial Systems Biomedicine, Oregon Health and Sciences University, Portland, OR, USA; Department of Biomedical Engineering, Oregon Health and Sciences University, Portland, OR, USA € NGST • Biochemistry Center Regensburg, University of Regensburg, Regensburg, GERNOT LA Germany CHARLES LIMOUSE • Department of Biochemistry, Stanford University School of Medicine, Stanford, CA, USA

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Contributors

ROSSELLA MAIONE • Department of Molecular Medicine, Sapienza University of Rome, Rome, Italy RODRIGO MALDONADO • Laboratorio de Biologı´a Celular y Molecular Aplicada, Universidad Mayor, Temuco, Chile MAIKA MALIG • Department of Molecular and Cellular Biology and Genome Center, University of California Davis, Davis, CA, USA; Integrative Genetics and Genomics Graduate Group, University of California Davis, Davis, CA, USA 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 JO¨RG MORF • Wellcome-MRC Cambridge Stem Cell Institute, Cambridge, UK JUN-ICHI NAKAYAMA • Division of Chromatin Regulation, National Institute for Basic Biology, Okazaki, Aichi, Japan; Department of Basic Biology, School of Life Science, The Graduate University for Advanced Studies (SOKENDAI), Okazaki, Aichi, Japan MICHEL NEDERLOF • Quantitative Imaging Systems, Inc., Pittsburgh, PA, USA MASAKI NISHIKAWA • School of Engineering, Chemical System Engineering, University of Tokyo, Tokyo, Japan EVGENIA NTINI • Max Planck Institute for Molecular Genetics, Berlin, Germany; Freie Universit€ at Berlin, Berlin, Germany ULF ANDERSSON VANG ØROM • Department for Molecular Biology and Genetics, Aarhus University, Aarhus C, Denmark ANNA POSTEPSKA-IGIELSKA • Division of Molecular Biology of the Cell II, German Cancer Research Center, DKFZ-ZMBH Alliance, Heidelberg, Germany MARIANNA NICOLETTA ROSSI • Division of Rheumatology, IRCCS Bambino Gesu` Children’s Hospital, Rome, Italy XIAOHUA SHEN • Department of Basic Medical Sciences, School of Medicine, Tsinghua University, Beijing, China OWEN K. SMITH • Department of Biochemistry, Stanford University School of Medicine, Stanford, CA, USA; Department of Chemical and Systems Biology, Stanford University School of Medicine, Stanford, CA, USA LOVORKA STOJIC • Cancer Research UK Cambridge Institute, University of Cambridge, Li Ka Shing Centre, Robinson Way, Cambridge, London, UK; Centre for Cancer Cell and Molecular Biology, Barts Cancer Institute, Queen Mary University of London, London, UK AARON F. STRAIGHT • Department of Biochemistry, Stanford University School of Medicine, Stanford, CA, USA; Department of Chemical and Systems Biology, Stanford University School of Medicine, Stanford, CA, USA NAOKI TANI • Liaison Laboratory Research Promotion Center, Institute of Molecular Embryology and Genetics, Kumamoto University, Kumamoto, Japan STEVEN W. WINGETT • Bioinformatics, Babraham Institute, Cambridge, UK NORIMOTO YANAGAWA • Medical and Research Services, Greater Los Angeles Veterans Affairs Healthcare System at Sepulveda, North Hills, CA, USA; David Geffen School of Medicine, University of California at Los Angeles, Los Angeles, CA, USA YAFEI YIN • Department of Basic Medical Sciences, School of Medicine, Tsinghua University, Beijing, China

Chapter 1 Tuning the Expression of Long Noncoding RNA Loci with CRISPR Interference Lovorka Stojic Abstract Long noncoding RNAs (lncRNAs) have emerged as important regulators of gene expression networks. Over 50,000 lncRNA loci have been annotated in the human genome, but only a subset has been involved in regulation of key cellular processes, organismal development, and diseases. Hence, the functional role for the majority of the lncRNA genes remains unknown. With the recent developments of different CRISPR/ Cas9 technologies, the function of lncRNAs can now be examined. CRISPR interference (CRISPRi) is one of these methods that can be used to inhibit the expression of any genomic locus including lncRNAs. This system utilizes catalytically inactive (d)Cas9 fused to KRAB repression domain and single guide RNA against targeted genomic locus. Since CRISPRi has negligible off-target effects and does not involve changes in the underlying genomic DNA sequence, it represents a valuable addition to the existing armamentarium used to investigate lncRNA biology. Key words Long noncoding RNAs, CRISPR/Cas9, CRISPR interference, Gene expression, Genome editing, Transcription, Single guide RNA

1

Introduction The twenty-first-century revolution of genomics revealed that only 2% of the human genomes encodes for proteins, while the majority is transcribed into noncoding RNA (ncRNAs). One group of these ncRNAs, called long noncoding RNAs (lncRNAs), represents the major transcriptional output of the mammalian genome. Until now, more than 50,000 lncRNA loci have been annotated in the human genome [1, 2]. LncRNAs are very heterogonous RNA molecules defined as transcripts longer than 200 nucleotides (nt) with low or no protein-coding potential. They can interact with other RNAs, DNA, and proteins through structural and sequence-specific elements and can be found in the cytoplasm as well as in the nucleus [3]. LncRNAs have been shown to regulate numerous cellular processes through diverse mechanisms including transcriptional and posttranscriptional gene regulation, chromatin

Ulf Andersson Vang Ørom (ed.), RNA-Chromatin Interactions: Methods and Protocols, Methods in Molecular Biology, vol. 2161, https://doi.org/10.1007/978-1-0716-0680-3_1, © Springer Science+Business Media, LLC, part of Springer Nature 2020

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organization, and posttranslational regulation of protein activities [4–6]. Importantly, deregulation of lncRNAs has been associated with many diseases including cancer [7, 8]. Some of the key questions in the emerging field of lncRNA research is how many and which lncRNA genes are functional, and what mechanisms lncRNAs use to regulate many cellular processes they are involved in. However, the identification of functional lncRNAs and their mechanism of action have proven so far a challenging task. LncRNAs can function through their RNA transcripts, the action of their transcription, or via regulatory DNA elements within lncRNA locus [9]. Although initial loss-off-function (LOF) studies were successful in identification of lncRNA functions [10, 11], they relied on methods such as RNA interference (RNAi) or antisense oligonucleotides (such as locked nucleic acids, LNAs) that suffer from large off-target effects and cannot distinguish among different potential mechanisms of action [12– 14]. Furthermore, many studies used only a single LOF method, which probably resulted in discrepancies between cellular and molecular phenotypes even for the same lncRNA [15, 16]. These data strongly suggest that multiple LOF approaches must be used in order to dissect the function of lncRNA genes [17, 18]. In addition to RNAi and LNAs, recent developments in CRISPR/Cas9 technologies allowed scientists to use this powerful genome editing system to manipulate lncRNA expression [19]. These methods include CRISPR/Cas9-mediated gene or promoter deletion, CRISPR/Cas9-mediated insertion of polyA termination signal to terminate lncRNA transcription [18], CRISPR interference (CRISPRi) [20], and CRISPR-Cas13 [21] including the latest CasRx system to cleave target RNA [22]. This chapter will describe a CRISPRi protocol as a tool to inhibit expression of lncRNAs, which can be used to downregulate expression of protein-coding genes as well. CRISPRi is based on deactivated Cas9 (dCas9) fused to a Kru¨ppel-associated box (KRAB) repression domain that can inhibit transcription by blocking RNA polymerase elongation and induce local deposition of heterochromatin marks, such as histone H3 lysine K9 trimethylation [20, 23, 24]. By designing different single guide RNAs (sgRNAs) between
50 and + 200 bp relative to the transcriptional start site (TSS), CRISPRi can downregulate expression of both coding and noncoding genes including lncRNAs [14, 20, 25–28]. Firstly, CRISPRi protocol starts with the production of lentiviruses expressing dCas9-KRAB vector, which can be fused to blue fluorescent protein (BFP) or mCherry, as well as lentiviruses expressing sgRNAs targeting lncRNA of interest. This is performed separately in human embryonic kidney (HEK) 293T cells. Secondly, after harvesting the lentivirus particles from 293T cells, the viruses are transduced to target cells (e.g., HeLa cells) in order to generate polyclonal CRISPRi population stably expressing

CRISPR Interference as a Tool to Inhibit Expression of Long Noncoding RNA Loci

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Fig. 1 Overview of the CRISPRi protocol. (a) Summary of the steps during the CRISPRi protocol. (b) Schematic overview of samples required to perform CRISPRi. (c) Example of CRISPRi-mediated repression of lncRNA SLC25A25-AS1 using four sgRNAs targeting the TSS of SLC25A25-AS1 relative to negative (non-targeting) guide RNA 1 in non-clonal HeLa cells. Expression level was normalized to the geometric mean of GAPDH and RPS18 and was determined with qPCR. n ¼ 3 biological replicates. Error bars, SEM. Statistical significance by two-tailed Student’s t-test; ∗∗P < 0.01 and ∗∗∗∗P < 0.0001

dCas9-KRAB. Since the dCas9-KRAB lentiviral vector contains BFP, the target cells can be sorted with flow cytometry. Thirdly, dCas9-KRAB-BFP-positive cells are expanded for 2–3 days, replated, and transduced with lentiviruses expressing sgRNAs against lncRNA of interest. These cells are then grown for additional 2–4 days before performing qPCR quantification to measure the knockdown efficiency as well as functional analysis, if the function of lncRNA is known (Fig. 1). Some authors have cultured the dCas9-KRAB-BFP-positive cells transduced with sgRNAs targeting lncRNAs for 10–20 days before analyzing the impact of lncRNAs on cell growth [25]. CRISPRi has a great potential in inhibiting expression of lncRNA genes, but it cannot distinguish cis- and trans-acting functions of lncRNAs, cis-mediated regulation due to act of lncRNA transcription, and/or enhancer like function of lncRNA loci

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[9]. These parameters should be taken into consideration when using CRISPRi to characterize lncRNA mode of action. Although minimal cis effects were reported with CRISPRi [20], CRISPRi could have some impact on cis regulatory elements such as enhancers, which are embedded within or near the lncRNA TSS. Indeed, CRISPRi was used to identify cis regulatory regions within MYC [29]. Furthermore, CRISPRi-mediated suppression of lncRNA PVT1 revealed that PVT1 promoter acts as a DNA element repressing MYC transcription, while PVT1 RNA product sustains MYC protein level [30], despite PVT1 initially being described as a transacting lncRNA [31]. Thus, through combinations of different LOF methods to manipulate lncRNA expression, it was demonstrated that lncRNAs could have dual DNA and RNA regulatory elements with opposing function [32, 33]. Furthermore, different cellular phenotypes were observed when depleting the lncRNA SLC25A25-AS1 with LNAs and CRISPRi, despite similar knockdown efficiency [14]. When compared to other LOF methods such as RNAi and LNAs [34, 28], CRISPRi is highly specific and has minimal off-target effects especially when used in non-clonal populations [14, 20]. CRISPRi was also successfully employed in a large-scale screening assay targeting more than 16,000 lncRNAs in 7 different human cell lines. In this approach, Lim et al. identified ~500 lncRNA loci involved in cell growth control in a cell-type-specific manner [25]. Attention should also be paid when targeting bidirectional promoters or lncRNAs near other transcriptional units as CRISPRi in these conditions might not be the best method of choice [27]. Ideally, CRISPRi should be used to manipulate expression of lncRNAs distal from protein-coding genes. Finally, since lncRNAs have multiple TSS, GENCODE and FANTOM consortium databases should be consulted in order to design sgRNAs targeting most of the lncRNA isoforms.

2

Materials When working with RNA, wear gloves and try to avoid RNase and DNase contamination. All surfaces and pipettes are treated with RNaseZap, an RNase decontamination solution. Perform viral handling after proper laboratory safety training and use dedicated cell culture hoods and incubators for the viral work.

2.1 Required Equipment

1. qPCR machine (e.g., QuantStudio 6 Flex). 2. Cell sorter (e.g., BD FACSAria III). 3. FlowJo v7.1 software. 4. Spectrophotometer (NanoDrop). 5. 37  C shaking incubator.

CRISPR Interference as a Tool to Inhibit Expression of Long Noncoding RNA Loci

5

6. Refrigerated and non-refrigerated microcentrifuge. 7. 15 ml and 50 ml centrifuge tubes. 8. Nuclease-free 1.7 ml microcentrifuge tubes. 9. 5 ml polysterene round bottom tubes with cell strainer cap. 10. 1.8 ml cryotube vials. 11. Category 2 cell culture facility. 12. qPCR tubes/microplates. 13. Sterile filters. 14. 6-well and 10-cm sterile cell culture dishes. 2.2 Required Reagents and Solutions 2.2.1 Cloning

1. T4 Polynucleotide Kinase. 2. Quick ligation kit. 3. Sterile and autoclaved LB agar plates containing 100 μg/ml ampicillin. 4. Competent DH5α E. coli. 5. BstXI and BlpI restriction enzymes. 6. Plasmid DNA purification kits. 7. Quick PCR Purification Kit. 8. Plasmid Gel extraction kit.

2.2.2 Required Vectors and Primers

1. pU6-sgRNA EF1Alpha-puro-T2A-BFP (Addgene, #60955). 2. pHR-SFFV-dCas9-BFP-KRAB (Addgene, #46911). 3. Second-generation packaging plasmid psPAX2 (Addgene, #12260) and the envelope plasmid pMD2.G (Addgene, #12259). 4. SgRNA oligonucleotides (e.g., Sigma), reconstituted in 1 low EDTA TE buffer at 100 μM. 5. mU6 forward primer, used for Sanger sequencing (Table 1). 6. Standard unmodified desalted oligonucleotides.

2.2.3 Cell Culture

1. HeLa, RPE1, and HEK293T cells were obtained from the American Type Culture Collection (ATCC). All cell lines were verified by short tandem repeat (STR) profiling and tested negative for mycoplasma contamination. 2. Trans-Lt1 transfection reagent. 3. 0.05% trypsin-EDTA. 4. Phosphate-buffered saline, pH 7.4. 5. Dulbecco’s Modified Eagle’s Medium F12 Nutrient Mixture used for RPE1 cells. 6. Dulbecco’s Modified Eagle’s Medium used for HeLa and HEK293T cells.

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Table 1 List of qPCR primer sequences Expression primers Forward primer (50 –30 )

Reverse primer (50 –30 )

GAPDH

CAACAGCCTCAAGATCATCAG

ATGGACTGTGGTCATGAGTC

RPS18

ATCCCTGAAAAGTTCCAGCA

CCCTCTTGGTGAGGTCAATG

ACTB (β-actin)

GTTACACCCTTTCTTGACAAA

GTCACCTTCACCGTTCCAGTT

H19

CTGGCTTGGCAGACA GTACA

TCCCTCCTGAGAGCTC ATTC

SLC25A25-AS1

CACCTAGGCCCAGCTT CTC

TTCAGACACGCTCCAG GTAA

mU6

GAGATCCAGTTTGGTTAGTACCGGG

7. Reduced serum medium. 8. 10% fetal bovine serum (FBS). 9. 5 μg/ml polybrene. 2.2.4 RNA Extraction and qPCR

1. RNaseZap. 2. RNeasy Kit. 3. DNase I. 4. RNase-free water. 5. Reverse Transcription Kit. 6. SYBR Green Master Mix. Conditions for qPCR: 95  C for 20 s followed by 40 cycles of 95 C for 1 s and 60  C for 20 s. At least two housekeeping genes (GAPDH, RPS18, β-actin) are used to normalize expression levels using the 2
ΔΔCT method. Table 1 includes all qPCR primer sequences. 

3

Methods The CRISPRi protocol describes all the steps from the design and cloning of sgRNAs targeting gene of interest to lentiviral production and analysis of gene expression (Fig. 1a, b). The protocol can be applied to lncRNAs as well as protein-coding genes and has been established in HeLa and hTERT-immortalized retinal pigment epithelial cell line, RPE1. In this protocol, I will use lncRNA SLC25A25-AS1 as an example of CRISPRi-mediated repression of lncRNAs in HeLa cells (Fig. 1c).

CRISPR Interference as a Tool to Inhibit Expression of Long Noncoding RNA Loci

3.1 Design and Cloning of sgRNA Guides

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1. Design sgRNA sequences using the online tools such as Benchling, CHOPCHOP, and/or Broad Institute GPP available at https://zlab.bio/guide-design-resources. We usually use the CRISPR Targets 10K Track settings on the USCS genome browser (GRCh37/hg19) (see Note 1). 2. Each sgRNA sequence is 20 nt in length and is designed against the genomic window of
50 to +200 bp relative to the TSS, as described previously [20]. We determine the TSS location using the NCBI RefSeq database and FANTOM CAGE database (see Note 2). 3. Use the Basic Local Alignment Search Tool to determine whether any of the sgRNA sequences are located within annotated genes other than the intended target. If so, discard these guide sequences. Potential off targets can be analyzed with CRISPR RGEN Cas-OFFinder web tool (http://www. rgenome.net/cas-offinder/). 4. Add additional sequences to the sgRNA sequences in order to clone them into the 50 BstXI-BlpI30 digested vector backbone of a pU6-sgRNA EF1Alpha-puro-T2A-BFP expression plasmid (Addgene, #60955, see Fig. 2 and Table 3). In case sgRNA does not start with a guanine (G), add a G at the 50 for its better targeting efficiency. 5. Order sgRNA oligonucleotides (e.g., Sigma) and reconstitute them in 1 low EDTA TE buffer at 100 μM. Store them at
20  C. By ordering already phosphorylated oligonucleotides, you can avoid Subheading 3.2.

3.2 Phosphorylation and Annealing of Oligonucleotides

1. Once the sgRNA oligonucleotides have been designed and ordered, you can set up the following reaction for their phosphorylation, annealing, and cloning into pU6-sgRNA EF1Alpha-puro-T2A-BFP expression plasmid (Addgene, #60955). Phosphorylation of sgRNA oligonucleotides using the M0201S NEB kit 1 μl sgRNA oligonucleotide forward (100 μM). 1 μl sgRNA oligonucleotide reverse (100 μM).

5’CCACCTTG TTG 3’GGTG GAACAAC

sgRNA (20nt) rev.compl.sgRNA (20nt)

TAAGC 3’ GTTTAAGAGC CG 5’ CAAATTCTCGATT

sgRNA Addgene plasmid (#60955)

Fig. 2 The cloning of sgRNA sequences into the pU6-sgRNA EF1Alpha-puro-T2A-BFP (Addgene, #60955). Additional overhangs at the 50 and 30 ends are added to the 20-nt guide sequence to enable their cloning into a 50 BstXI-BlpI30 digested backbone of #60955 vector

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1 μl T4 Polynucleotide Kinase reaction buffer (10). 0.5 μl T4 Polynucleotide Kinase (NEB). 6:5 μl RNase
f ree water 10 μl Total reaction volume 2. Seal the tubes with the parafilm and incubate in the water bath or thermomixer for 30 min at 37  C. 3. Place the phosphorylated oligonucleotides in a glass beaker with boiling ddH2O and leave them to gradually anneal while cooling to room temperature (RT). This usually takes between 2 and 4 h. Annealed oligonucleotides are briefly spun down at 14,000  g for 1 min. 3.3 Ligation of Oligonucleotides

1. Use Quick ligation kit (e.g., NEB) to ligate the annealed oligonucleotides into previously digested Addgene #60955 vector using the BstXI and BlpI enzymes. The annealed duplexes were diluted 1–200 in ddH2O, and 1 μl was used in the following reaction: 1 μl of diluted annealed oligonucleotides from Subheading 3.2 reaction (dilution 1–200). 50 ng (1 μl) of digested vector backbone #60955. 5 μl Quick ligase buffer (2, NEB). 1 μl Quick Ligase (NEB). 2 μl RNase
f ree water 10 μl Total reaction volume 2. Incubate the reaction for 20 min at RT. As a ligation control, we usually include a sample without any annealed oligonucleotides.

3.4 Transformation of Ligated Oligonucleotides into Bacteria

1. Thaw DH5α competent cells on ice and add ~2 μl of Subheading 3.3 ligation reaction to ~50 μl of E. coli by gently flicking the bottom of the tube. Place the mixture on ice for 30 min. 2. Perform the heat shock reaction by placing the tubes in water bath for 1 min at 42  C. 3. Immediately put the tubes back on ice for 2 min. 4. Recover the bacteria by adding ~900 μl Luria broth (LB) media to the mixture and transfer the tubes into a 37  C shaking incubator for 45 min. 5. Spin down the bacteria and resuspend them in ~100 μl LB media. 6. Plate the mixture on pre-warmed LB agar plates containing ampicillin (Amp, 100 μg/ml) and incubate overnight at 37  C.

CRISPR Interference as a Tool to Inhibit Expression of Long Noncoding RNA Loci

9

7. Next day, inoculate bacteria colonies for overnight culture in total volume of 3 ml LB media containing Amp. We usually inoculate at least four colonies/sgRNA oligonucleotide. 8. Purify DNA from bacterial cultures (e.g., Qiagen). 9. Elute the DNA. To increase the yield, we usually elute the plasmid DNA two times by adding ~10 μl 1 TE buffer or ddH20. 10. Quantify the DNA by NanoDrop and sequence the plasmid DNA to confirm insertion of the correct cloned sgRNA sequence. 11. For Sanger sequencing, dilute the DNA samples to a final concentration of ~100 ng/μl. We usually sent ~10 μl of each sample for Sanger sequencing including 3 μl of mU6 primer (10 μM). 12. Retransform the DNA samples with correct cloned sgRNA sequence into E. coli cells, as described above. 13. Take single bacteria colonies and culture them overnight for subsequent DNA plasmid preparation (e.g., Qiagen). 14. Elute the DNA two times in ~200 μl TE buffer or ddH2O.Purified DNA is stored at
20  C and ready to be used for the lentiviral production. 3.5 Lentiviral Production

1. Use second-generation lentiviral packaging system (psPAX.2 and pMD2.G, Addgene #12260 and #12259, respectively) to produce the lentiviruses. 2. One day prior to transfection, seed 4  106 HEK293T cells in a 10-cm dish in 10 ml of growing medium and incubate the cells overnight at 37  C with 5% CO2. 3. Transfect the cells with 15 μg of DNA, which is composed of 9 μg of the lentiviral vector DNA containing the transgene (e.g., dCas9-BFP-KRAB), 4 μg of the packaging vector psPAX.2, and 2 μg of the envelope vector pMD2.G. The final transfection volume is 1.5 ml (including Trans-Lt1 reagent), which was filled up with OptiMEM medium (see Note 3). 4. Vortex the pre-warmed Trans-Lt1 reagent and pipet ~45 μl of the Trans-Lt1 reagent to the DNA mixture from Subheading 3.5, step 2. Avoid any contact with the sides of the tube. 5. Incubate the transfection mixture at RT for 25 min. 6. Prior to transfection, replace the media by adding 14 ml of fresh growing media. 7. Add the transfection mixture dropwise to the cells, gently rock and incubate the cells for 24 h in a 37  C incubator with 5% CO2.

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8. The following day replace the old medium by adding 7 ml of fresh growing media and incubate the cells for another 24 h in a 37  C incubator with 5% CO2. 9. Collect the viral supernatant 48 and 72 h post transfection in a 15 ml Falcon tube. 10. Spin down the virus at 1800  g for 5 min at 4  C and filter the mixture through a 45 μm filter attached to a syringe. The virus can be stored at 4  C for up to a week. For long-term storage, viral supernatant is frozen in cryovial tubes and stored at
80  C. When you are ready to use the virus, thaw an aliquot on ice and bring all the reagents to RT. 3.6 Viral Transduction of HeLa Cells with dCas9-BFP-KRAB Lentiviral Vector and FACS Sorting

1. Culture HeLa cells in complete growing medium in a 37  C incubator with 5% CO2. 2. Simultaneously plate and transduce the cells with the lentivirus containing the pHR-SFFV-dCas9-BFP-KRAB to achieve higher infection rates. We usually plate 1.0  105 or 5.0  104 HeLa cells in 6-well or 12-well plates, respectively, and transduce the cells with the virus at 1–1 dilution with growing medium in the presence of polybrene (5 μg/ml) (see Note 4). 3. 24 h after lentivirus transduction remove the transduction mixture, and add the new HeLa medium for additional 48 h. 4. Wash the cells once with 1 PBS, trypsinize and resuspend in ~500 μl of 1 PBS. 5. Sort the dCas9-BFP-KRAB expressing cells using the BD FACSAria III cell sorter (CRUK Flow Cytometry Core Facility) (see Note 5). A representative histogram of dCas9-BFP-KRAB sorted cells is shown in Fig. 3. 6. Detect the expression of BFP fluorescent proteins using MACSQuant VYB and analyze the data using the FlowJo v7.1 software. 7. Plate sorted BFP-positive cells in 6-well plates to create a stable non-clonal cell population. We usually grow the cells for additional 72 h before replating them (see Note 6).

3.7 CRISPRiMediated Depletion of lncRNAs

1. Three to four days after FACS sorting, plate the dCas9-BFPKRAB transduced cells in 12-well or 6-well plates. 2. Add the lentivirus containing sgRNAs targeting lncRNA of interest or lentivirus containing two negative (non-targeting) control guide RNAs (see Note 7). 3. Dilute the lentivirus with HeLa medium at 1:1 dilution and add cationic polymer polybrene to facilitate viral transduction (5 μg/ml) (see Note 8).

CRISPR Interference as a Tool to Inhibit Expression of Long Noncoding RNA Loci

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Fig. 3 Representative histogram of untransduced (red) and transduced HeLa cells with dCas9-BFP-KRAB (blue) after sorting for BFP-positive cells

4. Remove the supernatant 24 h later and add fresh medium for another 48 h before RNA collection. We use non-transduced cells as a negative control (see Note 9). 5. Perform functional analysis if the function of lncRNA is known. In the case of Ch-TOG, we usually do live-cell imaging and analyze the timing of the mitotic progression (see Note 10). 3.8 RNA Extraction, cDNA, and Quantitative Real-Time PCR (qPCR)

1. Evaluate the knockdown of the target gene by extracting RNA using the RNeasy Kit (e.g., Qiagen). As an example of CRISPRi-mediated depletion of lncRNAs, we targeted SLC25A25-AS1 using four sgRNAs targeting TSS of SLC25A25-AS1. 2. Elute RNA in ~30 μl nuclease-free water (if working with HeLa cells in 6-well plates) and quantify RNA concentration with the NanoDrop. RNA is stored at
80  C after snap-freezing. 3. Perform cDNA synthesis using the QuantiTect Reverse Transcription Kit (e.g., Qiagen) and qPCR on a Quantstudio™ 6 Flex Real-Time PCR System (Applied Biosystems). The knockdown efficiency of SLC25A25-AS1 was verified by qPCR (Fig. 1c).

4

Notes 1. When establishing the CRISPRi protocol for the first time we recommend to standardize the method on positive control genes. We use two published negative control sgRNAs [20]

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and several positive control sgRNAs targeting lncRNAs (H19, SLC25A25-AS1) [20, 14] and protein-coding genes (Ch-TOG) [14]. Depletion of Ch-TOG leads to well-established cellular phenotype of mitotic delay, which can easily be identified as an increase in the number of mitotic cells [35]. Depletion of Ch-TOG and H19 usually leads to 80–90% reduction at their mRNA levels as shown with qPCR in HeLa cells. The primers and sgRNAs sequences are present in Tables 1 and 2. We also suggest to use the most potent sgRNA targeting B4GALNT1, as shown previously [20]. 2. We advise testing at least two sgRNAs targeting different regions of lncRNA TSS, as shown for lncRNA SLC25A25AS1 (Fig. 1c). For some lncRNAs, such as MALAT1, we tried five different sgRNAs but we were able to deplete MALAT1 only to 50% using one guide in HeLa cells [14]. 3. We routinely use pHIV-Zsgreen plasmid as a positive control for lentiviral transduction of 293T cells. 4. We have tried to use dCas9-mCherry-KRAB (Addgene, #60954) instead of dCas9-BFP-KRAB (Addgene, #46911) and did not observe any differences in lncRNA repression by qPCR. 5. During FACS we sort the top half of the BFP-positive cells based on BFP signal intensity. Our transduction efficiency is ~50–60% for HeLa cells (Fig. 3), and ~20–30% for RPE1 cells. 6. We don’t recommend the use of single cell cloning in CRISPRi as it introduces changes in the transcriptomic background even in the absence of any guide RNAs [14]. While it is possible to freeze several vials of CRISPRi non-clonal populations, we always prepared fresh population of cells before sgRNA transduction. Table 2 List of CRISPR guide RNA sequences Targeted lncRNA

Guide-ID

Guide sequence

References

SLC25A25-AS1

sgRNA 1

GATGGAGAATGTAAGGGTAC

Stojic et al. [14]

SLC25A25-AS1

sgRNA 9

TGCAGAGAACGGAGGCATGC

Stojic et al. [14]

SLC25A25-AS1

sgRNA 16

TGCAGGGATGGAGAATGTAA

SLC25A25-AS1

sgRNA 18

GCACAGGGATACAGGAATGG

Negative control

sgRNA 1

GCGCCAAACGTGCCCTGACGG

Gilbert et al. [20]

Negative control

sgRNA 2

GTGCGATGGGGGGGTGGGTAGC

Gilbert et al. [20]

Ch-TOG

sgRNA 94

TAAGCCGTTTGAAACCGCTT

Stojic et al. [14]

H19

sgRNA 2

GCTAGGACCGAGGAGCAGGGTG

Gilbert et al. [20]

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Table 3 List of sequences added to sgRNAs targeting gene of interest 50 end

sgRNA sequence

30 end

Forward primer 50 –30

TTG

20-nt guide sequence

GTTTAAGAGC

Reverse primer 50 –30

TTAGCTCTTAAAC

Reverse complement guide sequence

CAACAAG

7. We usually transduce cells with single sgRNAs, but it is possible to combine 2–4 sgRNAs in a single guide treatment. 8. We have tried different dilutions of lentivirus for infection of HeLa cells and found that 1–1 dilution yields the best results. To enhance viral transduction efficiency, we have tested ultracentrifugation and ViralBoost method but found no evidence of increased infection rate in HeLa cells. Based on these results, we decided to use an unconcentrated virus in combination with polybrene. 9. Although the lentiviral sgRNA vector contains puromycin resistance, we did not detect any differences in lncRNA repression with qPCR after further selecting the dCas9-BFP-KRABpositive cells with 0.5 μg/ml puromycin for 3 days. 10. For qPCR and functional analysis after lncRNA depletion, we advise to include cells without any transduction as well as the cells transduced with dCas9-BFP-KRAB in the absence of any guide RNAs (Fig. 1b). This will account for any possible off-target effects introduced by dCas9-KRAB transduction.

Acknowledgments I would like to thank Jasmin Mangei and Valentina Quarantotti for critically reading the manuscript. I also thank Richard Grenfell from the FACS Core Facility at the CRUK Cambridge Institute for help with the FACS sorting, and Alasdair Russel for providing us with the pHIV-Zsgreen vector. This work was supported by Cancer Research UK (C14303/A17197 to Fanni Gergely and Duncan Odom; A24455 to Fanni Gergely, and A20412 to Duncan Odom). We also acknowledge the support of the University of Cambridge, the Wellcome Trust (WT202878; Duncan Odom), European Research Council (615584; Duncan Odom), and Hutchison Whampoa Limited.

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Chapter 2 Identification of Chromatin Binding Sites for Long Noncoding RNAs by Chromatin Oligo-Affinity Precipitation (ChOP) Marianna Nicoletta Rossi

and Rossella Maione

Abstract Revealing the interactions of long noncoding RNAs (LncRNAs) with specific genomic regions is of basic importance to explore the mechanisms by which they regulate gene expression. Chromatin oligo-affinity precipitation (ChOP) technique was the first method developed to analyze the association of LncRNAs with genomic regions in the chromatin context. The first step of the procedure is cell cross-linking, aimed at stabilizing the RNA–protein–DNA complexes. Next, after chromatin fragmentation, the RNA complexes are pulled down through hybridization with antisense oligonucleotides tagged with biotin and purification with anti-biotin antibody. After extensive wash, the RNA-interacting chromatin is eluted by RNase treatment. Subsequent protein elimination and DNA purification allow to retrieve DNA fragments for following analyses such as qPCR or sequencing. In the present chapter, we describe the ChOP protocol, as used in our laboratory for investigating the interaction of the LncRNA Kcnq1ot1 with chromatin at specific regulatory regions of the Cdkn1c locus. Key words LncRNA, Chromatin, ChOP, Hybridization capture, Oligonucleotide precipitation

1

Introduction Long noncoding RNAs (LncRNAs) have emerged as a large class of regulators of gene expression, playing critical roles in many physiologic and pathologic processes. They act either in the nucleus or in the cytoplasm and can affect chromatin architecture, transcription, RNA maturation, stability, and translation, using various mechanisms of action still incompletely understood, which involve their interaction with genomic DNA, RNA, and proteins [1–3]. To gain further insight into the function of LncRNAs, a great effort is now being spent in the identification of their interaction partners. Since LncRNAs are frequently localized to the nucleus and, in most cases, to chromatin, a large number of studies are being focused on the development and improvement of technologies aimed at mapping and characterizing their chromatin interactions.

Ulf Andersson Vang Ørom (ed.), RNA-Chromatin Interactions: Methods and Protocols, Methods in Molecular Biology, vol. 2161, https://doi.org/10.1007/978-1-0716-0680-3_2, © Springer Science+Business Media, LLC, part of Springer Nature 2020

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Marianna Nicoletta Rossi and Rossella Maione

Chromatin oligo-affinity precipitation (ChOP) technique was the first method developed to analyze the interactions of LncRNAs with genomic regions in the chromatin context [4]. Along the lines of chromatin immunoprecipitation (ChIP), a ChOP assay involves the enrichment of a specific LncRNA from cross-linked chromatin, through hybridization with biotin-tagged antisense oligonucleotides, followed by qPCR analysis or deep sequencing of the isolated DNA. Other related methods, also based on the capture of the LncRNA of interest by complementary oligonucleotides, include chromatin isolation by RNA purification (ChIRP) [5], capture hybridization analysis of RNA targets (CHART) [6], and RNA antisense purification (RAP) [7]. It is worth mentioning that all the RNA hybridization-capture methods, in addition to allowing the study of the interactions of LncRNAs with genomic regions, can be also exploited for the characterization of other types of interacting partners of LncRNAs, such as RNAs and proteins, if used in combination with qRT-PCR or RNA sequencing and with western blot or mass spectrometry, respectively [8]. The originally published methods differed for several technical parameters, concerning the cross-linking agents, the criteria for designing oligonucleotides, and the elution conditions. For example, ChOP and CHART methods are recommended to perform cross-linking with formaldehyde, ChIRP with glutaraldehyde, and RAP with a combination of formaldehyde and disuccinimidyl glutarate. Moreover, ChOP and CHART were based on the use of a limited number of biotinylated oligonucleotides, while ChIRP and RAP on the tiling of the entire length of the LncRNA with capturing oligonucleotides. An additional difference was the length of the oligonucleotides: 20–25 nucleotides for ChOP, ChIP, and CHART and 90–120 nucleotides for RAP. However, in subsequent applications, the protocols were slightly modified, also through their mixing, which resulted in increased similarities between the methods [8, 9]. The choice of the optimal approach to a specific study is not so obvious since each of the above methods has both advantages and disadvantages and the efficiency in capturing the LncRNA as well as in preserving its particular interactions will be influenced by any of the structural and functional properties of specific transcript. The ChOP method was applied in most cases to analyze the interactions of LncRNAs with specific loci, as reported, for example, for Alu RNA [4], Kcnq1ot1 [10, 11], H3K4me2/WDR5 chromatin-enriched LncRNAs [12], and Scat7 [13]. However, slightly modified ChOP methods, combined with high-throughput sequencing, were also used for mapping the genome-wide occupancy of mrhl [14] and MEG3 LncRNAs [15]. The protocol described here was set up to investigate the interactions of the LncRNA Kcnq1ot1 with specific chromatin regions of the Cdkn1c/Kcnq1 locus [10]. Kcnq1ot1 is a macro-

Chromain Oligo Affinity Precipitation

19

LncRNA of about 90 kb, unspliced and exclusively localized to the nucleus [16]. It is involved in the silencing of the paternal Cdkn1c/ Kcnq1 imprinted domain and, as we have recently discovered, also in the repression of the maternal non-imprinted Cdkn1c (p57) allele [10]. This protocol, developed by modifying and implementing previously published ChOP protocols [4, 11], was used for differentiating mouse muscle cells as well as for mouse embryo fibroblasts, but it may well be applied also to cell types of different origins, grown either adherent or in suspension culture.

2

Materials General considerations: Always wear gloves and take care to avoid RNase and DNase contamination during all stages of this protocol. Treat all surfaces and pipettes with EtOH 70% before starting. Prepare all solutions for ChOP with sterile diethylpyrocarbonate (DEPC)-treated water.

2.1 Reagents and Equipment

1. 0.5 M EGTA. 2. 1 M MgCl2. 3. 1 M Tris–HCl (pH 7.4). 4. 1 M Tris–HCl (pH 7.9). 5. 1.25 M glycine. 6. 10% deoxycholate. 7. 10% NP-40. 8. 10% Triton-X-100. 9. 10% SDS. 10. 11% formaldehyde. 11. 1 M EDTA. 12. 1 M LiCl. 13. 20 μg/μl proteinase K solution, RNA grade. 14. 5 M NaCl. 15. 500 mM HEPES pH 8.0. 16. Agarose. 17. Agarose gel electrophoresis apparatus. 18. Anti-biotin antibody. 19. Aprotinin. 20. BSA 1 mg/ml. 21. DEPC. 22. DNA ladder. 23. Fume hood.

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24. Leupeptin. 25. Liquid nitrogen. 26. Multiblock heater and shaker. 27. NanoDrop spectrophotometer. 28. Nuclease-free water. 29. Orthovanadate. 30. pH indicator. 31. Phosphate-buffered saline (PBS). 32. PMSF. 33. Protein A Sepharose. 34. RNase A. 35. RNase H. 36. RNase inhibitor. 37. Rotating device. 38. Sonicator. 39. Syringe. 40. TAE buffer. 41. TE buffer. 42. Trypsin. 43. Yeast RNA. 44. Phenol:CHCl3:isoamyl alcohol saturated with 10 mM Tris pH 8.0, 1 mM EDTA. 45. Chloroform. 46. 2-Propanal. 47. 70% EtOH. 48. GlycoBlue co-precipitant. 49. Real-Time Thermal Cycler. 50. SYBR Green PCR Master Mix. 51. Standard unmodified desalted oligonucleotides. 2.2

Buffers

Use molecular biology grade reagents and nuclease-free water to make up all buffers and solutions. Solutions should be stored at 4
C unless stated otherwise. 1. Fixing Solution: 11% formaldehyde, 0.1 M NaCl, 1 mM EDTA, 0.5 mM EGTA, 50 mM HEPES pH 8.0. 2. Buffer A: 3 mM MgCl2, 10 mM Tris–Hcl (pH 7.4), 10 mM NaCl, 0.5% NP-40. 3. Buffer B: 50 mM Tris–HCl (pH 7.9), 10 mM EDTA, 1% SDS. Add protease and RNase inhibitors fresh before use.

Chromain Oligo Affinity Precipitation

21

4. Buffer C: 15 mM Tris–HCl (pH 7.9), 150 mM NaCl, 1 mM EDTA, 1% Triton-X-100. Add protease and RNase inhibitors fresh before use. 5. Buffer D: 15 mM Tris–HCl (pH 7.9), 150 mM NaCl, 1 mM EDTA, 0.5% NP-40. Add protease and RNase inhibitors fresh before use. 6. Low Salt Buffer: 0.1% SDS, 1% Triton-X-100, 2 mM EDTA, 20 mM Tris–HCl (pH 8.1), 150 mM NaCl. 7. High Salt Buffer: 0.1% SDS, 1% Triton-X-100, 2 mM EDTA, 20 mM Tris–HCl (pH 8.1), 500 mM NaCl. 8. LiCl Buffer: 0.25 M LiCl, 1% NP-40, 1% deoxycholate, 1 mM EDTA, 10 m Tris–HCl (pH 8.1). 9. TE Buffer: 10 mM Tris–HCl, 1 mM EDTA. 10. Elution Buffer: 1 TE, 0.5% SDS.

3

Methods

3.1 Cell Culture and Fixation

Perform all steps in sterile conditions. Grow cells in appropriate media (i.e., DMEM) supplemented with 10% fetal bovine serum and 100 U penicillin/0.1 mg/ml streptomycin. Grow cells at the appropriated confluence (70–90%) in tissue culture plates. Approximately 4–6  106 cells are needed for each oligo-precipitation (see Note 1). 1. On day 1, perform fixation in a fume hood. Add fixing solution directly to the growth medium (the fixing solution is 10X so we typically add 1 ml of fixing solution to 9 ml of growth media for one 10 cm dish) (see Note 2). Incubate for 10 min at 37
C (see Note 3). 2. Add glycine to a final concentration of 0.125 M in phosphatebuffered saline (PBS) and incubate for 5 min at 4
C and 5 min at room temperature with gentle shaking. 3. Remove supernatants and discard them (see Note 4). 4. Wash cells three times with cold PBS, discarding each wash as hazardous chemicals. 5. Add 1 ml of cold PBS supplemented with 100 mM PMSF for each 10 cm tissue culture dish. 6. Scrape cells and collect scraped cells in 15 ml conical tubes. 7. Centrifuge at 1200 rcf for 7 min. Aspirate the supernatant and immediately freeze down the cell pellet in liquid nitrogen. Cell pellets can be stored at 80
C.

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3.2 Cell Lysis and Sonication

Perform all steps on ice or at 4
C. 1. On day 2, thaw frozen cell pellets on ice. 2. Resuspend cell pellets in prechilled buffer A (consider 1 ml for each 10 cm tissue culture dish). 3. Incubate for 5 min on ice with frequent mixing and inverting of the tubes. 4. Centrifuge at 2600 rcf for 5 min in a refrigerated benchtop centrifuge. 5. Wash cell pellets with 500 μl of Buffer A. Centrifuge again as above. 6. Discard the supernatant. The pellets contain the cell nuclei. 7. Resuspend the nuclei in 500 μl of Buffer B, incubate for 10 min on ice, add an equal volume of Buffer C, and proceed to sonication (see Notes 5 and 6). 8. Centrifuge for 10 min at 4
C at 20,000 rcf. 9. Take 5 μl and add 995 μl 100 mM NaCl, incubate for 30 min at 37
C, and then quantify the chromatin for DNA content with a NanoDrop spectrophotometer. 10. Prepare aliquots of 150 μg of chromatin and store them at 80
C or proceed to incubation with biotin-labeled oligonucleotides.

3.3 RNA Precipitation

1. On day 3, take 50 μl of Protein A Sepharose for each sample (see Note 7). 2. Add 1 ml of Buffer D, invert several times until the complete resuspension of the resin, and then centrifuge at 1000 rcf for 2 min. Repeat twice. 3. Incubate Protein A Sepharose with 400 μg/ml yeast RNA and 800 μg/ml of bovine serum albumin (BSA) in buffer D for 4 h at 4
C in rotation to block aspecific sites of interaction. 4. Wash three times with 1 ml of Buffer D and incubate overnight at 4
C with 5 μg of anti-biotin antibody. 5. In parallel, incubate sonicated chromatins overnight at 4
C with 25 pmol of biotin-labeled oligonucleotides antisense to your RNA of interest or biotin-labeled scrambled oligonucleotide (see Note 8). 6. On day 4, wash Protein A Sepharose plus anti-biotin antibody three times with 1 ml of Buffer D (this step is necessary to eliminate unbound antibody). 7. Add 50 μl of beads with bound antibody to samples containing chromatins plus biotin-labeled oligonucleotides and incubate for 4 h at 4
C.

Chromain Oligo Affinity Precipitation

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8. Take the supernatant of the biotin-labeled scrambled oligonucleotide sample as input. 9. Wash steps: centrifuge each sample at 2600 rcf at 4
C for 3 min, remove the supernatants with a syringe, add 1 ml of wash buffers, incubate for 5 min in rotation at 4
C, and centrifuge again. Wash each sample twice with low salt buffer, twice with high salt buffer, once with LiCl buffer, and once with TE buffer (see Notes 9 and 10). 10. To elute the RNA–DNA–protein complexes, incubate the samples with 300 μl of elution buffer plus 10 U of RNase H and 10 μg of RNase A for 15 min on a vortex or in a shaker multiblock at room temperature. 11. Centrifuge at 20,000 rcf for 5 min at 4
C. Recuperate the supernatants. 12. Add 240 μg of proteinase K to each sample and incubate for 5 h at 62
C with gentle shaking (see Note 11). 13. Take the input samples and treat them with the same amount of RNase A and RNase H. 14. Incubate input samples as oligo-precipitated samples with proteinase K. 3.4 DNA Extraction and Amplification

1. Add to all samples NaCl 0.2 M final concentration. 2. Add an equal volume (300 μl) of PhOH/chloroform/isoamyl per sample. Shake vigorously for 10 min and spin down on a benchtop centrifuge at 16,000 rcf for 5 min at 4
C. 3. Take aqueous phase from the top. 4. Add an equal volume (300 μl) of 2-propanal, mix well, and store at 20
C overnight. 5. Spin samples at 16,000 rcf for 30 min at 4
C. 6. Remove supernatant carefully. Add 1 ml 70% EtOH and vortex to mix. Spin at 16,000 rcf for 5 min at 4
C. 7. Remove supernatant by pipetting. 8. Air dry. 9. Resuspend in 50 μl H2O. 10. Measure DNA concentration of each sample with a NanoDrop spectrophotometer. 11. DNA samples are ready for following analysis such as qPCR (see Notes 12 and 13). A schematic outline of the critical steps of the ChOP technique is shown in Fig. 1.

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Fig. 1 Schematic depiction of the ChOP workflow, from cross-linking of LncRNA–chromatin complexes to DNA fragment isolation

4

Notes 1. A single ChOP experiment consists in the oligo-precipitation with at least 1–2 specific oligonucleotides and a non-targeting control (scramble) oligonucleotide, so it is necessary to grow

Chromain Oligo Affinity Precipitation

25

up 2–3 times this number of cells depending on the number of reactions. Moreover, depending on LncRNA and cell types, it may be necessary to adjust the amount of starting material to improve the signal versus noise ratio. 2. Prepare each time a fresh fixation solution by adding the appropriate volume of 11% formaldehyde. 3. The correct time of fixation should be empirically determined. Too short or too long cross-linking times can lead to DNA loss and/or elevated background. Cross-linking can affect both efficiency of chromatin shearing and specificity of immunoprecipitation. In general, shorter cross-linking times (5–10 min) may improve shearing efficiency. On the other hand, shorter cross-linking times might reduce the efficiency of cross-linking for those RNAs that do not interact directly with the DNA and thus the yield of precipitated chromatin. In any case, do not cross-link for longer than 30 min as cross-links of more than 30 min cannot be efficiently sheared. 4. Formaldehyde and formaldehyde-containing solutions are hazardous chemicals and need to be disposed of in the appropriate manner. 5. The number and duration of sonication pulses depend on cell type and target LncRNA and need to be empirically determined. Please bear in mind that over-sonication of the samples can lead to degradation of the target LncRNA. Typically for fibroblasts and muscle cells, 6–10 pulses of 10 s each with at least 30 s recovery on ice are necessary. To optimize sonication, every four pulses, take 5 μl of lysates, and add 5 μl TE and 1 μl of proteinase K (1 μg/μl). Incubate for 45 min at 50
C. In the meanwhile, dissolve 100 g of agarose in 100 ml of TAE buffer to obtain a 1% agarose gel. Add the DNA loading buffer to your samples and load on a 1% agarose gel to check for the size of the fragments (remember to load also an appropriate DNA ladder marker). Usually, DNA fragments for a ChOP experiment should be between 500 and 1000 bp. 6. It is important, in this step, to avoid foam formation and to maintain the sample on ice, in order to preserve protein integrity. 7. It is possible to replace Protein A Sepharose beads and antibiotin antibody with Streptavidin Magnetic Beads. In this case, after blocking the beads, you can directly incubate them with the chromatin extracts plus biotin-labeled oligonucleotides. 8. The design of complementary DNA oligonucleotides is a key step of RNA precipitation. There are some online free tools to help you design oligonucleotides (e.g., www.singlemolecule. com). Several aspects, which can influence the stability and

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specificity of the interaction between the oligonucleotides and your LncRNA, have to be considered, such as length and GC content. A length of 30–40 bp with a GC% of around 45% can be considered optimal. Another key point is the target region. It is important to take into consideration if there is any information on the secondary structure of the LncRNA or on specific regions interacting with DNA or proteins. CHART protocols [6] use RNase H digestion to identify accessible regions of the LncRNA of interest [6, 17], but this assay does not indicate the portion of the LncRNA that mediates chromatin association, and in addition, this process can be expensive and time-consuming. If nothing is known about secondary structure or regions of interaction, it is reasonable to design the oligonucleotides complementary to the 50 or 30 end of the LncRNA. For example, due to its exceptional size (around 90 kbp), there was no information about the three-dimensional structure of Kcnq1ot1, nor it was conceivable to design a set of oligonucleotides tiling the entire length of the RNA. However, the use of a single oligonucleotide mapping at 50 [10, 11] gave us fairly good and specific results. Omit regions of repeats or extensive homology. All antisense oligonucleotides should be tested using BLAT searches to ensure that they uniquely align to the RNA target. If multiple hits along the genome are found, an oligonucleotide should not be used. It is recommended to design more than one biotinylated oligonucleotide. Indeed, using only one oligonucleotide increases the risk that the capture sites on the RNA could be not accessible or will fragment away from the RNA– chromatin site of interaction after sonication, resulting in poor yield of associated targets. Moreover, the comparison of ChOP experiments performed with different oligonucleotides targeting different regions of the LncRNA of interest will increase the robustness of the results. If necessary, two or more biotinylated oligonucleotides can be pooled together and used in a single ChOP experiments to increase the chance to precipitate the LncRNA of interest. Order the oligonucleotides biotinylated at the 50 . As a negative control, the scramble non-targeting oligonucleotide should have similar features (length and GC content) to the ones of targeting oligonucleotide. An oligonucleotide complementary to the target oligonucleotide will have same length and GC content and so can be considered an optimal negative control. 9. These steps are necessary to eliminate chromatin fragments aspecifically bound to the Protein A Sepharose or to the antibiotin antibody. 10. A good control of the ChOP efficiency is to verify the enrichment of the target LncRNA in the precipitated samples. To this

Chromain Oligo Affinity Precipitation

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end, instead of eluting samples with RNase A and RNase H, after the wash step, the RNA fraction can be extracted with a commercial kit (i.e., Trizol reagent), reverse transcribed, and qPCR amplified. Moreover, other additional controls are helpful to check for false-positive results. For example, to exclude that the enrichment of DNA fragments is a consequence of direct hybridization of the oligonucleotides with DNA, and not with the chromatin-bound RNA, we recommend to perform the hybridization reaction also in samples treated with RNase, which should not produce any signal. An even better control can be to do the ChOP assay in cells in which the LncRNA of interest is not expressed or is knocked down. 11. This step could be performed also overnight at 56
C and is necessary to revert the formaldehyde cross-link and to eliminate the proteins from samples. 12. Equal amounts of DNA (from 2 to 5 ng) of each sample, including input and scramble–biotin–oligonucleotide, should be used to amplify regions of interest as well as at least one negative non-target region. 13. qPCR analysis should be empirically optimized for the amount of DNA to be amplified. Sometimes, amplification of lower amounts of DNA gives better results. References 1. Chen LL (2016) Linking long noncoding RNA localization and function. Trends Biochem Sci 41(9):761–772. https://doi.org/ 10.1016/j.tibs.2016.07.003 2. Rinn JL, Chang HY (2012) Genome regulation by long noncoding RNAs. Annu Rev Biochem 81:145–166. https://doi.org/10.1146/ annurev-biochem-051410-092902 3. Vance KW, Ponting CP (2014) Transcriptional regulatory functions of nuclear long noncoding RNAs. Trends Genet 30(8):348–355. https://doi.org/10.1016/j.tig.2014.06.001 4. Mariner PD, Walters RD, Espinoza CA, Drullinger LF, Wagner SD, Kugel JF, Goodrich JA (2008) Human Alu RNA is a modular transacting repressor of mRNA transcription during heat shock. Mol Cell 29(4):499–509. https:// doi.org/10.1016/j.molcel.2007.12.013 5. Chu C, Qu K, Zhong FL, Artandi SE, Chang HY (2011) Genomic maps of long noncoding RNA occupancy reveal principles of RNA-chromatin interactions. Mol Cell 44 (4):667–678. https://doi.org/10.1016/j. molcel.2011.08.027 6. Simon MD, Wang CI, Kharchenko PV, West JA, Chapman BA, Alekseyenko AA, Borowsky

ML, Kuroda MI, Kingston RE (2011) The genomic binding sites of a noncoding RNA. Proc Natl Acad Sci U S A 108 (51):20497–20502. https://doi.org/10. 1073/pnas.1113536108 7. Engreitz JM, Pandya-Jones A, McDonel P, Shishkin A, Sirokman K, Surka C, Kadri S, Xing J, Goren A, Lander ES, Plath K, Guttman M (2013) The Xist lncRNA exploits threedimensional genome architecture to spread across the X chromosome. Science 341 (6147):1237973. https://doi.org/10.1126/ science.1237973 8. Chu C, Spitale RC, Chang HY (2015) Technologies to probe functions and mechanisms of long noncoding RNAs. Nat Struct Mol Biol 22 (1):29–35. https://doi.org/10.1038/nsmb. 2921 9. Machyna M, Simon MD (2018) Catching RNAs on chromatin using hybridization capture methods. Brief Funct Genomics 17 (2):96–103. https://doi.org/10.1093/bfgp/ elx038 10. Andresini O, Rossi MN, Matteini F, Petrai S, Santini T, Maione R (2019) The long non-coding RNA Kcnq1ot1 controls maternal

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p57 expression in muscle cells by promoting H3K27me3 accumulation to an intragenic MyoD-binding region. Epigenetics Chromatin 12(1):8. https://doi.org/10.1186/s13072019-0253-1 11. Pandey RR, Mondal T, Mohammad F, Enroth S, Redrup L, Komorowski J, Nagano T, Mancini-Dinardo D, Kanduri C (2008) Kcnq1ot1 antisense noncoding RNA mediates lineage-specific transcriptional silencing through chromatin-level regulation. Mol Cell 32(2):232–246. https://doi.org/10. 1016/j.molcel.2008.08.022 12. Subhash S, Mishra K, Akhade VS, Kanduri M, Mondal T, Kanduri C (2018) H3K4me2 and WDR5 enriched chromatin interacting long non-coding RNAs maintain transcriptionally competent chromatin at divergent transcriptional units. Nucleic Acids Res 46 (18):9384–9400. https://doi.org/10.1093/ nar/gky635 13. Ali MM, Akhade VS, Kosalai ST, Subhash S, Statello L, Meryet-Figuiere M, Abrahamsson J, Mondal T, Kanduri C (2018) PAN-cancer analysis of S-phase enriched lncRNAs identifies oncogenic drivers and biomarkers. Nat Commun 9(1):883. https://doi.org/10.1038/ s41467-018-03265-1

14. Akhade VS, Arun G, Donakonda S, Rao MR (2014) Genome wide chromatin occupancy of mrhl RNA and its role in gene regulation in mouse spermatogonial cells. RNA Biol 11 (10):1262–1279. https://doi.org/10.1080/ 15476286.2014.996070 15. Mondal T, Subhash S, Vaid R, Enroth S, Uday S, Reinius B, Mitra S, Mohammed A, James AR, Hoberg E, Moustakas A, Gyllensten U, Jones SJ, Gustafsson CM, Sims AH, Westerlund F, Gorab E, Kanduri C (2015) MEG3 long noncoding RNA regulates the TGF-beta pathway genes through formation of RNA-DNA triplex structures. Nat Commun 6:7743. https://doi.org/10.1038/ ncomms8743 16. Kanduri C (2011) Kcnq1ot1: a chromatin regulatory RNA. Semin Cell Dev Biol 22 (4):343–350. https://doi.org/10.1016/j. semcdb.2011.02.020 17. Vance KW (2017) Mapping long noncoding RNA chromatin occupancy using capture hybridization analysis of RNA targets (CHART). Methods Mol Biol 1468:39–50. https://doi.org/10.1007/978-1-4939-40356_5

Chapter 3 Knockdown of Nuclear lncRNAs by Locked Nucleic Acid (LNA) Gapmers in Nephron Progenitor Cells Masaki Nishikawa and Norimoto Yanagawa Abstract Despite recent advance in our understanding on the role of long noncoding RNAs (lncRNAs), the function of the vast majority of lncRNAs remains poorly understood. To characterize the function of lncRNAs, knockdown studies are essential. However, the conventional silencing methods for mRNA, such as RNA interference (RNAi), may not be as efficient against lncRNAs, partly due to the mismatch of the localization of lncRNAs and RNAi machinery. To circumvent such limitation, a new technique has recently been developed, i.e., locked nucleic acid (LNA) gapmers. This system utilizes RNase H that distributes evenly in both nucleus and cytoplasm and is expected to knock down lncRNAs of interest more consistently regardless of their localization in the cell. In this chapter, we describe the procedure with tips to silence lncRNAs by LNA gapmers, by using mouse nephron progenitor cells as an example. Key words ncRNA, lncRNA, Gapmer, Knockdown, Gymnosis, Nephron progenitor cells

1

Introduction With recent advance in sequencing technologies, we came to realize that much of the genome is transcribed into RNAs that are not translated into proteins, the so-called noncoding RNAs (ncRNAs). According to their functions, structures, and localizations, these ncRNAs are categorized into various classes, including housekeeping RNAs (tRNA, rRNA, etc.), microRNAs, Piwiinteracting RNAs (piRNAs), and long ncRNAs (lncRNAs) [1– 3]. The lncRNAs are defined as ncRNAs consisting of more than 200 nucleotides, such as Xist and Malat1, and they are found mostly within cell nucleus [4, 5]. Although an increasing number of lncRNAs have recently been found to exert functions that participate in various cellular and developmental processes, the majority of lncRNAs remain devoid of functional annotation [6, 7]. The involvement of lncRNAs in developmental processes represents an important aspect in the functionality of lncRNAs [8, 9]. For example, Xist is known to play a central role in the

Ulf Andersson Vang Ørom (ed.), RNA-Chromatin Interactions: Methods and Protocols, Methods in Molecular Biology, vol. 2161, https://doi.org/10.1007/978-1-0716-0680-3_3, © Springer Science+Business Media, LLC, part of Springer Nature 2020

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process of X chromosome inactivation during early developmental stage [10]. Many unannotated ncRNAs have also been found to be expressed in a tissue-specific fashion under epigenetic regulation by chromatin states, raising the possibility that these ncRNAs could have developmental functions like other regulatory genes [11]. However, it is often difficult to assess the functionality of lncRNAs in specific cells during development, partly due to the limited availability and stability of cells obtained from embryos. While loss-of-function studies are essential in characterizing the functions of lncRNAs, the conventional silencing methods for mRNA, such as RNA interference (RNAi), are found to be less efficient for lncRNAs. This is mainly due to the mismatch of the localization of lncRNAs and RNAi machinery, i.e., many lncRNAs locate in the nucleus, whereas the RISC complex, the RNAi machinery, is localized in the cytoplasm [12–15]. Another emerging method for RNA silencing is by using antisense oligonucleotides. This method utilizes RNase H, where the target RNA strand in heteroduplex with the designed antisense oligonucleotides can be degraded by RNase H. Since RNase H is distributed evenly in both nucleus and cytoplasm, a more consistent knockdown can be expected regardless of the localization of the target lncRNAs [16–18]. Among these antisense oligonucleotides, chemically modified chimeric DNA, such as locked nucleic acid (LNA) gapmers, which contains DNA nucleotides flanked by LNA, has been shown to be advantageous with high affinity and stability [19– 24]. Here we describe a method to silence lncRNAs by LNA gapmers, using nephron progenitor cells obtained from mouse embryonic kidneys as an example.

2

Materials The embryonic kidney develops from mainly three lineages of progenitor cells through their interactive regulatory mechanisms for differentiation, maturation, and proliferation [25, 26]. In our present discussion, we will focus on one of the three lineages, i.e., the nephron progenitor cells (NPCs), from which most of the nephron tubule structures derive. There have been multiple reports on the in vitro culture systems for NPCs [27–30], and we adopted some of these culture systems in our studies [27, 29].

2.1 Preparation of Cells from Mouse Embryonic Kidneys

1. Embryonic kidneys: Six2GCE/+ ((129/Sv x C57BL/6J)F1, Jax stock #009600)) mice can be obtained from Jackson Laboratory and maintained with C57BL/6J background to sort NPCs. Mouse embryonic kidneys should be obtained from E12.5 embryos for heterogenic aggregate culture and from E13.5 embryos for pure NPC culture.

Knockdown of Nuclear lncRNAs by Locked Nucleic Acid (LNA) Gapmers in. . .

31

2. For pure NPC culture, use NPEM medium, as previously described by Brown et al. [27]: APEL medium (1
) containing 1
Penstrep, 200 ng/ml FGF9, 10 μM Y27632, 20 ng/ml IGF1, 2 ng/ml IGF2, 30 ng/ml BMP4, 125 nM LDN-193189, 1.25 μM CHIR9902, 1 μg/ml heparin. 3. For heterogenic aggregate culture, use keratinocyte serum-free medium (KSFM; Invitrogen) containing 10% FBS, 10 μM Y27632, a ROCK inhibitor, and 110 μM 2-mercaptoethanol. 4. 70 μm cell strainer. 5. Ultralow attachment round bottom dish. 6. Matrigel. 7. Cell sorter (FACS). 2.2 LNA Gapmers and Lipofection

1. LNA gapmers: Screening grade negative control LNA gapmer with scrambled sequence and positive control LNA gapmer with complementary sequence to Malat1 (see Note 1). 2. Transfection reagent. 3. Reduced Serum Media. 4. Nuclease-free TE buffer: 10 mM Tris, pH 7.5, 0.1 mM EDTA.

2.3 Quantitative PCR Analysis

1. Trizol reagent. 2. Direct-zol RNA Purification Kits. 3. cDNA Synthesis Kit. 4. SYBR Green master mix. 5. Real-Time PCR Detection System. 6. Primers: Gapdh F: TGAACGGATTTGGCCGTATTG. Gapdh R: ACCATGTAGTTGAGGTCAATGAAG. Malat1 F: GAGCTCGCCAGGTTTACAGT. Malat1 R: AACTACCAGCAATTCCGCCA.

3

Methods The following protocols are for 96-well plate format. For other plates, adjust volumes accordingly.

3.1 Preparation and Storage of Gapmer Solutions

1. Briefly centrifuge the vial containing gapmer at low speed (maximum 4000
g). 2. Add nuclease-free TE buffer for reconstitution to obtain gapmer solution at 50 μM concentration. 3. Let the vial stand for a few minutes at room temperature. 4. Gently tap the vial to mix the solution.

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5. Repeat steps 3 and 4. 6. Centrifuge briefly. 7. Make aliquots and store in a constant-temperature freezer at 20  C or below (see Note 2). 3.2 Preparation of Cell Suspension from Mouse Embryonic Kidneys

1. Rinse the dissected embryonic kidneys in ice-cold PBS solution and soak them in 1 ml ice-cold TrypLE dissociation solution (Life Technologies) (see Note 3). 2. Leave at room temperature for 8 min, followed by pipetting rigorously with 1000p tip for 1–2 min. Do not overdigest. 3. Add ice-cold culture medium with 20% FBS and filter the cell suspension with 70 μm cell strainer. 4. Centrifuge at 650
g for 5 min.

3.3 Culture of Pure NPCs and Delivery of Gapmers

1. Dilute Matrigel 1:25 in cold DMEM/F12 and coat wells (50 μl/well for 96-well). 2. After Subheading 3.2, step 4, resuspend cells with NPEM medium and sort Six2-GFP+ NPCs by FACS. 3. Seed NPCs at 1
104 cells/cm2 and passage before becoming confluent. Use NPCs at passages 1–2 (see Note 4). 4. For knockdown experiments, seed NPCs at 1
105 cells/cm2, and incubate in a cell incubator at 37  C for 24 h. 5. Gapmers can be delivered to cells either directly without carriers or reagents, termed gymnosis, or assisted with lipid reagent, such as Lipofectamine (Subheading 2.2, item 2), termed lipofection [24, 31]. 6. For delivery by gymnosis, add gapmer directly to the fresh cell culture medium at 1 μM final concentration (see Note 5). 7. For delivery by lipid-assisted method in one reaction, prepare two 1.5 ml tubes. In Tube 1, gently mix 5 μl of Opti-MEM and 0.3 μl transfection reagent per well. In Tube 2, mix 5 μl of serum-free media and 0.2 μl of gapmer stock solution (50 μM) per well. Mix two tubes and keep at room temperature for 5 min. Add 100 μl of fresh culture medium into this mixture. Replace cell culture medium with the solution thus prepared immediately. The final gapmer concentration is 100 nM (see Notes 6 and 7). 8. Incubate cells from steps 6 and 7 in a cell incubator at 37  C for 24 h. 9. Rinse cells with PBS once and isolate RNA to conduct real-time qPCR to evaluate the knockdown experiment (Fig. 1).

Knockdown of Nuclear lncRNAs by Locked Nucleic Acid (LNA) Gapmers in. . .

33

Fig. 1 Knockdown of Malat1 in primary cultured pure mouse NPCs. The in vitrocultured pure NPCs were exposed to either the negative control gapmer with scrambled sequence (Neg) or the targeting gapmer against Malat1 (Malat1) for 24 h. Gapmers were delivered either without (gymnosis) or with transfection reagent (lipofection) at 100 nM or 1 μM, respectively. The expression level of Malat1 was significantly knocked down by the targeting gapmer (Malat1) when delivered by gymnosis, while the negative gapmer with scrambled sequence (Neg) was without effect. For NPCs, lipofection was not as effective as gymnosis

Fig. 2 Knockdown of Malat1 in heterogenic aggregates of mouse embryonic kidney cells. Heterogenic cell aggregates obtained from mouse embryonic kidneys were exposed to either negative control gapmer (Neg) or targeting gapmer against Malat1 (Malat1) at 100 nM for 24 h. Gapmers were delivered by lipofection. While negative control gapmer was without effect, the expression level of Malat1 was significantly knocked down by the targeting gapmer (Malat1). Thus, in contrast to primary cultured pure NPCs as shown in Fig. 1, lipofection was effective in heterogenic cell aggregates, demonstrating cell typespecific preference toward different delivery methods 3.4 Culture of Heterogenic Cell Aggregates and Delivery of Gapmers

1. After Subheading 3.2, step 4, resuspend cells with the medium described above (Subheading 2.1, item 3) at 2
105 cells/ml. 2. For delivery by lipid-assisted method, prepare two 1.5 ml tubes for each reaction. In Tube 1, gently mix 5 μl of serum-free media and 0.3 μl transfection reagent per well. In Tube 2, mix 5 μl of Opti-MEM and 0.1–0.2 μl of gapmer stock solution (50 μM) per well. Mix two tubes and keep at room temperature for 5 min. Add 100 μl of cell suspension from step 1 and mix well.

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3. Transfer all solution from step 2 into a well of ultralow attachment round bottom dish (2
104 cells with 50 nM or 100 nM gapmer per well). 4. Centrifuge at 650
g for 2 min and culture in a cell incubator at 37  C for 24 h. 5. Pick the cell aggregate and isolate RNA to conduct real-time qPCR to evaluate the knockdown experiment (Fig. 2) (see Note 8).

4

Notes 1. To design gapmers against target genes, it is important to choose different regions of the transcript and determine the efficiency experimentally. We noticed that the sequence of primary transcripts without splicing may work better for designing effective gapmers. 2. Do not use frost-free freezers with automatic thaw-freeze cycle. Avoid freeze-thaw cycles after the initial storage. Working solutions can be stored at 4  C for up to 2 weeks. 3. Twenty of E12.5–E13.5 embryonic kidneys can be digested in 1 ml TrypLE solution. Digestion should start with ice-cold TrypLE and raise the temperature gradually up to room temperature to achieve complete digestion throughout the tissues. 4. NPCs can be successfully passaged and maintained for up to three passages. Thereafter, the proliferation rate reduced. We noticed that the freshly seeded NPCs after sorting show more variation in cell morphology, as compared to the more homogenous populations on passages 1–2. 5. The effective gapmer concentrations for gymnosis should be determined experimentally for each gapmer and cell type. We found the typical range of gapmer concentration for gymnosis was from 100 nM to 5 μM. 6. The effective gapmer concentrations for lipid-assisted delivery should also be determined experimentally for each gapmer and cell type. We found the typical range of gapmer concentration for lipofection was from 0.1 nM to 100 nM. 7. Although much higher concentration of gapmers is required for gymnosis (typically 100 times more compared to lipidassisted delivery), we observed less toxicity to the cells by gymnosis than lipid-assisted delivery methods. The efficiency between gymnosis and lipofection may vary depending on the type of cells, as demonstrated by Figs. 1 and 2.

Knockdown of Nuclear lncRNAs by Locked Nucleic Acid (LNA) Gapmers in. . .

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8. The aggregates can be transferred onto a membrane filter (0.4 μm isopore polycarbonate filter; Millipore) and cultured in liquid–air interface. References 1. Jarroux J, Morillon A, Pinskaya M (2017) History, discovery, and classification of lncRNAs. Adv Exp Med Biol 1008:1–46. https://doi. org/10.1007/978-981-10-5203-3_1 2. Hombach S, Kretz M (2016) Non-coding RNAs: classification, biology and functioning. Adv Exp Med Biol 937:3–17. https://doi.org/ 10.1007/978-3-319-42059-2_1 3. Brosius J, Raabe CA (2016) What is an RNA? A top layer for RNA classification. RNA Biol 13:140–144. https://doi.org/10.1080/ 15476286.2015.1128064 4. Hutchinson JN, Ensminger AW, Clemson CM, Lynch CR, Lawrence JB, Chess A (2007) A screen for nuclear transcripts identifies two linked noncoding RNAs associated with SC35 splicing domains. BMC Genomics 8:39. https://doi.org/10.1186/1471-2164-8-39 5. Brown CJ, Hendrich BD, Rupert JL, Lafrenie`re RG, Xing Y, Lawrence J, Willard HF (1992) The human XIST gene: analysis of a 17 kb inactive X-specific RNA that contains conserved repeats and is highly localized within the nucleus. Cell 71:527–542. https://doi. org/10.1016/0092-8674(92)90520-m 6. Ivey KN, Srivastava D (2015) microRNAs as developmental regulators. Cold spring Harb Perspect biol 7:a008144. https://doi.org/10. 1101/cshperspect.a008144 7. Quinodoz S, Guttman M (2014) Long noncoding RNAs: an emerging link between gene regulation and nuclear organization. Trends Cell Biol 24:651–663. https://doi.org/10. 1016/j.tcb.2014.08.009 8. Sarropoulos I, Marin R, Cardoso-Moreira M, Kaessmann H (2019) Developmental dynamics of lncRNAs across mammalian organs and species. Nature 571:510–514. https://doi.org/ 10.1038/s41586-019-1341-x 9. Dela´s MJ, Hannon GJ (2017) lncRNAs in development and disease: from functions to mechanisms. Open Biol 7. https://doi.org/ 10.1098/rsob.170121 10. Duret L, Chureau C, Samain S, Weissenbach J, Avner P (2006) The Xist RNA gene evolved in eutherians by pseudogenization of a proteincoding gene. Science 312:1653–1655. https://doi.org/10.1126/science.1126316 11. Sachs M, Onodera C, Blaschke K, Ebata KT, Song JS, Ramalho-Santos M (2013) Bivalent

chromatin marks developmental regulatory genes in the mouse embryonic germline in vivo. Cell Rep 3:1777–1784. https://doi. org/10.1016/j.celrep.2013.04.032 12. Gagnon KT, Li L, Chu Y, Janowski BA, Corey DR (2014) RNAi factors are present and active in human cell nuclei. Cell Rep 6:211–221. https://doi.org/10.1016/j.celrep.2013.12. 013 13. Zeng Y, Cullen BR (2002) RNA interference in human cells is restricted to the cytoplasm. RNA 8:855–860. https://doi.org/10.1017/ s1355838202020071 14. Kawasaki H, Taira K (2003) Short hairpin type of dsRNAs that are controlled by tRNA(Val) promoter significantly induce RNAi-mediated gene silencing in the cytoplasm of human cells. Nucleic Acids Res 31:700–707. https://doi. org/10.1093/nar/gkg158 15. Chiu Y-L, Ali A, Chu C-Y, Cao H, Rana TM (2004) Visualizing a correlation between siRNA localization, cellular uptake, and RNAi in living cells. Chem Biol 11:1165–1175. https://doi.org/10.1016/j.chembiol.2004. 06.006 16. Crooke ST (1999) Molecular mechanisms of action of antisense drugs. Biochim Biophys Acta 1489:31–44. https://doi.org/10.1016/ s0167-4781(99)00148-7 17. Ideue T, Hino K, Kitao S, Yokoi T, Hirose T (2009) Efficient oligonucleotide-mediated degradation of nuclear noncoding RNAs in mammalian cultured cells. RNA 15:1578–1587. https://doi.org/10.1261/ rna.1657609 18. Vickers TA, Koo S, Bennett CF, Crooke ST, Dean NM, Baker BF (2003) Efficient reduction of target RNAs by small interfering RNA and RNase H-dependent antisense agents. A comparative analysis. J Biol Chem 278:7108–7118. https://doi.org/10.1074/ jbc.M210326200 19. Pendergraff HM, Krishnamurthy PM, Debacker AJ, Moazami MP, Sharma VK, Niitsoo L, Yu Y, Tan YN, Haitchi HM, Watts JK (2017) Locked nucleic acid Gapmers and conjugates potently silence ADAM33 , an asthma-associated Metalloprotease with nuclear-localized mRNA. Mol Ther Nucleic

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25. Rowan CJ, Sheybani-Deloui S, Rosenblum ND (2017) Origin and function of the renal Stroma in health and disease. Results Probl Cell Differ 60:205–229. https://doi.org/10. 1007/978-3-319-51436-9_8 26. Little MH, McMahon AP (2012) Mammalian kidney development: principles, Progress, and projections. Cold Spring Harb Perspect Biol 4: a008300. https://doi.org/10.1101/ cshperspect.a008300 27. Brown AC, Muthukrishnan SD, Oxburgh L (2015) A synthetic niche for nephron progenitor cells. Dev Cell 34:229–241. https://doi. org/10.1016/j.devcel.2015.06.021 28. Tanigawa S, Taguchi A, Sharma N, Perantoni AO, Nishinakamura R (2016) Selective in vitro propagation of nephron progenitors derived from embryos and pluripotent stem cells. Cell Rep 15:801–813. https://doi.org/10.1016/j. celrep.2016.03.076 29. Yuri S, Nishikawa M, Yanagawa N, Jo OD, Yanagawa N (2015) Maintenance of mouse nephron progenitor cells in aggregates with gamma-Secretase inhibitor. PLoS One 10: e0129242. https://doi.org/10.1371/journal. pone.0129242 30. Li Z, Araoka T, Wu J, Liao H-K, Li M, Lazo M, Zhou B, Sui Y, Wu M-Z, Tamura I, Xia Y, Beyret E, Matsusaka T, Pastan I, Rodriguez Esteban C, Guillen I, Guillen P, Campistol JM, Izpisua Belmonte JC (2016) 3D culture supports long-term expansion of mouse and human nephrogenic progenitors. Cell Stem Cell 19:516–529. https://doi.org/10.1016/ j.stem.2016.07.016 31. Stein CA, Hansen JB, Lai J, Wu S, Voskresenskiy A, Høg A, Worm J, Hedtj€arn M, Souleimanian N, Miller P, Soifer HS, Castanotto D, Benimetskaya L, Ørum H, Koch T (2010) Efficient gene silencing by delivery of locked nucleic acid antisense oligonucleotides, unassisted by transfection reagents. Nucleic Acids Res 38:e3–e3. https://doi.org/10.1093/nar/gkp841

Chapter 4 Design and Application of a Short (16-mer) Locked Nucleic Acid Splice-Switching Oligonucleotide for Dystrophin Production in Duchenne Muscular Dystrophy Myotubes Ce´lia Carvalho and Maria Carmo-Fonseca Abstract Splice-switching oligonucleotides (SSOs) have been used to modulate gene expression by interfering with pre-mRNA splicing with the intent to treat disease. For Duchenne muscular dystrophy, splicing modulation has been used to induce the skipping of exon 51 of the dystrophin transcript, allowing the production of a truncated but functional protein. Although oligonucleotide-based therapies are promising, the rapid degradation of oligonucleotides (ONs) by intracellular nucleases has been a major obstacle. Locked nucleic acid (LNA) substitution in SSOs protects oligonucleotides from nuclease degradation and enhances the hybridization properties of the oligo. However, the best optimum size of the oligo depends on the LNA substitution rate. Here we show that 16-mer DNA SSOs with 60% LNA substitution and full phosphorothioate (PS) linkage backbone efficiently induce exon 51 skipping in myogenic cells derived from a DMD patient, allowing expression of the dystrophin protein. Key words Splice-switching oligonucleotides, Locked nucleic acid, Duchenne muscular dystrophy, Splice modulation, RNA-targeted therapy

1

Introduction Splice modulation is a strategy recently developed to modulate gene expression by interfering with splicing with the intent to treat disease, an RNA-targeted therapy. The idea is to block either the RNA-RNA base pairing or the RNA-protein interactions normally required for splicing to occur. Short synthetic, antisense, modified nucleic acid sequences known as splice-switching oligonucleotides (SSOs) have been used to disrupt the splicing process and by that manipulate protein production. By using this strategy, a SSO to treat Duchenne muscular dystrophy (DMD), eteplirsen, had accelerated approval from FDA in 2016, and nusinersen was approved to treat spinal muscular atrophy (DMA), also in 2016 [1].

Ulf Andersson Vang Ørom (ed.), RNA-Chromatin Interactions: Methods and Protocols, Methods in Molecular Biology, vol. 2161, https://doi.org/10.1007/978-1-0716-0680-3_4, © Springer Science+Business Media, LLC, part of Springer Nature 2020

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Duchenne muscular dystrophy patients carry frame-shifting deletions or nonsense mutations in the dystrophin gene that result in a loss of functional protein production. The largest fraction of patients (approximately 13%) would benefit from exon 51 forced skipping therapy, allowing a restoration of the reading frame of coding sequence for dystrophin protein and the production of an internally truncated, but partially functional, protein. Although oligonucleotide-based therapies seem very promising, there are obstacles that must be overcome, including the rapid degradation of oligonucleotides (ONs) by intracellular nucleases. Different chemical modifications of nucleic acids have been developed and tested including morpholino (the chemistry of the approved drug eteplirsen) and 20 OMe RNA [2]. Locked nucleic acid (LNA)-based SSOs have been tested in cellular and animal models with encouraging results [3, 4]. LNA substitution protects oligonucleotides from nuclease degradation and enhances the hybridization properties of the synthetic LNA-containing oligonucleotides [4–7]. Phosphorothioate (PS) linkages are fully compatible with LNA modification and further increase LNA ON’s protection from nucleases [8, 9]. While it is known that LNA-modified ONs can be shorter than ONs modified by other chemistries, due to its enhanced duplex stabilization with the target RNA molecule, its optimum size is still a variable under investigation [10] that depends on the substitution rate but which needs to be determined on a case-bycase basis. We thus tested several options and found that a 16-mer DNA SSO with 60% LNA substitution and full phosphorothioate (PS) linkages carries the maximum efficiency to induce exon 51 skipping, allowing expression of the dystrophin protein [11]. We here show how skipping of exon 51 can be achieved by transfection of a short (16-mer) LNA-modified antisense oligonucleotide to modulate the splicing of dystrophin pre-mRNA in myogenic cells derived from a DMD patient carrying a deletion of DMD gene-exons 48–50 that benefits with the forced skipping of exon 51. By using this strategy, we observed, by RT-PCR, that the splicing pattern is effectively modulated and, by fluorescence microscopy, that the protein dystrophin is produced in differentiating cells and is localized at its expected location, which is in close proximity to the myotube membrane.

2

Materials Prepare all solutions using sterile deionized water: water with a resistivity of 18 MΩ cm at 25  C and filtered through a 0.22 μm filter. All reagents must be analytical grade unless indicated otherwise. Prepare and store all solutions at room temperature unless indicated otherwise. Perform all steps involving cell culture using sterile technique in a suitable hood.

Short 16-mer LNA-SSO Restore Dystrophin in DMD Myotubes

2.1 Cell Culture and Differentiation

39

1. The human myoblast cell lines have been previously described [12]: the Control cell line is derived from a healthy individual, and the DMD cell line is derived from a male patient with Duchenne muscular dystrophy, carrying a deletion in the dystrophin gene (Δ exons 48 to 50) (see Note 1). 2. Culture medium: supplemented (5% serum) Skeletal Muscle Cell Growth Medium. 3. Differentiation medium: add 2.5 mL of ITS (1 g/L Insulin; 0.55 g/L Transferrin; 0.5 mg/L sodium Selenite) to 250 mL of DMEM culture medium with high glucose (4.5 g/L) and sodium pyruvate (0.11 g/L) culture medium (no serum added) (see Note 2). 4. 12-well, flat bottom, tissue culture plates. 5. Tissue culture humidified incubator at 37  C with 5% CO2.

2.2

SSO Transfection

Please see Notes 3–5. 1. The LNA-SSOs described here (see Table 1) can be purchased from Exiqon (now in QIAGEN) using the GeneGlobe catalogue numbers provided (see Note 6). Dilute SSOs with sterile deionized water to a final concentration of 100 μM. Distribute in 5–10 μL aliquots and store at 20  C. 2. Oligonucleotide medium.

2.3 RNA Purification and Retrotranscription

transfection

reagent

and

transfection

1. RNA isolation reagent (e.g., PureZOL). 2. DNase I. 3. RiboSafe. 4. RNA precipitation buffer: 3 M Na acetate pH 5.2; glycogen blue (15 mg/mL). 5. Phenol:cloroform:isoamyl alcohol (25:24:1, v/v). 6. Absolute ethanol. 7. 75% ethanol diluted with sterile deionized water.

Table 1 List of antisense SSO sequences Identification Sequence

Chemistry

GeneGlobe catalogue number

51.1c

AGGAAGATGGCATTTC

LNA-DNA; PS

# YCO0116696

51.1d

GAAGATGGCATTTCTA

LNA-DNA; PS

# YCO0143765

51.3a

GTAAGTTCTGTCCAAG

LNA-DNA; PS

# YCO0149335

AO51

UCAAGGAAGAUGGCAUUUCU

20 OMe RNA

AO stands for antisense oligonucleotide

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8. High Fidelity cDNA Synthesis Kit. 9. Reverse primer for retrotranscription (here, complementary to exon 54: DMD_exon_54R (50 -GGAGAAGTTTCAGGGCC AAG-30 ). 2.4 Splice Pattern-Specific RT-PCR

1. Taq polymerase (Fast HotStart ReadyMix PCR Kit). 2. Splice pattern-specific primer pair (here, forward primer on exon 47: DMD_exon_47F (50 -ACCCGTGCTTGTAAGTG CTC-30 ); reverse primer complementary to exon 53: DMD_exon_53R (50 -TGACTCAAGCTTGGCTCTGG-30 ). 3. Thermocycler. 4. Tris Acetate-EDTA buffer (TAE) 50x: 242 g of Tris base dissolved in 650 mL of deionized water; add 57.1 mL of glacial acetic acid and 100 mL of 0.5 M EDTA solution, pH 8.0; complete to 1 L with deionized water. For working solution (1), dilute 20 mL up to 1 L with deionized water. 5. Agarose for gel preparation. 6. DNA stain. 7. DNA molecular marker. 8. Electrophoresis apparatus. 9. Gel imaging apparatus.

2.5 Immunofluorescence Microscopy

1. Sterilized glass coverslips: wash coverslips (18 mm diameter; VWR, ref. 631-0669) in ethanol and, using a curved tip forceps, spread the coverslips on a glass petri dish between layers of Whatman filter paper (cut to fit). Wrap the glass petri dish with aluminum foil and sterilize by autoclaving. 2. 0.1% (w/v) gelatin sterilized by autoclaving. 3. 7.4% (w/v) paraformaldehyde fixative (PFA, 2 concentrated): dissolve 7.4 g of paraformaldehyde in 100 mL of deionized water, heat at 70  C in a water bath for 4 h with gentle agitation and add 0.1 mL of 1 N NaOH to help dissolve, and then incubate for additional 1 h at 70  C. The final pH should not exceed 8. After cooling down, filter with a Whatman filter paper and store at 20  C in 5 mL aliquots. 4. 10% (w/v) Triton X-100 detergent. 5. 10 phosphate-buffered saline (PBS). 6. 10% BSA filter sterilized with a low protein binding syringe filter. 7. 1 mg/mL DAPI. 8. 3.7% paraformaldehyde in PBS. 9. Permeabilization solution: 0.5% Triton X-100 in PBS. 10. PBST: 0.05% Tween20 in PBS.

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11. Blocking solution: 1% BSA in PBST. 12. Antibodies: polyclonal rabbit anti-dystrophin antibody (Abcam, ab85302; dilution 1:100); Rhodamine (TRITC)conjugated AffiniPure Donkey anti-rabbit IgG (Jackson ImmunoResearch, ref. 711-025-152; dilution 1:200). Dilute antibodies in blocking solution and keep fluorophoreconjugated secondary antibody solution protected from light by wrapping it in aluminum foil. Keep at 4  C. Centrifuge for 5 min at 14,000 rpm before use. 13. DAPI DNA staining solution (1 μg/mL DAPI in PBS): dilute 1 μL of 1 mg/mL DAPI in 1 mL of PBS. Keep at 4  C protected from light. Use within 24 h. 14. Glass slides: NORMAX ground edges, frosted ends 20 mm, 1 mm thickness glass slides. 15. Mounting medium. 16. Microscope. For fluorescence microscopy, we used Zeiss LSM 710 Confocal Point-Scanning Microscope (ZEISS), lasers Diode 405-30 (405 nm), and DPSS 561-10 (561 nm); objective 63 magnification, Plan Apochromat, NA 1.40. 17. Image analysis software.

3

Methods

3.1 Cell Culture and Differentiation

1. Culture myoblast cell lines in Skeletal Muscle Cell Growth Medium, in a humidified, 37  C, 5% CO2 incubator. When cells reach a maximum of 60–70% confluence, split the culture in four, to avoid myoblast differentiation. Split culture every 3–5 days. 2. For differentiation assays, plate cells at high density, and 24 h after, when reaching 80% confluence, remove the medium, rinse once with differentiation medium, then add differentiation medium and incubate for the period of time needed for the assay. 3. If long incubation periods are needed, carefully replace half of the differentiation medium after 6 days and thereafter at every 2 or 3 days.

3.2

SSO Transfection

Please see Note 7 before starting. Use 12-well plates for transfection experiments. 1. Prepare a master dilution mix according to the number of wells you will use. Add 2 μL of transfection reagent to 28 μL of OptiMEM Medium per well. Mix well by pipetting up and down several times.

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Fig. 1 RT-PCR quantitative analysis of DMD exon 51 skipping reveals that SSO 51.1c induces exon 51 skipping with the highest efficiency and consistency. (a) Schematic representation of the analyzed stretch of DMD mRNA with exons represented as numbered boxes. At the top, the oligo primer used for retrotranscription is represented by a single arrow, and at the bottom the two arrows represent the primer pair used for RT-PCR analysis of exon 51 skipping; expected sizes of PCR products are indicated. (b) DMD myoblasts were transfected with the indicated SSOs at 50 nM and analyzed after 48 h of differentiation. Electrophoresis of PCR products in agarose gel shows unskipped and skipped transcripts. Note that all assays produced skipped and unskipped products in different proportions. (c) Univariate scatter dot of the percentage of exon skipping induced by each SSO. Values were obtained from quantification of the bands in the gel. The depicted mean and SD are from four independent experiments

2. Prepare a master SSO dilution mix by adding 0.33 μL of 100 μM SSO (kept on ice) to 30 μL of Opti-MEM Medium, per each well to assay. 3. Distribute the SSO dilution mix by dropping 30 μL in the bottom of each empty well in the cell culture plate and add 30 μL of transfection reagent master dilution to each drop. Mix by pipetting up and down several times. Incubate for 5–20 min. 4. Add 0.6 mL of trypsinized cell suspension (in Skeletal Muscle Cell Growth Medium, already in a density settled to reach 90% confluence after 24 h) to each well. Mix well by rocking the plate back and forth and repeat after rotating horizontally 90 . 5. Incubate the cells 24 h before switch to differentiation medium: remove all medium, rinse once with 0.5 mL of differentiation medium, add 1 mL of differentiation medium, and incubate for 48 h.

Short 16-mer LNA-SSO Restore Dystrophin in DMD Myotubes

3.3 RNA Purification and Retrotranscription

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Please see Note 8 before starting. 1. Discard all medium from cell culture plate and add 0.5 mL of PureZol RNA isolation reagent. Proceed with RNA purification according to manufacturer instructions and remove residual DNA with 30 min incubation with 0.5 U/μL DNase I in the presence of 1 U/μL RiboSafe, which will avoid RNA degradation. 2. Further purify RNA by acidic phenol extraction with equal volume phenol:cloroform: isoamyl alcohol (25:24:1, v/v) in the presence of 0.3 M sodium acetate pH 5.2 and 50 μg/mL GlycoBlue followed by ethanol precipitation by adding 2.5 the volume of absolute ethanol. Centrifuge at highest speed for 30 min, 4ºC. Wash twice with 75% ethanol. Dissolve RNA with sterile deionized water for 10 min at 60  C, and then keep it on ice. Keep purified RNA at 70  C. 3. For cDNA synthesis, add 0.6 μg of purified RNA, 2 μL of 10 μM reverse specific primer, and sterile deionized water up to 11.4 μL. Denature secondary structures of RNA by incubating at 65  C for 10 min, chill on ice, and keep it on ice for at least for 5 min, to avoid renaturation. Prepare the rest of the reaction on ice. 4. Prepare a master mix with buffer, RNase inhibitor, dNTPs, and DTT (no retrotranscriptase) according to manufacturer’s instructions. Mix well and keep it on ice. 5. Add the appropriate volume of master mix to the control tube RNA (RT-), and supplement with the volume of H2O that replaces retrotranscriptase. 6. Add retrotranscriptase to the master mix and mix well by gentle vortex and brief centrifugation. 7. Add 8.6 μL master mix to each of the RNA tubes, mix well by gentle vortex and spin down on centrifuge, and keep on ice. 8. Incubate it in the thermocycler for 10 min at 29  C, followed by 90 min at 55  C and 5 min at 85  C to denature the enzyme. 9. Dilute cDNA 1:7.5 with sterile deionized water and keep at 20  C.

3.4 Splice Pattern-Specific RT-PCR

Please see Note 9 before starting. 1. Prepare a master mix with 0.5 μL of each 10 μM primer, forward and reverse, and 5 μL of 2 Hot Fast ReadyMix with dye, per each reaction. Distribute 6 μL per each PCR tube or plate well. Add 4 μL of previously diluted cDNA. Mix well. 2. Incubate samples in a thermocycler for 6 min at 95  C, followed by 45 cycles of 15 s at 95  C, 30 s at 58  C, and 15 s at 72  C.

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3. Prepare a 2% agarose gel with TAE and DNA stain. 4. As the PCR mix already has the dye included, you can run your PCR products directly in the agarose gel without the need to add any loading dye. Run the marker in a side slot in the gel. Run the 2% agarose gel electrophoresis for 60 min at 80 V and image the result in a Chemidoc (Fig. 1b). 5. Use Image Lab software to quantitatively evaluate band intensity. This software allows you to quantify the fluorescence intensity of each band relative to a selected reference band. We further defined the percentage of exon skipping as the relative intensity ratio: unskipped/(skipped + unskipped) multiplied by 100 (Fig. 1c). Note that the % of exon skipping is a measure that only serves comparative purposes in a given experiment. See Note 4 for discussion. 3.5 Immunofluorescence Microscopy

1. Prepare a 12-well plate with gelatin-coated coverslips: in the tissue culture hood, lay one sterile coverslip per well using flamed forceps, add 0.5 mL of sterile 0.1% gelatin solution to each well, wait for 5 min, and quickly remove all the gelatin solution. Close the plate to prevent it from drying out. 2. Prepare the transfection mixture by adding transfection reagent dilution mix to SSO dilution mix (described in Subheading 3.2) on microtubes and incubate for 5–20 min. 3. Meanwhile, trypsinize the cells, resuspend it in culture medium, and count and make an appropriate dilution. Add 0.6 mL of cell suspension to each well over the gelatin-coated coverslip. 4. Subsequently, add 60 μL of transfection mixture to each well. Mix gently by rocking the plate back and forth and repeat after rotating horizontally 90 . Incubate for 24 h. 5. Switch to differentiation medium after confirming that cell confluence is above 80%. 6. After 4–6 days, check cell differentiation by phase-contrast inverted microscopy. You should see multinucleated myotubes formed by the fusion of multiple myoblasts. Proceed to immunodetection in the lab bench (step 7). 7. Discard culture medium and fix cells with 3.7% paraformaldehyde in PBS for 10 min at room temperature. 8. Permeabilize cells with 0.5% Triton X-100 in PBS for 10 min at room temperature, with gentle agitation. 9. Wash 3 5 min with 1 mL PBS. Remove all PBS. 10. Add 20 μL of blocking solution over the coverslip, carefully, to not disturb the cells. Incubate for 30 min at room temperature in a humid chamber. Remove blocking solution.

Short 16-mer LNA-SSO Restore Dystrophin in DMD Myotubes

45

11. Add 20 μL of primary antibody, polyclonal rabbit antidystrophin antibody. Incubate for 60 min at room temperature in a humid chamber. 12. Wash 3 5 min with 1 mL PBST. Remove all PBST. 13. Add 20 μL of secondary antibody, TRITC-conjugated antirabbit IgG antibody. Incubate for 60 min at room temperature in a humid chamber protected from light. 14. Wash 3 5 min with 1 mL PBST, protected from light. Remove all PBST. 15. To counterstain the nuclei, add 20 μL of DAPI DNA staining solution. Incubate for 10 min at room temperature in a humid chamber protected from light. 16. Wash with 1 mL PBST and mount the preparation over one drop of mounting medium on a glass slide. Seal the borders of coverslips with nail polish. 17. Image the result in a fluorescence confocal microscope (Fig. 2).

Fig. 2 Expression of dystrophin protein and its localization at the myotube membrane are restored after SSO-induced splicing modulation. A myoblast cell line derived from a DMD patient (DMD) was either transfected with SSO 51.1c at 50 nM or mock transfected; a myoblast cell line derived from a healthy individual was assayed as a control (Control). Cells were fixed 4 days after differentiation induction, immunostained with anti-dystrophin antibody, and nuclear counterstained with DAPI. Images were captured using an LSM 710 Confocal Point-Scanning Microscope (ZEISS), using the lasers Diode 405-30 (405 nm) for DAPI and DPSS 561-10 (561 nm) for TRITC

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Notes 1. The human myoblast cell line DMD is derived from a male patient with Duchenne muscular dystrophy, carrying a deletion in the dystrophin gene (DMD Δ exons 48–50), on the X chromosome. The absence of exons 48 to 50 in the dystrophin mature transcript leads to a frameshift that in turn introduces a premature stop codon (PTC) that will drive for degradation the incorrect mRNA, by the process of nonsense-mediated decay (NMD). The result is the lack of a functional protein. The forced skipping of exon 51, by hybridization with an antisense oligonucleotide, restores the reading frame of the coding sequence in the mRNA, allowing for the production of an internally truncated, shorter, but partially functional protein (see Fig. 3a). This is the molecular basis of the RNA-targeted therapeutic approach using splice modulation.

Fig. 3 The most evolutionarily conserved stretches of DMD exon 51 reveal regions that are candidates to be target for SSOs, with the aim of forcing exon 51 skipping and restore the reading frame of the DMD transcript. (a) In the DMD cell line, the genetic defect on DMD gene (Δ exons 48–50) generates a frameshift and a PTC (premature stop codon) that triggers NMD, resulting in no (or very low) production of protein. The forced exclusion of exon 51 restores the reading frame and allows production of an internally deleted but functional protein. (b) Comparison of several human DMD exon 51 orthologues drove the identification of three wellconserved regions that are shown highlighted. Region 51.1 is a stretch of 15 nt (68–82) conserved among all the species compared, from mammals to birds; the other stretches are region 51.3 spanning 15 nt (125–139) and region 51.2 of 11 nt (186–196), both conserved among all the mammals compared. Above the regions are depicted the SSOs used here to induce skipping of exon 51

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2. Human myoblast cell lines are maintained in culture in an undifferentiated state, always avoiding a density of cells over 80%, but, to study the expression of dystrophin, which only occurs in myotubes, we need to induce myoblast differentiation. Differentiation is achieved by culturing myoblasts at confluence density and serum deprivation. 3. The rationale for finding the best target sequences for the antisense SSOs in the exon 51 of human DMD gene was to look for the most conserved sequences among species. We collected the sequences for the several DMD gene orthologues available in Ensembl Genome Browser database (www. ensembl.org), identified the exons that correspond to the exon 51 in the human gene, and aligned all together using CLUSTALW tool (www.genome.jp). The most conserved stretches were named 51.1, 51.2, and 51.3, with 16, 11, and 15 nucleotides long, respectively. These stretches are shared among all the mammal genomes tested (see Fig. 3b). After a BLAST check for uniqueness in the cDNA database of the human genome, we selected the regions 51.1 and 51.3 to target. Remarkably, the region 51.1 is also targeted by the modified oligos already in clinical trials and recently approved for human therapy [2], corroborating our design approach. 4. LNA-modified oligos show higher affinity with the target sequence, and so a smaller antisense oligo can be used to efficiently interfere with gene expression in vitro and in vivo. A 14-mer LNA-modified oligonucleotide with phosphorothioate linkage backbone successfully silenced a liver-specific mRNA [3], and 8-mer fully LNA-substituted phosphorothioate oligonucleotides could silence microRNA families in a broad range of tissues in mouse [13]. We tested several oligonucleotides and found that efficiency depends on the target sequence and on the size of the SSO. The sequence 51.1 was found to be the better target for inducing exon 51 skipping, and among 13–16-mer oligos targeting this region, we observed that the 16-mer SSO was the most efficient oligo [11]. For region 51.3, among several different sizes, the 16-mer SSO induced the best rate of exon skipping, even though not so efficiently as when targeting sequence 51.1 (compare 51.3a and 51.1c lanes in Fig. 1b). Once the target region and oligo size are established, it is time to investigate the best sequence for the oligo. We tested fine-tuning variations of the sequence and found that 51.1c was more efficient than others in inducing exon skipping (compare 51.1c and 51.1d lanes in Fig. 1b). 5. Locked nucleic acid (LNA) synthetic chemistry [14, 15] is characterized by the inclusion of a methylene bridge connecting the 20 -oxygen and the 40 -carbon atoms in the furanose ring

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of ribose. The locking effect restricts the molecule conformation, leading to enhanced hybridization properties of the synthetic LNA-containing oligonucleotides [4, 7]. LNA is the nucleotide modification preferred for applications where high affinity is desirable [16]. Therapeutic oligonucleotides must be chemically modified in order to avoid rapid nuclease degradation in serum and renal filtration. Substituting three DNA monomers by LNA in both 30 and 50 ends of the ON improved the half-life in human serum more than ten times [5]. Two LNA monomers toward the 30 -end protected the ON from degradation at some level [17]. LNA internal positioning in addition to 30 and 50 end substitutions further increased protection from degradation [10]. Phosphorothioate modification consists of substitution of a non-bridging oxygen in phosphate linkages by sulfur, enhancing resistance to degradation by nucleases [18] and facilitating cellular uptake [19, 20]. These modifications generate ONs that are more suitable for in vivo studies. Phosphorothioate (PS) linkages, being fully compatible with LNA modification, further increase LNA ON’s protection from nucleases [8, 9]. 6. As for SSOs, we used 16-mer DNA oligonucleotides carrying a fully phosphorothioate-modified backbone and substituting LNA for DNA in 60% of the nucleotide residues, ensuring that the two terminal nucleotides at the 30 -end were obligatory LNAs, as well as the terminal nucleotide at the 50 -end. We designed the oligos with the help of Exiqon, and they have a policy to provide the sequence but not the LNA spike pattern of their designed ONs. However, readers are allowed to purchase the exact same oligonucleotide to reproduce our experiments by requesting the GeneGlobe catalogue number provided in Table 1. AO51, a 20 OMe RNA 20-mer antisense oligo, was a gently gift from Capucine Trollet (Centre de Recherche en Myologie, Sorbonne Universite´s, Paris) [12]. 7. We tested a range of different reagents for SSO transfection in our cell lines and had the best results when using the reverse transfection procedure with the Lipofectamine RNAiMAX transfection reagent in 12-well plates (3.8cm2). This is done by preparing the transfection mix directly on the bottom of the empty wells before adding the suspension of cells. Note that when preparing cells for microscopy, we plated cells on gelatincoated coverslips, and, in that case, the transfection mix was added immediately after adding the cells to avoid nonspecific trapping on the gelatin coat. You can use a scramble oligo sequence for control or just no oligo at all. 8. Performing cDNA synthesis using a reverse primer specific for the transcript you want to study will increase the sensitivity of the assay. Prepare an extra tube for the negative control (non-retrotranscribed control, RT-).

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9. For splice pattern-specific RT-PCR, we need to use a pair of primers that is specific to the constitutively included exons of the mRNA and that delimitates the region of alternative included exon. Preferentially the reverse primer should be different from the one used to prepare the cDNA, to increase specificity (see Fig. 1a). The Taq polymerase must be carefully chosen because the alternative isoforms in the cDNA will be competing for amplification and it is very tricky to get an informative result from the PCR. We had good results with KAPA2G Fast ReadyMix with dye. The negative control has the non-retrotranscribed control (RT-) as its target.

Acknowledgments We acknowledge the Association Franc¸aise Contre les Myopathies and the Platform for the Immortalization of Human Cells and Collaboration, Institut de Myologie, Paris, for the immortalized human cell lines, and Capucine Trollet (Centre de Recherche en Myologie, Sorbonne Universite´s, Paris) for oligonucleotide AO51. ˜es, Vanessa Borges Pires, and Kamel MamWe thank Ricardo Simo chaoui for collaboration and Ana de Jesus and Ana Margarida Nascimento for technical support. We would like to thank Marcia Triunfol for assistance in preparing this manuscript and Noe´lia Custo´dio for critical review. We further acknowledge funding from Fundac¸˜ao para a Cieˆncia e Tecnologia and FEDER/POR Lisboa 2020—Programa Operacional Regional de Lisboa, PORTUGAL 2020 (LISBOA-01-0145FEDER-016394; 007391). References 1. Yin W, Rogge M (2019) Targeting RNA: a transformative therapeutic strategy. Clin Transl Sci 12(2):98–112. https://doi.org/10.1111/ cts.12624 2. Aartsma-Rus A, Straub V, Hemmings R et al (2017) Development of exon skipping therapies for duchenne muscular dystrophy: a critical review and a perspective on the outstanding issues. Nucleic Acid Ther 27(5):251–259. https://doi.org/10.1089/nat.2017.0682 3. Lindholm MW, Elme´n J, Fisker N et al (2012) PCSK9 LNA antisense oligonucleotides induce sustained reduction of LDL cholesterol in nonhuman primates. Mol Ther 20(2):376–381. https://doi.org/10.1038/mt.2011.260 4. Kaur H, Babu BR, Maiti S (2007) Perspectives on chemistry and therapeutic applications of locked nucleic acid (LNA). Chem Rev 107

(11):4672–4697. https://doi.org/10.1021/ cr050266u 5. Kurreck J, Wyszko E, Gillen C, Erdmann VA (2002) Design of antisense oligonucleotides stabilized by locked nucleic acids. Nucleic Acids Res 30(9):1911–1918. https://doi. org/10.1093/nar/30.9.1911 6. Lundin KE, Højland T, Hansen BR et al (2013) Biological activity and biotechnological aspects of locked nucleic acids. Adv Genet 82:47–107. https://doi.org/10.1016/B9780-12-407676-1.00002-0 7. Petersen M, Nielsen CB, Nielsen KE et al (2000) The conformations of locked nucleic acids (LNA). J Mol Recognit 13(1):44–53. https://doi.org/10.1002/(SICI)1099-1352( 200001/02)13:13.0. CO;2-6

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8. Gupta N, Fisker N, Asselin MC et al (2010) A locked nucleic acid antisense oligonucleotide (LNA) silences PCSK9 and enhances LDLR expression in vitro and in vivo. PLoS One 5 (5):e10682. https://doi.org/10.1371/jour nal.pone.0010682 9. Stein CA, Hansen JB, Lai J et al (2010) Efficient gene silencing by delivery of locked nucleic acid antisense oligonucleotides, unassisted by transfection reagents. Nucleic Acids Res 38(1):e3. https://doi.org/10.1093/nar/ gkp841 10. Crinelli R, Bianchi M, Gentilini L, Magnani M (2002) Design and characterization of decoy oligonucleotides containing locked nucleic acids. Nucleic Acids Res 30(11):2435–2443. https://doi.org/10.1093/nar/30.11.2435 ˜ es R, Mamchaoui K, 11. Pires VB, Simo Carvalho C, Carmo-Fonseca M (2017) Short (16-mer) locked nucleic acid splice-switching oligonucleotides restore dystrophin production in Duchenne muscular dystrophy myotubes. PLoS One 12(7):e0181065. https:// doi.org/10.1371/journal.pone.0181065 12. Mamchaoui K, Trollet C, Bigot A et al (2011) Immortalized pathological human myoblasts: towards a universal tool for the study of neuromuscular disorders. Skelet Muscle 1:34. https://doi.org/10.1186/2044-5040-1-34 13. Obad S, dos Santos CO, Petri A et al (2011) Silencing of microRNA families by seedtargeting tiny LNAs. Nat Genet 43 (4):371–380. https://doi.org/10.1038/ng. 786 14. Obika S, Nanbu D, Hari Y et al (1997) Synthesis of 20 -O,40 -C-methyleneuridine and -cytidine. Novel bicyclic nucleosides having a fixed

C3, endo sugar puckering. Tetrahedron Lett 38(50):8735–8738. https://doi.org/10. 1016/S0040-4039(97)10322-7 15. Singh SK, Nielsen P, Koshkin AA, Wengel J (1998) LNA (locked nucleic acids): synthesis and high-affinity nucleic acid recognition. Chem Commun:455–456. https://doi.org/ 10.1039/A708608C 16. Lundin KE, Højland T, Hansen BR et al (2013) Biological activity and biotechnological aspects of locked nucleic acids. Adv Genet 82:47–107. https://doi.org/10.1016/B9780-12-407676-1.00002-0 17. Morita K, Takagi M, Hasegawa C et al (2003) Synthesis and properties of 20 -O,40 -C-ethylene-bridged nucleic acids (ENA) as effective antisense oligonucleotides. Bioorganic Med Chem 11(10):2211–2226. https://doi.org/ 10.1016/S0968-0896(03)00115-9 18. Stein CA, Subasinghe C, Shinozuka K, Cohen JS (1988) Physicochemical properties of phosphorothioate oligodeoxynucleotides. Nucleic Acids Res 16(8):3209–3221. https://doi. org/10.1093/nar/16.8.3209 19. Elme´n J, Lindow M, Schu¨tz S et al (2008) LNA-mediated microRNA silencing in non-human primates. Nature 452 (7189):896–899. https://doi.org/10.1038/ nature06783 20. Zhao Q, Matson S, Herrera CJ, Fisher E, Yu H, Krieg AM (1993) Comparison of cellular binding and uptake of antisense phosphodiester, phosphorothioate, and mixed phosphorothioate and methylphosphonate oligonucleotides. Antisense Res Dev 3(1):53–66. https://doi. org/10.1089/ard.1993.3.53

Chapter 5 Targeting Polyadenylation for Retention of RNA at Chromatin Evgenia Ntini and Ulf Andersson Vang Ørom Abstract The various steps of RNA polymerase II transcription, including transcription initiation, splicing, and termination, are interlinked and tightly coordinated. Efficient 30 end processing is defined by sequence motifs emerging in the nascent-transcribed RNA strand and the cotranscriptional binding of regulatory proteins. The processing of a mature 30 end consists of cleavage and polyadenylation and is coupled with RNA polymerase II transcription termination and the dissociation of the nascent RNA transcript from the chromatin-associated transcriptional template. The subcellular and subnuclear topological specificity of the various RNA species is important for their functions. For instance, the formation of RNA-binding protein interactions, critical for the final outcome of gene expression, may require the nucleoplasmic fully spliced and polyadenylated form of an RNA transcript. Thus, interfering with the critical step of transcription termination and 30 end formation provides a means for assaying the functional potential of a given RNA of interest. In this protocol, we describe a method for blocking 30 end processing of the nascent RNA transcript, by using RNase H-inactive antisense oligonucleotides targeting cleavage and polyadenylation, delivered via transient transfection in cell culture. Key words Transcription termination, 30 end formation, Polyadenylation signal, Nascent RNA transcript, 2-O-methyl phosphorothioate, 20 OMePS, Antisense oligonucleotide, Transcriptional readthrough

1

Introduction The processing of a mature 30 end in nascent RNA transcripts is tightly coupled with transcription termination of the transcriptionally engaged RNA polymerase II (Pol II) and is controlled by underlying sequence motifs emerging in the nascent RNA strand and the cotranscriptional binding of regulatory proteins termed transcription termination factors [1–3]. The most common pathway of 30 end processing and transcription termination in mammalian systems is the poly(A)-dependent termination, which involves the cleavage and polyadenylation stimulating factor (CPSF) cotranscriptionally interacting with Pol II. Once CPSF recognizes a

Ulf Andersson Vang Ørom (ed.), RNA-Chromatin Interactions: Methods and Protocols, Methods in Molecular Biology, vol. 2161, https://doi.org/10.1007/978-1-0716-0680-3_5, © Springer Science+Business Media, LLC, part of Springer Nature 2020

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polyadenylation signal (“pA signal” or “pAS,” usually the hexamer AAUAAA) that emerges in the nascent RNA strand, it induces a slowing down or transient pausing on the transcribing Pol II. Upon following exposure of a downstream GU-rich or U-rich sequence, CstF (cleavage stimulating factor) disengages CPSF allowing endonucleolytic cleavage of the nascent RNA transcript at the poly (A) site, which is about ~30 nucleotides downstream of the pA signal. The nascent cleaved RNA 30 end gets rapidly polyadenylated, leading to the formation of a mature 30 end, and the dissociation of the nascent RNA transcript from the chromatin-associated transcriptional template. In the accompanying torpedo model, a 50 –30 exoribonuclease (XRN2) attacks and degrades the downstream cleavage product following cleavage and polyadenylation at the poly(A) site, eventually causing displacement of Pol II from DNA and transcription termination [2]. Thus, Pol II transcription termination is tightly coupled with the efficiency of 30 end processing, and in cases where cleavage and polyadenylation of the nascent RNA transcript are impeded, Pol II transcriptional readthrough may occur for several kilobases downstream of the failed poly (A) site, while the nascent RNA transcript is (transiently) retained at chromatin. Upon chromatin dissociation, the nascent RNA transcripts will interact with RNA-binding proteins to perform nuclear functions or get exported to the cytoplasm for downstream processes including translation. The subcellular localization of RNAs is of great importance; especially for certain RNA species, like the long noncoding RNAs (lncRNAs), understanding their topological specificity may prove helpful in predicting functionality [4]. For instance, we recently showed that the nascent form of a chromatindissociated lncRNA transcript (A-ROD) is important for transcription regulation of its target gene (DKK1) within the 3D proximity space of a pre-established enhancer-promoter chromosomal loop [5]. In such studies, inhibiting the step of chromatin dissociation of the nascent RNA transcripts is a useful experimental strategy for assaying their potential functions. One way to achieve this is by genetic manipulation of the underlying sequences using the CRISPR/Cas9 system [6, 7], leading to the disruption of transcription termination motifs in genetically modified stable cell lines. Although the establishment of stable cell lines offers the advantage that downstream experiments can repetitively be performed in the same system at any time, it may require some effort to achieve the targeted effect in all present chromosomal alleles of the cell (some widely used human cell lines are polyploid). In addition, inhibiting transcription termination of a targeted locus in a stable cell line may generate secondary effects on the transcriptional state of the gene locus in the long term, causing the turning down of the associated transcription initiation step, eventually leading to overall transcriptional

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repression of the locus [8, 9]. This may comprise a drawback when trying to address complex biological questions, such as functionally uncouple and characterize the roles of the act of transcription itself versus its RNA products from a given locus. An alternative and overall faster experimental strategy is based on steric hindrance by blocking the sequences emerging in the nascent RNA strand recognized by transcription termination factors and the polyadenylation machinery. This is achieved by delivering 20 -O-methyl phosphorothioate antisense oligonucleotides (20 OMePS) in transient transfection experiments. The inclusion of 20 -O-methyl modified ribonucleotides in the antisense oligo ensures that no RNase H activity is triggered upon hybridization to the RNA target (which would be the case if the antisense oligonucleotide was DNA), securing that breakdown is not induced and the endogenous targeted RNA transcript remains intact (Fig. 1). The phosphorothioate (PS) bond confers resistance against nuclease degradation (cellular exo- and endonucleases), increasing the halftime of the delivered antisense oligonucleotides and thereby the efficiency of their blocking activity. Similar blocking activity via steric hindrance can be achieved with Morpholinos (MOs) [10], but 20 OMePS constitutes a cheaper alternative. 20 OMePS antisense oligonucleotides have been widely used in vivo to block splice sites and promote exon skipping, in noteworthy cases as a putative treatment of diseases arising from isoform misexpression, like the Duchenne muscular dystrophy [11, 12]. We recently used 20 OMePS antisense oligonucleotides in cell culture to block 30 end processing of the lncRNA A-ROD in vitro, in order to assess its

5’

Base

HO

O 2’O-Methyl O

S Phosphorothioate linkage

P

O CH3 Base

O

O

OH

O CH3

O S

P

O

OH

3’

Fig. 1 Structure of a 20 OMePS trinucleotide

Base O

OH

O CH3

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functional potential on target gene expression [5]. The effects can be evaluated within short time (1–3 days) after transfection.

2 2.1

Materials 30 RACE

1. Magnetic beads Oligo(dT)25. 2. Superscript III. 3. First-strand cDNA synthesis primer, AP_RT.30 RACE: GGCCACGCGTCGACTAGTACTTTTTTTTTTTTTTT TTTVN. 4. –AUAP primer GGCCACGCGTCGACTAGTAC. 5. Gene-specific forward primer: designed on the sense strand within ~200–1000 bp upstream of the putative poly(A) site. 6. Taq DNA polymerase. 7. PCR purification kit. 8. Gel extraction kit.

2.2 20 OMePS Antisense Oligonucleotide Transfection

1. Control oligo sequence: mC∗mC∗mU∗mC∗mU∗mU∗mA∗mC∗mC∗mU∗mC∗ mA∗mG∗mU∗mU∗mA∗mC∗mA∗mA∗mU∗mU∗mU∗ mA∗mU∗mA (∗phosphorothioate bond; m: 2-O-Methyl RNA base). Order the 20 OMePS oligos with HPLC purification (length, ~25 nt; GC content, ~40–60%). 2. Lipid-based transfection reagent. 3. Reduced-Serum Medium, for the formation of the transfection complexes (according to the transfection reagent enclosed protocol). 4. DMEM supplemented with 5% fetal bovine serum, for cell culture.

3

Methods

3.1 30 RACE to Validate Gene’s Major Poly(A) Site

In order to design 20 OMePS antisense oligonucleotides blocking 30 end processing of a nascent RNA transcript, the position of the mature 30 end where cleavage and polyadenylation (CPA) takes place should be known. This can be retrieved from available polyA+ RNA-seq data or from genome-wide 30 tag sequencing techniques [13, 14]. Since it is common for human transcription units to have more than one CPA site, stronger and weaker ones, it is advisable to experimentally verify a gene’s major 30 end by locusspecific 30 rapid amplification of cDNA ends (30 RACE).

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1. Extract whole-cell RNA or total nuclear RNA as in ref. 13 (see Note 1). 2. Enrich for polyadenylated RNA using commercial oligo(dT)25 Dynabeads according to the company’s enclosed protocol. 3. Subject 50 ng of poly(A)+ enriched RNA to first-strand cDNA synthesis with the primer AP_RT.30 RACE, using SuperScript III and the accompanying enclosed protocol. 4. Two microliter of the cDNA product is directly used in 30 RACE PCR reactions (50 μl) with 1.25 units Taq DNA polymerase and primer pair the gene-specific forward primer and AUAP. An indicative PCR program is (Initial denaturation step, 95  C 60 s, 30 cycles, 95  C 30 s, 63  C 20 s, 72  C 20 s, and final extension, 72  C for 10 min) (see Note 2). 5. Purify PCR products using a commercial kit (or perform DNA extraction using neutral-pH phenol and ethanol precipitation). Load the purified 30 RACE products on 1% agarose gel. 6. Excise the agarose bands and purify DNA using a commercial gel extraction kit. Subject the 30 RACE PCR products to sequencing with the gene-specific primer, according to the sequencing company’s instructions. 3.2 Design of 20 OMePS Antisense Oligonucleotides

Once the CPA site(s) is known and experimentally verified, 20 OMePS antisense oligonucleotides are designed to target it and block it in transient transfection. 1. Inspect the elements and sequence motifs around the CPA site. For genes terminating transcription via the poly(A)-dependent pathway (see Subheading 1), a pA signal (pAS) is found within ~10–30 nucleotides upstream of the CPA site. The pA signal is the hexamer A[A/T]TAAA or single substitution variants (“weak hexamers”) [12] (see Note 3). Downstream of the CPA site, GT-rich (“2GT/T”) or T-rich motifs are usually found, which constitute binding sites for the RNA-binding and 30 end processing factor CstF64 [12, 14–16] (see Note 4). Typical CPA flanking sequences for the host gene GAPDH and the lncRNA A-ROD are shown in Fig. 2.

GAPDH

TGTA pAS CPA 2GT/T T-rich 5’ CGCACCTTGTCATGTACCATCAATAAAGTACCCTGTGCTCAACCAGTTACTTGTCCTGTCTTATTCTAGGGTCTGGGGCAGAG 3’

A-ROD

pAS CPA T-rich 5’ GTTCTGGTTCTGTGCAAATTCTAATAAAAATGAAATGTAATTGATCCTCAAAAATATGATTCATCTATTTTAAGTCATCAGTCAACA 3’

Fig. 2 DNA sequences around the cleavage and polyadenylation site (CPA) of GAPDH and A-ROD. The polyadenylation signal (pAS) is the hexamer AAUAAA. The UGUA motif is found upstream of the pAS and recruits the cleavage factor I (CFI) [15]. 2GU/U and U-rich motifs are found downstream of the CPA and constitute binding sites for CstF64 [16]. Positions of the two 20 OMePS oligos used for blocking A-ROD 30 end formation [5] are underlined

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2. Design the 20 OMePS antisense oligonucleotides to target the actual CPA, the upstream pA signal, and/or the downstream GT-rich and T-rich elements. All blockers can be used in the same experiment to block 30 end processing, in co- or independent transfections, in accordance to the biological question of interest and the overall experimental design. As an example, the blockers used to inhibit A-ROD 30 end formation [5] are shown in Fig. 2 (see Note 5). 3.3 Transfection of 20 OMePS in Cell Culture

1. Grow cells to ~60–80% confluence (see Note 6). 2. Transfect the 20 OMePS antisense oligonucleotides at 500 nM final concentration using Hiperfect (see Note 7). 3. Harvest cells after 24 h (see Note 8) and proceed to downstream applications/experimental validation (see Note 9).

4

Notes 1. In the case of low expressed RNA transcripts or transcripts known to be enriched in the nucleus, it helps to start the 30 RACE protocol by subjecting isolated nuclear RNA (instead of whole-cell RNA) to poly(A) selection, as this will significantly reduce background and increase specificity in detecting the final 30 RACE products. 2. The 30 RACE PCR may require some optimization. If multiple or fuzzy bands appear after gel electrophoresis, this is an indication of high background. Try using less cDNA product as template in the PCR, increase the annealing temperature, decrease the number of PCR cycles, and design longer genespecific forward primers. It may also help to perform a “nested” 30 RACE, that is, remove 1–2 μl from the first 30 RACE PCR and use it as template in PCR with AUP and a second genespecific primer lying downstream of the first and closer to the expected 30 end. The sizes of the observed gel electrophoresis bands should match the distance between the position of the designed gene-specific forward primer(s) and the expected poly (A) site(s). 3. Sequence motifs are referring to the respective DNA strandtranscriptional template, but they are recognized by RNA-binding proteins (30 end processing and transcription termination factors) at the emerging nascent RNA strand, so the actual poly(A) signal is A[A/U]UAAA. 4. 2GT/T motifs are (GTGTT), (TGTGT), (GTTGT), and (TGT[GC]T). T-rich motifs are ([ATCG]TTTT), (T[ATCG] TTT), (TT[ATCG]TT), (TTT[ATCG]T), and (TTTT [ATCG]) (from ref. 12).

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5. The designed 20 OMePS antisense oligonucleotides are usually ~25 nt long, with ~40–60% GC content. It is advisable to check that they are not prone to forming strong secondary structures by using online available folding prediction and oligonucleotide properties tools. 6. We have transfected 20 OMePS in adherent MCF-7 cells grown in DMEM supplemented with 5% FCS. Any cell line could be used according to its own growth and maintenance standards. 7. Titration experiments can be performed to obtain an optimal final 20 OMePS concentration where maximum blocking activity is achieved with a minimum cellular toxicity. Other lipidbased transfection reagents can be tested as well. 8. Blocking activity can already be assessed at 24 h posttransfection, by assaying the caused locus-specific transcriptional readthrough, but also monitored at later time points. In cases of low efficiency, a second hit may be performed 16 h after the first transfection to increase the 20 OMePS uptake. 9. 20 OMePS-mediated blocking activity targeting the CPA is assayed by measuring the caused transcriptional readthrough extended downstream of the blocked CPA site. Several stateof-the-art methodologies can be performed to precisely measure transcriptional readthrough. One of them is the nuclear run-on method which assays transcriptionally engaged RNA polymerase II [19]. This can be coupled with locus-specific real-time quantitative reverse transcription PCR (qRT-PCR) using amplicons spanning the regions upstream and downstream of the CPA site and extracting the relative signal ratio in control and CPA-blocking conditions. A second method is similarly measuring transcriptional readthrough by qRT-PCR on transcriptionally nascent RNA obtained after short-pulse metabolic labeling [20]. Transcriptional readthrough can also be assayed by performing chromatin immunoprecipitation (ChIP) for RNA polymerase II phosphorylated at Ser2 (P-Ser2 Pol II); this modification reflects transcriptionally engaged Pol II. qRT-PCR is subsequently performed on the chromatin-immunoprecipitated DNA fragments, again using short amplicons spanning upstream and downstream of the blocked CPA site [5].

Acknowledgments This study was supported by an Alexander-von-Humboldt postdoctoral research fellowship to EN.

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2’-O-methyl phosphorothioate and morpholino antisense oligonucleotides for Duchenne muscular dystrophy exon skipping. J Gene Med 11:257–66 12. Yang L, Niu H, Gao X, Wang Q, Han G, Cao L, Cai C, Weiler J, Yin H (2013) Effective exon skipping and dystrophin restoration by 2’-o-methoxyethyl antisense oligonucleotide in dystrophin-deficient mice. PLoS One 8: e61584 13. Pelechano V, Wilkening S, Jarvelin AI, Tekkedil MM, Steinmetz LM (2012) Genome-wide polyadenylation site mapping. Methods Enzymol 513:271–296. https://doi.org/10.1016/ B978-0-12-391938-0.00012-4 14. Ntini E, Jarvelin AI, Bornholdt J, Chen Y, Boyd M, Jorgensen M, Andersson R, Hoof I, Schein A, Andersen PR, Andersen PK, Preker P, Valen E, Zhao X, Pelechano V, Steinmetz LM, Sandelin A, Jensen TH (2013) Polyadenylation site-induced decay of upstream transcripts enforces promoter directionality. Nat Struct Mol Biol 20(8):923–928. https:// doi.org/10.1038/nsmb.2640 15. Conrad T, Orom UA (2017) Cellular fractionation and isolation of chromatin-associated RNA. Methods Mol Biol 1468:1–9. https:// doi.org/10.1007/978-1-4939-4035-6_1 16. Salisbury J, Hutchison KW, Graber JH (2006) A multispecies comparison of the metazoan 30 -processing downstream elements and the CstF-64 RNA recognition motif. BMC Genomics 7:55. https://doi.org/10.1186/14712164-7-55 17. Martin G, Gruber AR, Keller W, Zavolan M (2012) Genome-wide analysis of pre-mRNA 30 end processing reveals a decisive role of human cleavage factor I in the regulation of 30 UTR length. Cell Rep 1(6):753–763. https:// doi.org/10.1016/j.celrep.2012.05.003 18. Yao C, Biesinger J, Wan J, Weng L, Xing Y, Xie X, Shi Y (2012) Transcriptome-wide analyses of CstF64-RNA interactions in global regulation of mRNA alternative polyadenylation. Proc Natl Acad Sci U S A 109 (46):18773–18778. https://doi.org/10. 1073/pnas.1211101109 19. Gardini A (2017) Global run-on sequencing (GRO-Seq). Methods Mol Biol 1468:111–120. https://doi.org/10.1007/ 978-1-4939-4035-6_9 20. Louloupi A, Orom UAV (2018) Metabolic pulse-chase RNA Labeling for pri-miRNA processing dynamics. Methods Mol Biol 1823:33–41. https://doi.org/10.1007/9781-4939-8624-8_3

Chapter 6 Simultaneous Detection of RNAs and Proteins with Subcellular Resolution Sunjong Kwon, Koei Chin, and Michel Nederlof Abstract We describe the detailed methods of “immunoFISH” to analyze the expression level and the spatial localization of RNA transcripts and proteins on cultured cells and formal-fixed, paraffin-embedded (FFPE) tissue sections. On cultured cells, we detect specific transcripts using the Stellaris fluorescence in situ hybridization (FISH) probes labeled with fluorophores that target multiple regions along the desired transcripts and proteins combining the immunofluorescent staining. On FFPE tissue sections, we use the RNAscope FISH probes, modified branched DNA (bDNA) probes to amplify the RNA signals, followed by immunofluorescent staining for protein detection. The abundance, composition, and spatial distribution are determined by signals from fluorescently labeled proteins and individual transcripts of images acquired using high-resolution fluorescence microscopy. Key words immunoFISH, smFISH, Immunofluorescent, Subcellular localization

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Introduction The expression level and spatial distribution of diverse RNA transcripts and proteins are important contributing factors of the behavior of normal and cancer cells. Comprehensive and quantitative measures of transcripts and proteins are usually performed using mass spectrometry and RNA-seq. However, the information of spatially variable expression in cellular or tissue contexts for cancer heterogeneity is determined using imaging tools of in situ hybridization (ISH) and immunohistochemical staining (IHC), respectively [1]. IHC is performed, based on the specific binding between the proteins and their specific antibodies in cellular or tissue contexts, and immunofluorescence (IF) is a particular type of IHC that applies specific antibodies which are chemically conjugated to fluorescent dyes for protein imaging [2]. IF has been successfully applied for

Ulf Andersson Vang Ørom (ed.), RNA-Chromatin Interactions: Methods and Protocols, Methods in Molecular Biology, vol. 2161, https://doi.org/10.1007/978-1-0716-0680-3_6, © Springer Science+Business Media, LLC, part of Springer Nature 2020

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a Cultured cells fixation

smFISH probes secondary Ab

imaging

primary Ab

b

FFPE sections pretreatment

RNAscope probes

FISH signal amplification

Immunofluorescence (primary then secondary Ab)

imaging

Fig. 1 ImmunoFISH workflow. (a) On fixed cultured cells, fluorophore-labeled smFISH probes and primary antibody are incubated to specifically recognize target RNA and protein and then fluorophore-labeled secondary antibody, followed by three-dimensional wide field deconvolution imaging system. (b) On FFPE tissue sections, slides are deparaffinized and pretreated for target retrieval, incubated with target RNA-specific RNAscope probes, and then RNA signals are amplified by serial hybridizations, followed by immunofluorescent staining for co-imaging of RNAs and proteins

labeling multiple antigens in the same cellular or tissue contexts with multichannel fluorescence microscopy techniques [3]. ISH is performed, using hybridization procedures between the specifically labeled nucleic acid strand (the probe) and its complementary RNA sequences in fixed cells or tissues, followed by visualization of the target transcript through the labeled probes [4]. Fluorescence in situ hybridization (FISH) is a specific ISH that uses fluorescence-labeled probes and provides enhanced resolution, specificity, and speed. Recently, single-molecule fluorescence in situ hybridization (smFISH) has been developed to identify the total copy number of mRNAs in intact cells and tissues at the single-molecule/single-cell level [5, 6]. SmFISH uses multiple short (around 20 nucleotides) probes, which reduces potential off-target effects from a few probes to target RNA [7]. The short probes need less stringent conditions for hybridization and post-hybridization washing steps, which can be applicable with immunofluorescent co-staining for protein by adding antibodies together with FISH probes. Therefore, we can successfully detect proteins and RNA simultaneously by combining smFISH and IF—a process referred to as “immunoFISH” [1]. Using immunoFISH, we can simultaneously detect endogenous RNAs and proteins, in single molecules at the level of single cells on cultured cells (Fig. 1). The yield of prepared RNA is typically lower in FFPE tissues than cultured cells [8], and higher background noises are unavoidable in FFPE sections [9]. We need to enhance the sensitivity of FISH to increase the signal-to-noise ratio to amplify detection signals. We use RNAscope (Advanced Cell Diagnostics, USA) FISH, which uses a unique probe design strategy for sequential hybridizations for simultaneous signal amplification and background suppression to achieve single-molecule visualization while preserving tissue morphology [10]. However, the efficiency of simultaneous immunofluorescent co-staining was significantly reduced by the more stringent conditions needed for RNAscope FISH. We overcame this by performing RNAscope FISH first

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followed by immunofluorescence staining unlike in cultured cells as described above. We present detailed protocols in this chapter of simultaneous in situ analyses of location, composition, and expression levels of transcripts and proteins by combining aspects of RNA FISH and IF.

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Materials Prepare all solutions with ultrapure nuclease-free water, and perform all steps with nuclease-free tips, pipettes, and tubes. Prepare and store all reagents at room temperature (RT), unless otherwise pointed out. Any florescent reagent should be kept in the dark. Carefully follow institutional guidelines for working with, and disposing of, hazardous chemicals and biohazard materials.

2.1

smFISH

1. TE buffer: 10 mM Tris–HCl, 1 mM EDTA, pH 8.0. 2. 20
Saline Sodium Citrate (SSC): 3.0 M NaCl, 0.3 M sodium citrate, pH 7.0. 3. 10
phosphate-buffered saline (PBS) pH 7.4, RNase-free. 4. Formaldehyde solution for molecular biology, 36.5–38% in H2O. 5. DNase/RNase-free distilled water. 6. Formamide (deionized). 7. Dextran sulfate, average molecular weight >500,000. 8. Ethanol for molecular biology. 9. Hybridization solution (see Note 1): 10% dextran sulfate, 2
SSC, 10% formamide. 10. Wash buffer (see Note 1): 2
SSC, 10% formamide. 11. 40 ,6-diamidino-2-phenylindole (DAPI). 12. ProLong Gold Antifade Mountant. 13. 18
18 mm square #1½ coverslip. 14. Six-well cell culture plates. 15. Humidified chamber (or equivalent): pipette tip box covered with aluminum foil to protect from light, a single layer of Parafilm placed on the tip insert, and the bottom part halffilled with water to maintain humidity in the chamber. 16. Superfrost Plus Microscope slides. 17. 37  C laboratory oven.

2.2

RNAscope FISH

Most reagents are available from RNAscope® Fluorescent Multiplex Reagent Kit (ACD Cat# 320850: https://acdbio.com/ rnascope%C2%AE-fluorescent-multiplex-assay). 1. RNAscope Protease Plus.

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Table 1 Fluorescent dye options for RNAscope FISHa RNAscope probe channel ID

Amp 4 Alt A-FL

Amp 4 Alt B-FL

Amp 4 Alt C-FL

C1

Alexa 488

Atto 550

Atto 550

C2

Atto 550

Alexa 488

Atto 647

C3

Atto 647

Atto 647

Alexa 488

a

Adapted from the manuals of Advanced Cell Diagnostics (https://acdbio.com/technical-support/user-manuals)

2. RNAscope Target Retrieval Reagents. 3. HybEZ™ oven, humidity control tray, slide rack (20 slide capacity). 4. Hydrophobic barrier pen. 5. Tissue-Tek Staining Dishes. 6. Water bath or incubator, capable of holding temperature at 40  C. 7. Fume hood. 8. Paper towel or absorbent paper. 9. Glass beakers. 10. Hot plate. 11. Aluminum foil. 12. Forceps, large. 13. Thermometer. 14. Amp 1-FL, Amp 2-FL, Amp 3-FL. 15. Amp 4-FL–Alt A Display module, Amp 4-FL–Alt B Display module, Amp 4-FL–Alt C Display module (Table 1). 16. 50
RNAscope Wash Buffer. 2.3 Immunofluorescence

1. Specific primary antibodies (see Note 2). 2. Secondary antibodies, Alexa 488 conjugated or Alexa 647 conjugated. 3. PBS, pH 7.4. 4. PBST: 1
PBS, 0.025% Tween, pH 7.4. 5. Normal goat serum (see Note 3). 6. Bovine serum albumin. 7. Blocking solution: 10% goat serum, 1% BSA, PBS, pH 7.4. 8. Ab solution: 5% goat serum, 1% BSA, PBS, pH 7.4. 9. 50
22 mm coverslip. 10. SlowFade Gold Antifade Mountant with DAPI.

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2.4

FISH Probes

2.4.1 Design and Labeling of smFISH Probes

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SmFISH Probes are 18–22 mer DNA oligonucleotides that are fluorescently labeled with a fluorescent dye at 30 end, and targeted GC% is 45. The number of single-labeled DNA oligos is minimum 25 and maximum 48, and each oligo is separated by at least 2 nucleotides to accommodate fluorophore. We recommend to use the Stellaris probe design web tool (https://www. biosearchtech.com/support/tools/design-software/stellarisprobe-designer) and to order the Stellaris RNA FISH probes from Biosearch Technologies, USA. The probes are shipped dry and should be dissolved in nuclease-free TE buffer (10 mM Tris–HCl, 1 mM EDTA, pH 8.0) to create a probe stock concentration of 25 μM. We recommend to make aliquots and keep at 20  C in the dark for storage lasting longer than a month.

2.4.2 RNAscope FISH Probe

We recommend to search and order FISH probes of target genes in the RNAscope Catalog Probes List (https://acdbio.com/catalogprobes). If your gene of interest is not available in the catalog, you can order Made-to-Order New Targets Probes following instructions on https://acdbio.com/target-probes-made-order.

2.5 Coverslips for Cell Cultures

Coverslips are supplied as prewashed, and we use two ways to sterilize coverslips for cell cultures.

2.5.1 Autoclaving

Place around ten coverslips without overlapping in a self-seal sterilization pouch, 31/200 X 83/400 . Perform the dry cycle of an autoclave with a 30-min step. Open the autoclaved sterilization pouch in a tissue culture hood and use sterile forceps to take out each coverslip to a well of six-well plate.

2.5.2 Ethanol and UV Light

Put the coverslips on coverslip rack, dip in beaker filled with 100% ethanol, and then sonicate them for 5 min. Take out sonicated coverslips on racks from 100% ethanol in a tissue culture hood and dry coverslips in hood with UV for 30 min, and then put each coverslip to a well of six-well plate.

2.6 FFPE Tissue Sections

Preparing FFPE sections varies dependent on the sources of the samples. We prefer to freshly cut FFPE slides of 5 μm and to keep them at 4  C for maximum 2–3 months (see Note 4).

3

Methods

3.1 immunoFISH on Cultured SKBR3 Breast Cancer Cells

Here we describe the protocol to detect two different mRNAs and one protein, and it can be modified to detect one mRNA and two different proteins by applying one smFISH probe and two primary/secondary Ab sets.

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1. Seed around 0.5
106 SKBR3 cells on a sterilized #1½ coverslip (18
18 mm) in a six-well plate and grow cells in growth media for 48 h to be 70–80% confluent. 2. Aspirate off growth media, wash coverslip with 2 mL of PBS, and fix with 1 mL of 4% formaldehyde in PBS at RT for 10 min. 3. Wash twice with 2 mL of PBS for 5 min each. 4. Permeabilize with 3 mL of 70% ethanol at 4  C for at least an hour. Coverslips can be stored at 4  C in 70% ethanol up to a week before hybridization. 5. Put the coverslip with the cells facing up in a well of new six-well plate. 6. Prehybridize with 2 mL of wash buffer (2
SSC, 10% formamide) at RT for 10 min. 7. Put carefully the coverslip with the cells facing down over 50 μL the hybridization solution (10% dextran sulfate, 2
SSC, 10% formamide) containing smFISH probes, conjugated with Quasar® 570 Dye (Abs Max ¼ 548 nm, Em Max ¼ 566 nm), Quasar® 670 Dye (Abs Max ¼ 647 nm, Em Max ¼ 670 nm), and a primary Ab on Parafilm in humidified chamber. For smFISH probes concentration per target RNA, we initially try four different doses of 25, 50, 100, and 250 nM and determine the concentration which provides highest signal-to-noise ratio. 8. Incubate the coverslip in a dark humidified chamber at 37  C overnight. 9. Put the coverslip with the cells facing up in a well of six-well plate. 10. Wash the coverslip twice in 2 mL of wash buffer (2
SSC, 10% formamide) at 37  C for 15 min and 30 min, respectively, in the dark. 11. Put carefully the coverslip with the cells facing down over 50 μL the hybridization solution (10% dextran sulfate, 2
SSC, 10% formamide) containing Alexa 488 (Abs Max ¼ 490 nm, Em Max ¼ 525 nm)-conjugated secondary antibody on Parafilm in humidified chamber (see Note 5). 12. Incubate the coverslip in a dark humidified chamber at RT for 1 h. 13. Put the coverslip with the cells facing up in a well of six-well plate. 14. Wash coverslips in 2 mL of wash buffer (2
SSC, 10% formamide) at RT for 30 min in the dark. 15. Incubate coverslips in 2 mL of wash buffer (2
SSC, 10% formamide) with 10 ng/mL DAPI for nuclear counterstaining at RT for 30 min in the dark.

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16. Wash coverslips with 2 mL of 2
SSC at RT for 5 min. 17. Mount coverslips with ProLong Gold and cure for at least 24 h in the dark. 3.2 Imaging of immunoFISH on Cultured Breast Cancer Cells

Imaging can be performed on high-resolution wide field microscope and in order to require 0.2 μm optical sections, we use Deltavision CoreDV Widefield Deconvolution System (https:// www.ohsu.edu/advanced-light-microscopy-core/hi-res-widefield) equipped with the following: 1. Stand: Olympus IX71 wide field inverted microscope. 2. Objectives: 60
oil N.A. 1.42. 3. Seven-color solid-state illumination unit for exciting fluorophores across the visible spectrum. 4. Polychroic beam splitters, optimized for imaging with bluegreen-red-far red fluorophore combinations. 5. Filter Set: DAPI (457/50), FITC (528/38), TRITC (617/23), Cy5 (685/40). 6. Camera: Nikon CoolPix HQ cooled CCD camera. 7. Motorized stage controlled by XYZ nano-motors for accurate z-stack and point-visiting functions. 8. SoftWoRx™ image restoration software (GE Healthcare, USA).

3.3 RNA smFISH Spot Counting and Protein Immunostaining Measurement

1. The raw images are deconvolved with SoftWoRx™ image restoration software to remove focus blur or, more specifically, reassign out-of-focus light contributions to their proper location in the source image. 2. The images are then reconstructed into 3D visualizations using Imaris software (Bitplane, USA) (Fig. 2). The 3D images show RNA molecules as bright “particles,” with a size of ~0.25 μm and a distinct signal from background noise (Figs. 3 and 4, see Note 6). 3. RNA particles are counted using a Spot Detection program from Imaris software as follows: Select “spots” on the 3D View tab. Add new spot per each RNA particles. Click on “Default” on Favorite creation parameters. Choose Source channel matching the fluorophore of target probes. Provide an estimated diameter for the spots (0.25 μm). Click on “Background subtraction.” To subtract background signals, the 3D images are applied with an algorithm based on Mexican Hat Effect.

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Fig. 2 SmFISH RNA particles are counted using the Spot Detection program from Imaris software (Bitplane). (a, b) HER2 mRNA (green) and intronic RNA (red) channels from immunoFISH on SKBR3 breast cancer cells. (c, d) Each RNA spot detected in the Spot Detection program is denoted with a white dot. (e, f) From each image c and d, the total number of HER2 mRNA spots and intronic RNA spots is 17,041 and 271, respectively, counted by Imaris. An automatic threshold is applied by calculating intensities and sizes from all spots based on statistics supplied by Imaris

Add Filter of “Quality above automatic threshold” by choosing Filter Type: Quality. Imaris detects an automatic threshold to insert the spots by calculating intensities from all spots based on statistics. If automatic threshold does not correspond to RNA particles by eye, the threshold could be refined manually changing threshold up and down. Export the statistics text file to determine the number of total RNA spots. 4. The number of total spots in an image is divided by the number of nuclei to obtain the number of transcripts in single cells (see Note 7).

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Fig. 3 HER2 mRNA subcellular location by immunoFISH of HER2 mRNAs (green), intronic RNAs (red), nuclear lamin A/C protein (blue), and DAPI (gray) on SKBR3 breast cancer cells. RNAs and protein were detected by adding smFISH probes and primary antibody together followed by incubating with fluorophoreconjugated secondary antibody. (a) HER2 mRNAs are detected both in cytoplasm and in nuclei. (b) HER2 intronic RNAs are exclusively detected in nuclei and micronuclei. (c) Lamin A/C protein staining defines nuclei and micronuclei. (d) Intron containing HER2 mRNA signals are colocalized with exon containing HER2 mRNA, suggesting these colocalized particles (yellow) are HER2 pre-mRNAs. The inset image shows a magnified view of the micronuclei (arrow). Image was obtained using a Deltavision Automated Widefield microscopy (60
objective, NA ¼ 1.42). Bar is 5 μm. (This figure is adapted from [1] with permission in accordance with the Creative Commons Attribution 4.0 International (CC BY 4.0) license)

5. Protein intensities in immunoFISH level are determined using QiTissue software (Quantitative Imaging Systems, USA) as follows: Nuclei are automatically segmented with a model-driven algorithm, one of the standard available segmentation functions in QiTissue that uses the DAPI-labeled image channel to find objects that are bright and with acceptable size and roundness features. The cytoplasm of each cell is automatically segmented by growing the nuclear object out with a constraint that it may not extend beyond cytoplasmic signals. The segmentation masks were interactively

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Fig. 4 ImmunoFISH to analyze protein levels and mRNA expression in human formalin-fixed, paraffin-embedded (FFPE) tissue sections. Please note HER2 RNAs were detected by RNAscope FISH, followed by immunostaining of HER2 (a) or HER2/KRT14 proteins (b). (a) HER2 protein (green) and HER2 mRNA (red) in a DAPI stained (blue) HER2+ invasive breast cancer. Nine tile images (3
3) using a Zeiss Axio Imager.M2 with a Plan-Apochromat 20
, NA ¼ 0.8 objective, were obtained, stitched, and fused to make single image. Bar is 100 μm. (b) High-resolution co-imaging of HER2 protein (green), HER2 mRNA (white), and the myoepithelial protein, KRT14 (red), on HER2+ breast tumor nest. Image around a tumor nest was obtained using a Deltavision Automated Widefield microscopy (60
objective, NA ¼ 1.42). Please note that the KRT14-positive cells do not express HER2 mRNA. Bar is 10 μm. (This figure is adapted from [1] with permission in accordance with the Creative Commons Attribution 4.0 International (CC BY 4.0) license)

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checked and corrected for proper cell boundaries as needed. This was accomplished by drawing lines with the “scissors” tool to provide operator input where adjacent cell cytoplasms overlap and boundaries are unclear. Then, protein intensities are determined by calculating mean pixel intensities of specific Ab staining channel from each segmented cell, by using the intensity feature calculation function in QiTissue (see Note 7). 3.4 immunoFISH on FFPE Tissue Sections

3.4.1 Pretreatment of FFPE Tissue Sections

Here we describe the protocol to detect one mRNA and two different proteins, and it can be modified to detect two different mRNAs and one protein by applying two RNAscope FISH probes and one primary/secondary Ab set. The protocols described for Subheadings 3.4.1 and 3.4.2, are adapted from the manuals of Advanced Cell Diagnostics (https://acdbio.com/technical-sup port/user-manuals). 1. Put 5 μm slides in a 24-slide holder and bake them in a dry oven at 65  C for 1 h. 2. Deparaffinize slides: (a) Place slides in xylene, twice for 5 min each. (b) Place slides in 100% ethanol, twice for 2 min each. (c) Air-dry slides for 5 min at RT. 3. Boil the slides in 1
Target Retrieval solution at 95–100  C for 15 min. 4. Rinse the slides in the distilled water twice. 5. Wash slides in fresh 100% ethanol and then air-dry. 6. Draw a loop around tissue using the hydrophobic barrier pen. Let the barrier dry completely at RT at least 5 min. 7. Incubate slides with Protease Plus in the HybEZ™ oven at 40  C for 30 min. Please make sure Protease Plus solution to cover the tissue sections entirely (see Note 8). 8. Put the slides in a 24-slide holder and wash slides by moving the holder up and down several times in distilled water twice.

3.4.2 RNAscope FISH

1. Warm RNAscope FISH probes for 10 min at 40  C, and then cool down to RT. 2. Put the slides in HybEZ™ slide rack and hybridize RNAscope FISH probes for 2 h at 40  C in the HybEZ™ oven. Please make sure FISH probes to cover the tissue sections entirely. 3. Put the slides in a 24-slide holder and wash slides by moving the holder up and down several times in 1
RNAscope Wash Buffer at RT for 2 min twice. 4. Put the slides in HybEZ™ slide rack and hybridize Amp 1-FL for 30 min at 40  C in the HybEZ™ oven. Please make sure Amp 1-FL to cover the tissue sections entirely.

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5. Put the slides in a 24-slide holder and wash slides by moving the holder up and down several times in 1
RNAscope Wash Buffer at RT for 2 min twice. 6. Put the slides in HybEZ™ slide rack and hybridize Amp 2-FL for 15 min at 40  C in the HybEZ™ oven. Please make sure Amp 2-FL to cover the tissue sections entirely. 7. Put the slides in a 24-slide holder and wash slides by moving the holder up and down several times in 1
RNAscope Wash Buffer at RT for 2 min twice. 8. Put the slides in HybEZ™ slide rack and hybridize Amp 3-FL for 30 min at 40  C in the HybEZ™ oven. Please make sure Amp 2-FL to cover the tissue sections entirely. 9. Put the slides in a 24-slide holder and wash slides by moving the holder up and down several times in 1
RNAscope Wash Buffer at RT for 2 min twice. 10. Put the slides in HybEZ™ slide rack and hybridize Amp 4-FL-Alt B (to get target RNA labeled Atto 550) for 15 min at 40  C in the HybEZ™ oven. Please make sure Amp 4-FL to cover the tissue sections entirely (see Notes 9 and 10). 11. Put the slides in a 24-slide holder and wash slides by moving the holder up and down several times in 1
RNAscope Wash Buffer at RT for 2 min twice. 3.4.3 Immunofluorescent Staining

1. Right after RNAscope FISH, incubate the slides in PBS for 5 min three times. 2. Incubate the slides in blocking solution (10% goat serum, 1% BSA, PBS) at RT for 1 h (see Note 3). 3. Add two different primary antibodies diluted in Ab solution (5% goat serum, 1% BSA, PBS) to the slides and incubate at 4  C overnight (see Note 3). The primary antibodies should be raised in different species. 4. Wash slides with PBST (1
PBS, 0.025% Tween, pH 7.4) at RT for 5 min three times. 5. Add secondary antibodies conjugated with Alexa Fluor 488 and Alexa Fluor 647 diluted in Ab solution (5% goat serum, 1% BSA, PBS) to the slides and incubate at RT for 1 h. 6. Wash slides with PBST (1
PBS, 0.025% Tween, pH 7.4) at RT for 5 min three times. 7. Add 2–3 drops of SlowFade Gold Antifade Mountant with DAPI reagent directly onto each slide. 8. Slowly cover the slide with a 50
22 mm coverslip and make sure no bubbles are trapped on top of the section.

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Imaging can be performed on any wide field microscope system with CCD or CMOS camera. We image the regions of interest (or full tissue sections) from a fluorescence microscopy system (Carl Zeiss Microscopy, Germany) equipped with the following: 1. Stand: Axio Imager.M2 upright microscope. 2. Objectives: Plan-Apochromat 20
/0.8 objective. 3. Illuminant: Solid-state light source Colibri 7, type FR-R[G/Y] BV-UV. 4. Filter Sets: 96HE (DAPI), 38HE (GFP), 43HE (Cy3), 50 (Cy5). 5. Camera: Orca Flash 4.0 LT CMOS camera (Hamamatsu Photonics, Japan). 6. Stage: Stepper motor-controlled XYZ stage for multichannel tile and autofocus imaging capabilities. 7. Zen Blue edition imaging software. High-resolution imaging can also be obtained using a Deltavision Automated Widefield microscopy (60
objective, NA ¼ 1.42) as described in Subheading 3.2.

4

Notes 1. Hybridization and wash buffers for smFISH are commercially available from Biosearch Technologies: Stellaris® RNA FISH Hybridization Buffer, Stellaris RNA FISH Wash Buffer A, and Stellaris RNA FISH Wash Buffer B. We find that these commercial buffers also provide good signal to background ratio for RNA detection and immunofluorescence staining with several Abs. 2. We find that antibodies working well for immunofluorescent staining do not always work for immunoFISH buffer condition containing 10% dextran sulfate and 10% formamide because the antibody-antigen interactions are usually mediated by the weaker binding of fewer hydrogen bonds than annealing of oligonucleotide probes to RNAs in smFISH. It also has been reported that formamide and elevated temperature weaken binding of the antibodies [11]. Therefore, high-affinity antibodies are prerequisite and should be tested for immunofluorescent staining with hybridization and wash buffers before being applied in immunoFISH. 3. We recommend that a serum matching the species of the secondary antibody is raised in order to minimize nonspecific binding of antibodies to FFPE sections. If two secondary

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antibodies are from different species, we recommend to use blocking sera from the species of both secondary antibodies. 4. We find that RNA qualities on FFPE sections frequently vary due to differences in fixation and storing conditions between clinical samples. We recommend to confirm that FFPE sections could provide high positive control signal and no negative control background. We use 3-plex RNAscope positive control probes of POLR2A, PPIB, and UBC and negative control probes of dapB on adjacent sections to confirm RNA preservation. 5. Usually, most autofluorescent signals are found in the green channel which provides worst signal to background ratio for smFISH. Therefore, we use Ab detection on green channel on immunoFISH, avoiding the detection of smFISH signal on green channel. 6. As an internal RNA FISH standard when visualizing and quantifying RNAs on smFISH, we attempted to detect poly-A RNA using oligo-dT probes in cultured cells. However, the FISH signals were dense and spread throughout the cytoplasm and nucleoli. We found it is problematic to resolve and count RNA spots detected using oligo-dT probes. 7. For alternate RNA particle measurements, we used QiTissue software to train a spot detection algorithm and count RNA spots satisfying a minimum intensity threshold in each segmented 3D cell. Training was done interactively by selecting several spots that met minimal visual detection requirements. These spot counts were consistent with the Imaris results we described in Subheading 3.3. Therefore, we were able to measure transcript counts as well as protein expression levels, in both the nuclear and cytoplasm volumetric regions of each cell. 8. We recommend to adjust the protease conditions for optimal protein detection by immunofluorescent staining. For example, an antibody provided better signal with 20 min treatment than 30 min. 9. There are three options of Amp 4-FL–Alt A, B, or C available for alternate fluorescent color modules for Amp 4-FL (Table 1). Fluorescent label combination of Amp 4-FL–Alt A, B, or C should be selected not overlapped with immunofluorescent channels. 10. We applied RNAscope FISH probes on cultured cells with smFISH conditions, in which protease treatment is not included. However, RNA particles were significantly reduced compared to those when smFISH probes are applied. The big molecular size of preamplifier and amplifier probes might have

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difficulty approaching to the complementary sequences of target RNAs, which are typically present in the ribonucleoprotein (RNP) complexes.

Acknowledgments We thank Dr. Joe Gray (OHSU, USA) for providing his substantial support and conceptual guidance on our immunoFISH research. This work was partially supported by OHSU SOM Faculty Innovation Fund Pilot Program (S.K.). References 1. Kwon S, Chin K, Nederlof M, Gray JW (2017) Quantitative, in situ analysis of mRNAs and proteins with subcellular resolution. Sci Rep 7 (1):16459. https://doi.org/10.1038/ s41598-017-16492-1 2. Matos LL, Trufelli DC, de Matos MG, da Silva Pinhal MA (2010) Immunohistochemistry as an important tool in biomarkers detection and clinical practice. Biomark Insights 5:9–20 3. Eng J, Thibault G, Luoh SW, Gray JW, Chang YH, Chin K (2020) Cyclic multiplexedimmunofluorescence (cmIF), a highly multiplexed method for single-cell analysis. Methods Mol Biol 2055:521–562. https://doi.org/10. 1007/978-1-4939-9773-2_24 4. Kwon S (2013) Single-molecule fluorescence in situ hybridization: quantitative imaging of single RNA molecules. BMB Rep 46(2):65–72 5. Femino AM, Fay FS, Fogarty K, Singer RH (1998) Visualization of single RNA transcripts in situ. Science 280(5363):585–590 6. Raj A, van den Bogaard P, Rifkin SA, van Oudenaarden A, Tyagi S (2008) Imaging individual mRNA molecules using multiple singly labeled probes. Nat Methods 5(10):877–879 7. Raj A, Tyagi S (2010) Detection of individual endogenous RNA transcripts in situ using

multiple singly labeled probes. Methods Enzymol 472:365–386 8. Ludyga N, Grunwald B, Azimzadeh O, Englert S, Hofler H, Tapio S, Aubele M (2012) Nucleic acids from long-term preserved FFPE tissues are suitable for downstream analyses. Virchows Arch 460(2):131–140. https://doi.org/10.1007/s00428-011-11849 9. Saka SK, Wang Y, Kishi JY, Zhu A, Zeng Y, Xie W, Kirli K, Yapp C, Cicconet M, Beliveau BJ, Lapan SW, Yin S, Lin M, Boyden ES, Kaeser PS, Pihan G, Church GM, Yin P (2019) Immuno-SABER enables highly multiplexed and amplified protein imaging in tissues. Nat Biotechnol 37(9):1080–1090. https://doi. org/10.1038/s41587-019-0207-y 10. Wang F, Flanagan J, Su N, Wang LC, Bui S, Nielson A, Wu X, Vo HT, Ma XJ, Luo Y (2012) RNAscope: a novel in situ RNA analysis platform for formalin-fixed, paraffin-embedded tissues. J Mol Diagn 14(1):22–29 11. Kochan J, Wawro M, Kasza A (2015) Simultaneous detection of mRNA and protein in single cells using immunofluorescence-combined single-molecule RNA FISH. BioTechniques 59 (4):209–212, 214, 216 passim. https://doi. org/10.2144/000114340

Chapter 7 In Vivo Crosslinking of Histone and RNA-Binding Proteins Yong-Eun Kim, Kyoon Eon Kim, and Kee K. Kim Abstract Protein–protein interactions are essential in various cellular processes including regulation of gene expression, formation of protein complexes, and cellular signaling transduction. In particular, several proteins in the nucleus interact to regulate transcription and RNA splicing. These protein–protein interactions are short and weak and occur through transient processes, making it difficult to identify these interactions. In addition, detection of interacting partners in vitro using cell lysates cannot provide complete information due to the loss of spatial organization and changes in protein modification. Here we describe an in vivo crosslinking technique using disuccinimidyl suberate (DSS), which is useful to capture and stabilize proteins to analyze the interacting proteins. Key words In vivo crosslinking, Disuccinimidyl suberate, Protein–protein interaction, Histone, RNA-binding protein, Immunoprecipitation, Mass spectrometry

1

Introduction Protein–protein interactions play crucial roles in orchestrating various functions in cellular processes. Some proteins form rigid complexes such as structures of subcellular compartments, transport machineries across membrane, and chromatin, through stable and strong protein interactions [1–3]. On the other hand, proteins involved in intracellular signaling pathways and regulation of gene expression and degradation interact transiently [4]. These transient interactions occur temporarily, weakly, and rapidly in phenomena such as protein phosphorylation, conformational changes, and transfer to specific compartments [5, 6]. Therefore, identification of transient protein interactions is important for a fundamental understanding of biological processes at the molecular level. Although the yeast two-hybrid system and co-immunoprecipitation are commonly used to determine the binding partners of target proteins in cellular systems, the

Kyoon Eon Kim and Kee K. Kim contributed equally to this work. Ulf Andersson Vang Ørom (ed.), RNA-Chromatin Interactions: Methods and Protocols, Methods in Molecular Biology, vol. 2161, https://doi.org/10.1007/978-1-0716-0680-3_7, © Springer Science+Business Media, LLC, part of Springer Nature 2020

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techniques for identifying protein–protein interaction have advanced considerably over the recent years [7, 8]. Nowadays chemical crosslinking and fluorescence resonant energy transfer (FRET) have become useful tools for identification and characterization of binding partners of target protein [9]. Stable protein interactions are easy to isolate as complexes in physiological conditions using co-immunoprecipitation. However, transient protein interactions are difficult to isolate as protein complexes because of their weak and short binding. Therefore, a method to tightly bind and immobilize proteins for isolating complexes that interact momentarily and weakly is required. Chemical crosslinking forms covalent bonds between proteins that exist in close proximity, resulting in the joining of two or more molecules. Thus, chemical crosslinking can be used to identify transient interactions as well as stable interactions [10]. Crosslinking reagents have specific chemical groups that react with certain functional groups of proteins. Representative reactive chemical groups for the crosslinking of proteins include N-hydroxysuccinimide ester (NHS ester), imidoester, maleimide, and diazirine (Fig. 1a). NHS ester and imidoester react with the primary amines (–NH2) existing at the N-terminal of proteins and in the side chain of lysine residues [11]. Maleimide reacts with sulfhydryls (–SH) of cysteine residues. Diazirine is most common photo-reactive chemical group, which is activated by UV light. Photo-reactive crosslinkers are used for nonspecific binding in vitro and in vivo [12]. Crosslinkers are either homo- or hetero-bifunctional reagents with identical or non-identical reactive groups, respectively, at the end of spacer arms. The most widely used chemical crosslinker is the homobifunctional NHS ester that can be used in a one-step reaction procedure. The NHS ester reacts with primary amines in physiological conditions (pH 7.2–9) to yield amide bonds and to release N-hydroxysuccinimide (NHS) (Fig. 1b) [13]. In addition, at physiologic pH, the primary amine in lysine is positively charged, and so, it can be exposed to the surface of most proteins and crosslink with NHS esters with high efficiency. Some examples of homobifunctional NHS esters are disuccinimidyl suberate (DSS) and bis [sulfosuccinimidyl] suberate (BS3) with spacer arms of 11.4 A˚ (Fig. 1c) [13, 14]. DSS is water-insoluble and membranepermeable, whereas BS3 is water-soluble and has a charged group [15]. Therefore, DSS has been used to detect interacting proteins in vivo, allowing the identification of proteins that engage in transient interaction with a variety of other partners in a given pathway and in spatial organization. In general, covalently crosslinked proteins are identified through a series of processes such as immunoprecipitation, SDS-PAGE, in-gel digestion, and mass spectrometry. Here we describe the in vivo crosslinking methods using DSS to verify the

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Fig. 1 Chemical structure of crosslinker. (a) Reactive chemical group of a popular crosslinker used for protein conjugation. (b) NHS ester reaction scheme forming an amide bond with the primary amine of protein for crosslinking. R represents one end of a crosslinker having the NHS ester reactive group. (c) Homobifunctional crosslinker having the NHS ester reactive group

interaction between histones and RNA-binding proteins in the nucleus.

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Materials Prepare all solutions using distilled and ultrapure water.

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2.1 Crosslinking Using DSS

1. Crosslinker solution: Dissolve DSS in DMSO (dimethyl sulfoxide) at 25 mM. Prepare DSS immediately before use (see Note 1). 2. 1
phosphate-buffered saline (PBS): 20 mM sodium phosphate, 0.15 M NaCl, pH 8.0. Dissolve 2.65 g of Na2HPO4, 0.16 g of NaH2PO4, and 8.77 g of NaCl in 800 ml of water. Adjust the pH to 8.0 with HCl or NaOH, and then make up to 1 l with water. The solution is sterilized by autoclaving and stored at room temperature. 3. Quench solution: 1 M Tris, pH 7.5. Dissolve 12.11 g of Tris base in 80 ml of water. Adjust the pH to 7.5 with HCl and make up to 100 ml with water. The solution is sterilized by autoclaving and stored at room temperature. 4. Trypsin-EDTA solution.

2.2 Nuclear Extraction and Immunoprecipitation

1. Tris-buffered saline (TBS): 50 mM Tris–Cl, 0.15 M NaCl, pH 7.5. Dissolve 6.05 g of Tris and 8.76 g of NaCl in 800 ml of water. Adjust the pH to 7.5 with HCl. Make volume up to 1 l with water. The solution is sterilized by autoclaving and is stored at 4  C. 2. Cytoplasmic fraction buffer: 20 mM HEPES, 0.1 mM EDTA, 5 mM NaF, 0.01 mM Na2MoO4, pH 7.5. Dissolve 0.48 g of HEPES, 20 μl of 0.5 M EDTA, 10 μl of 0.1 M Na2MoO4, and 0.024 g of NaF in 80 ml of water. Adjust the pH to 7.5 with NaOH and make volume up to 100 ml with water. Sterilize by filtering through a 0.2 μm filter. Store the filtered solution at 4  C (see Note 2). 3. Nonidet P-40 (NP-40): Dissolve 1 ml of 100% NP-40 in 9 ml of water to prepare a 10% solution. 4. RIPA buffer: 20 mM Tris–HCl, 150 mM NaCl, 1% NP-40, 1% sodium deoxycholate, 0.5% SDS, 1 mM EDTA, pH 7.5. Add 2 ml of 1 M Tris–HCl (pH 7.5), 0.88 g of NaCl, 1 ml of 100% NP-40, 1 g of sodium deoxycholate, 0.5 g of SDS, and 0.2 ml of 0.5 M EDTA into 80 ml of water. When the solution is completely dissolved, adjust volume to 100 ml with water. Sterilize by filtering through a 0.2 μm filter. Store the filtered solution at 4  C (see Note 3). 5. EDTA-free protease inhibitor cocktail tablets: Dissolve one tablet in a suitable volume of extraction buffer such as cytoplasmic faction buffer and RIPA buffer. 6. 2 U/μl DNase. 7. RNase-A/T1 mix: 2 mg/ml of RNase A and 5000 U/ml of RNase T1. 8. Rabbit polyclonal anti-histone H3 antibody.

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9. Mouse monoclonal anti-myc antibody. 10. Normal rabbit-IgG. 11. Protein G magnetic beads for immunoprecipitation. 2.3 SDSPolyacrylamide Gel Electrophoresis (SDSPAGE)

1. Thirty percent of acrylamide/Bis-acrylamide solution (29:1): Dissolve 29 g of acrylamide and 1 g of Bis-acrylamide in 50 ml of water. Make volume up to 100 ml with water, and sterilize by filtering through a 0.2 μm filter. Store the filtered solution at 4  C in a dark bottle for no more than 1 month (see Note 4). 2. Separating gel: 0.375 M Tris–HCl, pH 8.8, 10% acrylamide/ Bis-acrylamide, 0.1% SDS, 0.1% APS, and 0.1% TEMED. Add 6.7 ml of 30% acrylamide/Bis-acrylamide solution, 5 ml of 1.5 M Tris–HCl (pH 8.8), 0.2 ml of 10% SDS, 0.2 ml of 10% APS, 0.01 ml of TEMED, and 7.9 ml of water into a 50 ml conical tube, and mix carefully (see Note 5). 3. Stacking gel: 0.125 M Tris–HCl, pH 6.8, 5% acrylamide/Bisacrylamide, 0.1% SDS, 0.1% APS, and 0.1% TEMED. Add 1.67 ml of 30% acrylamide/Bis-acrylamide solution, 2.5 ml of 0.5 M Tris–HCl (pH 6.8), 0.1 ml of 10% SDS, 0.1 ml of 10% APS, 0.01 ml of TEMED, and 5.63 ml of water into a 50 ml conical tube and mix carefully (see Note 6). 4. Ten percent of ammonium persulfate (APS): Dissolve 0.1 g of APS in 1 ml of water. Prepare APS solution freshly before use. 5. N,N,N,N0 -Tetramethyl-ethylenediamine (TEMED). 6. SDS-PAGE running buffer: 25 mM Tris–HCl, 192 mM glycine, 0.1% SDS. Dissolve 3.02 g of Tris base, 14.4 g of glycine, and 1 g of SDS in 900 ml of water. Make volume up to 1 l with water. Store the solution at room temperature. 7. 2
sample buffer: 65.8 mM Tris–HCl, pH 6.8, 26.3% glycerol, 2.1% SDS, 0.01% bromophenol blue. Before use, add 5% of β-mercaptoethanol. Store the solution at room temperature.

2.4 Immunoblot Analysis

1. Transfer buffer: 25 mM Tris–HCl, 192 mM glycine, 20% methanol. Dissolve 30.2 g of Tris base and 144 g of glycine in 900 ml of water. Adjust volume up to 1 l with water to prepare 10
transfer buffer solution. Sterilize by filtering through a 0.45 μm filter, and store the solution at room temperature. Add 100 ml of 10
transfer buffer and 200 ml of methanol to 700 ml of water. Store the solution at 4  C. 2. 1
PBST buffer: 137 mM NaCl, 2.7 mM KCl, 10 mM Na2HPO4, 1.8 mM KH2PO4, 0.05% Tween 20. Dissolve 80 g of NaCl, 2 g of KCl, 14.4 g of Na2HPO4, and 2.4 g of KH2PO4 in 800 ml of water, and adjust the pH to 7.4 with NaOH or HCl. Make volume up to 1 l with water to prepare 10
PBS solution. Sterilize the solution by autoclaving and

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store at room temperature. To prepare 1
PBST buffer, dilute 100 ml of 10
PBS solution in 800 ml of water, and add 500 μl of Tween 20. Make volume up to 1 l with water and store at room temperature. 3. Blocking buffer: Dissolve 5 g of skim milk powder in 80 ml of 1
PBST buffer. Make volume up to 100 ml with 1
PBST buffer. 4. Nitrocellulose membrane. 5. Filter paper. 6. Rabbit polyclonal anti-histone H3 antibody: Dilute antibody in blocking buffer at a ratio of 1:1000. 7. Mouse monoclonal anti-myc antibody: Dilute antibody in blocking buffer at a ratio of 1:5000. 8. Goat anti-rabbit IgG antibody and HRP conjugate: Dilute antibody in blocking buffer at a ratio of 1:5000. 9. Goat anti-mouse IgG antibody and HRP conjugate: Dilute antibody in blocking buffer at a ratio of 1:5000. 10. Enhanced chemiluminescent (ECL) substrate. 2.5 Coomassie Blue Staining

1. Staining solution: 50% methanol, 10% glacial acetic acid, 0.1% Coomassie Brilliant Blue R-250. Dissolve 1 g of Coomassie Brilliant Blue R-250 in 400 ml of water, and then add 500 ml of methanol and 100 ml of glacial acetic acid. Store the solution at room temperature. 2. Destaining solution: 40% methanol, 10% glacial acetic acid. Add 400 ml of methanol and 100 ml of glacial acetic acid in 500 ml of water. Store the solution at room temperature.

3

Methods To examine the interaction of histones and RNA-binding proteins, we performed in vivo crosslinking using DSS according to the procedure illustrated in Fig. 2.

3.1 In Vivo Crosslinking

1. Grow HeLa cells on 150 mm cell culture plate until the cells are 80–90% confluent. Ten 150 mm cell culture plates are required to identify the binding partners of histone H3 or overexpressed myc-tagged Rbfox2 and myc-tagged Rbfox3 (see Note 7). 2. Rinse the cells with PBS and trypsinize with trypsin-EDTA solution to obtain single-cell suspensions (see Note 8). 3. Wash the cells twice in PBS to remove the amine-containing components remaining in the media (see Note 9).

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Fig. 2 Procedures of in vivo crosslinking using disuccinimidyl suberate (DSS) for identifying the interactions of histone and RNA-binding protein in the nucleus

4. Resuspend the cells in 800 μl of PBS. Add 200 μl of 25 mM DSS (crosslinker solution) into a concentrated cell suspension to obtain a final concentration of 5 mM DSS (see Note 10). 5. Incubate the reaction mixture at room temperature for 30 min. 6. Add 20.4 μl of quench solution (1 M Tris) into the reaction mixture to a final concentration of 20 mM Tris (see Note 11). 7. Incubate the quenching reaction mixture for 15 min at room temperature. 3.2 Immunoprecipitation (IP)

1. Wash the cells three times in ice-cold TBS buffer to remove excessive crosslinker solution (see Note 12). 2. Gently resuspend cells in 1 ml of cytoplasmic fraction buffer supplemented with complete protease inhibitor cocktail. Allow the cells to swell on ice for 15 min. 3. Add 50 μl 10% NP-40 to a final concentration of 0.5%, and vortex the tube vigorously for 10 s at the highest setting (see Note 13). Centrifuge at 14,000
g and 4  C for 30 s and remove the supernatant (cytoplasmic faction).

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4. Resuspend the nuclear pellet in 800 μl of RIPA buffer containing complete protease inhibitor cocktail. Incubate the suspension for 20 min on ice. 5. Sonicate the lysate 15 times for 1 s each at 30% amplitude microns power setting to disrupt the nuclear membrane and to cleave the DNA and RNA. 6. Add 20 μl of DNase and 20 μl of RNase-A/T1 mix into the lysate, followed by incubation at 37  C for 5 min (see Note 14). Centrifuge at 14,000
g and 4  C for 10 min. 7. Transfer the supernatants to a new microcentrifuge tube. All supernatants are used for IP. 8. Add each ~20 μg of anti-histone H3 antibody or anti-myc antibody into the nuclear lysates of HeLa cells or myc-tagged Rbfox2 and myc-tagged Rbfox3 overexpressed in HeLa cells, respectively (see Note 15). Incubate at 4  C for 1.5 h with rotation. 9. During step 8, prepare Dynabeads™ protein G for use in capturing IgG. Resuspend Dynabeads™ protein G beads in the vial. Transfer 85 μl of Dynabeads™ protein G beads to a new microcentrifuge tube. Place the microcentrifuge tube on the magnetic rack to separate the beads from the solution, and remove the supernatant. After removing the microcentrifuge tube from the magnetic rack, add 200 μl of RIPA buffer to the microcentrifuge tube and gently resuspend the Dynabeads™ protein G using a pipette (see Note 16). 10. Add complexes of antibody and crosslinked proteins to a microcentrifuge tube containing Dynabeads™ protein G. Incubate at 4  C for 1 h with rotation. 11. Place the microcentrifuge tube on the magnetic rack, and remove the supernatant. Then, separate the microcentrifuge tube from the magnetic rack. 12. Wash the Dynabeads™ protein G-antibody-crosslinked protein complexes by pipetting using 1 ml of RIPA buffer. Place the microcentrifuge tube on the magnetic rack and remove the supernatant. 13. Repeat washing twice as in step 12. 14. Add 100 μl of 1
sample buffer diluted in RIPA buffer, and gently resuspend the Dynabeads™ protein G-antibody-crosslinked protein complexes using a pipette. 15. Heat the samples in a heat block for 5 min at 95  C, and then allow the sample to cool on ice. 3.3 Immunoblot Analysis

1. Prepare the 10% SDS-PAGE gel (see Note 17). 2. Gently pull the comb out of the glass plates in one smooth motion.

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3. Rinse the sample well with distilled water and remove the water completely. 4. Place SDS-PAGE gel inside the electrophoresis chamber. 5. Load 1% of the immunoprecipitated samples (see Note 18). 6. Pour the SDS-PAGE running buffer into electrophoresis chamber. Run the gel at 150 V for approximately 1 h, until the dye front reaches the bottom of the gel. 7. Separate the gel from the glass plates and remove the stacking gel. Equilibrate the gel in transfer buffer for wet transfer. 8. Place the mini gel holder cassette in the order sponge-filter paper-gel-nitrocellulose membrane-filter paper-sponge (see Note 19). 9. Relocate the mini gel holder cassette to the transfer apparatus, and add the transfer buffer so that cassette is covered with the buffer. 10. Run the transfer at a current of 200 mA for 4 h. Place the transfer apparatus on ice or in a cold room to maintain 4  C. 11. Block the membranes with blocking buffer containing 5% skim milk for 1 h at room temperature on a shaker. 12. Incubate membranes with anti-histone H3 or anti-myc antibody diluted in blocking buffer overnight at 4  C on a shaker. 13. Wash the membranes three times with 1
PBST buffer for 15 min each. 14. Incubate the membrane with anti-rabbit or anti-mouse IgG conjugated to horseradish peroxidase (HRP) diluted in blocking buffer for 1 h at room temperature. 15. Wash the membranes three times with 1
PBST buffer for 15 min each. 16. Apply ECL substrate to the membrane for chemiluminescent signal development. 17. Acquire the images using X-ray film exposure or an automated image acquisition system (Fig. 3a, b) [16]. 3.4 Preparation of Proteins for Mass Spectrometry Analysis

After verification of crosslinking by immunoblot analysis, perform SDS-polyacrylamide gel electrophoresis (PAGE) again according to the procedure in Subheading 3.3, step 1–6 to isolate proteins crosslinked with histone H3 or myc-tagged Rbfox2 and myc-tagged Rbfox3. 1. 99% of the immunoprecipitated sample is subjected to SDS-PAGE. 2. Separate the gel from glass plates and stain the gel in staining solution for 1 h with gentle agitation.

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Fig. 3 Immunoblotting to confirm the crosslinking of histone and RNA-binding protein. (a) Nuclear proteins extracted from HeLa cells treated with DSS were immunoprecipitated with anti-H3 antibody. IgG antibody was used as a negative control. One percent of the immunoprecipitated sample was used for immunoblotting analysis with anti-H3 antibody. IgG derived from anti-H3 antibody and histone H3 are shown. Histone H3 containing the crosslinked complexes labeled with the dotted boxes was analyzed by mass spectrometry using the remaining 99% of immunoprecipitated sample. (b) HeLa cells transfected with myc-tagged Rbfox2 or myc-tagged Rbfox3 were treated with DSS. After nuclear protein extraction and immunoprecipitation, 1% of the immunoprecipitated sample was used for immunoblotting with anti-myc antibody. Mock vector was transfected as a negative control. The crosslinked complexes labeled with dotted boxes were analyzed by mass spectrometry using the remaining 99% of immunoprecipitated sample. Dimer and monomer indicate dimers and monomers of Rbfox2 or Rbfox3, respectively. IgG(H), immunoglobulin G heavy chain, derived from the anti-myc antibody

3. Place the gel in destaining solution and agitate gently (see Note 20). 4. Change the destaining solution several times until the gel background is destained. 5. Cut and isolate the blue-stained protein bands from the gel to identify the crosslinking proteins (see Note 21). 6. Request in-gel digestion and analysis of LC-MS/MS.

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Notes 1. The NHS ester hydrolyzes readily and becomes non-reactive. Discard the unused crosslinker solution. The length of the spacer arm in homo- and hetero-bifunctional crosslinkers is useful in determining the distances between two crosslinked residues either in the same protein or between two proteins in close interaction. Crosslinkers with short spacer arms can easily cause intramolecular crosslinking. Choose a crosslinker suitable for interactions between neighboring proteins. 2. Dissolve 2.42 g of Na2MoO4·2H2O in 100 ml of water to make 0.1 M solution. Dissolve 18.61 g of disodium EDTA·2H2O in 80 ml of water. Adjust the pH to ~8.0 by adding NaOH to dissolve the salt. Make the volume up to 100 ml with water to prepare a 0.5 M solution. The solution is sterilized by autoclaving and stored at room temperature. 3. Dissolve 121.14 g of Tris base in 800 ml of water. Adjust the pH to 7.5 with HCl. Make volume up to 1 l with water. The solution is sterilized by autoclaving and stored at room temperature. 4. Acrylamide is a neurotoxic substance that damages the peripheral nervous system. Additionally, acrylamide is water soluble and absorbed through the oral cavity, respiratory tract, and skin. Wear gloves, eye protection, and a mask when weighing powdered acrylamide. Dissolve acrylamide solution in a fume hood. 5. Dissolve 181.7 g of Tris base in 900 ml of water. Adjust pH to 8.8 with HCl. Make volume up to 1 l with water to prepare 1.5 M Tris–HCl. The solution is sterilized by autoclaving and stored at room temperature. Dissolve 10 g of sodium dodecyl sulfate (SDS) in 80 ml of water. Make volume up to 100 ml with water. Sterilize by filtering through a 0.2 μm filter and store the solution at room temperature. 6. Dissolve 60.6 g of Tris base in 900 ml of water. Adjust pH to 6.8 with HCl. Make volume up to 1 l with water to prepare 0.5 M Tris–HCl. The solution is sterilized by autoclaving and stored at room temperature. 7. HeLa cells were transfected with myc-tagged RBFOX2 and myc-tagged RBFOX3 by electroporation using the Amaxa Nucleofector (Lonza). 5.0
106 HeLa cells were transfected with 10 μg of plasmid DNA according to the manufacturer’s instructions. Plasmids may be transfected using other commercially available transfection reagents. 8. Hydrolysis of the NHS ester occurs more easily in lessconcentrated protein solutions. Crosslinking may be carried

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out on adherent cells in culture plates as well as on cells in suspension. However, it is advantageous to use a small volume of crosslinker solution, because the crosslinking reaction is more effective in concentrated cell suspensions. In case of adherent cells, application of the crosslinker solution to all cell surfaces will be limited, and the efficiency of reaction will be decreased. 9. Amine components such as amino acids remaining in the media might quench the crosslinking reaction. 10. NHS ester crosslinker reacts with primary amines in physiological conditions of pH 7.2–9.0 to yield amide bonds. The NHS ester also undergoes hydrolysis under physiological conditions, competing with the primary amine reaction. Increasing the pH of the buffer results in an increase in the rate of hydrolysis. Thus, we used PBS with pH 8.0 in the crosslinking experiment. Optimize the final concentration of DSS between 1 and 5 mM. 11. Primary amine buffers such as Tris compete with the primary amine of lysine, thus avoiding its use as a buffer in the crosslinking reaction with NHS esters. Optimize the final concentration of quench solution between 10 and 20 mM. 12. TBS buffer is used to prevent unexpected crosslinking by the remaining crosslinker solution in the washing step. 13. Check for cell lysis under a microscope. The cell membrane should be completely lysed, while the nuclear membrane remains intact. 14. Nuclear lysates are treated with DNase and RNase to block DNA- or RNA-mediated interactions of histone and RNA-binding protein during the IP process. 15. Negative control sample or antibody should be used during IP. Normal rabbit-IgG is used as a control for anti-histone H3 antibody against nonspecific binding. Cells transfected with mock vector are used as a control for myc-tagged Rbfox2 and myc-tagged Rbfox3. 16. The type of magnetic bead used is determined by the immunoglobulin type and species of the primary antibody. Rabbit polyclonal IgG and mouse monoclonal IgG1 antibodies have strong affinity for protein A and G beads and protein G beads, respectively. Select Dynabeads™ magnetic beads depending on the antibodies used. Binding capacity of Dynabeads™ protein G is ~8 μg of IgG per mg of bead. Concentration of the commercially available Dynabeads™ protein G is 30 mg/ml. Accordingly, 2.5 mg of Dynabeads™ protein G are required to bind all 20 μg of IgG to the beads. Prepare ~85 μl of 30 mg/ml Dynabeads™ protein G per sample. Resuspend Dynabeads™ protein G beads in the vial. Transfer 85 μl of Dynabeads™

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protein G beads to a new microcentrifuge tube. Place the microcentrifuge tube on the magnet to separate the beads from the solution, and remove the supernatant. After removing the microcentrifuge tube from the magnet, add 200 μl of RIPA buffer to the microcentrifuge tube and gently resuspend the Dynabeads™ protein G beads using a pipette. 17. For single electrophoresis, pour ~5 ml of 10% separating gel into glass plates (thickness 1.0 mm). After solidification, fill up the 5% stacking gel to the limit of the glass plates on top of the separating gel, and place a gel comb in the stacking gel. Wait until the gel is solidified. The acrylamide percentage in SDS-PAGE gel depends on the size of the target proteins. Because the size of covalently crosslinked proteins is larger than that of non-crosslinked proteins, 4–12% acrylamide gels are recommended. 18. Immunoblotting is performed to confirm successful crosslinking. Use minimal immunoprecipitated samples in immunoblot analysis because a large amount of proteins is required to identify the crosslinked proteins using mass spectrometry. 19. Cut a nitrocellulose membrane and filter paper to the size of the gel and soak them in transfer buffer. Ensure that there are no air bubbles during the stacking of the gel, membrane, and filter paper. The membrane should be placed between the gel and positive electrode so that the negatively charged proteins migrate from the gel to the membrane. 20. Copper, zinc, and silver stains are compatible with Coomassie Blue stain for in-gel digestion and analysis of LC-MS/MS. 21. Crosslinked complexes are shown in a dotted box in Fig. 3. Place the gel pieces in a microcentrifuge tube containing water, and store it at 4  C. It is important to maintain the gel piece handling as clean as possible to avoid contamination with keratin and other proteins. Wear gloves and a lab coat during the entire preparation process. and use a fresh microcentrifuge tube from a sealing bag and clean apparatus for running the gels.

Acknowledgments This work was supported by Chungnam National University. References 1. Veres DV, Gyurko DM, Thaler B, Szalay KZ, Fazekas D, Korcsmaros T, Csermely P (2015) ComPPI: a cellular compartment-specific

database for protein-protein interaction network analysis. Nucleic Acids Res 43: D485–D493

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2. Akey CW, Luger K (2003) Histone chaperones and nucleosome assembly. Curr Opin Struct Biol 13(1):6–14 3. Hillebrand M, Gersting SW, Lotz-Havla AS, Schafer A, Rosewich H, Valerius O, Muntau AC, Gartner J (2012) Identification of a new fatty acid synthesis-transport machinery at the peroxisomal membrane. J Biol Chem 287 (1):210–221 4. Vaschetto LM (2017) Understanding the role of protein interaction motifs in transcriptional regulators: implications for crop improvement. Brief Funct Genomics 16(3):152–155 5. Watanabe N, Osada H (2016) Small molecules that target phosphorylation dependent protein-protein interaction. Bioorg Med Chem 24(15):3246–3254 6. Silberberg Y, Kupiec M, Sharan R (2014) A method for predicting protein-protein interaction types. PLoS One 9(3):e90904 7. Fields S, Song O (1989) A novel genetic system to detect protein-protein interactions. Nature 340(6230):245–246 8. Snider J, Kotlyar M, Saraon P, Yao Z, Jurisica I, Stagljar I (2015) Fundamentals of protein interaction network mapping. Mol Syst Biol 11(12):848 9. Xing S, Wallmeroth N, Berendzen KW, Grefen C (2016) Techniques for the analysis of protein-protein interactions in vivo. Plant Physiol 171(2):727–758 10. Kluger R, Alagic A (2004) Chemical crosslinking and protein-protein interactions-a

review with illustrative protocols. Bioorg Chem 32(6):451–472 11. Madler S, Bich C, Touboul D, Zenobi R (2009) Chemical cross-linking with NHS esters: a systematic study on amino acid reactivities. J Mass Spectrom 44(5):694–706 12. Suchanek M, Radzikowska A, Thiele C (2005) Photo-leucine and photo-methionine allow identification of protein-protein interactions in living cells. Nat Methods 2(4):261–267 13. Gaucher SP, Hadi MZ, Young MM (2006) Influence of crosslinker identity and position on gas-phase dissociation of Lys-Lys crosslinked peptides. J Am Soc Mass Spectrom 17 (3):395–405 14. Merkley ED, Rysavy S, Kahraman A, Hafen RP, Daggett V, Adkins JN (2014) Distance restraints from crosslinking mass spectrometry: mining a molecular dynamics simulation database to evaluate lysine-lysine distances. Protein Sci 23(6):747–759 15. Shi JM, Pei J, Liu EQ, Zhang L (2017) Bis (sulfosuccinimidyl) suberate (BS3) crosslinking analysis of the behavior of amyloid-beta peptide in solution and in phospholipid membranes. PLoS One 12(3):e0173871 16. Kim YE, Park C, Kim KE, Kim KK (2018) Histone and RNA-binding protein interaction creates crosstalk network for regulation of alternative splicing. Biochem Biophys Res Commun 499(1):30–36

Chapter 8 Proteomics-Based Systematic Identification of Nuclear Proteins Anchored to Chromatin via RNA Kyoko Hiragami-Hamada, Naoki Tani, and Jun-ichi Nakayama Abstract Chromatin serves as a platform for a multitude of biological processes, including transcription and co-transcriptional RNA processing. Consequently, chromatin is likely to be covered with many RNA molecules. Here we describe a simple, reliable, and cross-link-free method for the systematic identification of chromatin-associated RBPs that exhibit RNA-dependent chromatin association. Key words RNA, Protein, Chromatin

1

Introduction Recent studies of chromatin-associated RNAs (caRNAs) revealed that the majority of caRNAs are in fact the products of ongoing transcription, from both coding and non-coding genomic regions [1, 2]. The development of cross-linking and immunoprecipitation (CLIP)-mass spectrometry (MS) and various versions of the methodology has greatly accelerated the identification of RNA-bound proteins (RBPs) [3–7]. Currently, over 1000 proteins have been identified as RBP candidates in mouse and human cultured cells [8]. Nevertheless, it remains unclear how many of these RBPs actually associate with and function on chromatin. The CLIP-MSbased methods are powerful tools for the discovery of novel RBPs. However, they are generally laborious and expensive and do not directly address the chromatin-associated RBPs (caRBPs). Our method is basically an adaptation of Werner et al.’s protocol to isolate caRNAs [2]. By applying this method to HeLa S3 cells, we identified approximately 160 proteins as caRBP candidates. This method is highly applicable to different types of cells or organisms and thus will potentially enable the examination and exploration of caRNA-caRPB interactions in diverse biological contexts. Note

Ulf Andersson Vang Ørom (ed.), RNA-Chromatin Interactions: Methods and Protocols, Methods in Molecular Biology, vol. 2161, https://doi.org/10.1007/978-1-0716-0680-3_8, © Springer Science+Business Media, LLC, part of Springer Nature 2020

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that the following sections are mainly focused on the preparation of “RNase-solubilized” proteins and only briefly mention the sample preparation and the actual protein measurement by mass spectrometry. Our results obtained using the method described in this chapter are available as a preprint on the bioRxiv website [9].

2 2.1

Materials Buffer Solutions

Note: Use nuclease-free H2O and chemicals/reagents to prepare these buffers. 1. Buffer A: 10 mM Tris–HCl [pH 7.5], 10 mM KCl, 10% glycerol, 340 mM sucrose, 4 mM MgCl2, 1 mM dithiothreitol [DTT], 1
protease inhibitor cocktail. 2. Buffer NRB: 20 mM Tris–HCl [pH 7.5], 50% glycerol, 75 mM NaCl, 1 mM DTT, 1
protease inhibitory cocktail. 3. Buffer NUN: 20 mM Tris–HCl [pH 7.5], 300 mM NaCl, 1 M urea, 1% NP-40, 10 mM MgCl2, 1 mM DTT. 4. Buffer A for RNase III digestion (for RNase III treatment only): 10 mM Tris–HCl [pH 7.5], 10 mM KCl, 5% glycerol, 340 mM sucrose, 10 mM MgCl2, 1 mM dithiothreitol [DTT], 1
protease inhibitor cocktail. 5. RNase III storage buffer (for RNase III treatment only): 10 mM Tris–HCl [pH 8.0], 0.5 M NaCl, 0.5 mM EDTA, 1 mM DTT, 50% glycerol. 6. SDS loading buffer (4
–6
concentrate). 7. D-PBS (without Ca2+ and Mg2+). 8. SDS-PAGE running buffer. 9. Chromatography-grade H2O.

2.2 Enzymes and Reagents

1. RNase A (DNase-free, 10 mg/mL). 2. RNase III. 3. Benzonase. 4. RNase decontamination spray (e.g., Ambion/Sigma-Aldrich RNaseZap, Nacalai-Tesque RNase Quiet).

2.3

Cells

1. 2
106 to 1
107 cells of your choice (we tested the method with human HeLa S3 cells and K562 cells). Do not use frozen cell/nuclear pellets for the following procedures. Since the chromatin-associated protein and RNA levels vary among cell types, the minimal number of cells required to obtain sufficient amounts of “RNase-solubilized” proteins for the downstream applications (e.g., mass spectrometry, western blotting) must be empirically determined.

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2.4 Other Consumables

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1. Protein LoBind 1.5 mL tubes or siliconized 1.5 mL tubes. 2. Nuclease-free, filtered 10, 200, 1000 μL pipette tips. 3. Precast 10 cm
10 cm gradient gels (e.g., 4–20%). 4. Mass spectrometry-compatible CBB staining solution (e.g., GelCode Blue staining solution from Thermo Fisher). 5. Clean OHP transparency film sheets. 6. Sterile, disposable razor blades. 7. Clean plastic/glass dish designated for staining SDS-PAGE gels for MS sample preparation.

2.5 Laboratory Equipment

1. Heating/cooling block. 2. Temperature controlled centrifuge. 3. Overhead rotator. 4. Electrophoresis tank and power pack for SDS-PAGE. 5. Orbital shaker.

3

Methods The overall flow of the procedures is shown in Fig. 1. Please be aware of the following points before starting: 1. Once the experiment is started, it cannot be stopped until the end of Subheading 3.2, step 7 (approximately 2 h). 2. Clean the bench surface and pipettes, etc. with RNase decontaminating spray. 3. Handle all equipment and reagents with gloved hands.

3.1 Cell Fractionation (See Note 1)

1. Remove the culture medium.

3.1.1 For Adhesive Cells

3. Harvest cells with a cell scraper.

2. Wash cells 2
with 5 mL chilled D-PBS. Leave approx. 1 mL D-PBS in the culture dish. 4. Pipette up and down until no cell clumps are observed. 5. Collect the cells in a 15 mL tube and bring the volume to 5 mL with chilled D-PBS. 6. Centrifuge at 500
g for 5 min at 4  C. 7. Carefully but completely remove the D-PBS.

3.1.2 For Nonadhesive Cells

1. Harvest cells in a 15 mL tube. 2. Centrifuge at 500
g for 5 min at 4  C. 3. Remove the culture medium. 4. Resuspend cells in 5 mL chilled D-PBS.

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Harvest cells

~ 15 min

Cell fractionation

~ 1 hr

RNase treatment of chromatin faction SDS-PAGE & CBB staining of RNase-solubilized proteins

Preparation of gel pieces

~ 40 min

~ 2.5 - 3 hrs

~ 1 hrs

Dehydration, alkylation & ingel digestion of gel pieces

LC-MS/MS

Raw data processing & protein identification Quantification of identified proteins & further analysis

Fig. 1 The flowchart of the procedures

5. Centrifuge at 500
g for 5 min at 4  C. 6. Remove the D-PBS. 7. Repeat the D-PBS wash step once more. After the wash, carefully but completely remove the D-PBS from the cell pellet. 8. Add 2.5
cell pellet volume of Buffer A, and resuspend the cells well by pipetting up and down. 9. Transfer the cell suspension into a 1.5 mL Protein LoBind tube. 10. Add an equal volume of Buffer A containing 0.2% Triton X-100, and mix well by inverting the tube several times. 11. Incubate for 15 min on ice or for 12 min with slow rotation on an overhead rotator at 4  C. 12. Centrifuge at 1200
g for 5 min at 4  C. 13. Remove the supernatant.

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14. Wash by resuspending the resulting nuclear pellet in 1 mL Buffer A and centrifuging the mixture at 1200
g for 5 min at 4  C. 15. Remove the supernatant. 16. Resuspend the nuclear pellet in 250 μL Buffer NRB. 17. Centrifuge at 500
g for 5 min at 4  C. 18. Remove the supernatant. 19. Resuspend the nuclear pellet in 250 μL Buffer NRB. 20. Add 250 μL Buffer NUN to the nuclear suspension, and mix well by gently pipetting up and down several times. 21. Incubate the mixture on ice for 5 min. 22. Centrifuge at 1200
g for 5 min at 4  C. 23. Remove the supernatant. 24. Wash the “chromatin” pellet with 1 mL Buffer A. 25. Centrifuge at 1200
g for 5 min at 4  C. 26. Remove the supernatant. 27. Repeat the wash step at least once more. 28. Resuspend the “chromatin” pellet in 220 μL Buffer A (at room temperature). For RNase III digestion, resuspend in 220 μL Buffer A containing 5% glycerol and 10 mM MgCl2. In our experience, resuspension and RNase digestion in buffer without a sucrose/ glycerol cushion or in buffer containing [Na+/K+] ≧50 mM caused a marked release of proteins in the negative controls (i.e., RNase-untreated samples). Thus, we do not recommend the use of the reaction buffer provided with the RNase III by the manufacturer. 3.2 RNase A/III Treatment (See Note 2)

1. Aliquot 100 μL portions of the chromatin suspension into two 1.5 mL Protein LoBind tubes. 2. Dilute 10 mg/mL RNase A to 5 mg/mL with nuclease-free H2O immediately prior to the RNase A treatment. 3. Add 1 μL nuclease-free H2O to one of the 100 μL aliquots (untreated control) and 1 μL 5 mg/mL RNase A to the other (RNase A-treated sample). For the RNase III treatment, add 10–20 U RNase III to one of the 100 μL aliquots (RNase III-treated sample) and an equal volume of the RNase III storage buffer to the other (untreated control). Mix well by gently pipetting up and down several times. 4. Incubate at room temperature or at 25  C for 30 min for the RNase A treatment. Incubate at 30  C for 30 min for the RNase III treatment. Gently finger-vortex several times every 10 min during the incubation.

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5. Centrifuge at 1200
g for 5 min at 4  C. 6. Carefully transfer 75 μL of the supernatant containing the freely dissociated or RNase A-solubilized proteins to new 1.5 mL Protein LoBind tubes. 7. Add 15 μL 6
SDS sample buffer or an appropriate amount of SDS sample buffer to each sample from step 6. Mix well by vortexing. (The following steps are optional to analyze the chromatinbound proteins after RNase A treatment.) 8. Resuspend the remaining pellet in 1 mL Buffer A. 9. Centrifuge at 1200
g for 5 min at 4  C. 10. Remove the supernatant. 11. Repeat the wash step (steps 8–10) once more. 12. After the final wash step, resuspend the pellet in 100 μL Buffer A. 13. Add 20 μL 6
SDS sample buffer or an appropriate volume of SDS sample buffer. Mix well by vortexing. Note that the solution will be very viscous. For thorough mixing, it is best to vortex each sample immediately after the addition of the SDS sample buffer. 14. Add 2 μL Benzonase (25 U/μL) to the chromatin-bound fractions, and incubate at 37  C for 15 min on a shaking platform (set at 850 rpm). Make sure that the solution is no longer viscous after the Benzonase treatment. Otherwise, it will be very difficult to load the samples onto the gel. If the digestion is insufficient, add an additional 1–2 μL Benzonase and incubate at 37  C for 10 min. 3.3 Resolving RNase-Solubilized Proteins by SDS-PAGE

1. Assemble a precast gradient gel in an electrophoresis tank and fill the tank with SDS-PAGE running buffer. 2. Incubate the RNase A-solubilized protein samples and the control samples at 95  C for 5 min. 3. Load 5 μL of the molecular weight markers and 20–25 μL of the samples onto the precast gradient gel. Leave 1–2 wells between each sample/marker to avoid cross-contamination. Note: for western blotting, 5–10 μL sample/lane is normally sufficient, and there is no need to leave blank wells between samples/marker. As “positive controls” for western blotting, load 1–2 μL whole nuclear lysate samples and/or chromatin-bound fractions. 4. Run the gel at 100 V for 10 min and then at 200 V for 60 min. 5. Remove the gel from glass plates and rinse it with chromatography-grade H2O.

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kDa 250 150 100 75 50 37 25 20 15 10

Fig. 2 A CBB-stained gel image of the RNase-treated and control samples. The gray and black arrows indicate the position of RNase III and RNase A/T1, respectively

6. Wash the gel several times with chromatography-grade H2O (e.g., 3
10 min). 7. Stain the gel with a mass spectrometry-compatible CBB solution, according to the manufacturer’s instructions. 8. Perform 1–2
destaining steps with chromatography-grade H2O, according to the manufacturer’s instructions, until the background staining is sufficiently reduced. 9. Capture images of the stained gel. An example of a CBB-stained gel of RNase A/III-treated and control samples is shown in Fig. 2. 3.4 Sample Processing for Mass Spectrometry (See Note 3)

Preparation of gel pieces should be performed as recommended by the mass spectrometry facility of your institute or company. We generally place the destained gel on a clean OHP sheet; cut each lane of the stained 10 cm
10 cm gel into 13–16 gel pieces with sterile, disposable razor blades; mince the gel pieces further into 1–1.5 mm-sized cubes (for efficient in-gel trypsin digestion); and submit the samples to the mass spectrometry facility for further

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processing, including dehydration, alkylation, and in-gel trypsin digestion. For LC-MS/MS, we are currently using an Advanced UHPLC system (AMR/Michrom Biosciences) coupled to a Q Exactive mass spectrometer (Thermo Fisher Scientific). However, any other HPLC-coupled mass spectrometer used for general proteomics studies (e.g., one of the Thermo Fisher Orbitrap series) should be sufficient. 3.5 Protein Identification from LC-MS/MS Raw Data and Data Analysis (See Note 4)

The raw mass spectral data are generally processed with the software package accompanying the mass spectrometer. For example, we use Xcaliber for the raw mass spectral data processing and Proteome Discoverer with the Mascot search engine for the protein identification. For the false discovery rate (FDR) estimation and the evaluation of false positives, a decoy database composed of either randomized or reversed sequences in the target database and the Percolator algorithm can be used, respectively, to filter against the 1% global FDR for a high confidence level. The resulting datasets can be analyzed further, using the software of your choice. For example, we used Scaffold 4 for our analysis with the following cutoff values: minimum number of peptides ¼ 2, peptide threshold ¼ 95%, and protein threshold ¼ 1% FDR. The values from RNase-solubilized samples and untreated control samples from two to three independent experiments were divided into the “RNase-treated” and “untreated” groups, respectively. Quantification was performed using the total spectrum counts (TSC) or the Top3 precursor intensities (Top3) for each protein. Any proteins in the RNase-treated samples that showed 2-fold enrichment over the untreated samples and passed a statistical test (Fisher’s exact test in combination with the BenjaminiHochberg procedure for the TSC, or t-test for the Top3) were regarded as RNase-solubilized proteins or caRBP candidates. The cutoff values, quantification methods, and/or type of statistical tests can be altered. However, it must be noted that different quantification methods may result in different outcomes (e.g., only 70% overlap when the same datasets were quantified by the TSC or Top3 method). Therefore, we recommend testing several quantification methods and cutoff values to obtain the maximum number of highly reproducible caRBP candidates. After a list of caRBP candidates is obtained, the candidate proteins can be analyzed further, using software such as String, DAVID, Reactome, and so on, to investigate the enrichment of proteins involved in particular biological functions, molecular functions, and biological pathways and/or the enrichment for specific groups of protein domains among the candidates. Most of the software is available for free online.

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-

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RNase A

PurA hnRNP U Matrin-3 XRCC5/Ku80 DNA-PKcs

ILF3

HP1 TIF1

Fig. 3 An example of the mass spectrometry result validation by western blotting. The supernatants recovered after RNase A treatment were analyzed by western blotting. Asterisks indicate the proteins which were not identified as caRBP candidates by the mass spectrometry analysis 3.6 Evaluation of the Mass Spectrometry Results

4

The results from the mass spectrometry analysis should be validated by an alternative method, such as western blotting with reliable antibodies against the candidate proteins, at least for a subset of the caRBP candidates. An example of the validation by western blotting is shown in Fig. 3. Repeated, marked inconsistencies between the mass spectrometry results and the western blotting results (especially for those with high enrichment and statistical significance) may indicate that the cutoff values and/or quantification methods are not appropriate.

Notes 1. During the pilot experiments, the fractionation efficiency should be tested by western blotting analyses of each fraction, using antibodies against known cytoplasmic, nucleoplasmic, and chromatinproteins. We noticed that in the case of HeLa S3 cells, the Triton X-100 treatment released some abundant, soluble nuclear proteins into the cytoplasmic fraction during

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the cell lysis step. This is not a problem for our purpose, as we would like to remove the soluble nuclear proteins to obtain a relatively pure chromatin fraction. However, to fractionate intact nuclei, please refer to the paper by Sun and Fang [10]. In addition, some adhesive cell types may require a higher concentration of the detergent and/or a longer incubation time. The detachment of cells from a culture dish using an enzyme-free cell detachment reagent (e.g., Gibco Cell Dissociation Buffer) may facilitate homogeneous and efficient cell lysis. 2. If possible, titrate the amount of RNase A/III (e.g., 5, 50, 500 μg/mL RNase A) during the pilot experiments to determine the optimal concentration of RNase A/III for the cell type of interest. 3. For mass spectrometry, we recommend resolving the protein samples to some degree by SDS-PAGE, as less abundant proteins are likely to be “masked” by highly abundant proteins in the samples, such as ribosomal proteins and splicing factors. For cost reduction, run the samples to ~1/2 of the gel length and cut each lane into six to seven pieces. 4. Although we have used the Proteome Discoverer, Scaffold 4, and R software for our analyses, the MaxQuant and accompanying Perseus software are now more commonly used for the raw data processing, protein identification, quantitative analysis, and data visualization. These software programs are available for free online.

Acknowledgement This work was supported by the Joint Usage/Research Center for Developmental Medicine, IMEG, Kumamoto University. References 1. Mondal T, Rasmussen M, Pandey GK et al (2010) Characterization of the RNA content of chromatin. Genome Res 20:899–907 2. Werner MS, Ruthenburg AJ (2015) Nuclear fractionation reveals thousands of chromatintethered noncoding RNAs adjacent to active genes. Cell Rep 12:1089–1098 3. Castello A, Fischer B, Eichelbaum K et al (2012) Insights into RNA biology from an atlas of mammalian mRNA-binding proteins. Cell 149:1393–1406 4. Conrad T, Albrecht A-S, de Melo Costa VR et al (2016) Serial interactome capture of the human cell nucleus. Nat Commun 7:11212

5. Baltz AG, Munschauer M, Schwanh€ausser B et al (2012) The mRNA-bound proteome and its global occupancy profile on protein-coding transcripts. Mol Cell 46:674–690 6. He C, Sidoli S, Warneford-Thomson R et al (2016) High-resolution mapping of RNA-binding regions in the nuclear proteome of embryonic stem cells. Mol Cell 64:416–430 7. Bao X, Guo X, Yin M et al (2018) Capturing the interactome of newly transcribed RNA. Nat Methods 15:213–220 8. Hentze MW, Castello A, Schwarzl T, Preiss T (2018) A brave new world of RNA-binding proteins. Nat Rev Mol Cell Biol 19:327–341

Identifying Proteins Anchored to Chromatin via RNA 9. Hiragami-Hamada K, Tani N, Nakayama J (2018) Proteomic analysis of RNA-dependent chromatin association of nuclear proteins. bioRxiv. https://doi.org/10.1101/391755

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10. Sun L, Fang J (2016) Macromolecular crowding effect is critical for maintaining SIRT1’s nuclear localization in cancer cells. Cell Cycle 15(19):2647–2655

Useful Websites STRING (https://string-db.org/) DAVID (https://david.ncifcrf.gov) Reactome (https://reactome.org)

MaxQuant (https://www.maxquant.org/maxquant/) Perseus (https://www.maxquant.org/perseus/)

Chapter 9 2D Saturation Transfer Difference NMR for Determination of Protein Binding Sites on RNA Guanine Quadruplexes Ewan K. S. McRae, David E. Davidson, and Sean A. McKenna Abstract Saturation transfer difference (STD) NMR is a technique that provides information on the intermolecular interfaces of heterogenous complexes by cross-saturation from one molecule to the other. In this case, selective saturation of protein protons is applied, and the cross-relaxation to the RNA sample results in a reduction of the peak intensities in the measured H1–H1 NOESY spectrum. This allows for a relatively rapid and simple method of identifying the protein binding interface of an RNA with assigned chemical shift data. Key words Saturation transfer difference, NMR, RNA–protein interactions, Quadruplex, TERRA

1

Introduction Protein–nucleic acid interactions are fundamentally important for many cellular processes, yet our understanding of these complex systems is hampered by the difficulty of obtaining structural information on them. Often it is far simpler to solve the structure of an individual component of a protein–nucleic acid complex than the complex itself. Once this has been achieved, a frequently overlooked NMR methodology can be used to probe the binding interface of the complex. Identifying such contact surfaces can aid in analyzing mutagenesis experiments, docking simulations, design of disruptive small molecules for the binding interface and even provide a structural basis for the function of a complex. Saturation transfer difference (STD) NMR experiments were first developed as method to screen small molecule libraries for binding to proteins [1] and were quickly adapted to look at the protein epitope involved in protein–protein [2], protein–sugar [3, 4], and protein–nucleic acid [5, 6] binding. Similarly, though restricted by the limited number of assigned nucleic acid NMR structures, RNA epitopes have been investigated for small molecule interactions [7, 8] and more recently RNA–protein interactions

Ulf Andersson Vang Ørom (ed.), RNA-Chromatin Interactions: Methods and Protocols, Methods in Molecular Biology, vol. 2161, https://doi.org/10.1007/978-1-0716-0680-3_9, © Springer Science+Business Media, LLC, part of Springer Nature 2020

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[9, 10]. STD NMR offers multiple advantages compared to other methods, such as chemical shift perturbation, namely: 1. Low protein concentration requirements. 2. No labeling requirements. 3. No size limitations for protein of interest. 4. Simple data interpretation. 5. Signal intensity reflects interaction strength. In the STD experiment, the protein concentration is typically 1/100th that of the RNA concentration, such that at any given time the majority of RNA are unbound. The protein of interest is selectively saturated with a radio-frequency (RF) pulse at a frequency where the RNA does not absorb. During the saturation pulse, a bound RNA will receive saturation to its binding site via intermolecular H1–H1 cross-relaxation from the protein and dissociate, allowing another RNA molecule to bind (Fig. 1a). Over the course of the experiment, each protein molecule interacts with multiple RNA, resulting in a buildup of saturation at the protein binding site and reduced signal intensity when measured by Nuclear Overhauser Effect Spectroscopy (NOESY). This reduced signal spectrum is then subtracted from a reference spectrum, resulting in a difference spectrum that shows only the peaks that received saturation transfer (Fig. 1b)). The use of a 2D detection method allows for a relatively simple assignment of the peaks compared to the 1D method.

2

Materials Prepare all solutions using 0.22 μm filtered ultrapure water (MilliQ or equivalent to attain a resistivity of 18.2 MΩ cm (at 25  C). Latex or nitrile gloves should be worn when handling instruments and/or materials that will come into direct or indirect contact with RNA in order to prevent RNase contamination. Glassware used should be thoroughly cleaned, rinsed, and then subjected to a 245  C dry heat for 6 h to deactivate any contaminating RNases. All chemicals and reagents should be molecular biology grade and RNase-free and should be stored, handled, and disposed of according to the manufacturer’s instructions and institutional policy. Keep all buffers at room temperature unless otherwise mentioned.

2.1 Expression and Purification of RNA Binding Proteins

1. BL21(DE3) chemically competent E. coli strain (see Note 1). 2. PET vector encoding RNA binding protein of interest (see Note 2). 3. Sterile agar plates supplemented with antibiotic. 4. Sterile plastic spreader.

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Fig. 1 Schematic of the STD NMR method. (a) Protein (blue) and RNA (green) are in equilibrium, with multiple dissociation/association events occurring over the course of the experiment. RF pulse applies saturation (red) to the protein that is transferred through intermolecular cross-relaxation to the RNA. (b) Reference spectra are collected with no protein saturating RF pulse applied, and the saturated spectrum is then subtracted from the reference spectrum resulting in the STD spectrum, which only shows the peaks affected by the saturation transfer

5. Sterile LB broth supplemented with antibiotic. 6. Shaker incubator. 7. Baffled 2 L and 250 mL culture flasks. 8. Centrifuge capable of spinning 1 L and 50 mL tubes. 9. 1 M isopropyl β-D-1-thiogalactopyranoside (IPTG) dissolved in water. 10. Sonic Dismembrator. 11. Ni-NTA resin for gravity flow column or prepacked column. 12. Lysis buffer: 50 mM Tris–Cl pH 7.5 at 4  C, 150 mM NaCl, 500 mM KCl, 1 mM PMSF, 3 mM DTT (see Note 3). 13. Wash buffer 1: 50 mM Tris–Cl pH 7.5 at 4  C, 300 mM KCl, 2 mM imidazole. 14. Wash buffer 2: 50 mM Tris–Cl pH 7.5 at 4  C, 300 mM KCl, 20 mM imidazole. 15. Elution buffer: 50 mM Tris–Cl pH 7.5 at 4  C, 300 mM KCl, 20 mM imidazole. 16. Bradford reagent. 2.2 Preparation of RNA and Protein Samples for NMR

1. Sterile, RNase-free, filter-tip plastic pipettes. 2. Desalted, lyophilized RNA.

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3. 10 NMR buffer: Dependent upon experimental conditions of the assigned NMR spectrum. For TERRA 10mer RNA, 200 mM potassium phosphate buffer pH 7.0 at 25  C, 700 mM KCl. 4. Dialysis membrane/cassette. 5. Deuterium oxide (D2O). 6. Standard NMR tube. 7. Temperature controlled block heater. 2.3 Assignment of STD Peaks and Determination of STD Amplification Factor

3

1. Desktop computer with NMRPipe [11] and Sparky [12] and CCP4mg [13] installed (see Note 4). 2. Assigned spectra for the RNA of interest.

Methods

3.1 Expression and Purification of RNA Binding Proteins

The PET28b(+) vector containing the DDX21 C-terminal 209 amino acid (C209) cDNA with an N-terminal 6-His tag and thrombin cleavage site is available upon request or can be generated by PCR from commercially available cDNA libraries. The following protocol has been optimized for expression of C209 but is easily adapted to other RNA binding proteins of interest. 1. Chemically competent BL21(DE3) cells are transformed with the PET28b(+) vector according to manufacturer’s protocol, spread on an agar plate containing the appropriate antibiotic, and incubated overnight at 37  C. 2. A single, isolated, colony should be picked from the agar plate and used to inoculate 50 mL of autoclaved and antibiotic supplemented LB broth in a baffled 250 mL cell culture flask. Incubate the flask at 37  C in a shaker incubator at 200 rpm (see Note 5). 3. After approximately 12–16 h of growth, use 5 mL of starter culture to inoculate 500 mL of pre-warmed, antibiotic supplemented LB broth in a baffled 2 L flask. Incubate the flask at 37  C in a shaker incubator at 200 rpm. 4. Monitor the optical density at 600 nm with a spectrophotometer, and once it has reached 0.4, add 0.5 mL of 1 M IPTG to each flask (1 mM final IPTG concentration) (see Note 6). 5. Five hours after induction with IPTG, the cells should be collected by centrifugation at 4000  g for 10 min. 6. Resuspend the cells in 10 mL of ice-cold lysis buffer per 500 mL flask of cell culture and keep on ice.

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7. Sonicate the cell suspension on ice using 10-s pulses, at 50% max amplitude, with 30 s of rest between pulses for a total cumulative sonication time of 4 min (see Note 7). 8. Pellet the insoluble cell debris by centrifugation at 30,000  g for 45 min at 4  C. 9. During the centrifugation step, prepare 1 mL of fresh Ni-NTA resin in a gravity flow column by passing 5 mL of elution buffer through it, followed by 25 mL of lysis buffer. 10. Apply the clarified cell lysate to the Ni-NTA resin and allow it to flow through. 11. Pass 10 mL of Lysis buffer through the column. 12. Pass 10 mL of wash buffer 1 through the column. 13. Pass 10 mL of wash buffer 2 through the column. 14. Pass 10 mL of elution buffer through the column, collecting the eluent in a new tube. 15. Repeat step 14 until the eluent no longer reacts with Bradford reagent (see Note 8). 3.2 Preparation of RNA and Protein Samples for NMR

1. Thoroughly dialyze the protein sample into 1 NMR buffer (see Note 9). 2. Concentrate the protein sample until it is ten times the final concentration that will be used in the NMR experiment. The final concentration should be 1/100th that of the RNA, in this case 10 μM. 3. Dissolve the RNA to a concentration of 1.25 mM in 1.111 NMR buffer (see Note 10). 4. Heat the RNA in a block heater set to 95  C for 3 min, followed by a slow cool to room temperature (1  C/min). 5. Slowly add 100 μL of the 10 protein sample (or dialysis buffer for the RNA only sample) to 800 μL of the RNA (see Note 11). 6. Slowly add 100 μL of D2O to the protein–RNA mixture (see Note 12). 7. Filter the NMR sample through a 0.1 μm spin filter to remove aggregates.

3.3 Setup and Optimization of Initial NMR Parameters (TopSpin)

1. Insert the sample into the spectrometer and equilibrate to the desired experimental temperature. Tune, lock, and shim the NMR sample, and then acquire a conventional 1H NMR experiment (see Note 13). 2. Note the chemical shift of the solvent; ~4.78 ppm is expected in aqueous buffer. Set O1P to this value. 3. Create a new dataset, and retrieve parameters for an excitationsculpting solvent-suppression experiment by typing the rpar

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command. Select the ZGESGP parameter set [14] (see Note 13). 4. In the command line, use the GETPROSOL to set the default safe probe power levels. Calibrate the 1H 90 hard pulse for the sample using the pulsecal command, copying the power determined. Load the power levels for the experiment by using the GETPROSOL 1H command (see Note 14). 5. Set the shape to be used for the selective excitation pulse SPNAM1 to Squa100.1000. Set pulse length P12 to 2000 μs, set pulse power SPdB1 to a value 6 dB less than the pulse calculated in step 5 (this will be a 180 pulse), and set recycle delay D1 to 1.5 s. Ensure that gradient GPZ1 is set to 31% and GPZ2 is set to 11%. 6. Record a 1H 1D NMR experiment with solvent suppression. 7. Inspect the 1D NMR spectrum to determine the appropriate saturation frequency. Choose a frequency which is nearby the protein peaks to be saturated, but separated (at least 1.8 ppm) from other peaks; record this value. Choose an off-resonance saturation frequency distant from both protein and other peaks (often 30 ppm or 50 ppm is sufficient) (see Note 15). 8. Create a frequency list for saturation transfer experiments. In the command line, enter .freqlist, and then enter a name for the frequency list. Ensure “Don’t sort frequencies” is selected, and then click OK. Place the cursor on the protein peak to saturate and left-click the mouse. Then, place the cursor on the off-resonance frequency and left-click the mouse. Save the frequency list by entering .sret in the command line, and click “NO.” In the command line, enter FQLIST and edit the newly created list. Remove the first line of the list and save the list (see Note 16). 3.4 Setup of Saturation Transfer NMR Experiment (TopSpin)

1. Create a new dataset, and retrieve parameters for a saturation transfer difference (STD) experiment by typing the rpar command. Select one of the STDDIFF parameter sets from the list such as STDDIFFESGP.3 [1, 14, 15] (see Note 17). 2. In the command line, use the GETPROSOL to set the default safe probe power levels. Set the relaxation delay D1 to 1.5 s, and ensure the solvent suppression pulse P40 is set to 2000 μs. Duplicate settings for offset O1 (both values) and power level SPdB1 from values used in Subheading 3.3, step 5. 3. In this sequence, set PLW10 and set shaped pulse SPNAM10 to values used previously for PLW1 and SPNAM1 in Subheading 3.3, step 5. Set spinlock time D29 to 10 ms to suppress T2 relaxation while allowing T1ρ to occur.

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4. Set the saturation pulse shape SPNAM9 to Gaussian1.1000 and the saturation pulse length P42 to 50,000 μs. In the command line, enter FQ2LIST and set the desired frequency list saved in Subheading 3.3, step 8. Set the saturation time D20 as 0.5 s. 5. On the parameter screen (ased), make a note of the computed value of SPW9; this will need to be entered in the 2D NOESY STD experiment (see Note 17). 6. Ensure that GPZ1 is 40%, GPZ2 is 31%, and GPZ3 is 11%. 7. In the command line, enter “rga” to adjust the receiver gain. 8. Record the 1D saturation transfer NMR experiment with solvent suppression. 3.5 Setup of 2D NOESY Saturation Transfer NMR Experiment (TopSpin)

1. Create a new dataset, and set up a 2D NOESY saturation transfer difference (STD) experiment by typing the rpar command, and select the STDNOESYESGPPH parameter set [1, 14–17]. 2. In the command line, use the GETPROSOL to set the default safe probe power levels. From the saturation transfer NMR experiment (Subheading 3.4), enter values for D1, P40, O1, SPdB1, PLW10, SPNAM10, D20, D29, SPNAM9, P42, and FQ2LIST. Enter the computed value of SPW9 recorded in Subheading 3.4, step 5 (see Note 17). 3. Set 1 FnMODE to States-TPPI, set TD to 2048, and set 1 TD to 256. This corresponds to a 2D experimental array with 2048 points in the direct observation dimension and 256 points in the indirect dimension. 4. Ensure that SW in F1 matches the value of SW in F2. 5. Ensure that GPZ1 is 40%, GPZ2 is 40%, GPZ3 is 31%, and GPZ4 is 11%. 6. Set NOE mixing time D8 to 150 ms.

3.6 Setup and Optimization of Initial NMR Parameters (VnmrJ BioPack)

1. Insert the sample into the spectrometer and equilibrate to the desired experimental temperature. Tune, lock, and shim the NMR sample, and then acquire a conventional 1H NMR experiment (see Note 12). 2. Note the chemical shift of the solvent; ~4.63 ppm is expected in aqueous buffer. Set the carrier to this value in Acquire(tab)/ Channels/Offset(ppm). 3. Create a new experiment, and retrieve parameters for a WATERGATE solvent-suppression experiment in Experiments (menu)/Setup BioPack Experiment . . .Water Suppression Experiments/1D/3919 Watergate. 4. Determine an optimized 90 pulse. In Acquire(tab)/Channels/ Observe/90 Degree Pulse Width, note the initial value and vary

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the pulse width around the initial value (e.g., try 1 μS < initial pw < +1 μs in 0.1 μs increments). Run the array and choose a value which provides highest peak intensity away apart from the solvent peak. 5. Check the optimized 90 pulse. In Acquire(tab)/Channels/ Observe/90 Pulse Width, set the best value from Subheading 3.6, step 4, as well as two times pw (180 null), three times pw (270 negative peak), and four times pw (360 null) to verify the correct pw setting. Record the pw value as well as the power value. Set pw and pw90 to this value. 6. Optimize solvent suppression in Acquire(tab)/WATERGATE/ Selective pulse and select the checkbox for Automatic. Then click the button for Optimize Parameters. Once complete, record and save the 1D water-suppressed NMR spectrum. 7. Choose frequencies for Saturation and Reference. Place the cursor on the protein peak to saturate and note the chemical shift value in PPM. Then, to set the frequency scale in Hz, set Display(tab)/Axis/Hertz(button). Left-click the spectrum in the same location to read off the “cursor” position in the window in Hertz, and note the saturation frequency. Repeat this process for the off-resonance reference frequency. 3.7 Setup Saturation Transfer NMR Experiment (VnmrJ BioPack)

1. Create and join a new NMR experiment in Experiments (menu)/Setup BioPack Experiment . . .Water Suppression Experiments/DPFGSE 1D/Saturation Transfer 1D-3 [4, 14, 18] (see Note 18). 2. Set up the pulse power for the experiment. In Acquire(tab)/ Channels/Observe/90 Pulse Width, set the carrier at 4.63 ppm and set the best pw value and power from Subheading 3.6, step 5. 3. Set up the acquisition parameters for the experiment. In Acquire(tab)/Acquisition/Excitation, set relaxation delay d1 to 1.5 s, and observe pulse pw90 to the calibrated pw value from Subheading 3.6, step 5. Set Scans Requested to nt 256 and set steady-state ss to 8. Set Receiver Gain to Auto. 4. Set up water suppression. In Acquire(tab)/Pulse Sequence, ensure that Double PFG spin echo is selected, and then click the button to Recreate water refocusing shape. 5. Set up saturation transfer. Under Saturation, set transfer delay satdly to 2 s, and choose a saturation frequency based upon the data in the water-suppression spectrum collected in Subheading 3.6, as well as a reference frequency (see Note 19). 6. Once complete, record and save the 1D saturation transfer NMR spectrum.

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1. Create and join a new NMR experiment, and then select Experiments(menu)/Setup BioPack Experiment . . .Water Suppression Experiments/DPFGSE 2D/Saturation Transfer Noesy. 2. Set up a 1D experiment in Acquire(tab)/Basic by clicking the button 1D. 3. Set up the pulse power for the experiment. In Acquire(tab)/ Channels/Observe/90 Pulse Width, set the carrier at 4.63 ppm and set the best pw value and power from Subheading 3.6, step 5. 4. Set up water suppression. In Acquire(tab)/Pulse Sequence, ensure that Double PFG spin echo is selected, and then click the button to Recreate water refocusing shape. Select Flipback option, and then click the button to Recreate flipback shape. 5. Set up saturation transfer. Under Saturation, set transfer delay satdly to 2 s, and choose a saturation frequency from the watersuppression spectrum collected in Subheading 3.6 (see Note 19). 6. Set up NOESY mixing. Under NOESY mixing time, set 0.150 s. 7. In Acquire(tab)/Acquisition/Acquisition in F2, set Scans Requested nt to 16, Receiver Gain to Auto, and run a test 1D spectrum. 8. Set up 2D experimental parameters. In Acquire(tab)/Basic, click the F1F2 2D button. In Acquire(tab)/Acquisition/Acquisition in F1, the Acquisition Mode should have automatically been set to Hypercomplex 2D. Set spectral width to match the spectral width in F2, and set increments in t1 to 128. 9. Record and save the 2D NOESY saturation transfer NMR spectrum.

3.9 Assignment of STD Peaks and Determination of STD Amplification Factor

1. Convert the STD NMR spectrum processed in NMRPipe [11] to UCSF format using the pipe2ucsf program that comes with the Sparky distribution [12]. 2. To pick peaks, switch to peak picking mode in Sparky [12] using F8 and select the area to pick peaks in with the cursor. 3. When finished picking peaks, assign them by cross-referencing to the previously assigned dataset. Switch back the cursor mode to select (F1) and open the assignment window with the shortcut “at.” Manually assign the picked peaks by selecting individual peaks and entering the assignment. 4. When finished assigning the peaks, select all peaks with the shortcut “pa.” 5. Integrate the selected peaks with the shortcut “pi.”

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6. The peak list can be viewed using the shortcut “lt” and then saved as a “.list” or “.txt” file. 7. The same processing should now be applied to the reference NMR spectrum. 8. To calculate the STD amplification factor, take the ratio of the S/N of each peak in the STD spectrum to that of the S/N for the same peak in the reference spectrum (STD/Ref), and divide this ratio by the ratio of the peak with the maximal value. 9. For each peak with the same w1 assignment, the normalized ratio is averaged to obtain a representative STD value for that proton. 10. Finally this averaged, normalized ratio is multiplied by the RNA/protein excess, in this case 100. 11. A convenient way of representing the STD amplification factor on the 3D structure of the RNA, we recommend editing the “. pdb” file of the structure and changing the B-factor value to the STD amplification value for each proton. 12. Using CCP4mg13 to open the modified .pdb file, one can then colorize the protons by B-factor to achieve a pictorial representation of the protons that received the most saturation transfer from the protein.

4

Notes 1. We use low background strain (LoBStr) BL21(DE3) cells from Kerafast, which seems to provide a clean enough purification from only Ni-NTA affinity chromatography. Any BL21(DE3) strain should be fine, but may require further polishing steps to attain a pure sample (i.e., size exclusion chromatography). 2. For C209 we use PET28b(+) and utilize the N-terminal hexaHis tag and thrombin cleavage site and include a stop codon in the primers used to generate the insert to prevent tagging of the C-terminus. This works well for C209, but other RNA binding proteins may work better with a C-terminal tag. 3. It is important to pH the buffer at 4  C, since Tris has a significant variance of pH at different temperatures. It is also important to add the PMSF and DTT immediately prior to use, as they have short half-lives in aqueous solution. We recommend preparing 100 mM stocks of PMSF in 100% ethanol to allow remain in solution upon dilution into the lysis buffer. 4. For operation on a Windows PC, we recommend running the latest version of NMRPipe_CentOS11 virtual machine using VMware Workstation Player. It is then possible to install Sparky on this virtual machine. CCP4mg is used for visualization of

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the STD amplification factors on the structure. All the abovelisted software is open source for noncommercial use and can be found at the following links: https://www.vmware.com/ca/products/workstation-player/ workstation-player-evaluation.html https://www.ibbr.umd.edu/nmrpipe/install.html https://www.cgl.ucsf.edu/home/sparky/ http://www.ccp4.ac.uk/MG/ 5. We recommend to pre-warm LB broth and add antibiotics immediately prior to induction. 6. For C209 a rapid induction with 1 mM IPTG works best, and for other larger RNA binding proteins, we have found that lower IPTG concentrations (as low as 0.001 mM) and lower incubation temperatures (as low as 15  C) result in greater yield of soluble protein. 7. These are the settings for a Fisher Scientific Model 500 Sonic Dismembrator. If the cell suspension is warming during sonication, you may need to reduce the intensity of your sonicator or sonicate for less time. 8. We use 100 μL of Bradford in a flat 96 well dish and add 10uL of eluent to check for the presence of protein. 9. We recommend dialyzing in 200 times the volume of sample (i.e., 10 mL sample in 2 L of dialysis buffer), twice for 1 h and then one more time overnight. 10. For purchased RNA it is recommended to spin down the sample briefly to ensure it is all at the bottom of the tube and then rely on the reported mass of RNA provided by the manufacturer to determine the correct volume of buffer to dissolve in. This is the most accurate and consistent way, unless the hypochromicity of the folded RNA has been accurately determined, and then one could use an extinction coefficient to determine concentration. 11. The importance of adding both the protein and the D2O SLOWLY to the RNA cannot be overstated. Add a couple μL at a time and stir with the pipette tip in between additions of protein or D2O to the RNA sample. 12. This guide assumes the user is familiar with the basic operation of the NMR spectrometer, including how to safely load samples, lock, shim, and collect simple 1D datasets, Fourier transform, and phase data. 13. If using a pulse sequence containing a solvent-suppression scheme, the solvent suppression will need to be optimized prior to performing the STD experiment. A detailed practical guide to solvent suppression by excitation sculpting is provided in the Bruker TopSpin manual’s library, in the “Acquisition—

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User Guides” section of the manual titled “1D and 2D Stepby-Step—Advanced” in Sections 7.2 (pp. 146–150) and 7.6 (pp. 157–158). These experimental parameters will need to be copied to the STDDIFFESGP.3 experiment for effective solvent suppression. 14. The pulse duration and power determined by the PULSECAL command will need to be appended to the GETPROSOL command (e.g., if the duration of the pulse was determined to be 8.5 μS at power 1.0 dB, use GETPROSOL 1H 8.5–1.0). 15. It may be necessary to conduct 1H 1D experiments on separate NMR samples of ligand/buffer and protein/buffer to determine which peaks are associated with each molecule. 16. The first line of the frequency list defines the 1H resonance of the list and will not be required. There should be two remaining entries in the list: the desired on-resonance frequency, followed by a new line containing the desired off-resonance frequency. 17. The value of SPW9 is computed by the STDDIFFESGP.3 pulse sequence but is not computed by the NOESY experiment and must be manually entered in the 2D NOESY STD experiment. This value should be non-zero. 18. BioPack contains multiple versions of the saturation transfer experiment, some of which provide automatic difference spectra (Saturation Transfer 1D-2) and others which do not and must be collected and processed to determine difference spectra manually after the completion of the experiment (Saturation Transfer 1D-3). We recommend manual processing of individual spectra. The appearance of a variable named reference frequency depends on sequence. 19. If using the Saturation Transfer 1D-3 experiment, the saturation and reference frequencies will need to be collected and saved as separate experiments. Remember to repeat the experiment for each frequency.

Acknowledgments Funding is provided by an NSERC CGS-D scholarship (E.M), Cancer Research Society (Canada) [20085], and Canadian Cancer Society Research Institute [703809]. References 1. Mayer M, Meyer B (1999) Characterization of ligand binding by saturation transfer difference NMR spectroscopy. Angew Chem Int Ed Engl 38:1784–1788

2. Takahashi H, Nakanishi T, Kami K, Arata Y, Shimada I (2000) A novel NMR method for determining the interfaces of large proteinprotein complexes. Nat Struct Biol 7:220–223

2D Saturation Transfer Difference NMR for Determination of Protein Binding. . . 3. Haselhorst T, Weimar T, Peters T (2001) Molecular recognition of sialyl Lewis and related saccharides by two lectins. J Am Chem Soc 123:10705–10714 4. Mayer M, Meyer B (2001) Group epitope mapping by saturation transfer difference NMR to identify segments of a ligand in direct contact with a protein receptor. J Am Chem Soc 123:6108–6117 5. Ramos A, Kelly G, Hollingworth D, Pastore A, Frenkiel T (2000) Mapping the interfaces of protein-nucleic acid complexes using crosssaturation. J Am Chem Soc 122:11311–11314 6. Lane AN, Kelly G, Ramos A, Frenkiel TA (2001) Determining binding sites in proteinnucleic acid complexes by cross-saturation. J Biomol NMR 21:127–139 7. Mayer M, James TL (2002) Detecting ligand binding to a small RNA target via saturation transfer difference NMR experiments in D2O and H2O. J Am Chem Soc 124:13376–13377 8. Mayer M, James TL (2004) NMR-based characterization of phenothiazines as a RNA binding scaffoldt. J Am Chem Soc 126:4453–4460 9. Harris KA, Shekhtman A, Agris PF (2013) Specific RNA-protein interactions detected with saturation transfer difference NMR. RNA Biol 10:1307–1311 10. McRae EKS, Davidson DE, Dupas SJ, McKenna SA (2018) Insights into the RNA quadruplex binding specificity of DDX21. Biochim Biophys Acta Gen Subj 1862:1973–1979

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11. Delaglio F, Grzesiek S, Vuister GW, Zhu G, Pfeifer J (1995) NMRPipe: a multidimensional spectral processing system. J Biomol NMR 6:277–293 12. Lee W, Tonelli M, Markley JL (2015) NMRFAM-SPARKY: enhanced software for biomolecular NMR spectroscopy. Bioinformatics 31:1325–1327 13. McNicholas S, Potterton E, Wilson KS, Noble MEM (2011) Presenting your structures: the CCP4mg molecular-graphics software. Acta Crystallogr D Biol Crystallogr 67:386–394 14. Hwang TL, Shaka AJ (1995) Water suppression that works. Excitation sculpting using abritrary waveforms and pulsed field gradients. J Magn Reson Ser A 112:275–279 15. Mayer M, Meyer B (1999) Charakterisierung von Ligandenbindung durch S€attigungstransfer-Differenz-NMR- Spektroskopie. Angew Chemie 111:1902–1906 16. Jeener J, Meier BH, Bachmann P, Ernst RR (1979) Investigation of exchange processes by two-dimensional NMR spectroscopy. J Chem Phys 71:4546–4553 17. Wagner R, Berger S (1996) Gradient-selected NOESY—A fourfold reduction of the measurement time for the NOESY experiment. J Magn Reson Ser A 123:119–121 18. Dalvit C (1998) Efficient multiple-solvent suppression for the study of the interactions of organic solvents with biomolecules. J Biomol NMR 11:437–444

Chapter 10 Mapping Transcriptome-Wide and Genome-Wide RNA–DNA Contacts with Chromatin-Associated RNA Sequencing (ChAR-seq) Charles Limouse, David Jukam, Owen K. Smith, Kelsey A. Fryer, and Aaron F. Straight Abstract RNAs play key roles in the cell as molecular intermediates for protein synthesis and as regulators of nuclear processes such as splicing, posttranscriptional regulation, or chromatin remodeling. Various classes of non-coding RNAs, including long non-coding RNAs (lncRNAs), can bind chromatin either directly or via interaction with chromatin binding proteins. It has been proposed that lncRNAs regulate cell-statespecific genes by coordinating the locus-dependent activity of chromatin-modifying complexes. Yet, the vast majority of lncRNAs have unknown functions, and we know little about the specific loci they regulate. A key step toward understanding chromatin regulation by RNAs is to map the genomic loci with which every nuclear RNA interacts and, reciprocally, to identify all RNAs that target a given locus. Our ability to generate such data has been limited, until recently, by the lack of methods to probe the genomic localization of more than a few RNAs at a time. Here, we describe a protocol for ChAR-seq, an RNA–DNA proximity ligation method that maps the binding loci for thousands of RNAs at once and without the need for specific RNA or DNA probe sequences. The ChAR-seq approach generates chimeric RNA–DNA molecules in situ and then converts those chimeras to DNA for next-generation sequencing. Using ChAR-seq we detect many types of chromatin-associated RNA, both coding and non-coding. Understanding the RNA–DNA interactome and its changes during differentiation or disease with ChAR-seq will likely provide key insights into chromatin and RNA biology. Key words Chromatin, Proximity ligation, RNA, ChAR-seq, Transcriptome, Non-coding RNA, Next-generation sequencing

1

Introduction Much of the mammalian genome is transcribed, yet we do not understand the biological functions of most of the RNAs encoded by the genome [1]. Many RNAs interact with chromatin at the

Electronic supplementary material: The online version of this chapter (https://doi.org/10.1007/978-1-07160680-3_10) contains supplementary material, which is available to authorized users. Ulf Andersson Vang Ørom (ed.), RNA-Chromatin Interactions: Methods and Protocols, Methods in Molecular Biology, vol. 2161, https://doi.org/10.1007/978-1-0716-0680-3_10, © Springer Science+Business Media, LLC, part of Springer Nature 2020

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source of transcription or via association with nucleoprotein complexes that can bind far from the locus of transcription [2]. Some well-studied non-coding RNAs (ncRNAs) have revealed that these interactions may modulate gene expression and chromatin state. For example, XIST is a long intergenic non-coding-RNA (lincRNA) which coats one of the X chromosomes in female mammals to repress X-linked genes and establish dosage compensation [3]. The lincRNA HOTAIR has been proposed to serve as an RNA scaffold to coordinate the recruitment of two distinct histonemodifying complexes to target genes [4]. Alpha-satellite RNAs may recruit histone methyltransferases at pericentromeres to stabilize heterochromatin structure at this locus [5–8]. Yet, at the genome-wide level, we do not know what RNAs are associated with what genomic loci or the functional consequences of these associations. A broad open question remains: how do chromatinassociated RNAs regulate histone modifications, chromatin folding, and transcription [2, 9]? Multiple methods have been developed to map the interactions between individual RNAs and chromatin. These methods include Chromatin Isolation by RNA Purification (ChIRP-seq) [10], Capture Hybridization Analysis of RNA Targets (CHART-seq) [11], and RNA antisense purification sequencing (RAP-seq) [12]. These methods rely on the isolation of a specific RNA target in a setting that preserves RNA–DNA–protein contacts to identify the genomic target loci of this RNA. Yet, despite their power, these methods all require a priori knowledge of the RNA target. Thus, they cannot be used for the de novo identification of RNAs that may modulate a given chromatin locus. Importantly, there are thousands of uncharacterized lncRNAs in the human genome, and transcription can occur outside of traditional gene regions, for instance, at enhancers [13–15] or at transposable elements [16]. Therefore, a multiplexed and unbiased approach to identify chromatin-associated RNAs and their binding sites will accelerate our understanding of chromatin regulation by RNAs. Recently, several methods have been designed to systematically capture and sequence all RNA-DNA contacts in the cell: ChAR-seq [17, 18], GRID-seq [19], MARGI [20], and RADICL-seq [21]. Here we describe a protocol for ChAR-seq [17], a method that uses in situ proximity ligation between RNA and DNA to identify chromatin-associated RNAs and map their target DNA loci (Fig. 1a). The method can be thought of as a massively parallel RNA–DNA interactome sequencing assay. ChAR-seq enables the generation of chromatin localization maps for thousands of RNAs, without a priori knowledge of their sequence. A central component of the method is the use of a short 24 bp linker dsDNA molecule, henceforth referred to as the “bridge,” which contains three key features (Fig. 1b). (1) One end of the bridge molecule contains a 50 -adenylated ssDNA overhang, allowing specific ligation to the 30

ChAR-Seq 0. Crosslink cells, isolate & permeabilize nuclei (steps 3.2, 3.3) RNA

1. Fragment RNA, dephosphorylate 3’ ends (steps 3.4, 3.5)

Protein

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7. Reverse crosslink, isolate DNA, and sheer (steps 3.13 - 3.15)

+ Bridge + T4KQ RNA ligase

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6. 2nd Strand Synthesis (step 3.12)

+ T4 DNA Ligase

+ DpnII

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Fig. 1 (a) ChAR-seq library preparation: overview of protocol steps and corresponding steps in Subheading 3. Steps 0–9 are performed in situ. In brief: (0) cells are fixed and nuclei are isolated and swollen in hypotonic buffer. (1) RNAs are fragmented to generate free 30 ends by exposure to heat and magnesium, and 30 ends are dephosphorylated using T4 PNK making them compatible for ligation in step 2. (2) ChAR-seq bridge is added to the nuclei and ligated to the 30 ends of fixed RNAs using the T4 RNA ligase 2 truncated R55K K227Q. (3) First strand synthesis is performed with Bst3.0 DNA polymerase using the “bottom strand” bridge of the bridge as a primer and the RNA as a template, to convert bridge-ligated RNAs to RNA–DNA hybrids. (4) Chromatin and bridge are digested with DpnII. (5) Genomic DNA is ligated to RNA-bridge molecules using DpnII overhangs and T4 DNA ligase. (6) Second strand synthesis is performed with E. coli DNA Pol I to convert RNA-bridge-DNA hybrids into cDNA-bridge-DNA chimeras. (7) Cross-links are reversed, and DNA is ethanol precipitated and sheared to produce fragments of ~200 bp. (8) Bridge-containing molecules are isolated by biotin pulldown. (9) Sequencing adapters are ligated to the streptavidin bead-bound fragments, and these fragments are amplified by PCR. (10) Size selection is performed to remove primer dimers and molecules that are too long for Illumina sequencing, and the library is sequenced. (b) Schematic of the ChAR-seq bridge sequence showing the major features of the bridge. The 50 adenylation allows the bridge to be ligated to the 30 end of RNAs using the mutated T4KQ ligase. The DpnII site is cleaved during DpnII digestion, and the overhang is subsequently ligated to the genomic DpnII overhangs. The 30 C3 spacer prevents nonspecific ligation of bridge molecules that were not DpnII digested. A biotin moiety serves as a molecular handle for isolation of cDNA-bridge-DNA chimeras. The unique molecular identifier (UMI) allows one to distinguish PCR duplicates from true biological duplicates. The PacI site is used after Illumina adapter ligation to remove molecules for which the bridge was not DpnII digested. The full bridge sequence up to the DpnII site (AANNNAAACCGGCGTCCAAGGATC) can be identified in the sequencing reads and marks the cDNA–DNA junction DpnII

Adenylation UMI

PacI

3’ C3 spacer

5’-App-AANNNAAACCGGCGTCCAAGGATCTTTAATTAAGTCGCAG-3’-Sp3 TTGGCCGCAGGTTCCTAGAAATTAATTCAGCGTCTAG-5’ Biotin

Fig. 1 (continued)

end of RNAs using a mutated T4 RNA ligase (T4 RNA ligase 2, truncated, R55K K227Q) [22]. In addition, the complementary strand serves as a primer for reverse transcription which allows, after second strand synthesis, the conversion of the ssRNA-bridge hybrid

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to a cDNA-bridge chimeric DNA. (2) The other end of the bridge contains a DpnII restriction site, which allows its ligation to the DpnII-digested genome and formation of a cDNA-bridge-DNA chimera. (3) Finally, the bridge contains a biotin molecule, which allows for selective isolation of cDNA-bridge-DNA molecules for next-generation sequencing. The bridge has a defined non-palindromic sequence, not present in the human genome, that can be identified in each sequencing read to delineate the junction between the cDNA and the DNA. Due to the RNA-to-ssDNA ligation, the method preserves the orientation of the original RNA and thus enables stranded alignment of RNA with the genome. In ChAR-seq, the biochemistry steps to generate cDNAbridge-DNA chimeric molecules are performed in situ, in fixed nuclei. Thus, the native chromosome architecture and spatial proximity between RNA and DNA are preserved. The steps of the ChAR-seq protocol are detailed in the methods section and outlined in Fig. 1. From a sequenced ChAR-seq library, one obtains the interaction frequency between each chromatin-associated RNA and each genomic locus. With this dataset in hand, one can ask questions such as: what RNAs bind a locus of interest? Reciprocally, with what genomic loci does an RNA of interest interact? Furthermore, the chromatin-association pattern of an RNA can be correlated with chromatin features, such as histone modifications and presence of specific chromatin binding proteins. ChAR-seq also makes it possible to investigate the relationship between the binding of specific RNAs and local chromatin folding into topological domains and promoter-enhancer interactions. This type of analysis should allow one to make predictions on the potential function of an unknown ncRNA and potentially discover new ncRNAs involved in controlling gene expression or chromatin state. In this chapter, we describe the ChAR-seq protocol for human cells. We use the RPE-1 cell line as an example for the cell culture and harvesting parts of the protocol, but the protocol is general and the RPE-1 line can be substituted for another human cell line of choice. The same protocol can likely be applied without any major modification to any mammalian cell type or even to cells from other species or isolated from tissue samples. We have successfully applied this protocol to map the RNA-DNA interactome in human cell lines (RPE-1, K562, THP-1, and H9 hESCs), Xenopus laevis A6 cells, and Drosophila melanogaster cell lines (cl.8+ and Kc167).

2

Materials

2.1 Stable Buffers and Solutions

The buffers and solutions listed below can be stored at room temperature or 4  C and reused for multiple ChAR-seq library preparations. All solutions must be RNase-free. Water and PBS are used in large quantities throughout the protocol. We thus

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prepare these solutions in-house and use diethyl pyrocarbonate (DEPC) and autoclaving to inactivate nucleases. Alternatively, RNase-free water and PBS can be purchased from commercial suppliers. Refer to the method section for practical recommendations on how to handle RNase-free solutions. We suggest that these solutions are used exclusively for ChAR-seq and stored separately to avoid mishandling and RNase contamination. 1. Diethyl pyrocarbonate (DEPC). 2. DEPC-treated water: add 1 mL of fresh DEPC to 1 L of ddH2O (sterile filtered or molecular biology grade) for a final concentration of 0.1% DEPC. Allow DEPC to inactivate nucleases by incubating at room temperature 2 hrs or overnight. Inactivate DEPC by autoclaving for 15 min. 3. DEPC-treated 1 PBS, pH 7.4. Mix in a 1 L beaker 800 mL ddH2O, 8 g NaCl, 200 g KCl, 1.44 g Na2HPO4·2H2O, and 0.24 g KH2PO4. Adjust pH to 7.4. Add ddH2O to 1 L. Filter sterilize. Then add 1 mL of fresh DEPC for a final concentration of 0.1% DEPC. Allow DEPC to inactive nucleases by incubating at room temperature overnight. Inactive DEPC by autoclaving for 15 min. 4. 1 M Tris–HCl pH 8, RNase-free. 5. 1 M MgCl2, RNase-free. 6. 0.5 M EDTA, RNase-free. 7. 10% SDS (w/v), RNase-free. 8. 5 M NaCl, RNase-free. 9. 2 M KCl, RNase-free. 10. 3 M sodium acetate, pH 5.5, nuclease-free. 11. 1 M (NH4)2SO4, prepared with DEPC-treated water. 12. cDNA buffer: 10 mM Tris-Cl, pH 7.6–8.0, 90 mM KCl, 50 mM (NH4)2SO4. Prepare using the RNase-free stock solutions. Store up to 6 months at 4  C. 13. RNase decontamination solution (Thermo Fisher Scientific RNaseZAP or equivalent). 2.2 Cell Culture and Harvesting

1. Cultured cells of cell type of interest (here RPE-1). 2. Cell culture media. Here for RPE-1, DMEM:F12. 3. Heat inactivated fetal bovine serum (FBS). 4. 10,000 (μg/mL) streptomycin, 10,000 IU/mL penicillin mixture (100 solution to add to cell culture medium). 5. 16% (w/v) methanol-free formaldehyde in ampules. 6. 2.5 M glycine. 7. 15 cm round petri dishes.

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8. 50 mL conical tubes. 9. Hemocytometer. 10. Centrifuge with swinging-bucket rotor and adapters for 50 mL tubes. 11. 1.5 mL nuclease-free microcentrifuge tubes. 12. Benchtop centrifuge. 13. Liquid nitrogen. 14. Rocking platform shaker. 2.3 Preparation of ChAR-seq Bridge

1. ChAR-seq bridge “top strand”: order HPLC purified and lyophilized ssDNA oligonucleotide modified with a 50 adenylation (/5rApp/) and a 3’ C3 spacer (/3SpC3/) and with the following sequence:

/5rApp/AANNNAAACCGGCGTCCAAGGATCTTTAATTAAGTCGCAG/3SpC3/ where NNN are random nucleotides that serve as a unique molecular identifier (UMI). 2. ChAR-seq bridge “bottom strand”: order HPLC purified and lyophilized ssDNA modified with an internal biotin group (/ iBiodT/) and with the following sequence: /5Phos/GATCTGCGACTTAATTAAAGATCCTTGGACGCCGG/iBiodT/T. 3. Bridge annealing buffer (BAB): 20 mM Tris–HCl, pH 8.0, 1 mM EDTA, 100 mM NaCl. Prepare using RNase-free 1 M Tris–HCl, 0.5 M EDTA, 5 M NaCl, and DEPC-treated water. 4. TE buffer: 10 mM Tris–HCl pH 8.0, 1 mM EDTA. Prepare using RNase-free 1 M Tris–HCl, 0.5 M EDTA, and DEPCtreated water. 5. 1.5 mL tubes, 94  C heat block, or thermocycler. 2.4 In Situ Generation and Isolation of RNA(cDNA)-Bridge-DNA Hybrid Molecules

In addition to the reagents and buffers in Subheading 2.3. 1. Protease inhibitor cocktail, make a 10 protease inhibitor (PI) stock. 2. 1 M dithiothreitol (DTT). Store at 20  C. 3. 10 mM Tris–Cl, pH 8.0. Prepare using DEPC-treated water and RNase-free 1 M Tris–HCl. 4. 40 U/μL RNase inhibitor. 5. 10 T4 RNA ligase buffer. 6. 10 polynucleotide kinase buffer. 7. T4 polynucleotide kinase (PNK).

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8. 200 U/μL T4 RNA ligase 2, truncated KQ (T4 Rnl2tr R55K, K227Q). 9. Polyethylene glycol (PEG 8000). 10. 50 μM dsDNA ChAR-seq bridge (see Subheading 2.1). 11. 120 U/μL Bst3.0 polymerase. 12. dNTP mix, diluted to 10 mM (each nucleotide) in DEPCtreated water. 13. 50 U/μL DpnII restriction enzyme. 14. T4 DNA ligase buffer. 15. 400 U/μL T4 DNA ligase. 16. 10 U/μL Escherichia coli DNA polymerase I. 17. 5 U/μL RNase H. 18. 20 mg/mL Proteinase K. 19. Bench centrifuge with fixed angle rotor. 20. Bench centrifuge with swinging bucket rotor. 21. Thermomixer with shaking ability for 1.5 mL microcentrifuge tubes. 2.5 Preparation of Library for Deep Sequencing

1. 100% ethanol (molecular biology grade). 2. 70% (v/v) ethanol: prepare fresh using 100% ethanol and DEPC-treated water. 3. 5 mg/mL glycogen. 4. Ultrasonicator. 5. Appropriate tubes for the ultrasonicator in use. 6. Magnetic streptavidin T1 beads. 7. Magnetic rack for multiple (6–12) 1.5–2.0 mL tubes. 8. Tween wash buffer (TWB): 5 mM Tris–Cl, pH 7.6–8.0, 0.5 mM EDTA, 1 M NaCl, 0.05% Tween 20. Prepare fresh. 9. 2 bead binding buffer (BBB): 10 mM Tris–Cl, pH 7.6–8.0, 1 mM EDTA, 2 M NaCl. 10. NEBNext Ultra II DNA Library Prep Kit for Illumina (NEB, cat. no. E7645S) including: (a) End Prep Reaction Buffer. (b) End Prep Enzyme Mix. (c) Ligation Enhancer. (d) Ultra II Ligation Master Mix. (e) NEBNext Ultra II Q5 Master Mix. 11. NEBNext Ultra II Q5 Master Mix (NEB, cat. no. M0544S; this is required in addition to amount included in kit E7645S).

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12. 10 μM Universal Primer NEBNext Multiplex Oligos for Illumina (Set 1) (kit: NEB, cat. no. E7335S). 13. 10 μM Indexing Primer; NEBNext Multiplex Oligos for Illumina (Set 1) (kit: NEB, cat. no. E7335S; note that there are multiple indexing primers available in sets of 12) including: (a) NEBNext Adaptor. (b) NEB USER Enzyme. 14. CutSmart Buffer (NEB, cat. no. B7204S). 15. 10 U/μL PacI restriction enzyme. 16. PCR thermocycler. 17. 200 μL PCR tubes or strips. 18. 1.5–2.0 mL nuclease-free, low-bind microcentrifuge tubes. 19. AMPure XP SPRI (solid-phase reversible immobilization) beads (5 mL; Beckman Coulter, cat. no. A63880). 20. 100 Sybr Green (10,000 concentrate, diluted 1:100). 21. Agilent Bioanalyzer automated electrophoresis system or equivalent. 22. Real-time PCR machine for qPCR. 23. KAPA Library Quantification Kit for Illumina platforms (Kapa Biosystems, cat.no. KK4854).

3

Methods It is critical that all procedures are performed in an RNase-free environment. To avoid RNase contamination, wipe the bench and all the handled equipment (pipettes, sharpies, racks, timers, centrifuges, etc.) with RNase decontamination solution (RNaseZAP or equivalent), and rinse with DEPC-treated water before starting the protocol. Lightly spray gloves with RNase decontamination solution when starting the preparation and every time a non-RNasefree surface is touched. However, one should be cautious not to transfer RNase decontamination solution into tubes, reagents, or onto the skin. Gloves should be dried with a clean wipe after using the solution. Use RNase-free reagents exclusively, as listed in Subheading 2. Throughout the protocol until reverse cross-linking (Subheading 3.13), the H2O is always DEPC-treated ddH2O, and the PBS is DEPC-treated PBS. Carry out all spins at 2500  g at room temperature unless otherwise specified. For every centrifugation step, we recommend spinning the samples for 90 s in a swinging bucket rotor and then transferring the tubes to a fixed rotor centrifuge to spin for an additional 90 s (see Note 1).

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For all reaction mixes, volumes indicated in parenthesis are for one sample. If multiple samples are processed in parallel, prepare each reaction mix as a master mix by multiplying the indicated volumes by the number of samples. Refer to the “ChAR-seq buffer and reaction mixes worksheet” (Supplementary Table 1) for a list with the composition of all the reaction mixes (and for an example protocol for six samples). For all the reaction mixes, water and buffer should be added first unless otherwise noted. Generating ChAR-seq libraries ready for sequencing takes ~4–5 days. Table 1 shows a timeline of the protocol. If multiple samples are pooled for sequencing, a qPCR should be performed with a PhiX standard curve or the KAPA Library Quantification Kit to accurately determine the concentration of each individual library and the proper volume ratios for pooling. Allocate an extra ~½ day for this quantification. Stable buffers (see Subheading 2.1) and dsDNA ChAR-seq bridge (Subheading 3.1) should be prepared ahead of time (day 0 or before). 3.1 Preparation of dsDNA ChAR-seq Bridge

dsDNA ChAR-seq bridge will be used in Subheading 3.6, but we recommend preparing it ahead of time. 1. Resuspend lyophilized “top bridge strand” in the appropriate volume of TE to obtain a 200 μM stock. Repeat the same procedure with “bottom bridge strand.” Store both tubes at 20  C or proceed to step 2. 2. Anneal the top and bottom strands to make dsDNA ChAR-seq bridge: in a 1.5 mL microcentrifuge tube, mix 2 BAB, 200 μM bridge top strand, and 200 μM bridge bottom strand in a volume ratio of 2:1:1 to make a stock solution of dsDNA ChAR-seq bridge at 50 μM in 1 BAB. Place the tube in a 94  C heatblock and incubate for 3 min. Leave the tube in the block, and turn off the heat to allow the tube to slowly cool to room temperature (see Note 2). Do not place the tube on ice until room temperature has been reached. 3. Optional: verify that the bridge strands have fully annealed by running a small aliquot on a 12% polyacrylamide gel. In separate lanes, run a similar amount of the top and bottom strands as a control. 4. Store the 50 μM dsDNA bridge at 20  C until use.

3.2 Harvesting of Cells and Formaldehyde Fixation

1. Grow cells in appropriate culture media. A ChAR-seq library can be prepared using ~10 million cells (see Note 3). Here, for RPE-1, grow cells in a 15 cm plate to ~80% confluency in DMEM-F12 supplemented with 10% FBS and 1x streptomycin/penicillin mixture. 2. Trypsinization: remove media from the plate and add 10 mL PBS to wash the cells. Remove PBS and add 5 mL trypsin-

DNA ligation 2nd strand synthesis Reverse crosslinking 2nd strand synthesis Reverse crosslinking

2h O/N

2h 6.5 h O/ N

1st strand synthesis DpnII digestion

1st strand synthesis DpnII digestion DNA ligation

Option 2 Same as option 1

Option 3 Same as option 1

1.5 h O/ N

4.5 h 1.5 h O/ N

2h 1.5 h 2h 0.75 h 1.5 h 2h 1.5 h

3h 1h 1.5 h 2–4 h

Same as day 3 option 1

Same as day 3 option 1

Side-qPCRb Off-bead PCR Size selectionb Bioanalyzer, qPCRb Send for sequencing

Day 4

Same as day 4 option 1

Same as day 4 option 1

Day 5

The three options correspond to whether the genomic digestion and DNA-bridge ligation are done overnight (see Note 10). For each step, the indicated duration is approximate and includes the enzymatic step and preceding pellet wash steps. Allocate more time if more than four samples are prepared at once. O/N overnight a Cells can be harvested and fixed at the beginning of day 1 or ahead of time and stored as frozen pellets at 80  C b Indicates a possible stopping point in the protocol, where the DNA can be safely stored at 20  C

Option 1 Cell fixationa Bench cleanup Cell lysis RNA fragmentation Dephosphorylation RNA-bridge ligation DNA precipitation Shearingb Bioanalyzerb Biotin isolation Adapters ligation PacI digestion On-bead PCR, SPRIb

Day3 2h 6.5 h 4.5 h 1.5 h O/N

3h 1h 1.5 h 0.5 h 1h O/N

Day2 1st strand synthesis DpnII digestion DNA ligation 2nd strand synthesis Reverse crosslinking

Day 1

Table 1 Timeline of the ChAR-seq protocol

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EDTA, and then incubate at 37  C for a few minutes to let the cells detach off the plate. Quench the trypsin with 15 mL media. 3. Fixation: transfer the cells to a 50 mL Falcon tube and spin at ~500  g for 5 min. Remove media and resuspend the cell pellet in 16.25 mL serum-free media. Add 3.75 mL of fresh 16% formaldehyde (3% final formaldehyde concentration). Incubate for 10 min at room temperature on a rocking platform. 4. Quench the formaldehyde by adding 6.25 mL of 2.5 M glycine (0.6 M final concentration). Incubate for 5 min at room temperature on a rocking platform, and then incubate for an additional 15 min on ice with intermittent mixing. 5. Centrifuge the cells for 5 min at ~500  g at 4  C, remove the supernatant, and wash the cells by resuspending the pellet in 10 mL ice-cold DEPC-treated PBS. 6. Take a 10 μL aliquot of cells for counting using a hemocytometer. Aliquot may need to be diluted 1:2 to 1:10 for accurate counting. Using the hemocytometer count, estimate the total amount of cells in the Falcon tube. 7. Centrifuge the cells again at ~500  g at 4  C, discard the supernatant, resuspend the pellet in 1 mL DEPC-treated PBS, and transfer the cells to a 1.5 mL microcentrifuge tube. If the number of cells (as estimated in step 6) is far from 10 million cells, split or pool cells into the appropriate number of 1.5 mL microcentrifuge tubes to obtain ~10 million cells per tube. 8. Centrifuge the cells for 5 min at ~500  g at 4  C. Discard the supernatant and flash freeze the cells in liquid nitrogen or proceed immediately to Subheading 3.3. Flash frozen cells can be stored at 80  C for several months, until use for Subheading 3.3, step 2. 3.3 Cell Lysis and Nuclei Preparation

1. Before taking the frozen cells out of the 80  C, prepare 500 μL of fresh cell lysis buffer with protease inhibitors and RNaseOUT: 10 mM Tris–HCl pH 8

(5 μL of 1 M)

10 mM NaCl

(1 μL of 5 M)

0.2% Igepal-CA630

(5 μL of 20% stock)

1 mM DTT

(0.5 μL of 1 M)

1 U/μL RNaseOUT

(12.5 μL of 40 U/μL)

1 protease inhibitor

(50 μL of 10 stock)

H2O

(426 μL)

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Also prepare 500 μL of the same buffer without Igepal detergent, protease inhibitors, and RNaseOUT for the upcoming wash step (replace detergent, RNaseOUT, and protease inhibitor cocktail volumes with DEPC-treated water). 2. Gently resuspend the cell pellet until the suspension is homogenous in 500 μL ice-cold cell lysis buffer containing detergent, protease inhibitors, and RNaseOUT. If starting with a frozen pellet, add the ice-cold lysis buffer directly to the frozen pellet, and thaw the pellet by pipetting up and down until it is fully resuspended. Place the sample immediately on ice (see Note 4). 3. Incubate the sample on ice for 15 min to complete the cell lysis. 4. During incubation, prepare the 0.5% SDS buffer described and used at Subheading 3.3, step 6. Store at room temperature until use (do not put on ice as SDS may precipitate). 5. Wash: centrifuge the sample to pellet the nuclei using the two-step centrifugation as described above, pipette, and discard the supernatant. Resuspend the pellet in 500 μL lysis buffer without Igepal, RNaseOUT, or protease inhibitor (see Note 5). Centrifuge the sample, and discard the supernatant. 6. SDS treatment: the goal of this step is to loosen the nuclear mesh and increase accessibility of small molecules for the subsequent enzymatic reactions. Resuspend the nuclei in 400 μL 0.5% SDS buffer, prepared ahead of time at Subheading 3.3, step 4: 10 mM Tris–HCl pH 8

(4 μL of 1 M)

10 mM NaCl

(0.8 μL of 5 M)

1 mM DTT

(0.4 μL of 1 M)

0.5% SDS

(20 μL of 10%)

1 U/μL RNaseOUT

(10 μL of 40 U/μL)

H2O

(364.8 μL)

Incubate for 10 min at 37  C. 7. SDS quenching: allow the sample to cool for 2 min to room temperature. Do not place on ice. Quench the SDS by adding 60 μL of 10% Triton X-100 (1.3% final concentration), and mix well by pipetting. Incubate for 15 min at 37  C. 8. During the incubation, prepare the fragmentation buffer (described and used at Subheading 3.4, step 3) and the dephosphorylation mix (described and used at Subheading 3.5, step 3), and store them on ice until use.

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At Subheading 3.6, RNAs will be ligated at their 30 ends to the adenylated 50 end of the ChAR-seq bridge. The goal of RNA fragmentation is to generate more RNA 30 ends available for ligation. RNA fragmentation is achieved by heating up the sample at 70  C in the presence of magnesium (2.5 mM). Fragmentating RNA also increases the library complexity because multiple distinct RNA-bridge junctions can be generated for a given RNA. In addition, fragmentation is important for reducing the representation of 30 poly(A) tails in the RNA reads. 1. Wash the nuclei to remove the detergents: add 800 μL PBS, centrifuge the sample, and discard the supernatant. At this step, it is normal for the appearance and consistency of the pellet to change (see Note 6). 2. Wash the nuclei a second time: resuspend the pellet in 800 μL 1 RNA ligase buffer. Centrifuge the sample, and discard the supernatant (see Note 7). 3. RNA fragmentation: resuspend the pellet in 150 μL fragmentation buffer, prepared ahead of time at Subheading 3.3, step 8: 0.25 T4 DNA ligase buffer

(3.75 μL of 10)

1 U/μL final RNaseOUT

(3.75 μL of 40 U/μL)

H2O

(142.5 μL)

Incubate for 4 min at 70  C. Then immediately place the samples on ice to stop the fragmentation reaction. Fragmentation time may need to be optimized for each cell type (see Note 8). 3.5 RNA 30 Dephosphorylation

1. Wash the nuclei to remove the RNA fragments that are not cross-linked: add 800 μL PBS to the sample, pipette up and down, centrifuge the sample, and discard the supernatant. 2. Wash the nuclei a second time: resuspend the pellet in 800 μL 1 RNA ligase buffer. Centrifuge the sample, and discard the supernatant. 3. Dephosphorylation: resuspend the pellet in 150 μL dephosphorylation mix prepared ahead of time at Subheading 3.3, step 8: 1 T4 PNK buffer

(15 μL of 10)

1 U/μL T4 PNK

(7.5 μL of 20 U/μL)

1 U/μL RNaseOUT

(3.75 μL of 40 U/μL)

H2O

(123.15 μL)

Incubate for 30 min at 37  C with agitation.

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4. During the incubation, prepare the RNA-bridge ligation mix described and used at Subheading 3.6, step 3, and store on ice until use. 3.6 RNA-Bridge Ligation

1. Wash the nuclei once to wash out the PNK: add 800 μL PBS, pipette up and down, centrifuge the sample and discard the supernatant. 2. Wash the nuclei a second time: resuspend the pellet in 800 μL 1 RNA ligase buffer. Centrifuge the sample, and discard the supernatant. 3. RNA-bridge ligation: resuspend the nuclei in 200 μL RNA-bridge ligation mix prepared ahead of time at Subheading 3.5, step 4: 1 T4 ligase buffer

(20 μL of 10)

20% PEG-8000

(80 μL of 50% PEG, see Note 9)

1.5 U/μL RNaseOUT

(7.5 μL of 40 U/μL)

2.5 μM annealed bridge

(10 μL at 50 μM)

10 U/μL T4KQ ligase

(10 μL of 200 U/μL)

H2O

(72.5 μL)

Incubate overnight in a thermomixer at 23  C with shaking at 900 rpm. 3.7 First Strand Synthesis

1. Before starting the wash steps below, prepare 250 μL of first strand synthesis mix, described and used at Subheading 3.7, step 4. 2. Wash the nuclei to remove the unbound bridge molecules, enzymes, and PEG: add 800 μL PBS, pipette up and down, centrifuge the sample, and discard the supernatant. 3. Wash the nuclei a second time: resuspend the pellets in 800 μL 1 RNA ligase buffer. Centrifuge the sample, discard the supernatant. 4. First strand synthesis: resuspend the pellet in 250 μL first strand synthesis mix prepared ahead of time at Subheading 3.7, step 1: 1 T4 RNA ligase buffer

(25 μL of 10)

1 mM DTT

(0.25 μL 1 M)

1 U/μL RNaseOUT

(6.25 μL 40 U/μL)

1 mM (each) dNTP mix

(25 μL of 10 mM)

8 U/μL Bst3.0

(16.7 μL of 120 U/μL)

H2O

(176.8 μL)

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Incubate: (a) 15 min at 23  C. (b) 10 min at 37  C. (c) 20 min at 50  C. 3.8 Denature and Wash out Bst3.0

1. Add 8 μL of 0.5 M EDTA (15 mM final) and 14 μL of 10% SDS (0.5% final) to the sample. Incubate for 10 min at 37  C to inactivate Bst3.0. 2. Add 43 μL of 10% Triton X-100 (for 1.3% final concentration) to quench the SDS, and mix well. Incubate for 15 min at 37  C. 3. During the Triton X-100 quench step, prepare the DpnII digestion mix, described and used at Subheading 3.9, step 1. Store the mix on ice until use. 4. Wash the nuclei to remove Bst3.0 and dNTPs: add 800 μL PBS, pipette up and down, centrifuge the sample, and discard the supernatant. 5. Wash the nuclei a second time: resuspend the pellet in 800 μL 1 RNA ligase. Centrifuge the sample, and discard the supernatant.

3.9 Genomic Digestion

1. Resuspend the pellet in 250 μL DpnII digestion mix, prepared ahead of time at Subheading 3.8, step 3:

1 T4 RNA ligase buffer

(25 μL of 10)

1 mM DTT

(0.25 μL of 1 M)

1 U/μL RNaseOUT

(6.25 μL of 40 U/μL)

3 U/μL DpnII

(15 μL of 50 U/μL)

H2O

(203.5 μL)

Incubate for 6 h at 37  C (see Note 10). 3.10 Denature and Wash Out DpnII

1. Add 8 μL of 0.5 M EDTA (15 mM final) and 14 μL of 10% SDS (0.5% final) to the sample. Incubate for 10 min at 37  C to inactivate DpnII. 2. Add 43 μL of 10% Triton X-100 (for 1.3% final concentration) to quench the SDS, and mix well. Incubate for 15 min at 37  C. 3. During the Triton X-100 quench step, prepare the bridgeDNA ligation mix described and used at Subheading 3.11, step 1. Store the mix on ice until use.

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4. Wash the nuclei to remove DpnII: add 800 μL PBS, pipette up and down, centrifuge the sample, and discard the supernatant. 5. Wash the nuclei a second time: resuspend the pellet in 800 μL 1 T4 RNA ligase buffer. Centrifuge the sample, and discard the supernatant. 3.11 Bridge-DNA Ligation

1. Resuspend the pellet in 250 μL bridge-DNA ligation mix, prepared ahead of time at Subheading 3.10, step 3:

1 T4 DNA ligase buffer

(25 μL of 10)

1 U/μL RNaseOUT

(6.25 μL of 40 U/μL)

10 U/μL T4 DNA ligase

(6.25 μL of 400 U/μL)

H2O

(212.5 μL)

Incubate for 4 h at 23  C (see Note 11). 3.12 Second Strand Synthesis

1. Before proceeding, prepare the second strand synthesis mix described and used at Subheading 3.12, step 5. Save the mix on ice until use. 2. T4 ligase inactivation: add 8 μL 0.5 M EDTA (15 mM final concentration) to the sample to quench the Mg2+. Incubate for 2 min at room temperature. 3. Wash the nuclei to remove the T4 ligase and buffer: add 800 μL PBS, pipette up and down, centrifuge the sample, and discard the supernatant. 4. Wash the nuclei a second time: resuspend the pellet in 800 μL 1 cDNA buffer (see Subheading 2.1, step 12, Supplementary Table 1). Centrifuge the sample, and discard the supernatant. 5. Resuspend the pellet in 250 μL second strand synthesis mix prepared ahead of time at Subheading 3.12, step 1: 10 mM Tris–HCl pH 8

(2.5 μL of 1 M)

90 mM KCl

(22.5 μL of 1 M)

10 mM MgCl2

(2.5 μL of 1 M)

50 mM (NH4)2SO4

(12.5 μL of 1 M)

1 mM DTT

(1.25 μL of 1 M)

1 mM dNTPs (each)

(25 μL of 10 mM)

0.025 U/μL RNaseH

(1.25 μL of 5 U/μL)

0.5 U/μL E. coli DNA PolI

(12.5 μL of 10 U/μL)

H2O

(164.25 μL)

Incubate for 1.5 h at 37  C.

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Add to each sample:

3.13 Reverse Cross-Linking

1% SDS

(31.25 μL of 10% SDS)

0.5 M NaCl

(31.25 μL of 5 M)

0.6 mg/mL Proteinase K

(9 μL of 20 mg/mL).

Incubate overnight at 68  C. 1. Add 31.25 μL 3 M sodium acetate to the sample and mix; then add 930 μL 200 proof ethanol. A flocculent DNA is typically visible at this step after inverting the tubes a few times. If no flocculent is observed, add 2 μL 5 mg/mL glycogen as a carrier for DNA precipitation. Incubate for 30 min at 20  C.

3.14 DNA Precipitation

2. Centrifuge the sample for 20 min at 20,000  g at 4  C. 3. Discard the supernatant and wash the pellet by adding 1 mL 70% ice-cold ethanol (freshly prepared) to the tube and briefly vortexing. Centrifuge the sample for 10 min at 20,000  g, and discard the supernatant. 4. Repeat step 3 to further wash the DNA (see Note 12). 5. Let the pellet air-dry for 5 min, and then resuspend it in 132 μL TE buffer. 6. Save 2 μL to assay the fragment length distribution prior to shearing if desired (Fig. 2). The remaining 130 μL will be sheared. At this stage, DNA can be stored frozen at 20  C, if necessary, until ready to proceed. 3.15

1. Shear the purified DNA samples (containing the desired cDNA-bridge-DNA chimeras as well as genomic DNA) using a Covaris S220 acoustic shearer or equivalent system to obtain a peak fragment size of 200 bp. For the Covaris S220, we found that the following parameters reliably provide a 200 bp peak fragment size: 175 peak incident power, 10% duty factor, and 200 cycles/burst, for 180 s (see Note 13).

Shearing

2. Proceed to the next step or save the sheared DNA at 20  C until ready to proceed.

A)

B)

C)

[FU]

[FU] 300

[FU]

300

100

200

200 100 0

50

100

0

0 35

150

300

500

10380

[bp]

35

150

300

500

10380

[bp]

35

150

300

500

10380

[bp]

Fig. 2 Example of bioanalyzer traces quantifying the fragment length distribution of the library. (a) Before shearing, aliquot saved at Subheading 3.14, (b) after Covaris shearing at Subheading 3.15, and (c) after final PCR amplification and size selection at Subheading 3.27

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3.16 Assessment of Fragment Length Distribution

3.17 Isolation of Biotinylated Molecules

Use 0.5 μL of the sheared sample and dilute it in 2.5 μL TE buffer. Measure the DNA concentration of this aliquot by Qubit dsDNA HS assay. Further dilute the aliquot if necessary to obtain a DNA concentration of ~1–5 ng/μL, and measure the fragment size distribution using an Agilent High Sensitivity DNA bioanalyzer. Figure 2 shows an example of a bioanalyzer trace for the same sample pre-shearing (from aliquot saved at Subheading 3.14) and post-shearing. This trace indicates shearing to the desired fragment size distribution, with a peak fragment size around ~200 bp. When starting with ~10 million cells, expect a post-shearing DNA concentration around 500 ng/μL. Most of this DNA is genomic DNA. The goal of the next step is to isolate and amplify bridge-containing molecules. 1. Streptavidin bead wash: for each sample, pipette 150 μL MyOne Streptavidin T1 dynabeads in a 1.5 mL microcentrifuge tube. Place the tubes on a magnetic rack and wait for ~1–2 min for the beads to collect on the side of the tubes, and then discard the supernatant. Remove the tubes from the magnetic rack and wash the beads by resuspending them in 400 μL TWB (see Subheading 2.5, item 8 or Supplementary Table 1). 2. Binding of biotinylated molecules to the beads: collect the washed beads using a magnetic rack. Discard the supernatant, remove the tubes from the magnetic rack, and resuspend the beads in 130 μL 2 BBB (see Subheading 2.5, item 9 or Supplementary Table 1). Add 130 μL of sheared DNA and mix well. Incubate for 15 min at room temperature with agitation (see Note 14). 3. Wash out unbound molecules: collect the beads using a magnetic rack, remove the supernatant, and resuspend the beads in 750 μL TWB. Incubate for 2 min at 50  C without agitation. The supernatant should only contain genomic DNA but can be saved for troubleshooting. 4. Wash the beads a second time: collect the beads, discard the supernatant, and resuspend them in 750 μL TWB. 5. Collect the beads, discard the supernatant, and resuspend the beads in 40 μL TE buffer.

3.18 On-Bead Sequencing Adaptor Ligation

This step uses the NEBNext library preparation kit, but other standard library preparation kits can be substituted if desired. 1. End repair and dA tailing: add. (a) 7 μL NEBNext End Prep Buffer. (b) 3 μL NEBNext End Prep Enzyme Mix.

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Incubate the sample for 20 min at room temperature with agitation and then 30 min at 65  C with agitation and then cool to room temperature. 2. Adaptor ligation: add the following components to each sample, in the order indicated and mix vigorously by pipetting: (a) 2.5 μL NEBNext Adaptor. (b) 1 μL Ligation Enhancer. (c) 30 μL NEBNext UltraII Ligation Master Mix. (d) Incubate for 15–20 min at room temperature with agitation. (e) Add 3 μL NEB USER Enzyme and incubate for 15 min at 37  C with agitation. 3. During incubation, prepare ahead of time the PacI digestion mix described and used at Subheading 3.19, step 1 (if PacI digestion performed). 4. Wash the beads to remove extra adapters and enzymes: add 750 μL TWB to the sample, and then incubate for 2 min at 50  C without agitation. 5. Wash the beads a second time: collect the beads using the magnetic rack, discard the supernatant, and resuspend the beads in 750 μL TWB. 6. Collect the beads, discard the supernatant, and proceed immediately to PacI digestion if desired or directly to on-bead PCR. 3.19 (Optional) PacI Digestion (See Note 15)

1. Resuspend the beads in 100 μL PacI digestion mix prepared ahead of time at Subheading 3.18, step 3: 1 CutSmart buffer

(10 μL 10)

0.4 U/μL PacI

(4 μL of 10 U/μL)

H2O

(86 μL)

Incubate for 30 min at room temperature with agitation. 2. Wash the beads to remove PacI: add 750 μL TWB to the sample, and pipette up and down. 3. Wash the beads a second time: collect the beads using the magnetic rack, discard the supernatant, and resuspend the beads in 750 μL TWB. 4. Collect the beads, discard the supernatant, and proceed immediately to on-bead PCR. 3.20 Library First Amplification (On-Bead PCR)

1. Resuspend the beads in 50 μL PCR amplification mix: (a) 25 μL 2 NEBNext High-Fidelity Master Mix.

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(b) 2.5 μL 10 μM universal primer. (c) 2.5 μL 10 μM indexing primer. (d) 20 μL H2O. Transfer the sample to a PCR tube. 2. Amplify the library in a thermocycler using the following program (see Note 16): (a) 1 cycle: 98  C for 30 s (initial denaturation). (b) 5–7 cycles: 98  C for 10 s (denaturation), 65  C for 75 s (annealing and extension). 3. Collect the beads using the magnetic rack and transfer the supernatant to a clean 1.5 mL microcentrifuge tube. The supernatant contains the amplified library. Save the beads in 50 μL TE buffer, as they can be used to rescue the library if any of the downstream step fails (see Note 17). 3.21 Remove Adapter Dimers with SPRI Bead-Based Size Selection

The goal of this step is to remove any adapter dimers, which may have amplified during the on-bead PCR. These adapter dimers can preferentially amplify during the side qPCR Subheading 3.22 and bias the estimation of the number of additional off-bead PCR. 1. Binding of the DNA library to SPRI beads: warm the SPRI beads to room temperature. Add 1:1 volume of beads to sample and mix well by pipetting. The exact volume of beads should be accurately determined by measuring the volume of the supernatant from Subheading 3.20 (which should be ~50 μL). Incubate for 5 min at room temperature. 2. Collect the SPRI beads using a magnetic rack, wait for the beads to settle (~1–2 min), and remove the supernatant. The supernatant contains the undesired adapter dimers and fragments shorter than ~200 bp, but can be saved for troubleshooting. 3. Wash the beads by resuspending them in 250 μL of fresh 70% ethanol. Collect the beads again and discard the supernatant. 4. Elute the library by resuspending the beads in 31 μL 10 mM Tris–Cl, pH 8 (see Note 18). Incubate for 5 min at room temperature. Collect the beads using a magnetic rack, and transfer the supernatant to a clean 1.5 mL microcentrifuge tube. The supernatant contains the library DNA. 5. Save 1 μL for quantification of fragment size distribution if desired (see Note 19). Use the remaining 30 μL for the next step of the protocol.

ChAR-Seq

The goal of the qPCR at this step, which we call “side qPCR,” is to evaluate how many extra PCR cycles can be done for each library while remaining in the “exponential amplification range” of the PCR (see Note 16). 1. Mix in a qPCR well: (a) 5 μL library DNA. (b) 6 μL 2 NEBNext High-Fidelity Master Mix. (c) 0.5 μL 10 μM universal primer. (d) 0.5 μL 10 μM indexing primer. (e) 0.12 μL 100 Sybr Green. 2. Perform a qPCR with the following parameters: (a) 1 cycle: 98  C for 30 s (initial denaturation). (b) 25 cycles: 98  C for 10 s (denaturation), 65  C for 75 s (annealing and extension). 3. Determine the number of off-bead PCR cycles to perform at the next step by plotting on a linear scale for each sample the fluorescence intensity versus cycle number, as shown Fig. 3. The number N of off-bead PCR cycles is the number of cycles such that the fluorescence intensity is about 1/3 the plateau intensity at the PCR saturation (relative to the baseline intensity). This number is typically N ¼ 1–10 when using ~10 million human cells, for total number of PCR cycles of 6–17 (see Note 20).

80000 Fluorescence [a.u.]

3.22 Side qPCR to Determine How Many Additional Cycles of PCR Amplification to Perform

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40000 Sample 1 (N=4) Sample 2 (N=9) Sample 3 (N=11)

20000

0

0

2

4

6

8 10 12 14 Side qPCR cycle #

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18

20

Fig. 3 Example of side qPCR for three samples that each require a different number of additional off-bead PCR cycles (N ¼ 4 for sample 1, N ¼ 9 for sample 2, N ¼ 11 for sample 3). Vertical bars indicate selected number of extra off-bead PCR cycles for each sample. These numbers were chosen to obtain a fluorescence of ~1/3 the plateau fluorescence

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3.23 Off-Bead Library Amplification

1. Mix in a normal PCR tube: (a) 25 μL of library DNA (the remainder of the sample from Subheading 3.21). (b) 25 μL 2 NEBNext High-Fidelity Master Mix. (c) 2.5 μL 10 μM universal primer. (d) 2.5 μL 10 μM indexing primer. 2. Run a PCR with the following parameters: (a) 1 cycle: 98  C for 30 s (initial denaturation). (b) N cycles: 98  C for 10 s (denaturation), 65  C for 75 s (annealing and extension). (c) Transfer the samples to 1.5 mL microcentrifuge tubes.

3.24 Removal of High Molecular Weight DNA Fragments

1. Bind high molecular weight fragments to the SPRI beads: pre-warm SPRI beads to room temperature, and add to each sample from Subheading 3.23 a volume of beads equal to 0.6 the volume of the PCR sample (see Note 21). Incubate for 5 min at room temperature with agitation. 2. Collect DNA fragments of interest: place the tubes on a magnetic rack and wait until the beads settle (~1–2 min). Collect the slurry and transfer it to a clean 1.5 mL microcentrifuge tube. The bead pellet contains DNA fragments larger than ~500 bp and can be discarded or stored in 70% ethanol at 4  C for troubleshooting. The supernatant contains the fragments of interest as well as primer dimers, which needs to be removed in the next step.

3.25 Final Removal of Primer Dimers and Low Molecular Weight DNA

1. Binding of low molecular weight DNA fragments to SPRI beads: for each sample, carefully measure the volume Vsup of the supernatant collected at Subheading 3.24. Add to each sample a volume of beads Vbeads equal to Vbeads ¼ 0.1875  Vsup to obtain a final ratio of bead slurry to sample equal to 0.9 (see Note 21). Incubate for 5 min at room temperature without agitation. 2. 1st wash: place the tubes on a magnetic rack and wait until the beads settle (~1–2 min). Discard the supernatant which contains molecules 70% of the reads with a single bridge, 0)—that is, binary indicators of whether each Poisson variable is greater than zero. Given the maximum likelihood estimate of λX is log(1  mean(IX)), and similar for λY, the number of beads with unique barcodes, which is P (X + Y ¼ 1) ¼ (λX + λY)exp.(λX  λY), since X + Y ~ Pois (λX + λY), can be estimated. Similarly, the number of beads with multiple barcodes is P(X + Y > 1) ¼ 1  P(X < 2) ¼ 1  (1 + λX + λY)exp.((λX + λY)). The ratio of single-to-multiple

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barcodes can be estimated as P(X + Y ¼ 1)/P(X + Y > 1). This assumption can be verified with the chi-squared test of independence with two degrees of freedom, since the total number of beads is not fixed and two parameters are estimated from the data. 5. It is recommended when setting up barcoding of beads to control barcode length and purity on a 12% urea/polyacrylamide gel before and after random tailing. Single-stranded barcodes are released from beads in formamide-containing loading buffer at 90  C for 5 min and run on a gel following standard procedures. 6. Longer crosslinking results in the co-purification of cytoplasmic content with nuclei as indicated by, for example, an increase in the fraction of rRNA and exonic sequencing reads. Implications on biological results and the interpretation thereof are expected to be substantial if the fraction of cytoplasmic RNA increases considerably. Also, long crosslinking resulting in increased cytoplasmic RNA contamination cannot be counterbalanced with harsher sonication, as the RNA composition in the sample remains unchanged by sonication. Significantly shorter crosslinking or lower concentrations of fixatives are expected to reduce the efficiency and sensitivity of the assay as the fraction of singly barcoded and sequenced RNA in the sample might increase. However, the Proximity RNA-seq analysis pipeline will still accurately identify RNA proximities in under-crosslinked conditions, but many actual RNA contacts will lack statistical power. 7. The optimal amount of input material, i.e., subnuclear particles, corresponds to an equivalent between 50 and 100 ng RNA. Below 50 ng the library quality decreases, for example, we sequence more unmappable reads, presumably due to the low quantity and the increased library amplification required. With 100 ng input material, we observe smaller divergence in valency values for transcripts. The assignment of valency, i.e., how often a transcript is found alone or in proximity to other transcripts, is hampered with more frequent encapsulation of multiple particles into individual droplets. As for bead barcoding, a Poisson model underlies particle encapsulation (see also Subheading 3.2). 8. We chose a cyclic temperature program for reverse transcription, which we found to increase the amount of synthesized cDNA. We assume that repeated cycles of low temperature for annealing followed by high temperature for synthesis increase the chance to capture RNA on beads in a given droplet. Possibly a fraction of crosslinks can be reversed throughout the high temperature phases, and RNA might become fragmented at

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high temperature and by magnesium cations present. Both processes potentially liberate RNA in order to anneal to random primers on beads. 9. Library amplification consists of two PCR reactions. G homopolymers required for PCR amplification are added to unused barcodes as well as to cDNA ends. Therefore, after the first PCR of five cycles, products are size-selected. The second PCR is performed in emulsion to reduce amplification biases. The resulting library is again selected against small fragments consisting only of a barcode. 10. To aid the interpretation of the results, a slight modification is made to the reference genome. Firstly, the ribosomal RNA sequences are replaced in the genome with Ns (representing bases whose identity could not be determined). The rRNA sequence is then incorporated into the genome as a separate chromosome. References 1. Stahl PL et al (2016) Visualization and analysis of gene expression in tissue sections by spatial transcriptomics. Science 353:78–82 2. Lee JH et al (2014) Highly multiplexed subcellular RNA sequencing in situ. Science 343:1360–1363 3. Chen KH, Boettiger AN, Moffitt JR, Wang S, Zhuang X (2015) RNA imaging. Spatially resolved, highly multiplexed RNA profiling in single cells. Science 348:aaa6090 4. Shah S et al (2018) Dynamics and spatial genomics of the nascent transcriptome by intron seqFISH. Cell 174(2):363–376 5. Xia C, Fan J, Emanuel G, Hao J, Zhuang X (2019) Spatial transcriptome profiling by MERFISH reveals subcellular RNA compartmentalization and cell cycle-dependent gene expression. Proc Natl Acad Sci U S A 116 (39):19490–19499 6. Weidmann CA, Mustoe AM, Weeks KM (2016) Direct duplex detection: an emerging tool in the RNA structure analysis toolbox. Trends Biochem Sci 41:734–736 7. Nguyen TC et al (2016) Mapping RNA–RNA interactome and RNA structure in vivo by MARIO. Nat Commun 7:12023

8. Kudla G, Granneman S, Hahn D, Beggs JD, Tollervey D (2011) Cross-linking, ligation, and sequencing of hybrids reveals RNA-RNA interactions in yeast. Proc Natl Acad Sci U S A 108:10010–10015 9. Ramani V, Qiu R, Shendure J (2015) Highthroughput determination of RNA structure by proximity ligation. Nat Biotechnol 33:980–984 10. Sugimoto Y et al (2015) hiCLIP reveals the in vivo atlas of mRNA secondary structures recognized by Staufen 1. Nature 519:491–494 11. Morf J et al (2019) RNA proximity sequencing reveals the spatial organization of the transcriptome in the nucleus. Nat Biotechnol 37:793–802 12. Langmead B, Salzberg SL (2012) Fast gappedread alignment with bowtie 2. Nat Methods 9:357–359 13. Kim D, Langmead B, Salzberg SL (2015) HISAT: a fast spliced aligner with low memory requirements. Nat Methods 12:357–360 14. Wingett SW, Andrews S (2018) FastQ screen: a tool for multi-genome mapping and quality control. F1000Res 7:1338

Chapter 14 Immunoprecipitation of DNA:RNA Hybrids Using the S9.6 Antibody Hunter R. Gibbons and Thomas M. Aune Abstract Formation of DNA:RNA hybrids or R-loops contributes to numerous biologic processes. The development of the S9.6 antibody makes the analysis of R-Loops (DNA:RNA hybrids) possible through immunoprecipitation. Here, we describe the isolation of DNA:RNA hybrid structures using the S9.6 antibody. Using this protocol, both the DNA and RNA binding partners of the R-loop can be analyzed via qPCR, whole genome sequencing, or other methods. Key words DRIP, RNA, DNA, ChIP, R-loop, S9.6, Immunoprecipitation

1

Introduction DNA:RNA hybrids (or R-loops) are important regulators of transcription and impact RNA processing at initiation, elongation, and termination [1–6]. R-loops are triple stranded nucleic acid structures including a DNA:RNA hybrid strand and one displaced DNA strand (Fig. 1) [7]. R–loops can regulate transcription by blocking transcriptional silencing by preventing binding of DNA methyltransferases and subsequent DNA methylation of CpG islands [8], induce transcriptional termination by RNA polymerase pausing [9], and repress transcription by promoter occlusion [4]. Ranging in size from 50 to 2000 base pairs, R-loops are abundant around promoters and modify transcription to both induce and repress impacted genes [10]. Antisense and divergent long noncoding RNAs can also form R-loops to regulate transcription of neighboring mRNA genes [1, 11]. R-loops can also cause replicative stress [10], genome instability [11], and chromatin alterations [12] that may be associated with cancer [9, 13]. Effective analysis of R-loop formation and location is critical to understand transcriptional dynamics.

Ulf Andersson Vang Ørom (ed.), RNA-Chromatin Interactions: Methods and Protocols, Methods in Molecular Biology, vol. 2161, https://doi.org/10.1007/978-1-0716-0680-3_14, © Springer Science+Business Media, LLC, part of Springer Nature 2020

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R-Loop Formation

Standard Transcription

G-Rich DNA C-Rich DNA RNA RNA-Pol II

Fig. 1 Model of R-loop formation. Following standard transcription, an RNA is normally trafficked away from its gene locus to be further processed as shown on the left. Sometimes, for example, if the region is C-G rich, it stays bound to its own locus, forming a DNA:RNA hybrid, also known as an R-loop, as shown on the right

R-loops were previously studied using a variety of techniques including mobility shift assays, in situ hybridization, and native bisulfite modification. Analysis of R-loops greatly expanded following the development of the S9.6 antibody that specifically recognizes these DNA:RNA hybrids [14]. The S9.6 antibody binds to DNA:RNA hybrids in a sequence independent manner, making immunoprecipitation of the R-loop (DNA:RNA immunoprecipitation or DRIP) a standard detection method [15]. The DRIP technique, first introduced in 2010, made the analysis of R-loops more accessible and comprehensive through qPCR analysis and whole genome sequencing [16]. The DRIP technique does have limitations. The S9.6 antibody binds double stranded RNA, although not as effectively as it binds DNA:RNA hybrids [2]. Additionally, formaldehyde crosslinking can increase the number of false positives detected [17]. Despite this, the DRIP assay is still by far the most effective tool to analyze R-loop locations and size. Here we describe an immunoprecipitation procedure to analyze both the DNA and RNA fractions involved in R-loop formation (Fig. 2). In this protocol, cells are fixed with formaldehyde, lysed, and sonicated similar to a standard ChIP assay. R-loops are immunoprecipitated using the S9.6 antibody, followed by extraction of either the RNA or DNA fraction from the immunoprecipitate. DNA or RNA fractions can be analyzed via qPCR or sequencing to provide effective analysis of R-loop formation around a single gene or throughout the entire genome.

2 2.1

Materials Nuclei Isolation

1. Hypotonic solution: 20 mM Tris–HCl, pH 7.4, 10 mM NaCl, 3 mM MgCl2. 2. 10% NP-40.

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Fig. 2 DNA:RNA immunoprecipitation protocol flowchart. Each step is uniform until after the S9.6 antibody immunoprecipitation, in which the DNA and RNA fractions are processed separately so both can be isolated from the immunoprecipitate

3. R-Loop Digestion Buffer: 100 mM NaCl, 10 mM Tris pH 8, 25 mM EDTA pH 8, 0.5% SDS, 0.2 mg/ml Proteinase K (add just before use) (see Note 1). 4. Eppendorf 5810 R refrigerated centrifuge (see Note 2). 2.2 Nucleic Acid Isolation

1. FA Lysis Buffer: 20 mM Tris–HCl pH 7.5, 150 mM NaCl, 1 mM Na2EDTA, 1 mM EGTA, 1% Triton, 2.5 mM sodium pyrophosphate, 1 mM β-glycerophosphate, 1 mM Na3VO4, 1 μg/ml leupeptin (see Note 3).

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Sonication

1. Diagenode Bioruptor Sonicator. 2. New England Biolabs 100 bp DNA ladder. 3. New England Biolabs 6 loading dye. 4. New England Biolabs RNase H.

2.4 Immunoprecipitation

1. Anti-Mouse IgG antibody. 2. S9.6 DNA-RNA Hybrid antibody. 3. Pierce Protein A/G Magnetic Beads. 4. DynaMag2 magnetic tube rack. 5. Bead Wash Buffer: TBS containing 0.05% Tween 20.

2.5 RNA Extraction and cDNA Conversion

1. Tri-Reagent (RNA/DNA/Protein) Isolation reagent. 2. QIAGEN RNeasy MinElute Cleanup Kit. 3. Tris-EDTA Buffer: 1 M Tris pH 8, 0.5 M EDTA pH 8, H2O. 4. Invitrogen Superscript 3 First Strand Synthesis System. 5. QIAGEN Qiaquick PCR purification kit.

2.6 DNA Extraction and PCR

1. Elution Buffer: 100 mM NaHCO3, 1% SDS. 2. ABI Quantstudio 3 or equivalent qPCR platform. 3. ABI PowerUP SYBR Green Reaction Mix. 4. PCR primers for each unique target region.

3

Methods Carry out all methods with samples on ice unless otherwise specified.

3.1

Fixation

1. Using no more than ten million cells, add formaldehyde directly to the media, dropwise to a final concentration of 0.75% of total volume, and rotate for 10 min. (Example: a 3 ml culture would receive 60.8 μl of formaldehyde). 2. Add glycine to a final concentration of 125 mM to the media, and rotate again for 5 min at room temperature. 3. Isolate cells by centrifuging at 4  C and 300  g for 10 min. 4. Pour off the supernatant from the sample. 5. Wash once in ice cold PBS at an equivalent volume to the previous media volume. 6. Repeat step 3 and remove as much PBS as possible without disturbing the cell pellet (see Note 4).

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Nuclei Isolation

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1. Resuspend cell pellet in 500 μl of Hypotonic Solution and pipette up and down to mix. 2. Incubate for 15 min. 3. Add 25 μl of 10% NP40 detergent and vortex on highest setting for 10 s. 4. Centrifuge at 4  C at 720  g for 10 min. 5. A pellet should be visible at the bottom of the tube containing cell nuclei. Remove the supernatant from the tube without disturbing the pellet. 6. Resuspend pellet in 1 ml of R-loop Digestion Buffer, and leave overnight in a 65  C water bath (approx. 12–16 h). Remove samples from the water bath, and immediately place on ice. 7. Add 500 μl of 25:24:1 Phenol:Chloroform:Isoamyl Alcohol to the sample, pipetting up and down to mix effectively. 8. Centrifuge sample at 4  C at maximum speed (~21,000  g) for 10 min. 9. Isolate the top aqueous layer, which will contain the nucleic acids from the sample. 10. Combine aqueous layer with ¼ volume of 9 M sodium acetate (if aqueous layer is 500 μl, add 125 μl of sodium acetate). 11. Add equivalent volume of ice-cold isopropanol to the sample (if combined volume from step 5 is 625 μl, add 625 μl of isopropanol). 12. Add 3 μg of glycogen and invert tube five times to effectively mix (see Note 5). 13. Place on ice for 10 min. 14. Centrifuge sample at 4  C and ~21,000  g for 10 min. 15. Pour off supernatant and resuspend in 1 ml of 70% ethanol, followed by another centrifugation at 4  C and ~21,000  g for 10 min. 16. Pour off supernatant, and resuspend the sample in 500 μl of FA Lysis Buffer.

3.3

Sonication

1. Nucleic acids should be effectively resuspended to dissolve into the FA Lysis Buffer before sonication. Sonication should be completed to form an average fragment size of ~500 bp of DNA (see Note 6). 2. Sonication in a Diagenode Bioruptor can be completed in two sessions of 15 min each, at the medium setting with on and off pulses of 30 s (see Note 7). 3. Prepare a 1.5% agarose gel to analyze the size of DNA fragments of your sonicated nucleic acid.

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4. Load nucleic acid into the gel with a 6 loading dye and a separate 100 bp DNA ladder to determine fragment size. 5. Electrophorese at 100 V until the ladder reaches 2/3 of the way down the gel. 6. Analyzed gel should indicate an average fragment size of ~500 bases, this is ideal to identify R-loop formation. 7. Before beginning immunoprecipitation, reserve 25 μl of chromatin from each sample to use as an input control, or around 5% of the total sample. Store this chromatin for use in final analysis (see Note 8). 8. Each sample will be analyzed using three reactions: A non-specific antibody control (IgG), a sample with the S9.6 antibody that has been treated by RNaseH (negative control), and a sample with the S9.6 antibody (experimental). 9. From your sonicated chromatin, isolate 40 μg of nucleic acid for each of these three reactions (see Note 9). 10. To your negative control, add 5 μl of RNase H. To all 3 samples, add the salt solution for the RNase H reaction. Incubate all three samples at 37  C on a heat block for 1 h. RNase can be heat inactivated at 65  C for 20 min. 11. To each sample, add FA lysis buffer to raise the total volume to 500 μl. 3.4 S9.6 Immunoprecipitation

1. Add 10 μg of anti-mouse IgG to the isotype control, and 10 μg of S9.6 antibody to the negative and experimental samples. 2. For each sample including controls, prepare 25 μl of Pierce Protein A/G magnetic beads. Wash with 25 μl of bead wash buffer. 3. Place tubes on a Dynamag 2 magnetic tube rack (or equivalent magnet) to isolate beads, then remove and discard supernatant while saving the beads (see Note 10). 4. Repeat wash with 500 μl of Bead wash buffer, and isolate beads on magnetic rack. 5. Resuspend beads in 25 μl of FA lysis buffer. 6. Add beads to each sample, and rotate at 4  C for 4 h. 7. After 4-h rotation, place each tube on the magnetic rack, and pipette away the supernatant, saving for future use (see Note 11). 8. Remove tube from magnetic rack, and softly pipette 500 μl of wash buffer over the beads containing the immunoprecipitate. Without vortexing, mix beads to wash. 9. Place tubes back on magnetic rack and remove the wash buffer without disturbing the bead pellet on the side of the tube.

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10. Repeat steps 7 and 8 two more times. 11. Wash 1 with 500 μl of ice-cold PBS. 12. If you plan to isolate the RNA fraction, proceed to RNA Extraction. If you plan to isolate the DNA fraction, proceed to DNA Extraction. 3.5

RNA Extraction

1. Resuspend beads in 1 ml of TRI-Reagent, and freeze the sample at 80  C to extract RNA from beads (see Note 12). 2. Thaw sample completely before moving on to the rest of the RNA isolation, usually ~15 min at room temperature (see Note 13). 3. Add 200 μl of chloroform, followed by vigorous vortexing and mixing by inversion. Vortex multiple times, during which two layers will begin to form. 4. Samples should incubate at room temperature for 10 min, after which the two phases should be clearly visible. 5. Centrifuge at ~21,000  g for 10 min at 4  C. 6. Transfer the aqueous phase to new eppendorf tubes, and add 500 μl of ice-cold isopropanol and 2 μl of glycogen to each tube (see Note 14). 7. Mix by inversion and incubate at room temperature for 10 min. 8. Centrifuge at 21,000  g for 10 min at 4  C. Following this spin, a pellet should be visible at the bottom of the tube. 9. Pour off supernatant from each tube, and wash with 1 ml ice-cold 75% ethanol. 10. Centrifuge at 21,000  g for 10 min at 4  C. 11. Pour off supernatant, and let sample air dry for 10 min to remove excess ethanol. 12. Resuspend in 100 μl of Tris-EDTA Buffer. 13. Using the RNeasy MinElute Cleanup Kit, add 350 μl of Buffer RLT to each sample. 14. Add 250 μl of 100% ethanol to each sample, and mix by pipetting up and down. 15. Transfer each sample to a Minelute column, and centrifuge at 13,500  g for 30 s. Discard the flow thru following centrifugation. 16. Add 500 μl of Buffer RPE to the Minelute column, and again centrifuge at 13,500  g for 30 s. Again, discard the flow thru. 17. Add 500 μl of 80% ethanol to each column, and centrifuge for 2 min at 13,500  g. Discard the flow thru. 18. Centrifuge the dry column with the cap open at 13,500  g for 5 min.

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19. Move to a new eppendorf tube for collection, and dispose of the previous collection tube. 20. Add 18 μl of ultrapure water directly to the column. Let the column sit at room temperature for 5 min, then centrifuge at 13,500  g for 1 min (see Note 15). 21. Discard the column and proceed to cDNA conversion (see Note 16). 3.6

cDNA Conversion

To convert RNA to cDNA to complete qPCR, use the Superscript 3 first strand synthesis system. To use our whole sample, we will run a double reaction from the first-strand kit. 1. Place 18 μl of RNA from previous step to a strip cap tube for use in a thermal cycler. Combine with 1 μl of dNTPs, and 1 μl of Oligo(dt) if your RNA of interest is polyadenylated, or 1 μl random hexamers if it is not or you do not know. 2. Incubate at 65  C in a thermal cycler for 5 min. Then place on ice, or at 0  C for at least 1 min. 3. To each sample, add 4 μl of 10 RT buffer, 8 μl of 25 mM Magnesium Chloride, 4 μl of 1 M DTT, 2 μl of RNaseOUT, and 2 μl of Superscript III RT (see Note 17). (a) If using Oligo dT, go directly to step 5. If using random hexamer, samples should be incubated at 25  C for 10 min before proceeding. 4. Incubate at 50  C for 50 min, followed by 85  C for 5 min. 5. Add 1 μl of RNase H to each sample, and incubate at 37  C for 20 min. 6. Using the QIAquick PCR purification kit, add 105 μl of Buffer PB to each reaction. 7. Place the mixture of sample and buffer to a QIAquick column and centrifuge at 13,500  g for 1 min. Discard the flow thru. 8. Add 750 μl of Buffer PE to each column, and centrifuge at 13,500  g for 1 min. Discard the flow thru. 9. Centrifuge the dry column with cap open at 13,500  g for 1 min. 10. Place the column in a fresh eppendorf tube for collection, and add 50 μl of Buffer EB to the center of the column. 11. Centrifuge at 13,500  g for 2 min. 12. Proceed to quantitative PCR or other method of analysis.

3.7

DNA Extraction

1. Add 120 μl of elution buffer to the beads and rotate for 15 min at room temperature.

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2. The antibody/DNA complex is now present in the supernatant and separate from the beads. Place tubes on the magnetic rack, and isolate supernatant in a new eppendorf tube. 3. Add 5 μl of proteinase K to each tube, and allow at least 4 h (or overnight) of incubation in a 65  C water bath. 4. The following morning, add 500 μl of phenol chloroform and vortex vigorously. 5. Centrifuge for 5 min at 4  C at 21,000  g. 6. Remove the top aqueous layer and move to a new eppendorf tube. 7. Add 125 μl (or 1/4th of total volume of aqueous layer) of 9 M sodium acetate. 8. Add 625 μl (or combined volume of sample and sodium acetate) of 100% isopropanol to each sample. 9. Add 2 μl of glycogen to each sample. 10. Vortex vigorously and place on ice for 10 min. 11. Centrifuge for 10 min at 4  C at 21,000  g. 12. Pour off supernatant and let the pellet air dry. 13. Resuspend in 50 μl Ultrapure water, and proceed to qPCR evaluation. 3.8

Quantitative PCR

1. Dilute samples uniformly to the volume necessary for your PCR reactions using 5 μl of sample per reaction (50 μl provides 10 reactions, so if you plan to run 20 reactions, dilute all samples to 100 μl total). 2. Each 25 μl reaction should include: (a) 12.5 μl of SYBR Green reaction mix. (b) 6.25 μl of Ultrapure water. (c) 1.25 μl of PCR primers at 10 μM concentrations (final concentration of each primer should be 0.5 μM). (d) 5 μl of sample per reaction. 3. Each sample will run at the following temperatures and cycles to complete the qPCR. (a) 2 min at 50.0  C. (b) 10 min at 95.0  C. (c) 40 cycles repeating l

15 s at 95.0  C.

l

60 s at 60.0  C.

4. Change qPCR settings according to your reagents and qPCR instrument, but these settings are applicable to the ABI qPCR system.

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Notes 1. Proteinase K activity can degrade over time, so prepare the R-loop digestion buffer in large quantities without the proteinase K included. Before using the digestion buffer, combine the premade buffer with the proteinase K for the most effective digestion. 2. On many of descriptions of this protocol, a speed of ~21,000  g is referenced for phase separations, including phenol:chloroform and Tri-Reagent extractions. This is the top speed of the refrigerated centrifuge we use. Most centrifugations could be completed at slower speeds and produce similar results, but the most important part is to keep the samples chilled during these centrifugations. No matter what centrifuge you have, spin at top speed available while keeping samples chilled at 4  C. 3. Cell Signaling provides a 10 cell lysis buffer that we used for our FA lysis buffer that works great, as opposed to preparing all the reagents. Combine this with the New England Biolabs Proteinase K, which is the total buffer used in these reactions. 4. Excess PBS left in can really affect the function of the NP40 detergent, so pour off the supernatant PBS first, and then pull as much off as possible with a pipette to limit how much is left in the sample. 5. This isolation should contain a large amount of nucleic acid, so a visible pellet should be present. If not, attempt the isolation again from the original sample. 6. It is never going to be perfect 500 base pair fragments, but keeping your fragments almost entirely below the 1200–1500 base pair range will significantly reduce your background and the number of false positives you will find. Hybrid DNA:RNA formations are usually larger than 50 bases, so if your fragments are mostly around 500 base pairs, you will have a good sample to work with for the immunoprecipitation. 7. You absolutely must use a bath sonicator for this step (Fig. 3). Using a needle sonicator results in a significant decrease in the overall yield recovered. You can even just use a cocktail of endonucleases for sonication, but a bath sonicator works most effectively and efficiently. The Bioruptor works great at the settings presented, but this step does take optimizing, because every sonicator is a little bit different. 8. The input is necessary for the final calculation for the qPCR determinations, just like a standard ChIP experiment. If used for the RNA fraction, it needs to be incorporated during RNA isolation immediately Subheading 3.5 RNA extraction. If used

DNA:RNA Hybrid Immunoprecipitation

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Fig. 3 A bath sonicator should be used for fragmentation in DRIP assays, as a needle sonicator results in loss of R-loops and lack of signal for the final analysis

for the DNA determination, it can be incorporated at Subheading 3.8, step 3. Otherwise, treat as you would any input for a standard ChIP assay. 9. Another step that could need optimizing: For most ChIP experiments, 25 μg works well. With immune cells, I scaled up to 40 μg, and this produced an effective qPCR measurement. If you end up with too much background, or not enough signal, possibly scaling up or down the initial chromatin per reaction may help. 10. Pierce Protein A/G magnetic beads work great for this assay, although other brands of magnetic A/G beads may also be effective. Agarose beads did not allow for a clean enough immunoprecipitation and resulted in too high background Ct values. 11. Saving the supernatant is not incredibly important, but it can help you verify when the experiment failed. If the supernatant nor your experimental sample tests positive for your gene of interest, then something went wrong before this step. If it is present in the supernatant, but not your experimental, then it is most likely negative for DNA:RNA hybrid formation. 12. Tri-Reagent, or another alternative, Trizol, are both available to complete the RNA extraction. No matter which you use, RNA extraction utilizing guanidinium-thiocyanate-phenolchloroform will provide the most efficient and accurate results for this protocol.

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13. I would advise freeze/thawing three times before beginning the experiment, as additional freeze thaws could help increase RNA yields by a small amount. At this point, you are working with a small total amount of nucleic acid, so every bit should be preserved as much as possible. 14. A dyed glycogen, like Glycoblue, makes this step much easier. Unlike the previous isolation, this step does not have much material present in comparison. Thus, using the dyed glycogen makes it easier to see if there is any nucleic acid present. 15. The instructions for the MinElute kit do not include the incubation step with ultrapure water on top of the column, but we have found that soaking the column really helps maximize RNA recovery. 16. This is a good time to quantitate your RNA, possibly using a nanodrop to test if sample is present or not. It will never be a very high concentration, but this is a good step to make sure whether anything has gone wrong in your previous steps. 17. It is by far easier to prepare a master mix for these reagents as opposed to pipetting them all individually. I would advise combining before adding to the samples.

Acknowledgements This Research was supported by National Institute of Health grants R01 AI044924 and T32 DK07563. References 1. Boque-Sastre R, Soler M, Oliveira-Mateos C et al (2015) Head-to-head antisense transcription and R-loop formation promotes transcriptional activation. Proc Natl Acad Sci U S A 112 (18):5785–5790 2. Chen P, Chen H, Acharya D et al (2015) R loops regulate promoter-proximal chromatin architecture and cellular differentiation. Nat Struct Mol Biol 22(12):999 3. Grunseich C, Wang IX, Watts JA et al (2018) Senataxin mutation reveals how R-loops promote transcription by blocking DNA methylation at gene promoters. Mol Cell 69:426–437. e7. https://doi.org/10.1016/j.molcel.2017. 12.030 4. Hage EA, French S, Beyer A et al (2010) Loss of topoisomerase I leads to R-loop-mediated transcriptional blocks during ribosomal RNA synthesis. Genes Dev 24(14):1546–1558. https://doi.org/10.1101/gad.573310

5. Hage EA, Webb S, Kerr A et al (2014) Genome-wide distribution of RNA-DNA hybrids identifies RNase H targets in tRNA genes, retrotransposons and mitochondria. PLoS Genet 10.10(2014):e1004716 6. Hala´sz L, Kara´nyi Z, Boros-Ola´h B et al (2017) RNA-DNA hybrid (R-loop) immunoprecipitation mapping: an analytical workflow to evaluate inherent biases. Genome Res 27 (6):1063–1073. https://doi.org/10.1101/gr. 219394.116 7. Hu Z, Zhang A, Storz G et al (2006) An antibody-based microarray assay for small RNA detection. Nucleic Acids Res 34(7): e52–e52. https://doi.org/10.1093/nar/ gkl142 8. Santos-Pereira J, Aguilera A (2015) R loops: new modulators of genome dynamics and function. Nat Rev Genet 16(10):583 9. Nakama M, Kawakami K, Kajitani T et al (2012) DNA–RNA hybrid formation mediates

DNA:RNA Hybrid Immunoprecipitation RNAi-directed heterochromatin formation. Genes Cells 17(3):218–233 10. Ponaro KT, Mitter R et al (2014) RECQL5 controls transcript elongation and suppresses genome instability associated with transcription stress. Cell 157(5):1037–1049 11. Gibbons H, Shaginurova G, Kim L et al (2018) Divergent lncRNA GATA3-AS1 regulates GATA3 transcription in T-helper 2 cells. Front Immunol 9:2512 12. Roberts RW, Crothers DM (1992) Stability and properties of double and triple helices: dramatic effects of RNA or DNA backbone composition. Science 258:1463–1466. https://doi.org/10.1126/science.1279808 13. Sanz L, Hartono S, Lim Y et al (2016) Prevalent, dynamic, and conserved R-loop structures

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associate with specific epigenomic signatures in mammals. Mol Cell 63(1):167–178 14. Shivji M, Renaudin X, Williams C et al (2018) BRCA2 regulates transcription elongation by RNA polymerase II to prevent R-loop accumulation. Cell Rep 22(4):1031–1039 15. Stork C, Bocek M, Crossley M et al (2016) Co-transcriptional R-loops are the main cause of estrogen-induced DNA damage. elife 5: e17548 16. Sun Q, Csorba T, Skourti-Stathaki K et al (2013) R-loop stabilization represses antisense transcription at the Arabidopsis FLC locus. Science 340(6132):619–621 17. Whalen S, Truty R, Pollard K et al (2016) Enhancer–promoter interactions are encoded by complex genomic signatures on looping chromatin. Nat Genet 48(5):488

Chapter 15 Characterization of R-Loop Structures Using Single-Molecule R-Loop Footprinting and Sequencing Maika Malig and Frederic Chedin Abstract R-loops are three-stranded structures that form during transcription when the nascent RNA hybridizes with the template DNA resulting in a DNA:RNA hybrid and a looped-out single-stranded DNA (ssDNA) strand. These structures are important for normal cellular processes and aberrant R-loop formation has been implicated in a number of pathological outcomes, including certain cancers and neurodegenerative diseases. Mapping R-loops has primarily been performed using DRIP (DNA:RNA immunoprecipitation) based methods that are dependent on the anti-DNA:RNA hybrid S9.6 antibody and short-read sequencing. While DRIP-based methods are robust and report R-loop formation genome-wide, they only do so at the population average level; interrogating R-loop formation at the single molecule level is not feasible with such approaches. Here we present single molecule R-loop footprinting (SMRF-seq), a method that relies on the chemical reactivity of the displaced ssDNA strand to non-denaturing sodium bisulfite and single molecule long-read sequencing as a readout, to characterize R-loops. SMRF-seq can be used independently of S9.6 to generate high resolution, strand-specific, maps of individual R-loops at ultra-deep coverage on kilobases-length DNA fragments. Key words R-loop, DNA:RNA hybrid, Non-denaturing bisulfite conversion, SMRT sequencing, Transcription

1

Introduction R-loops are three-stranded nucleic acid structures consisting of a DNA:RNA hybrid and a displaced single stranded DNA. R-loops are abundant non-B DNA structures in genomes, covering 5–10% of the genomic space in organisms ranging from yeasts, plants, and mammals [1–6]. While evidence exists to support that R-loops can form in trans [7–9], nuclear R-loops are predominantly understood to form co-transcriptionally from the re-invasion of the nascent transcript into the duplex DNA template. This is evidenced from the overwhelming genic distribution of R-loops, their co-directionality and correlation with transcription, and conclusions from in vitro transcription reactions [10]. Investigations into the possible roles of R-loops have suggested that these

Ulf Andersson Vang Ørom (ed.), RNA-Chromatin Interactions: Methods and Protocols, Methods in Molecular Biology, vol. 2161, https://doi.org/10.1007/978-1-0716-0680-3_15, © Springer Science+Business Media, LLC, part of Springer Nature 2020

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structures influence a range of cellular processes [11–13]. Under normal conditions, R-loops participate in the regulation of gene expression [2, 14, 15], class switch recombination in B cells [16, 17], and efficient transcription termination [18, 19]. Deregulation of R-loop metabolism has also been linked to genome instability [20–22] ultimately contributing to pathological conditions such as certain cancers and neurological disorders [23, 24]. R-loops have been mapped genome-wide using DNA:RNAimmunoprecipitation (DRIP) [1], a method that relies on the S9.6 anti-DNA:RNA hybrid antibody [25] and high-throughput shortread sequencing. Since its initial introduction, DRIP has been widely adopted and several variants of the method have been introduced to improve its resolution and strand-specificity [2, 6, 15]. While robust, these methods nonetheless possess few limitations. A first complication is due to the significant residual affinity of the S9.6 antibody for double-stranded RNA (dsRNA) [26]. This residual affinity can result in inaccurate R-loop maps when RNA strands are directly sequenced unless additional steps are taken to remove contaminating dsRNAs [5, 27]. Secondly, DRIP, like ChIP approaches, only provides a population average view of R-loop formation derived from a large cell population. The use of highthroughput short-read sequencing technologies as read-outs further precludes obtaining any information on individual R-loops including their lengths and individual start and stop positions. To overcome these limitations and provide an S9.6-independent method to profile individual R-loops at ultra-deep coverage, we developed Single-Molecule R-loop Footprinting and sequencing (SMRF-seq) [28]. SMRF-seq relies on non-denaturing bisulfite conversion [16] to catalyze the efficient conversion of unpaired cytosines (C) located on the displaced ssDNA strand of R-loops to uracils (U). After long-range locus-specific PCR amplification, molecules amplified from the tagged displaced strand will carry patches of cytosine to thymine (T) conversions flanked by unmodified DNA. These patches represent R-loop “footprints”. The use of Pacific Bioscience’s Single-Molecule Real Time sequencing (SMRT-seq) on long PCR amplicons permits the characterization of collections of individual footprints at high coverage. Overall, SMRF-seq permits R-loop mapping at high resolution, in a strand-specific manner, on kilobases-long single-molecule reads and at high coverage. SMRF-seq, together with its accompanying analysis package Gargamel, will fuel new waves of investigations into the formation and function of R-loop structures.

2

Materials Prepare all reagents and dilutions using molecular biology-grade nanopure water (ddH2O).

Characterization of R-Loop Structures Using Single-Molecule R-Loop. . .

2.1

General

211

1. Tris-EDTA (TE) buffer: 10 mM Tris–HCL pH 8.0 and 5 mM EDTA. 2. 200 proof Ethanol. 3. Elution Buffer (EB): 10 mM Tris–HCl pH 8.5. 4. 1.5 mL and 2 mL Microcentrifuge tubes. 5. 8-strip PCR tubes. 6. Bench top Microcentrifuge. 7. DynaMag-2 beads (Life Technologies, 12321D). 8. AMPure XP beads (Beckman Coulter). Set at room temperature before use. 9. Mini tube rotator.

2.2 Nucleic Acid Purification

1. 15 mL conical tubes. 2. Proteinase K. Dilute to 20 mg/mL working solution. 3. Sodium dodecyl sulfate (SDS). Dilute to 20% working solution. 4. 5PRIME Phase Lock Gel Light 2 mL tubes. 5. Phenol:Choloroform:Isoamyl alcohol (25:24:1). 6. 3 M Sodium Acetate (NaOAc) pH 5.2. 7. Wide bore 200 and 1000 μL barrier tips.

2.3 Non-denaturing Bisulfite Treatment

1. DNA Methylation-Lightning.

2.4 Site-Specific Amplification

1. Thermocycler (optional: gradient-enabled block).

2. Rotisserie oven.

2. Phusion Hot Start DNA Polymerase. 3. 5
Phusion HF Buffer. 4. 5
Phusion GC Buffer. 5. dNTPs: Equimolar amounts of ATP, CPT, GTP, TTP. Dilute to 10 mM working solution. 6. 5 M Betaine.

2.5 SMRT-Bell Library Preparation

1. SMRTbell Template Prep Kit 1.0 (Pacific Biosciences, 100-259-100). 2. AMPure PB 5 mL (Pacific Biosciences, 100-265-900).

2.6

Quality Check

1. Spectrophotometer. 2. 2100 Bioanalyzer. 3. GelRed (Phenix Research Products, RGB-4102). 4. 50
TAE buffer: For a 1 L stock solution, dissolve 242 g Tris base in 600 mL ddH2O. Add 57.1 mL glacial acetic acid and

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100 mL 0.5 M EDTA pH 8. Adjust volume to 1 L with ddH2O. Dilute stock to 1
working concentration. 5. Agarose gel: dissolve molecular biology grade agarose to a final 0.7% (weight vol.) concentration in 1
TAE by heating. Cool to ~65  C. Add 1
final concentration of GelRed. Cast gel and insert comb.

3

Methods Overview. SMRF-seq exploits the presence of long stretches of ssDNA on the displaced strands of three-stranded R-loop structures. The intrinsic ssDNA character of this strand can be revealed upon treatment with sodium bisulfite under non-denaturing conditions [16]. This will lead to the conversion of susceptible cytosines to uracils within the displaced strand of R-loops. This genetic tag can subsequently be read out after PCR amplification and library construction by long read, single-molecule sequencing (SMRT-seq) (Fig. 1). Downstream data analysis steps are performed using the available Gargamel pipeline [28] to seek and capture single-molecule R-loop footprints (SMRFs). True R-loop footprints are expected to be strand-specific and sensitive to pre-treatment with Ribonuclease H [2, 16], an enzyme that specifically degrades RNA in the context of DNA:RNA hybrids [29]. By contrast, spontaneous DNA breathing or strand-separation events Experimental

Analysis

Gentle DNA extraction

CCS read generation

Non-denaturing bisulfite conversion

Remove PCR duplicates

Site specific PCR amplification

Map reads and strand assignemnt

SMRTbell library construction

Call C to T conversion tracks (peak calling)

SMRT sequencing

Cluster peak calls

Visualization

Fig. 1 Overall workflow for both experimental procedures and computational analysis

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will lead to conversion on both DNA strands in a manner insensitive to Ribonuclease H activity. Minimal length thresholds enforced during analysis serve to reinforce the distinction between characteristically long R-loops and short DNA breathing events [30]. Note that SMRF-seq can be carried out with or without preliminary R-loop enrichment with the S9.6 antibody without distortion of footprints [28]. The protocol described here does not include any S9.6 enrichment. Detailed step-by-step instructions for performing DRIP have been recently published [31]. 3.1

Cell Harvest

This section is optimized for the human Ntera-2 and HEK293 cell lines grown in culture. The following steps can be easily adapted to other cell lines. Note that SMRF-seq can be easily performed on R-loops generated upon in vitro transcription of plasmids carrying R-loop prone sequences [32, 33]. If interested in such application, carry out in vitro transcription as described [32] and proceed directly to non-denaturing bisulfite treatment (Subheading 3.3) using the products of in vitro transcription reactions as initial material. 1. Passage cells 16 h before harvesting. 2. After 16 h of growth, count cells and measure cell viability. Optimal cells count should be around 5–6 million cells per sample with >90% viable counts. Cells should be no more than 80–90% confluent at harvest. 3. Add 1.5 mL trypsin to dislodge cells and incubate for 2–5 min at 37  C until cells come off the plate. 4. Add 5 mL culture medium and pipette mix to remove clumps. 5. Transfer to a new 15 mL conical tube and spin for 3–5 min at 1000 rpm (~200 RCF) to pellet cells. 6. Aspirate media carefully without disturbing cell pellet. 7. Add 10 mL 1
DPBS and pipette mix to resuspend cells. 8. Re-spin at 200 RCF for 3–5 min to pellet cells. 9. Aspirate supernatant off without disturbing pellet.

3.2 Nucleic Acid Purification

1. Add 875 μL TE to the pelleted cells. Pipette mix ~10
to resuspend. 2. Transfer 200 μL of resuspended cells into a new 1.5 microcentrifuge tube. Up to four DNA extractions can be performed from the cell material collected. 3. Cell lysis: add the following volumes to each tube: 12.5 μL

20% SDS

10 μL

Proteinase K

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4. Invert 4–5
gently until lysed. The solution should appear clear and viscous. 5. Incubate in a 37  C water bath for 2 h to permit proteinase K digestion. Invert mix and do a brief spin down every hour to remove condensation that forms on the top of the tube. 6. Spin down 2 mL Phase Lock tube (PLT) for 1 min at 12,000 RCF. 7. Add 177.5 μL TE to each Proteinase K treated sample for a total volume of 400 μL. 8. Transfer (pouring or pipetting) Proteinase K treated cell lysate into PLT directly. 9. Add 400 μL Phenol:Choloroform:Isoamyl alcohol to lysate. Invert gently a few times (see Note 1). 10. Spin down at 12,000 RCF for 5 min at room temperature or at 4  C if possible (see Note 2). 11. Pour top aqueous layer containing nucleic acids (~400 μL) from PLT into new 2 mL tube. 12. Add the following to the 400 μL sample to precipitate nucleic acid: 40 μL

3 M sodium acetate (1:10 by volume)

1.0 mL

100% ethanol (2.5
volume)

13. Invert gently until DNA precipitate (5–10 min) (see Note 2). 14. Spool DNA using wide bore 1000 μL barrier tips and transfer to a new 1.5 mL tube while taking care not to carry over residual supernatant. Otherwise, remove as much supernatant as possible. 15. Add 1.5 mL cold 70% ethanol. Incubate 5–10 min on ice. 16. Remove supernatant and repeat 70% ethanol wash. Do not spin between washes. 17. Remove excess ethanol and air dry while tubes are inverted. This may take ~1 h depending on the size of the pellet. 18. Add 50 μL TE and incubate on ice for 30 min to resuspend DNA. Do not pipette to mix during that time as R-loops are sensitive to mechanical stress. 19. Using a wide bore 200 μL tip, pipette mix 3
. 20. Measure nucleic acid concentration using a spectrophonometer (i.e., Nanodrop). Proceed to bisulfite treatment within 24 h, keeping DNA at 20  C (see Note 3). 3.3 Non-denaturing Bisulfite Treatment (See Note 4)

1. Pipette 1.5 μg of DNA from previous isolation bring volume to 20 μL with PCR grade water. 2. Add 130 μL Lightning conversion reagent to 20 μL template. Invert mix ~3
.

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3. Incubate at 37  C in a rotisserie oven for 2 h, protected from light. 4. Add 600 μL M-binding buffer to spin column. 5. Load 150 μL bisulfite-treated sample to column with buffer. Gently invert mix a few times. 6. Spin samples at 10,000 RCF for 30 s. Discard flow-through. 7. Add 100 μL M-wash buffer to column. Spin for 30 s. 8. Add 200 μL L-Desulphonation buffer to column. 9. Incubate for 20 min at room temperature. Then spin for 30 s. Discard flow-through. 10. Add 200 μL M-wash buffer to column. Spin for 30 s. 11. Repeat wash. Air dry for 1 min. 12. Place columns in new 1.5 mL tubes. 13. Add 12.5 μL EB to elute sample. Wait 2 min. Spin for 30 s to collect samples. 14. Quantify DNA using spectrophotometer. Recovery should be around 80%. 3.4 Site-Specific Amplification (See Note 5)

1. Assemble PCR reaction in 8-strip PCR tubes as follows (see Note 6): Reagent

Vol. per reaction (μL)

ddH2O

14.75

Final concentration

5
Phusion HF/GC buffer

5.0

1


10 mM dNTPs

0.5

200 μM

Template (10 ng/μL)

2.5

25 ng

Forward + reverse primers (10 μM each)

2.0

0.8 μM

Phusion U DNA polymerase (2 U/ μL)

0.25

0.02 U/μL

Total

25.0

For difficult templates, use reaction conditions below:

Reagent

Vol. per reaction (μL)

Final concentration

ddH2O

9.75

5
Phusion HF buffer

5.0

1


10 mM dNTPs

0.5

200 μM

Template (10 ng/μL)

2.5

25 ng (continued)

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Vol. per reaction (μL)

Reagent

Final concentration

Forward + reverse primers (10 μM each)

2.0

0.8 μM

5 M Betaine

5.0

1


Phusion U DNA polymerase (2 U/ μL)

0.25

0.02 U/μL

Total

25.0

2. Perform gradient PCR using program below: (see Note 7). Stage

Temp ( C)

Time

Cycles

1. Denaturation

98

30 s

1

2. Amplification Denature Annealing Extension

98 50–72 72

10 s 30 s 2.5 min

3. Final extension

72

5 min

4. Hold

4

1

25–35

1

3. Check PCR products via agarose gel electrophoresis. The goal is to obtain a single PCR band of the correct size (Fig. 2). If a single band cannot be achieved, we advise the redesign and optimization of primers. In the event that contaminating bands cannot be completely reduced, then the dominant correct band can be excised from gels post PCR and the DNA purified subsequently. 4. Amplification of converted template. Use optimized PCR conditions as determined above. Double total reaction volume to 50 μL to increase recovery. 5. Purify products using AMPure Magbeads (see Note 8). 6. Transfer the 50 μL PCR product from the previous step into a new 1.5 mL tube. 7. Add 50 μL AMPure XP beads (1
) to the template. 8. Gently tap to mix. Briefly spin down. 9. Place samples in mini tube rotator and incubate for 10 min. 10. Briefly spin down then place tubes on DynaMag-2 magnetic rack. Wait ~1 min to allow beads migrate to the magnet and the supernatant to clear. 11. Remove supernatant carefully without pulling any beads.

Characterization of R-Loop Structures Using Single-Molecule R-Loop. . . sample sample 72°C 71.2°C

sample 69.7°C

sample 67.5°C

sample 64.8°C

sample sample 62.8°C 61.2°C

217

NTC 60°C

3.0 kb

1.0 kb

Fig. 2 Example gradient PCR with 2990 bp product. Agarose gel stained with 1
GelRed. Run for 20 min at 150 V

12. Add 1.5 mL 70% ethanol. Cap tubes and invert a few times while on the magnetic rack. Remove supernatant. 13. Repeat ethanol wash. 14. Air dry for ~1 min. Do not let the bead dry excessively, which is evident when the beads begin to crack. 15. Resuspend in 30 μL EB, remove the tubes from the magnetic rack and gently mix by tapping or flicking. Incubate with rotation for 5 min. 16. Place tubes back on the DynaMag-2 rack and wait ~1 min. Recover DNA in supernatant and place in new 1.5 mL tube. 17. Check sample concentration using a spectrophotometer. 18. Calculate number of molecules per amplicon: 0 1 mass of dsDNA ð g Þ
 A, mole dsDNA ðmolÞ ¼ @ g length of dsDNA ðbpÞ
650 mol:bp where the average weight of a DNA base-pair is 650 g/mol. 19. If pooling non-overlapping amplicons, pool equimolar amounts of each amplicon together with a total final mass no less than 500 ng for library preparation. Suggested starting for library preparation input is 500–700 ng.

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3.5 SMRT-Bell Library Preparation (See Note 9)

1. Concentrate pooled amplicons (see Note 10). 2. Add 0.8
AMPure PB magnetic beads to pooled amplicons (i.e., if starting with 100 μL pooled sample, use 80 μL AMPure PB beads). 3. Perform ethanol purification as described in previous section for AMPure bead clean-up, steps 8–16. 4. Elute in 22 μL EB. Proceed to end repair step or store in 20  C until ready to proceed to end-repair. 5. Assemble end-repair reaction on ice as follows: Reagent

Volume (μL)

Concentrated pooled amplicons

22.0

Final Conc.

10
template prep buffer

3.0

1


10 mM ATP high

3.0

1 mM

10 mM dNTP

1.2

0.4 mM

20
end repair mix

1.5

1


Total volume

30.0

6. Mix samples by gentle tapping or flicking the tube and briefly spin down. 7. Incubate samples at 25  C for 15 min in thermocycler, and then hold reaction at 4  C. 8. Purify end-repaired template using 0.8
AMPure PB beads (24 μL) and elute in 32 μL EB. This is a good stopping point. End-repaired samples can be stored at 20  C for a few days. If continuing, proceed directly to adaptor ligation through the end of library preparation. 9. For blunt end SMRTbell adaptor ligation assemble reaction on ice as follows to avoid adapter-adapter ligation: Reagent

Volume (μL)

Final Conc.

Repaired template

32.0

20 μM blunt adapter

1.0

0.5 μM

10
template prep buffer

4.0

1


1 mM ATP low

2.0

0.05 mM

Ligase (30 U/μL)

1.0

0.75 U/μL

Total volume

40.0

Gently mix before next step

Gently mix before next step

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10. Gently flick or tap to mix and spin down briefly. 11. Incubate for 1 h at 25  C in a thermocycler. Hold at 4  C for at least 1 min, up to overnight (see Note 11). 12. Inactivate ligase by incubating at 65  C for 10 min in a thermocycler. Hold at 4  C for at least 1 min. Proceed directly to exonuclease treatment. 13. Exonuclease treatment. This will remove unligated and damaged template. Set reaction on ice as follows: Reagent

Volume (μL)

Ligated template

40.0

Exo III (100 U/μL)

0.5

Exo VII (100 U/μL)

0.5

Total volume

41.0

14. Gently flick or tap to mix reaction then briefly spin down. 15. Incubate for 1 h at 37  C then hold at 4  C in a thermocycler for at least 1 min. Proceed directly to clean-up step. 16. Library Clean-up, first purification. Perform purification using 0.8
AMPure PB magnetic beads (32.8 μL). 17. Elute in 50 μL EB then proceed directly to second purification. 18. Second purification, perform the last purification using 0.8
AMPure PB magnetic beads (40 μL). 19. Elute in 12 μL EB. This will be the library to be sent for sequencing. 20. Check concentration on a spectrophotometer. Check with your PacBio sequencing provider the requirements for library submission. If the concentration is too low, repeat library preparation from pooled amplicons. 21. Proceed to quality check. Use 1 μL of library to run on a BioAnalyzer High-Sensitivity chip (see Fig. 3 for example). 3.6 SMRF-Seq Analysis Using Gargamel Pipeline (See Note 12)

1. Download SMRT Link package from https://www.pacb.com/ support/software-downloads/ (seeNote 13) and instructions (https://www.pacb.com/wp-content/uploads/SMRT_Link_ Installation_v600.pdf). CCS generation will be performed on the command line, thus GUI interface is not necessary. Perform all system checks to be able to call CCS tool. 2. Load smrtlink after opening a terminal for Circular Consensus Sequence (CCS) generation. module load smrtlink/6.0

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[FU] 450 400 350 300 200

26 48

50 0 35

100

200

300

400

500

700

2000

75 91

100

35

150

10 38 0

55 35

250

10380

[bp]

Fig. 3 Example BioAnalyzer trace of a SMRTbell library. Pooled amplicons range from 2938 to 4379 bp

3. Run CCS tool. Depending on file size, this may take several hours to several days depending on compute capacity and file size. Input file will be under Primary Analysis folder annotated as “subreads.bam”. Default parameters are --minPasses ¼ 3 and --minPredictedAccuracy ¼ 0.9. ccs

4. Removing PCR duplicates using dedupe2.sh from BBMap package v37.90 (https://sourceforge.net/projects/bbmap/). Thresholds had been optimized for this application specifically. dedupe2.sh in=ccs.fq out=deduplicated.fq e=30 mid=98 k=31 nam=4

5. Analyze data using Gargamel pipeline. This is an open source software designed to call R-loop peaks and visualize patterns of C to T conversions. General usage for this pipeline is available at: https://github.com/srhartono/footLoop/blob/master/ README.md 6. Download Gargamel pipeline: git

clonehttps://github.com/srhartono/

footLoop

Required packages for this pipeline are listed in the README.md document. 7. Map and assign reads using footLoop (Bismark) (see Note 13). This will require an index file and the reference genome to be mapped to (see Fig. 4 for example). footLoop.pl

–r

-n

-l -i

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Fig. 4 Example index file. Columns are as follows: chr, start, end, gene, 0, strand. Tab delimited and no header format

8. Call C to T conversion tracks (footPeak) (see Note 14). footPeak.pl –n –o –w 20 –t 0.55–l 100

9. Generate peak clusters (footClust). footClust.pl –n

10. Visualize tracks by generating png files (footPeak_graph) (see Note 15). footPeak_graph.pl –n -r 1

(See Fig. 5 for an example) 11. Generate Genome browser tracks (footPeakGTF). footPeak_GTF.pl –n

This script will generate GTF files that can be directly uploaded to USCS Genome Browser. Example output is shown in Fig. 6.

4

Notes 1. Wear appropriate PPE when handling Phenol:Choloroform: Isoamyl alcohol as it is considered a hazardous material. Dispense in a chemical hood. Dispose of waste in designated sealed container. 2. Keeping reagents and spins cold will help with nucleic acid precipitation. When precipitating nucleic acids, DNA should be transparent after ethanol addition. If nucleic acid looks “milky”, continue to invert gently and incubate for another

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A

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Fig. 5 Example footprinted region with PNG output after footPeak_graph.pl run. (a) SNRPN70 footprinted region (hg19) and DRIPc-seq data with (+) and () strands shown in red and blue, respectively. (b) Example output showing random 200 peaks reads on the non-template strand. (c) Example output showing random 100 reads for template strand. For panels (b) and (c), each horizontal line corresponds to one read. Red tick lines indicate converted Cs within a called peak, yellow indicates non-converted Cs, green indicates converted Cs not called a peak; gray is missing/ambiguous sequence

5–10 min. If the cloudiness does not go away, repeat the phenol organic extraction or repeat isolation. Do not vortex samples. 3. To preserve R-loop structures, keep temperatures low. Handle with care, avoid vortexing and unnecessary pipetting. R-loops are sensitive to mechanical stress and can spontaneously fall apart, reducing yields as a result. It is particularly essential to avoid any nicking or fragmentation of the displaced ssDNA strand as this will prevent the subsequent PCR amplification of that strand and compromise the recovery of information derived from that strand.

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Fig. 6 Genome Browser snapshot of same SNRPN70 terminal region in previous figure. Converted cytosines within called peaks are annotated as red tick marks on the positive strand (blue tick marks for footprints called in the negative strand). Each horizontal line is a single molecule read mapping to the region

4. Conditions specified in the methylation kit were modified to enable non-denaturing R-loop footprinting. These modifications include omitting the initial denaturation step to ensure that only intrinsically single-stranded regions are targeted, and lowering the temperature to 37  C during bisulfite treatment. In addition, treatment time was lowered to 2 h. These modifications may lead to a slight reduction in the efficiency of conversion compared to that typically required for CpG methylation profiling. We encourage users to use denatured spike-in controls to directly measure the conversion efficiency. Based on our data [28], conversion efficiencies are routinely in the 80–90% range, which is sufficient to allow for R-loop footprinting. Conversion efficiency is also influenced by the amount of nucleic acids to be treated, with an optimal recommended amount of 200–500 ng of input DNA, up to a maximum of 2 μg. The amount of input DNA also determines the number of PCR reactions that can be carried out post-treatment. Each PCR reaction optimally uses 25–50 ng of bisulfite-converted DNA to ensure the recovery of a diverse set of R-loop footprints. The amount of input DNA recommended here to go into the bisulfite reaction (1.5 μg) ensures good conversion efficiency and a strong recovery yield that permits at least 20 downstream PCR reactions. 5. Native primers are designed to hybridize to dsDNA regions flanking the candidate R-loops under study to ensure that no selection or distortion of R-loop patterns is introduced (Fig. 7). This is distinct from past protocols [16], where

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CpG Islands SNRNP70 SNRNP70 80 –

DRIPc-seq (+)

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Fig. 7 Example target region in SNRPN70. Red bar indicates possible target region to amplify. Blue arrow indicates forward and reverse primer sites flanking peaks of DRIPc-seq signal (below)

converted primers that hybridize directly to the C to T converted strand were used to increase the recovery of R-loop structures. Typical amplicon lengths range from 2 to 5 kb, which permits efficient PacBio sequencing. For long-range amplicons (>1 kb), PCR optimization may be necessary especially for high GC content or repetitive regions. 6. Due to the high-throughput capacity of current PacBio sequencing instruments (4–500,000 reads on PacBio Sequel), it is recommended to pool multiple samples in one sequencing reaction. When pooling amplicons, we recommend that amplicons be in the same size range and suggest pooling non-overlapping genomic regions that can be uniquely mapped. In this way, barcoding is not necessary as mapping can easily be performed against the reference amplicons. In case users interested in mapping R-loops observed the same amplicon under varied conditions, barcoding is required. Barcoding is not covered in this protocol and users are referred to available technical information from PacBio (www.pacb.com). 7. When optimizing primers, use DNA from cell line to be used. In general, use HF buffer. For high GC content regions, use GC buffer. For difficult templates, use HF or GC buffer with betaine as a PCR additive. For gradient PCR, set a range of 7  C from the expected primer annealing temperatures to determine the optimum in terms of efficiency and specificity. It is preferable to perform the least possible number of cycles to decrease PCR duplicates and increase molecular diversity. 8. Avoid unnecessary pipetting to prevent damaging or fragmenting PCR products, especially for longer amplicons. Make sure that AMPure beads are at room temperature to avoid sample loss. Use freshly made 70% ethanol when performing bead washes. 9. The SMRTbell template library preparation presented here is for version 1 kits and adapted from PacBio’s “Procedure & Checklist—2 kb Template Preparation and Sequencing” (PN 001-143-835-08). Estimated recovery after library prep is 20–30%. If preparing the library for the first time, we suggest starting with ~1 μg template and adjust adapter concentration (see Note 11). Newer version kits have different steps and may

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use a different set of primers for sequencing. Make sure to notify your PacBio sequencing provider which version of template prep kit you are using. For amplicons less than 5 kb in length, 10-h movie times should suffice to get at least 3 minimum DNA polymerase pass. Note that longer movie times can yield higher quality sequencing reads. 10. Optimize amplicons to avoid wide ranges in lengths when pooling amplicons for sequencing. In general, smaller fragments are more efficiently sequenced, leading to underrepresentation of longer fragments. Small fragment contaminations, such as from primer dimers, will soak up sequencable reads. Performing an additional AMPure clean-up, reducing the ratio of AMPure beads to DNA to 0.6–0.8
can help remove smaller fragments. See a recent protocol for more detailed instructions on how to perform AMPure cleanups [31]. Be sure to use AMPure BP beads during SMRTbell library preparation. Using generic AMPure beads may result in sequencing failure. 11. In the original PacBio protocol, ligation incubation can be extended to overnight. Extended incubation and increased input DNA may result in increased chimeric ligation products (double-insert templates), which can be checked on BioAnalyzer trace. If this happens, adjust adapter concentration to 30
(up to 50
) molar excess. 12. Further description of this analysis can be found here [28] and the GitHub page (listed in Subheading 3.6, step 1). This analysis pipeline can be computationally intensive depending on the size of data. We suggest using a high-performance computer cluster to submit jobs. It is highly suggested to use terminal multiplexer such as Screen or TMUX before starting a job. A newer version of SMRTLink v7 is available for download; this version should be compatible with previous versions. Software downloads can be requested from this link: https:// www.pacb.com/support/software-downloads/ 13. Thresholds are derived from [28]. The pipeline only considers reads if they are 95% of the total expected amplicon length. This may be too stringent for some purposes and can be reduced. 10 bp buffer flanking the start and end of amplicons are used to improve mapping as indicated by –x and –y. For the –l option, use descriptive alphanumerical values like ‘Experiment1’ to clearly annotate the outputs of successive data analyses. 14. Peak calling is further described [28] and the GitHub page (listed in Subheading 3.6, step 1). The default threshold calls for at least 55% C to T conversion per window composed of 20 cytosines. A minimum 100 bp length was imposed. These

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thresholds can be varied by users. A key indicator that the threshold is working is that conversion patterns should be strand-specific for R-loops. 15. This script can create PNG and PDF files by using –r 1 and –R 1 options respectively. Option 1 creates files for the relevant data each parsed in its own appropriate directory (i.e., ‘peak’ and ‘no peak’ reads, ‘template’ and ‘non-template’ strand reads). Four relevant directories are generated. The ‘PEAK’ directory contains files for peak-containing reads from the non-template strand (i.e., converted looped-out DNA strand). The ‘NOPK’ directory contains no-peak reads from the non-template strand. The ‘PEAK_TEMP’ and ‘NOPK_TEMP’ contain the template strand information. File names will contain “CH” and “GH” annotations indicating positive and negative strand, respectively.

Acknowledgements We thank Chedin lab members for useful discussions, Dr. Lionel A. Sanz for constructive comments on the manuscript, and Dr. Stella R. Hartono for developing the Gargamel analysis pipeline. This work was funded by the National Institutes of Health (Grant R01 GM120607 to F.C.) and was supported, in part, by National Science Foundation Graduate Research Fellowship (Grant 1650042 to M.M.) and National Institute of General Medical Sciences Biomolecular Technology Predoctoral T32 Training Program (Grant T32-GM008799 to M.M.). References 1. Ginno PA, Lott PL, Christensen HC, Korf I, Chedin F (2012) R-loop formation is a distinctive characteristic of unmethylated human CpG island promoters. Mol Cell 45(6):814–825. https://doi.org/10.1016/j.molcel.2012.01. 017 2. Sanz LA, Hartono SR, Lim YW, Steyaert S, Rajpurkar A, Ginno PA, Xu X, Chedin F (2016) Prevalent, dynamic, and conserved R-loop structures associate with specific epigenomic signatures in mammals. Mol Cell 63 (1):167–178. https://doi.org/10.1016/j. molcel.2016.05.032 3. Wahba L, Costantino L, Tan FJ, Zimmer A, Koshland D (2016) S1-DRIP-seq identifies high expression and polyA tracts as major contributors to R-loop formation. Genes Dev 30 (11):1327–1338. https://doi.org/10.1101/ gad.280834.116

4. El Hage A, Webb S, Kerr A, Tollervey D (2014) Genome-wide distribution of RNA-DNA hybrids identifies RNase H targets in tRNA genes, retrotransposons and mitochondria. PLoS Genet 10(10):e1004716. https://doi.org/10.1371/journal.pgen. 1004716 5. Hartono SR, Malapert A, Legros P, Bernard P, Chedin F, Vanoosthuyse V (2018) The affinity of the S9.6 antibody for double-stranded RNAs impacts the accurate mapping of R-loops in fission yeast. J Mol Biol 430 (3):272–284. https://doi.org/10.1016/j. jmb.2017.12.016 6. Xu W, Xu H, Li K, Fan Y, Liu Y, Yang X, Sun Q (2017) The R-loop is a common chromatin feature of the Arabidopsis genome. Nat Plants 3(9):704–714. https://doi.org/10.1038/ s41477-017-0004-x

Characterization of R-Loop Structures Using Single-Molecule R-Loop. . . 7. Zaitsev EN, Kowalczykowski SC (2000) A novel pairing process promoted by Escherichia coli RecA protein: inverse DNA and RNA strand exchange. Genes Dev 14(6):740–749 8. Wahba L, Gore SK, Koshland D (2013) The homologous recombination machinery modulates the formation of RNA-DNA hybrids and associated chromosome instability. elife 2: e00505. https://doi.org/10.7554/eLife. 00505 9. Kasahara M, Clikeman JA, Bates DB, Kogoma T (2000) RecA protein-dependent R-loop formation in vitro. Genes Dev 14(3):360–365 10. Chedin F (2016) Nascent connections: R-loops and chromatin patterning. Trends Genet 32(12):828–838. https://doi.org/10. 1016/j.tig.2016.10.002 11. Santos-Pereira JM, Aguilera A (2015) R loops: new modulators of genome dynamics and function. Nat Rev Genet 16(10):583–597. https:// doi.org/10.1038/nrg3961 12. Costantino L, Koshland D (2015) The yin and Yang of R-loop biology. Curr Opin Cell Biol 34:39–45. https://doi.org/10.1016/j.ceb. 2015.04.008 13. Crossley MP, Bocek M, Cimprich KA (2019) R-loops as cellular regulators and genomic threats. Mol Cell 73(3):398–411. https://doi. org/10.1016/j.molcel.2019.01.024 14. Chen L, Chen JY, Zhang X, Gu Y, Xiao R, Shao C, Tang P, Qian H, Luo D, Li H, Zhou Y, Zhang DE, Fu XD (2017) R-ChIP using inactive RNase H reveals dynamic coupling of R-loops with transcriptional pausing at gene promoters. Mol Cell 68(4):745–757 . e745. https://doi.org/10.1016/j.molcel. 2017.10.008 15. Chen PB, Chen HV, Acharya D, Rando OJ, Fazzio TG (2015) R loops regulate promoterproximal chromatin architecture and cellular differentiation. Nat Struct Mol Biol 22 (12):999–1007. https://doi.org/10.1038/ nsmb.3122 16. Yu K, Chedin F, Hsieh CL, Wilson TE, Lieber MR (2003) R-loops at immunoglobulin class switch regions in the chromosomes of stimulated B cells. Nat Immunol 4(5):442–451. https://doi.org/10.1038/ni919 17. Wiedemann EM, Peycheva M, Pavri R (2016) DNA replication origins in immunoglobulin switch regions regulate class switch recombination in an R-loop-dependent manner. Cell Rep 17(11):2927–2942. https://doi.org/10. 1016/j.celrep.2016.11.041 18. Skourti-Stathaki K, Proudfoot NJ, Gromak N (2011) Human senataxin resolves RNA/DNA hybrids formed at transcriptional pause sites to

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promote Xrn2-dependent termination. Mol Cell 42(6):794–805. https://doi.org/10. 1016/j.molcel.2011.04.026 19. Proudfoot NJ (2016) Transcriptional termination in mammals: stopping the RNA polymerase II juggernaut. Science 352(6291): aad9926. https://doi.org/10.1126/science. aad9926 20. Stork CT, Bocek M, Crossley MP, Sollier J, Sanz LA, Chedin F, Swigut T, Cimprich KA (2016) Co-transcriptional R-loops are the main cause of estrogen-induced DNA damage. Elife 5:e17548. https://doi.org/10.7554/ eLife.17548 21. Sollier J, Cimprich KA (2015) Breaking bad: R-loops and genome integrity. Trends Cell Biol 25(9):514–522. https://doi.org/10.1016/j. tcb.2015.05.003 22. Aguilera A, Garcia-Muse T (2012) R loops: from transcription byproducts to threats to genome stability. Mol Cell 46(2):115–124. https://doi.org/10.1016/j.molcel.2012.04. 009 23. Richard P, Manley JL (2017) R loops and links to human disease. J Mol Biol 429 (21):3168–3180. https://doi.org/10.1016/j. jmb.2016.08.031 24. Groh M, Gromak N (2014) Out of balance: R-loops in human disease. PLoS Genet 10(9): e1004630. https://doi.org/10.1371/journal. pgen.1004630 25. Boguslawski SJ, Smith DE, Michalak MA, Mickelson KE, Yehle CO, Patterson WL, Carrico RJ (1986) Characterization of monoclonal antibody to DNA.RNA and its application to immunodetection of hybrids. J Immunol Methods 89(1):123–130 26. Phillips DD, Garboczi DN, Singh K, Hu Z, Leppla SH, Leysath CE (2013) The sub-nanomolar binding of DNA-RNA hybrids by the single-chain Fv fragment of antibody S9.6. J Mol Recognit 26(8):376–381. https://doi.org/10.1002/jmr.2284 27. Vanoosthuyse V (2018) Strengths and weaknesses of the current strategies to map and characterize R-loops. Noncoding RNA 4(2): E9. https://doi.org/10.3390/ncrna4020009 28. Malig M, Hartono SR, Giafaglione JM, Sanz LA, Chedin F (2019) High-Throughput Single-Molecule R-loop Footprinting Reveals Principles of R-loop Formation. bioRxiv:640094 https://doi.org/10.1101/ 640094 29. Cerritelli SM, Crouch RJ (2009) Ribonuclease H: the enzymes in eukaryotes. FEBS J 276 (6):1494–1505

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30. Kouzine F, Wojtowicz D, Baranello L, Yamane A, Nelson S, Resch W, Kieffer-Kwon KR, Benham CJ, Casellas R, Przytycka TM, Levens D (2017) Permanganate/S1 nuclease Footprinting reveals non-B DNA structures with regulatory potential across a mammalian genome. Cell Syst 4(3):344–356.e347. https://doi.org/10.1016/j.cels.2017.01.013 31. Sanz LA, Chedin F (2019) High-resolution, strand-specific R-loop mapping via S9.6-based DNA-RNA immunoprecipitation and highthroughput sequencing. Nat Protoc 14 (6):1734–1755. https://doi.org/10.1038/ s41596-019-0159-1

32. Stolz R, Sulthana S, Hartono SR, Malig M, Benham CJ, Chedin F (2019) Interplay between DNA sequence and negative superhelicity drives R-loop structures. Proc Natl Acad Sci U S A 116(13):6260–6269. https://doi. org/10.1073/pnas.1819476116 33. Carrasco-Salas Y, Malapert A, Sulthana S, Molcrette B, Chazot-Franguiadakis L, Bernard P, Chedin F, Faivre-Moskalenko C, Vanoosthuyse V (2019) The extruded non-template strand determines the architecture of R-loops. Nucleic Acids Res 47 (13):6783–6795. https://doi.org/10.1093/ nar/gkz341

Chapter 16 Analysis of RNA–DNA Triplex Structures In Vitro and In Vivo Anna Postepska-Igielska, Alena Blank-Giwojna, and Ingrid Grummt Abstract RNA can bind within the major groove of purine-rich DNA via Hoogsteen base pairing and form a triple helical RNA–DNA structure that anchors the RNA to specific DNA sequences, thereby targeting RNA-associated regulatory proteins to distinct genomic sites. Here we present methods to analyze the potential of a given RNA to form triplexes in vitro and to validate these structures in vivo. Key words Noncoding RNA, RNA–DNA triplexes, Electrophoretic mobility shift assay, Biotinylated RNA, Triplex capture, Psoralen, Pull-down

1

Introduction Thousands of long-noncoding RNAs (lncRNAs) are prevalent in metazoan transcriptomes. Yet, while the importance of lncRNAs as versatile regulators of cellular processes in health and disease is increasingly recognized, fundamental aspects regarding their mode of action remain poorly understood. The ability of RNA to interact with proteins, DNA, or other RNAs provides an immense regulatory potential. Serving as molecular signals, guides, or scaffolds, lncRNAs target transcription regulators to specific genomic sites. Base complementarity allows RNA to interact with doublestranded DNA either by canonical Watson-Crick pairing with one of the DNA strands, forming R-loop structures, or by direct binding to DNA, forming RNA–DNA triplexes. RNA can bind within the major groove of DNA via forward (parallel) or reverse (antiparallel) Hoogsteen hydrogen bonds between the purine-rich strand of duplex DNA and single-stranded RNA (Fig. 1), [1]. Although the ability of DNA to engage in triplex structures with RNA is well established, the in vivo existence and biological relevance of these structures remain largely unknown. Notably, sequences with triplex-forming potential are overrepresented at regulatory gene regions, such as promoters and enhancers, suggesting that RNA–DNA triplex formation may represent a general

Ulf Andersson Vang Ørom (ed.), RNA-Chromatin Interactions: Methods and Protocols, Methods in Molecular Biology, vol. 2161, https://doi.org/10.1007/978-1-0716-0680-3_16, © Springer Science+Business Media, LLC, part of Springer Nature 2020

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Fig. 1 Purine-rich double-stranded DNA is able to accommodate RNA in the major groove, forming a triple helical structure. Depending on the sequence, RNA can bind either in parallel orientation (UC or UG motifs in the RNA, left) or in anti- parallel orientation (AG or UG motifs, right)

mechanism for lncRNA-mediated recognition of genomic target sites [2, 3]. Examples of RNA–DNA triplexes formed by lncRNA with specific DNA sequences include the nucleolar lncRNAs pRNA and PAPAS, which repress transcription of rRNA genes (rDNA) by targeting DNMT3b [4] and the CHD4/NuRD co-repressor [5] to the rDNA promoter, respectively. Other examples include Fendrr, which regulates transcription of developmental genes by recruiting the PRC2 complex [6]; MEG3, which guides PRC2 to TGF-β-responsive genes [7]; and KHPS1, which activates expression of SPHK1 (Sphingosine kinase 1) via recruitment of KHPS1associated p300 [8, 9]. Furthermore, PARTICLE and HOTAIR as well as some miRNAs were shown to bind to DNA and regulate expression of specific target genes [10–12]. These studies support that tethering RNA to specific genomic sites guides RNA-associated proteins to regulatory sequences to establish an epigenetic landscape that facilitates or inhibits gene expression [13]. Rigorous proof for the existence and physiological relevance of RNA–DNA triplexes requires experimental set-ups that enable isolation and functional validation of these structures. In this chapter we provide detailed methods and notes for assessment of the triplex forming potential of a given RNA in vitro and for subsequent validation in vivo [4, 5, 8, 9]. Together with a recently developed method to isolate and sequence RNA–DNA triplexes genome-wide [14], we now have an experimental toolbox to catalogue and characterize cellular RNA–DNA triplexes, which is a prerequisite for unraveling their function. The electrophoretic mobility shift assay (EMSA) relies on the altered electrophoretic mobility of double-stranded DNA upon binding to RNA. After incubation of purine-rich radiolabelled DNA with a molar excess of complementary RNA, the formation of RNA–DNA complexes is monitored by electrophoresis in a polyacrylamide gel. RNA–DNA triplexes migrate slower than unbound double-stranded DNA (Fig. 2). Importantly, as single-

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Fig. 2 (a) Cartoon illustrating retardation of the electrophoretic mobility of purine-rich duplex DNA by triplex formation with a complementary RNA. (b) Increasing amounts of radiolabelled synthetic RNA were incubated with a double-stranded DNA oligonucleotide, resolved on a polyacrylamide gel, and analyzed by autoradiography (reproduced from [8]). (c) Fluorescently labelled RNA (green) and duplex DNA (red) were used to form triplex structures as in (b)

stranded DNA (ssDNA)–RNA hybrids also retard the electrophoretic mobility, negative controls and enzymatic digestions should be included to substantiate that the interaction observed is due to triplex formation. In vitro triplex pull-down assay relies on binding of biotinylated RNA to DNA followed by capture of biotin-modified RNA-DNA triplexes on streptavidin beads. Triplex formation is validated by incubating the same RNA with a 7-deaza-purine modified DNA fragment that does not support Hoogsteen base pairing (Fig. 3). The in vivo triplex capture assay is used to examine triplex formation of the RNA of interest with specific DNA sequences in cells. To this end, biotinylated RNA is transfected into cells, and binding to genomic target sequence is monitored upon deproteinization, DNA fragmentation, and binding to streptavidin beads (Fig. 4). The major challenge for reliable detection of triplex formation in vivo is to prove that the interaction between DNA and RNA is direct. In the UV-assisted triplex capture assay, cells are transfected with oligoribonucleotides that carry a psoralen moiety at the 50 -end and a biotin residue at the 30 -end. The presence of psoralen allows for fixation of direct nucleic acid interactions by photoactivation without crosslinking proteins to DNA or RNA (Fig. 5).

2 2.1

Materials EMSA

1. Commercially synthesized gene-specific forward primers and reverse primers fused to the T7 promoter sequence (see Note 1). 2. PCR reaction mix containing a thermostable DNA polymerase.

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Synthetic purine-rich DNA

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Fig. 3 Outline of the in vitro triplex pull-down assay. Biotinylated RNA is incubated with either a DNA fragment containing a purine-rich triplex forming sequence (left) or with a 7-deaza-modified DNA fragment (right). RNA–DNA triplexes are captured on streptavidin-coated magnetic beads. Upon RNase A digestion, recovered DNA is analyzed by qPCR

3. PCR clean-up kit. 4. Gel extraction kit. 5. Synthetic RNA Transcription kit.

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6. RNA clean-up kit. 7. 100 μM HPLC-purified DNA oligonucleotides, commercially synthesized, dissolved in DNase-free H2O. Store in aliquots at 20  C. 8. 3000 Ci/mmol [γ-32P] labelled ATP. 9. 10 U/μl T4 polynucleotide kinase (PNK). 10. Nucleotide removal kit.

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Fig. 4 Outline of the in vivo triplex capture assay. Biotinylated RNA is transfected into cells. After nuclear lysis, deproteinization, and DNA fragmentation, RNA-associated DNA is captured on streptavidin-coated magnetic beads. Upon RNase A digestion, recovered DNA is analyzed by qPCR Transfection of a 5‘-psoralen and 3‘-biotin-labelled RNA oligonucleotide

Irradiation of cells with UV light (365 nm) to crosslink psoralen to DNA

Bio

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Fig. 5 Scheme outlining the UV-assisted triplex capture assay

11. Phosphatase inhibitor. 12. 5 triplex buffer: 200 mM Tris–acetate, pH 7.5, 100 mM KCl, 50 mM magnesium–acetate, 50% glycerol, 5 phosphatase inhibitor (see Notes 2 and 3). 13. TE buffer (10 mM Tris–HCl, 1 mM EDTA, pH 8.0) (see Note 3). 14. 40% acrylamide/bisacrylamide stock solution (37.5:1). 15. 2 M Tris–acetate, pH 7.5.

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16. 2 M magnesium acetate. 17. 10% sodium dodecyl sulphonate (SDS). 18. 10% ammonium persulfate (APS). 19. Tetramethyl-ethylenediamine (TEMED) stored at 4  C. 20. Buffers for RNA and DNA electrophoresis, DNA and RNA staining reagent. 21. Spectrophotometer. 22. Thermoblock. 23. Thermocycler. 24. Standard horizontal electrophoresis apparatus. 25. Table top centrifuge. 26. 1.5 ml DNA low-binding tubes. 27. UV table for preparative nucleic acid work. 28. Glass plates, spacers, vertical electrophoresis equipment, binder clips. 29. Gel dryer. 30. Phosphorimager. 31. Phosphor screen and storage cassette. 32. Scintillation counter. 2.2 In Vitro Triplex Pull-Down Assay

1. Biotin-16-UTP. 2. Commercially synthesized gene-specific forward primers and reverse primers fused to the T7 promoter sequence (see Note 1). 3. PCR reaction mix containing a thermostable DNA polymerase. 4. PCR clean-up kit. 5. Gel extraction kit. 6. Synthetic RNA generated with MEGAscript T7 Transcription kit (see Note 4). 7. RNA clean-up kit. 8. 7-deaza-dATP. 9. 7-deaza-dGTP. 10. PwoSuperYield DNA polymerase Kit. 11. Ribonuclease Inhibitors. 12. 10 mg/ml Ribonuclease A (RNase A). 13. MyOne Streptavidin C1 Dynabeads (see Note 5). 14. 20 U/μl Exonuclease I (Exo I). 15. Oligonucleotides for validation of RNA-associated DNA by qPCR (see Note 6).

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16. SYBR Green PCR kit (see Note 7). 17. Forward and reverse primers that flank the region involved in the DNA–RNA interaction to generate the DNA fragments (see Note 8). 18. 10 triplex buffer: 100 mM Tris–HCl, pH 7.5, 200 mM KCl, 100 mM MgCl2, 0.5% Tween 20, ribonuclease inhibitors 10x (see Notes 2 and 3). 19. Washing buffer 1: 150 mM KCl, 10 mM Tris–HCl, pH 7.5, 5 mM MgCl2, 0.5% NP-40, ribonuclease inhibitors (see Note 3). 20. Washing buffer 2: 15 mM KCl, 10 mM Tris–HCl, pH 7.5, 5 mM MgCl2, ribonuclease inhibitors (see Note 3). 21. TE buffer: 10 mM Tris–HCl, pH 8, 1 mM EDTA (see Note 3). 22. Buffers for RNA and DNA electrophoresis. 23. DNA staining reagent. 24. RNA staining reagent. 25. Spectrophotometer. 26. Thermoblock. 27. Thermocycler. 28. Standard horizontal electrophoresis apparatus. 29. Table top centrifuge. 30. 1.5 ml DNA low-binding tubes. 31. UV table for preparative nucleic acid work. 32. Magnetic separation rack. 33. Tube rotator with shaking function. 34. Real-time PCR ´ınstrument. 2.3 In vivo Triplex Capture Assay with Biotinylated RNA

1. TransIT-mRNA Transfection kit for Large RNA transfections (Mirus Bio); (see Note 9). 2. 10 mg/ml Proteinase K. 3. 10 mg/ml Ribonuclease A (RNase A). 4. Streptavidin C1 Dynabeads (see Note 5). 5. PCR purification kit. 6. SYBR Green PCR kit (see Note 7). 7. Oligonucleotides for validation of RNA-associated DNA by qPCR (see Note 6). 8. Biotinylated RNA generated with MEGAscript T7 transcription kit (see Notes 1 and 4). 9. 10 PBS: 2.7 M NaCl, 53.7 mM KCl, 20 mM Na2HPO4, 29.4 mM KH2PO4, pH 7.4. Prepare 1 working solution with autoclaved millipore water. Cool down to 4  C before use.

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10. Nuclei isolation buffer: 10 mM KCl, 10 mM Hepes, pH 8.0, 5 mM MgCl2, 0.34 M sucrose, 10% glycerol, ribonuclease inhibitors. Ice-cold (see Note 3). 11. Triplex buffer: 10 mM Tris–HCl, pH 7.5, 20 mM KCl, 10 mM MgCl2, ribonuclease inhibitors (see Notes 2 and 3). 12. Washing buffer 1: 150 mM KCl, 10 mM Tris–HCl, pH 7.5, 5 mM MgCl2, 0.5% NP-40, ribonuclease inhibitors (see Note 3). 13. Washing buffer 2: 15 mM KCl, 10 mM Tris–HCl, pH 7.5, 5 mM MgCl2, ribonuclease inhibitors (see Note 3). 14. TE buffer: 10 mM Tris–HCl, pH 8.0, 1 mM EDTA, pH 7.5 (see Note 3). 15. Buffers for RNA and DNA electrophoresis. 16. DNA staining reagent. 17. RNA staining reagent. 18. Spectrophotometer. 19. Thermoblock. 20. Thermocycler. 21. Standard horizontal electrophoresis apparatus. 22. Table top centrifuge. 23. 1.5 ml DNA low-binding tubes. 24. UV table for preparative nucleic acid work. 25. Magnetic separation rack. 26. Tube rotator with shaking function. 27. Real-time PCR machine. 28. Sonicator and corresponding sonication tubes. 29. Cell culture incubator (37  C, 5% CO2). 2.4 UV-Assisted Triplex capture Assay

1. TransIT-mRNA Transfection kit for Large RNA transfections (Mirus Bio); (see Note 9). 2. 10 mg/ml Proteinase K. 3. 10 mg/ml Ribonuclease A (RNaseA). 4. Streptavidin C1 Dynabeads (see Note 5). 5. PCR clean-up kit. 6. QuantiTect SYBR Green PCR kit (see Note 7). 7. Oligonucleotides for validation of RNA-associated DNA by qPCR (see Note 6). 8. Commercially synthesized 50 -psoralen and 30 -biotin-modified RNA oligonucleotides (see Note 10).

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9. 10 PBS: 2.7 M NaCl, 53.7 mM KCl, 20 mM Na2HPO4, 29.4 mM KH2PO4, pH 7.4. Prepare 1 working solution with autoclaved millipore water. 10. Nuclei isolation buffer: 10 mM KCl, 10 mM Hepes, pH 8.0, 5 mM MgCl2, 0.34 M sucrose, 10% glycerol, ribonuclease inhibitors (see Note 3). 11. Nuclei crosslinking buffer: 10 mM Tris–HCl, pH 7.5, 0.88 M sucrose, 5 mM MgCl2, ribonuclease inhibitors (see Note 3). 12. Triplex buffer: 10 mM Tris–HCl, pH 7.5, 50 mM NaCl, 10 mM MgCl2, ribonuclease inhibitors (see Note 3). 13. Washing buffer 1: 10 mM Tris–HCl, pH 7.5, 150 mM NaCl, 5 mM MgCl2, 0.5% NP-40, ribonuclease inhibitors (see Note 3). 14. Washing buffer 2: 10 mM Tris–HCl, pH 7.5, 15 mM NaCl, 5 mM MgCl2, ribonuclease inhibitors (see Note 3). 15. TE buffer: 10 mM Tris–HCl, pH 8, 1 mM EDTA, pH 7.5 (see Note 3). 16. Buffers for DNA electrophoresis, DNA staining reagent. 17. Appliances listed in Subheading 2.3, steps 18, 21, 22 and 24–28. 18. Longwave Ultraviolet Crosslinker (365 nm).

3 3.1

Methods EMSA

3.1.1 Preparation of Transcription Templates

For all steps, use ultra-pure DNase and RNase-free H2O. 1. Carry out standard Notes 1, 4 and 11):

PCR reactions

as indicated

(see

2 GoTaq PCR master mix

100 μl

Forward primer, 20 μM

4 μl

Reverse primer, 20 μM

4 μl

Genomic DNA (cDNA or a linearized vector harboring a cloned DNA fragment can also be used)

30 ng

RNase and DNase-free H2O

to 200 μl

2. Resolve the PCR electrophoresis.

product

by

standard

agarose

gel

3. Excise and purify the DNA fragment with a gel extraction kit. 4. Determine DNA concentration by spectrophotometry (see Note 12).

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3.1.2 RNA Synthesis

1. Set up 20 μl transcription reactions using MEGAscript T7 Transcription Kit containing: 10 transcription buffer

2 μl

ATP, CTP, GTP, UTP

2 μl each

T7 RNA Polymerase Enzyme Mix

2 μl

DNA template

30 ng

RNase and DNase-free H2O

to 20 μl

2. Incubate in a thermocycler for 6–12 h at 37  C. 3. Digest the DNA template by incubation with 2 μl of TURBO DNase for 15 min at 37  C. 4. Purify RNA with an RNA clean-up kit. 5. Determine RNA concentration by spectrophotometry. 6. Assess the size and purity of RNA by standard agarose gel electrophoresis. 3.1.3 Radiolabelling of DNA Oligonucleotides

All procedures should be carried out in a restricted area allowing work with radioactive material (see Note 13). 1. A 20 μl reaction contains (see Note 14): DNA oligonucleotide (20 pmol/μl)

2 μl

10 PNK reaction buffer

2 μl

ATP-(γ- P) (3000 Ci/mmol)

3 μl

T4 polynucleotide kinase (10 U/μl)

1 μl

RNase and DNase-free H2O

to 20 μl

32

2. Incubate the reaction for 1 h at 37  C in a thermoblock. 3. Purify the DNA oligonucleotide with a nucleotide removal kit, elute in 50 μl. 4. Determine specific activity with a scintillation counter; between 5  105 and 1  106 cpm/μl should be achieved. 5. Proceed immediately or store aliquots at 20  C. 3.1.4 Oligonucleotide Duplex Formation

1. Anneal 30 μl of purified radiolabelled oligonucleotide (about 24 pmol) with equimolar amounts of the complementary unlabelled oligonucleotide in 100 μl of 10 mM Tris–acetate, pH 7.5. Mix by gentle pipetting. 2. Incubate for 10 min each at 95  C, 65  C, 55  C, 45  C, 35  C, 25  C, 15  C, and hold at 4  C. 3. Place on ice or store aliquots at 20  C.

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1. Each reaction contains 1 μl 5 triplex buffer, 0.1 μl phosphatase inhibitor 50x, and 0.9 μl H2O. Prepare master mix and pipette 2 μl into 1.5 ml DNA low-binding tubes on ice. 2. Add 1 μl (about 0.3 pmol, equivalent of 3–5  105 cpm) of labelled duplex oligonucleotide and 2 μl of synthetic RNA corresponding to a 10–100 molar excess. The total reaction volume is 5 μl. 3. Incubate at 23  C for 1 h (see Note 15). 4. Load reaction on a vertical 10% Tris–acetate/MgCl2 polyacrylamide gel (see Note 16). Run the gel at room temperature for about 2 h at 100 V and dry at 85  C for 1 h (see Note 17). 5. Expose dried gel on a screen for at least 12 h and visualize by phosphorimaging.

3.2 In Vitro Triplex Pull-Down Assay

1. Prepare transcription templates by PCR as described in Subheading 3.1.1.

3.2.1 Synthesis of Biotinylated RNA

2. Perform in vitro transcription reactions as described in Subheading 3.1.2, steps 1–5, but replace 40% of total UTP by biotin-16-UTP (see Note 4). 3. Assess the quality of RNA by standard agarose gel electrophoresis. Only RNA of the correct size should be used. RNA aliquots (1 pmol/μl) should be stored at 80  C (see Note 18).

3.2.2 Generation of DNA Fragments

1. Prepare DNA fragments by PCR as indicated below (see Note 19). To synthesize 7-deaza-purine modified DNA fragments, replace dATP and dGTP by 7-deaza-dATP and 7-deaza-dGTP (see Note 20). 10 concentrated PCR buffer, containing MgSO4

5 μl

10 mM dCTP

1 μl

10 mM dTTP

1 μl

10 mM dATP or 7-deaza-dATP

1 μl

10 mM dGTP or 7-deaza-dGTP

1 μl

Forward primer, 20 μM

0.7 μl

Reverse primer, 20 μM

0.7 μl

GC-RICH solution, 5 concentrated

10 μl

Genomic DNA

15–30 ng

Pwo SuperYield DNA polymerase, 5 U/μl

0.5 μl

RNase and DNase-free H2O

up to 50 μl

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2. Check the quality of PCR products by gel electrophoresis. Only intact and pure DNA fragments should be used (see Note 21). 3. Purify DNA fragments using a PCR clean-up kit. 3.2.3 In Vitro Triplex Pull-Down

1. Prepare two 15 μl reactions in 1x Exonuclease I (Exo I) reaction buffer containing 150 fmol of unmodified or 7-deazamodified PCR-fragments and 15 units of Exo I. Mix by gentle pipetting (see Notes 22 and 23). 2. Incubate at 37  C for 30 min. 3. In parallel, in a DNA low-binding tube, assemble a 5 μl reaction containing 1.5 μl of 10 triplex buffer and 1 pmol of biotinlabelled synthetic RNA. 4. Add 10 μl of Exo I-digested DNA. Remaining 5 μl of Exo I-digested DNA should be saved as input. 5. Incubate for 20–40 min at room temperature (see Note 24) with gentle shaking in a Thermomixer (300 rpm). 6. Add 10 μl of MyOne Streptavidin C1 Magnetic Dynabeads washed three times with 600 μl of 1 triplex buffer. 7. Shake for 40 min at room temperature (see Note 25). 8. After short spinning, place the tube on a magnetic rack until the supernatant becomes clear (1–2 min). Remove the supernatant. 9. Add 600 μl of washing buffer 1 and rotate the tube for 1 min at room temperature. 10. Repeat steps 8 and 9. 11. Add 600 μl of washing buffer 2 and rotate the tube for 1 min at room temperature. 12. Repeat steps 8 and 11. 13. Add 100 μl of TE buffer containing RNase A (50 ng/ml). Include input samples from step 4. 14. Incubate samples at 37  C for 30 min with shaking in a Thermomixer (1000 rpm). 15. Spin down using a table top centrifuge. 16. Place the tube on magnetic rack and wait until the supernatant becomes clear (1–2 min). 17. Transfer the supernatant to a fresh tube. 18. Purify DNA with a PCR clean-up kit. 19. Analyze undiluted samples containing eluted RNA-associated DNA and 1:100 1:1000 diluted input DNA by qPCR (see Note 6). 20. Normalize RNA-associated DNA to input DNA.

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1. Seed 1  106 HeLa cells per 6 cm dish (see Note 26). 2. Transfect cells with 8 pmol of biotinylated RNA using TransITmRNA transfection reagent according to the manufacturer’s instructions. Include a sample that contains no RNA but only the transfection reagent (for the synthesis of biotinylated RNA, refer to the Subheadings 3.1.1 and 3.1.2); (see Note 4). 3. Harvest cells on ice 12–16 h after transfection by scraping in cold 1 PBS. Collect cells by centrifugation at 4  C (see Note 27). 4. Resuspend cell pellet in 200 μl of ice-cold nuclei isolation buffer and incubate on ice for 10 min. 5. Lyse cells by adding 0.5% Triton X-100 and incubation on ice for 5–8 min (see Note 28). 6. Collect nuclei by centrifugation at 1300  g for 3 min at 4  C. 7. Wash nuclei with 200 μl of ice-cold nuclei isolation buffer to remove residual detergent. 8. Resuspend nuclei in 100 μl of cold triplex buffer. Disrupt nuclei by short sonication (1–2 pulses); (see Note 2). 9. Transfer lysate to sonication tubes, add 100 ng proteinase K, and incubate for 20 min at room temperature. 10. Sonicate to obtain DNA fragments of about 500 bp (see Note 29). 11. Spin at 16,000  g at 4  C for 5 min. Transfer supernatant to a fresh tube (see Note 30).

3.3.2 Capture and Analysis of RNA-Associated DNA

1. Mix 95 μl of nuclear lysate in a DNA-low binding tube with 10 μl of streptavidin-coated magnetic beads (washed three times with 600 μl of triplex buffer). Incubate with shaking for 30 min at room temperature. Keep 5 μl lysate (~5%) as input (see Note 25). 2. Separate beads with a magnetic rack. Discard supernatant. 3. Wash beads as described in Subheading 3.2.3 (steps 8–12). 4. Elute RNA-bound DNA from beads by incubation for 30 min  at 37 C with 100 μl TE containing 50 ng/μl RNase A. Include input sample (add 95 μl TE with RNase A to 5 μl input material). 5. Separate the beads on a magnetic rack, save the supernatant. 6. Purify DNA with a PCR clean-up kit, elute in 30 μl. 7. Analyze 2 μl of each sample (no RNA, positive RNA, negative RNA, and input RNA) by qPCR using at least two different primer pairs. One primer pair should amplify the region where

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the assayed RNA forms the triplex structure. The other primer pair should be designed for a region where no homopurine stretches are present (see Note 6). 3.4 UV-Assisted Triplex Capture Assay 3.4.1 Transfection of RNA Oligonucleotides

1. Seed 1.5  106 HeLa cells per 6 cm dish (see Note 26). 2. Transfect cells with 1 pmol/μl of modified RNA oligonuclotide (negative and positive, see Note 10). We used the TransITmRNA transfection reagent according to the manufacturer’s instructions. Include a sample that contains the transfection reagent but no RNA. Perform each reaction in duplicates to harvest crosslinked cells and controls that have not been exposed to UV-crosslinking. 3. Incubate cells for 6 h.

3.4.2 Nuclei Isolation and Crosslinking

1. Harvest cells and isolate nuclei as described in the Subheading 3.3.1, steps 3–7. 2. Resuspend nuclei in 20 μl of ice-cold nuclei crosslinking buffer. 3. Cover a glass plate with parafilm and place it on ice. 4. Drop 20 μl of nuclei suspension on the parafilm, place the tray in the crosslinker, and irradiate with UV-light (365 nm) for 10 min. 5. Transfer the nuclei suspension back to tubes. Add 180 μl of crosslinking buffer to all samples (crosslinked and not crosslinked). 6. Snap-freeze and store at 80  C.

3.4.3 Lysate Preparation, Capture, and Analysis of RNA–Bound DNA

1. Thaw nuclei on ice. Collect by centrifugation at 1800  g for 5 min at 4  C. Remove the supernatant. 2. Resuspend nuclei in 100 μl of cold triplex buffer, disrupt by short sonication (1–2 pulses); (see Note 31). 3. Transfer lysate to sonication tubes. Add 0.5% SDS and 100 ng of proteinase K. Incubate for 20 min at room temperature. 4. Sonicate to obtain DNA fragments of about 500 bp (see Note 29). 5. Spin at 16,000  g at 4  C for 5 min. Transfer supernatant into a fresh tube. 6. Capture DNA as described in Subheading 3.3.2, steps 1–6. Note that the composition of the washing buffers is different. 7. Use 2 μl of each sample for qPCR using at least two different primer pairs. One primer pair should amplify the potential triplex-forming region, the other one should amplify a control region where no homopurine stretches are present (see Note 32).

RNA-DNA Triplex Structures

4

243

Notes 1. To synthesize sense RNA, the forward primer should be fused to the T7 promoter sequence. For the synthesis of antisense RNA, the reverse primer should be fused to the T7 promoter sequence. 2. The composition of the triplex buffer should be optimized for a given DNA and RNA. One may replace KCl with NaCl and titrate the final salt concentration from 20 mM to 50 mM (we observed considerable differences within this small range). MgCl2 might be reduced to 5 mM. 3. All buffers should be freshly prepared with DNase- and RNasefree autoclaved water. Stock solutions are stored in small aliquots at 20  C. Dilutions should be used just once. Washing buffers should be cooled down to 4  C before use. 4. At least two different transcripts need to be prepared for this experiment. Apart from the RNA to be assayed (positive RNA), one needs a negative control, that is, an RNA which should not interact with DNA (negative RNA). We usually use sequences adjacent to the predicted triplex forming region at the gene of interest. Importantly, a sample without any RNA should be included in every assay as a technical negative control. 5. It is worthwhile to test different streptavidin beads. We have tried Dynabeads M280 which are more affordable and more convenient in handling but give lower yield. Streptavidin Sepharose High Performance (GE Healthcare) increased the yield but also the background. The ultimate choice depends on the capture efficiency for a given RNA. 6. Primers should not anneal to the DNA template used to synthesize biotinylated RNA to avoid amplification of residual DNA template. 7. The QuantiTect SYBR Green PCR kit from Qiagen has worked best in amplification of CG-rich sequences. 8. The DNA fragments should not entirely overlap with the transcription templates used for RNA synthesis. The non-overlapping parts should be used as priming sites for qPCR validation. In addition to the amplicon containing the triplex-forming region, an amplicon serving as negative control should be assayed. 9. This transfection reagent outperforms other transfection reagents in terms of low toxicity and high efficiency for both long and short RNA species. 10. Two different RNA oligonucleotides should be designed. Apart from the RNA oligonucleotide to be assayed (positive RNA), a negative control is required, that is, an RNA oligonucleotide which—based on its sequence composition—should

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not form Hoogsteen bonds (negative RNA). For instance, one could use sequences from the same gene adjacent to the predicted triplex-forming region. A more stringent control would be an RNA oligonucleotide which can form triplexes at a different locus. Importantly, a sample without any RNA oligonucleotide should be included in every assay as a technical negative control. 11. For optimal yield, assemble 4  50 μl PCR. 100 μM primer stocks are prepared in TE. 20 μM working solutions are diluted with H2O. PCR conditions, such as the annealing temperature (between 58  C and 65  C) and the number of cycles (between 25 and 35), need to be optimized for each amplicon. 12. DNA concentration of at least 8–10 ng/μl is required for in vitro transcription. 13. Fluorescent labelling can be used instead of radiolabelling [15] (Fig. 2c). 14. The DNA oligonucleotide which does not hybridize to the assayed RNA must be labelled. 15. To prove that retardation of electrophoretic mobility of DNA is due to triplex formation, one should include RNase A and/or RNase H treatment after step 3 (Subheading 3.1.5). For this, 5 μl containing 0.5 ng of RNase A in H2O or 0.5 U RNase H in RNase H buffer is added and incubated for additional 30 min at 23  C before gel loading [8]. 16. Glass plates should be cleaned thoroughly before each casting, and all buffers should be prepared fresh. Gels should polymerize for at least 2 h and pre-run for about 30 min before sample loading. 17. Electrophoresis is performed in Tris–acetate buffer. The pH and the Mg++ molarity should be identical as the triplex buffer. 18. Do not use RNA that has been repeatedly thawed. 19. This protocol has been used for the generation of PCR fragments with the Pwo SuperYield DNA Polymerase Kit. Other polymerases could be used as well. In any case, the number of cycles and the annealing temperature need to be optimized for a given DNA fragment. 20. Depending on the sequence composition of the triplexforming region, either both 7-deaza-dATP and 7-deazadGTP or just one of them should be used to generate the modified DNA fragment. 21. As the deaza modification reduces the intercalating efficiency of ethidium bromide, the staining is weaker with deaza-modified DNA compared to unmodified DNA [16]. 22. As this assay yields low amounts of DNA, it is essential to assure RNase- and DNase-free conditions and avoid contamination with foreign DNA. The procedure should be performed on a

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bench where no DNA work is done, ideally using a dedicated set of pipettes. No PCR signal should be present in control samples that contained H2O or did not contain any biotinylated RNA. 23. Exonuclease I is used to eliminate single-stranded DNA that could form heteroduplexes with RNA. Heteroduplexes would yield false-positive qPCR signals. 24. The incubation time has to be optimized to increase the specificity and efficiency of triplex formation. 25. Samples should be shaken to avoid sedimentation of the beads. For best results, a tube rotator with shaking function should be used. 26. Transfection should be performed at 70–80% cell confluency. 27. Wash cells 2–3 times with cold 1 PBS before scraping to remove the culture medium completely. Use sterile disposable cell scrapers for each dish. 28. Conditions for cell lysis and nuclei release must be carefully optimized, especially the amount and type of detergent as well as the incubation time. Check microscopically that intact nuclei are released. 29. Sonication conditions must be optimized. It is critical to properly fragment the DNA (about 500 bp) as the presence of long genomic DNA will increase the background in PCR. The size of DNA should always be checked on agarose gels. 30. As an alternative approach, one can incubate isolated nuclei with biotinylated RNA rather than transfect the cells. For this, 8 pmol of RNA is incubated in DNA low-binding tubes with nuclei from 1  106 HeLa cells in 100 μl triplex buffer for 30–60 min at room temperature. The incubation time should be optimized as longer incubation will increase background signal while too short incubation will reduce the yield of triplexes. Samples should be tapped gently every 5 min to prevent sedimentation of nuclei. After incubation, 100 μl of nuclei suspension is spun for 10 min at 16,000  g at room temperature through a 1 ml sucrose cushion containing 880 μl 1 M sucrose, 10 μl 1 M Tris–HCl, pH 7.5, and 110 μl H2O. Proceed with sonication and proteinase K digestion, as described in Subheading 3.3.1, steps 8–11. 31. As SDS is used to lyse nuclei, the triplex buffer must contain NaCl rather than KCl to avoid formation of insoluble potassium SDS salts. Salt concentrations between 20 and 50 mM NaCl and 5 and 10 mM MgCl2 are optimal. 32. Alternatively, the UV-assisted capture can be performed using genomic DNA rather than chromatin lysate. For this, cells are transfected and harvested as described in Subheadings 3.4.1

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and 3.4.2. DNA isolated from thawed nuclei (using DNeasy blood and tissue kit) is fragmented by sonication in 100 μl triplex buffer. 6 μg of sonicated DNA is put in 100 μl of triplex buffer and captured as described in Subheadings 3.4.3, steps 6 and 7.

Acknowledgements Work in the Grummt lab has been funded by the DFG (GR475/ 22-2; SFB1036), CellNetworks (EcTop Survey 2014), the BadenWu¨rttemberg Stiftung, and the Deutsches Krebsforschungszentrum (DKFZ). References 1. Felsenfeld G, Davies DR, Rich A (1957) Formation of a three-stranded polynucleotide molecule. J Am Chem Soc 79:2023–2024 ˜ i JR, de la Cruz X, Orozco M (2004) 2. Gon Triplex-forming oligonucleotide target sequences in the human genome. Nucleic Acids Res 32:354–360 3. Buske FA, Bauer DC, Mattick JS, Bailey TL (2012) Triplexator: detecting nucleic acid triple helices in genomic and transcriptomic data. Genome Res 22:1372–1381 4. Schmitz KM, Mayer C, Postepska A, Grummt I (2010) Interaction of noncoding RNA with the rDNA promoter mediates recruitment of DNMT3b and silencing of rRNA genes. Genes Dev 24:2264–2269 5. Zhao Z, Sentu¨rk N, Chen C, Grummt I (2018) LncRNA PAPAS tethered to the rDNA enhancer recruits hypophosphorylated CHD4/NuRD4 to repress rRNA synthesis at elevated temperature. Genes Dev 32:836–848 6. Grote P, Wittler L, Hendrix D, Koch F, W€ahrisch S, Beisaw A et al (2013) The tissuespecific lncRNA Fendrr is an essential regulator of heart and body wall development in the mouse. Dev Cell 24:206–214 7. Mondal T, Subhash S, Vaid R, Enroth S, Uday S, Reinius B et al (2015) MEG3 long noncoding RNA regulates the TGF-β pathway genes through formation of RNA-DNA triplex structures. Nat Commun 6:7743 8. Postepska-Igielska A, Giwojna A, GasriPlotnitsky L, Schmitt N, Dold A, Ginsberg D et al (2015) LncRNA Khps1 regulates expression of the proto-oncogene SPHK1 via triplexmediated changes in chromatin structure. Mol Cell 60:626–636 9. Blank-Giwojna A, Postepska-Igielska A, Grummt I (2019) LncRNA KHPS1 activates

a poised enhancer by triplex-dependent recruitment of epigenetic regulators. Cell Rep 26:2904–2915 10. O’Leary VB, Ovsepian SV, Carrascosa LG, Buske FA, Radulovic V, Niyazi M et al (2015) PARTICLE, a triplex-forming long ncRNA, regulates locus-specific methylation in response to low-dose irradiation. Cell Rep 11:474–485 11. Kalwa M, H€anzelmann S, Otto S, Kuo CC, Franzen J, Joussen S et al (2016) The lncRNA HOTAIR impacts on mesenchymal stem cells via triple helix formation. Nucleic Acids Res 44:10631–10643 12. Paugh SW, Coss DR, Bao J, Laudermilk LT, Grace CR, Ferreira AM et al (2016) MicroRNAs form triplexes with double stranded DNA at sequence-specific binding sites; a eukaryotic mechanism via which microRNAs could directly alter gene expression. PLoS Comput Biol 12(2):e1004744 13. Li Y, Syed J, Sugiyama H (2016) RNA-DNA triplex formation by long noncoding RNAs. Cell Chem Biol 23:1325–1333 14. Sentu¨rk Cetin N, Kuo C-C, Ribarska T, Li R, Costa IG, Grummt I (2019) Isolation and genome-wide characterization of cellular DNA:RNA triplex structures. Nucleic Acids Res 47:2306–2321 15. Maldonado R, Schwartz U, Silberhorn E, Langst G (2019) Nucleosomes stabilize ssRNA-dsDNA triple helices in human cells. Mol Cell 73:1243–1254 16. Brown WT, Nolin S, Houck G Jr, Ding X, Glicksman A, Li SY, Stark-Houck S, Brophy P, Duncan C, Dobkin C, Jenkins E (1996) Prenatal diagnosis and carrier screening for fragile X by PCR. Am J Med Genet 64:191–195

Chapter 17 Analyzing RNA–DNA Triplex Formation in Chromatin Rodrigo Maldonado and Gernot L€angst Abstract A significant fraction of non-coding RNAs (ncRNAs) is associated with chromatin, shown to regulate gene expression and to organize nuclear architecture. Mechanisms of direct and indirect RNA-chromatin interactions have been described, including the sequence-specific formation of triple helix structures. Triplexes are formed by the sequence-specific binding of RNA to the bases located in the major groove of DNA. We recently showed that triplexes do exist in the context of cellular chromatin and that these structures are stabilized by the histone H3 tail of adjacent nucleosomes. The in vitro characterization of the specificity and binding affinity of triplex sequences next to nucleosomes are essential parameters to identify potential sites of RNA-chromatin interaction in vivo. Here we provide a detailed protocol to determine the influence of nucleosome positioning on triple helix formation. This assay allows the comparative quantification of triplex formation and specificity for triplex targeting sequences relative to the spatial nucleosome position. Key words Non-coding RNA, Chromatin, Nucleosome, Triple helix structures, Triplex Targeting site, Triplex Forming Oligo, Electrophoretic Mobility Shift Assay

1

Introduction The DNA of eukaryotic cells is packaged into chromatin, a compact nucleoprotein structure with the nucleosome being the basic packaging unit. The wrapping of the DNA around the core of eight histone proteins and the folding of the nucleosomal arrays into higher order structures of chromatin restricts the accessibility of DNA for sequence-specific binding of proteins and RNA. The sequence-specific targeting of ncRNAs to chromatin has been shown for several examples and targeting is essential for their regulatory activity. RNA binding to chromatin was shown to induce local changes in chromatin structure and its posttranslational modifications, and hence modifying gene expression. The sequence-specific binding of an RNA to the major groove of a Triplex Targeting Sequence (TTS) according to the Hoogsteen and reverse Hoogsteen base-pairing rules results in a triple helix, one kind of site specific RNA targeting mechanism [1–3]. Functional

Ulf Andersson Vang Ørom (ed.), RNA-Chromatin Interactions: Methods and Protocols, Methods in Molecular Biology, vol. 2161, https://doi.org/10.1007/978-1-0716-0680-3_17, © Springer Science+Business Media, LLC, part of Springer Nature 2020

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triple helix formation has been described for a number of long non-coding RNAs, including MEG3, HOTAIR, SPHK1, promoter-associated ncRNAs [4–8], and has been suggested for a subset of micro-RNAs [9, 10]. Based on the nuclease accessibility assays (DNase I, exonuclease III, and restriction enzymes), it was demonstrated that triple helices cannot form at central positions of the nucleosome. In contrast, nucleosomes harboring the TTS positions close to the nucleosomal DNA entry-exit site are compatible with the triple helix formation [11–13]. Chemically-modified Triplex-Forming Oligos (TFOs) have been used in vivo to study the cellular binding and functional roles of triplex formation in yeast, mouse, and human cells [14–17]. These studies showed that triple helices can be formed in the context of cellular chromatin. Moreover, we recently showed that nucleosome positioning and the alignment of TTS sites next to nucleosomes greatly stabilize the formation of triple helices in vitro and in vivo [18]. This chapter provides a detailed protocol for the analysis of triplex formation on fluorescently-labeled nucleosomal DNA templates. TTS sequences are placed at different positions, relative to the nucleosome positioning sequence (NPS) by PCR, using fluorescently labeled primers (Fig. 1). Nucleosomes are reconstituted

Fig. 1 Preparing fluorescent nucleosomal substrates with triplex targeting sites. (a) An AvaI flanked 601 sequence is obtained after restriction enzyme digestion of the Widom 601 template. (b) This template is used for PCR amplification using a common fluorescently-labeled reverse primer (primer marked with a star) and different forward primers carrying the TTS (primers I, II, and III). The PCR fragments are used for nucleosome assembly and served as control template (primer IV). (c) The different DNA templates (I, II, and III) are incubated with increasing amounts of histone octamers, following the salt dialysis method for nucleosome assembly. The fluorescent nucleosomes are analyzed on a 6% native polyacrylamide gel, where the nucleosomes migrate slower than the free DNA

Analyzing RNA–DNA Triplex Formation in Chromatin I

II

- +

- +

III - +

249

Triplex efficiency: TFO-RNA

Channel fluorophore B

Channel fluorophore A

( Triplex on Nucleosome signal / Nucleosome signal ) < Nucleosomes

Triplex on control template

< CTL DNA

< Triplex on Nucleosomes

< Triplex on control template

Fig. 2 Quantification of triplex binding. Each nucleosomal template (I, II, and III, as shown in Fig. 1) and the control template (labeled with fluorophore A), were incubated with (+) or without (
) the RNA-TFO (fluorophore B-labeled). The binding reactions were analyzed on native TA polyacrylamide gels and visualized on a fluorescent scanner for each fluorophore (left panel). The quantification and efficiency of triple helix formation on the nucleosomal DNA is given on the right side

on these templates and triplex binding is quantified with respect to control DNA containing the same TTS site. The templates labeled with one fluorophore and the TFO labeled with a distinct fluorophore are mixed and then separated by native electrophoresis (see Note 1). When analyzing the samples on native polyacrylamide gels, it is possible to clearly separate the nucleosomes and the control template. However, due to the size of the nucleosome a change in mobility of the nucleosome bound triplex cannot be observed (see Note 2). Gels are scanned to detect the fluorophores of the nucleosome/control DNA and the TFO separately. TFO signals appearing at the height of nucleosomes and triplexes formed with the control template are quantified, and the nucleosomal signal is normalized to the signal of the control DNA (Fig. 2). Therefore, this experimental setup allows to monitor and to quantify the efficiency of triplex formation, with respect to nucleosome positions and TFO/TTS sequence motifs. This analysis is an essential prerequisite for the identification of TFO binding sites in vivo.

2

Materials Prepare all solutions with deionized water and analytical grade reagents. Solutions can be stored at room temperature unless indicated.

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2.1 Nucleosomal Templates, Control Sequence, and TFOs for Triplex Formation

1. For nucleosome positioning, we used the 601 NPS (146 bp) identified by Widom and colleagues [19] (Fig. 1a) (see Note 3). 2. PCR reagents. 3. Ethanol. 4. TE buffer: 10 mM Tris–HCl [pH 8.1], 1 mM EDTA. 5. PCR clean-up column.

2.2

Triplex Reactions

1. Fluorescently-labeled nucleosomes (see Note 4). 2. Fluorescently-labeled triplex control template (100 bp) (see Note 5). 3. 1 Tris–acetate buffer: 40 mM Tris [pH 7.4], 5 mM Magnesium Acetate (see Note 6). Prepared as a 10 solution (1 L) and store at 4  C. 4. BSA 200 ng/μl final concentration, from a stock 10 mg/ml in ddH2O. 5. Orange G loading dye: 50% w/v glycerol, 10 mM Tris–HCl [pH 8.0], 0.25% w/v Orange G. 6. Deionized water. 7. Low binding reaction tubes. 8. Thermoblock. 9. Fluorescence scanner. 10. Ammonium Persulfate (APS) 20% w/v. 11. Acrylamide 30%, Bis-acrylamide at a ratio 37, 5:1. 12. TEMED. 13. Deionized water.

2.3 Analysis and Quantification of the Triplex Formation Efficiency

3

1. ImageJ software.

Methods A detailed protocol for the nucleosome assembly was published previously and used as described [20]. The triplex assembly reactions must be maintained on ice unless indicated.

3.1 NucleosomeTriplex Mobility Shift Assays

1. Before preparing the triplex reactions, perform a pre-run of the 6% Polyacrylamide TA gels in 1 TA buffer for 30 min at 90 V and 4  C (Table 1, see Note 7). 2. During the pre-run, prepare the triplex binding reactions. Use low binding tubes (1.5 ml) to avoid unspecific binding of the

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Table 1 Preparation of the Polyacrylamide TA gel 6% Polyacrylamide TA gel 1 Gel

6 Gels

Acrylamide 30%

2 ml

12 ml

10 Tris–acetate buffer

1 ml

6 ml

APS 20%

42 μl

252 μl

Water

6.95 ml

41.7 ml

TEMED

8 μl

48 μl

nucleosomes and the RNA-TFOs to the reaction tube. A standard reaction has a final volume of 10 μl containing: 50–100 nM (see above) of fluorescently-labeled nucleosomes (Fluorophore A), 50 nM of fluorescently-labeled triplex control template (Fluorophore A), 0.1 to 4 μM of the fluorescently-labeled RNA-TFO (Fluorophore B) (see Note 8), 200 ng/μl of BSA, 1 μl of 10 TA buffer, and water. 3. Incubate the samples in a thermoblock for 30–60 min at 37  C without agitation, and then place the samples on ice. 4. To each sample, add 3 μl of the Orange G loading dye and mix gently. Load the samples onto the pre-run TA gel. Electrophorese the samples at 15 V/cm until the Orange G reaches the bottom of the gel. Nucleosomes and the triplex control template must be well separated for proper quantification (Fig. 2). 5. Scan the gel with a fluorescence reader, appropriate filters and lasers to detect both fluorophores individually (Fig. 2). 6. To quantify and to compare triplex formation efficiencies between different nucleosomal templates, open the image files with ImageJ and follow the recommendations provided to quantify signal intensities (https://di.uq.edu.au/commu nity-and-alumni/sparq-ed/sparq-ed-services/using-imagejquantify-blots). For each sample to be analyzed, quantify the signal intensities of the nucleosome, the triplexes formed on nucleosomes, and the signals of the triplexes on the control template. The efficiency is calculated following two normalizations. First, the samples are normalized to the nucleosome template, dividing the nucleosome-triplex signal by the signal of the nucleosome. This value is then normalized with respect to the control template, by dividing it by the triplex signal on the control template (Fig. 2).

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Notes 1. To avoid fluorescence cross-bleeding, choose fluorophores that have widely separated spectral profiles. We experienced no problems when combining Carboxyfluorescein (FAM) and Cyanine-5 (Cy5) as fluorophores. 2. When working with RNA-TFOs to form triplexes (lengths between 20 and 40 bases), generally (depending on the sequence) no difference in mobility between the control DNA and its triplex bound form can be observed on 12–15% TA gels. Also for the nucleosomes, no change in migration can be observed upon triplex binding. This is due to the large size of the nucleosomes when compared to the TFOs. Therefore, it is required to study the triplex formation on nucleosomes (180–146 bp depending on the template) and control template (100 bp) in the same gel, and to use different fluorescent labels for the TFO and the TTS-containing template. To separate nucleosomes and control triplex in a single gel, 6% polyacrylamide TA gels were used. For an example showing a comparison between DNA- and RNA-TFOs, please refer to [18]. Triplex formation is determined from RNA signals overlapping with the nucleosomal positions (Fig. 2). 3. The 601 sequence served as the core template for PCR with oligonucleotides encoding the TTS. Using one common fluorescently-labeled reverse primer (arrow with star) and different forward primers carrying a TTS sequence at their 50 ends (arrows I, II, and III), we incorporated the TTS sequences at different positions relative to the nucleosomal binding site (Fig. 1b). A short DNA fragment, not reconstituted into nucleosomes, served as internal control for quantification (Triplex control template). For nucleosome assembly, we used the salt dialysis method as previously described [20]. The histones have to be titrated, relative to the template DNA, to estimate the optimal (DNA:histone) assembly ratios. The slower migration of the reconstituted nucleosomes compared to the freeDNA can be easily visualized on a native 0.4 TBE 6% polyacrylamide gel (Fig. 1c). Finally, the nucleosomes are mixed at equimolar amounts with the short DNA fragment containing the TTS, to allow subsequent quantification. The different forward primers (determining the location of the TTS) must have at least 12–15 bases complementary to the 601 core template to obtain specific DNA fragments by PCR. 4. After the nucleosome assembly reaction, these are stored at 4  C in 10 mM Tris–HCl [pH 7.6], 200 mM NaCl, 1 mM EDTA, 1 mM 2-mercaptoethanol, 0.05% Igepal CA-630 [20]. The concentration of the nucleosomes is calculated

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from the DNA amount used for the assembly, taking into account the final reaction volume after dialysis. Use 50-100 nM of fluorescently-labeled nucleosomes per triplex assembly reaction. 5. Use 50 nM per triplex reaction. The amount can differ from the concentration of nucleosomal templates, but it is important to maintain the same amount of control DNA in all the reactions to be analyzed. Fluorescently-labeled RNA-TFOs: These DNA or RNA oligonucleotides must have a different fluorophore than the nucleosomes and triplex control template. The oligos have to be efficiently modified with the fluorophore to obtain consistent results. 6. For the triplex formation on nucleosomes, we use a maximum of 5 mM Magnesium ions, as higher concentrations would cause the precipitation of nucleosomes. 7. Clean the electrophoretic chambers thoroughly before running the samples. When using TA buffer, we observed precipitations on the wires, affecting the running behavior of the gels. 8. It is recommended to perform titration series of the TFO, starting with a nucleosome:TFO ratio of about 1:1 and then increasing the amount of TFO until a clear signal of triplexes binding can be observed with accessible sites. We observed clear triplex formation on accessible TTS sites within nucleosomes at ratios of about 1:4 to 1:20 depending on TFO sequences. References 1. Hoogsteen K (1959) The structure of crystals containing a hydrogen-bonded complex of 1-methylthymine and 9-methyladenine. Acta Crystallogr 12:822–823. https://doi.org/10. 1107/S0365110X59002389 2. Rajagopal P, Feigon J (1989) Triple strand formation in the homopurine:homopyrimidine DNA oligonucleotides d(GA)4 and d(TC)4. Nature 339:637–640. https://doi.org/10. 1038/339637a0 3. Rinn JL, Chang HY (2012) Genome regulation by long noncoding RNAs. Annu Rev Biochem 81:145–166. https://doi.org/10.1146/ annurev-biochem-051410-092902 4. Mondal T, Subhash S, Vaid R, Enroth S, Uday S, Reinius B, Mitra S, Mohammed A, James AR, Hoberg E, Moustakas A, Gyllensten U, Jones SJM, Gustafsson CM, Sims AH, Westerlund F, Gorab E, Kanduri C (2015) MEG3 long noncoding RNA regulates the TGF-β pathway genes through formation of RNA–DNA triplex structures. Nat Commun

6:7743. https://doi.org/10.1038/ ncomms8743 5. Kalwa M, H€anzelmann S, Otto S, Kuo C-C, Franzen J, Joussen S, Fernandez-Rebollo E, Rath B, Koch C, Hofmann A, Lee S-H, Teschendorff AE, Denecke B, Lin Q, Widschwendter M, Weinhold E, Costa IG, Wagner W (2016) The lncRNA HOTAIR impacts on mesenchymal stem cells via triple helix formation. Nucleic Acids Res 44: gkw802. https://doi.org/10.1093/nar/ gkw802 6. Postepska-Igielska A, Giwojna A, GasriPlotnitsky L, Schmitt N, Dold A, Ginsberg D, Grummt I (2015) LncRNA Khps1 regulates expression of the proto-oncogene SPHK1 via triplex-mediated changes in chromatin structure. Mol Cell 60:626–636. https://doi.org/ 10.1016/j.molcel.2015.10.001 7. Martianov I, Ramadass A, Serra Barros A, Chow N, Akoulitchev A (2007) Repression of the human dihydrofolate reductase gene by a

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non-coding interfering transcript. Nature 445:666–670. https://doi.org/10.1038/ nature05519 8. Schmitz K-M, Mayer C, Postepska A, Grummt I (2010) Interaction of noncoding RNA with the rDNA promoter mediates recruitment of DNMT3b and silencing of rRNA genes. Genes Dev 24:2264–2269. https://doi.org/10. 1101/gad.590910 9. Paugh SW, Coss DR, Bao J, Laudermilk LT, Grace CR, Ferreira AM, Waddell MB, Ridout G, Naeve D, Leuze M, LoCascio PF, Panetta JC, Wilkinson MR, Pui C-H, Naeve CW, Uberbacher EC, Bonten EJ, Evans WE (2016) MicroRNAs form triplexes with double stranded DNA at sequence-specific binding sites; a eukaryotic mechanism via which microRNAs could directly Alter gene expression. PLoS Comput Biol 12:e1004744. https:// doi.org/10.1371/journal.pcbi.1004744 10. Toscano-Garibay JD, Aquino-Jarquin G (2014) Transcriptional regulation mechanism mediated by miRNA–DNA·DNA triplex structure stabilized by Argonaute. Biochim Biophys Acta 1839:1079–1083. https://doi.org/10. 1016/j.bbagrm.2014.07.016 11. Westin L, Blomquist P, Milligan JF, Wrange O (1995) Triple helix DNA alters nucleosomal histone-DNA interactions and acts as a nucleosome barrier. Nucleic Acids Res 23:2184–2191 12. Espina´s ML, Jime´nez-Garcı´a E, Martı´nez-Bal´ , Azorı´n F (1996) Formation of tripleba´s A stranded DNA at d(GA·TC)(n) sequences prevents nucleosome assembly and is hindered by nucleosomes. J Biol Chem 271:31807–31812. https://doi.org/10.1074/jbc.271.50.31807 13. Brown PM, Fox KR (1998) DNA triple-helix formation on nucleosome-bound poly(dA). poly(dT) tracts. Biochem J 333(2):259–267

14. Barre FX, Ait-Si-Ali S, Giovannangeli C, Luis R, Robin P, Pritchard LL, Helene C, Harel-Bellan A (2000) Unambiguous demonstration of triple-helix-directed gene modification. Proc Natl Acad Sci U S A 97:3084–3088. https://doi.org/10.1073/pnas.050368997 15. Luo Z, Macris MA, Faruqi AF, Glazer PM (2000) High-frequency intrachromosomal gene conversion induced by triplex-forming oligonucleotides microinjected into mouse cells. Proc Natl Acad Sci U S A 97:9003–9008. https://doi.org/10.1073/ pnas.160004997 16. Macris MA, Glazer PM (2003) Transcription dependence of chromosomal gene targeting by triplex-forming oligonucleotides. J Biol Chem 278:3357–3362. https://doi.org/10.1074/ jbc.M206542200 17. Besch R, Giovannangeli C, Schuh T, Kammerbauer C, Degitz K (2004) Characterization and quantification of triple helix formation in chromosomal DNA. J Mol Biol 341:979–989. https://doi.org/10.1016/j. jmb.2004.05.079 18. Maldonado R, Schwartz U, Silberhorn E, L€angst G (2019) Nucleosomes stabilize ssRNA-dsDNA triple helices in human cells. Mol Cell 73:1243–1254. https://doi.org/10. 1016/j.molcel.2019.01.007 19. Lowary PT, Widom J (1998) New DNA sequence rules for high affinity binding to histone octamer and sequence-directed nucleosome positioning. J Mol Biol 276:19–42. https://doi.org/10.1006/jmbi.1997.1494 20. L€angst G (2016) Chapter 9: preparation of chromatin templates to study RNA polymerase I transcription in vitro. Methods Mol Biol 1455:109–119

Chapter 18 Mapping RNA–Chromatin Interactions In Vivo with RNA-DamID Seth W. Cheetham and Andrea H. Brand Abstract Long-noncoding RNAs (lncRNAs) are emerging as regulators of development and disease. lncRNAs are expressed in exquisitely precise expression patterns in vivo and many interact with chromatin to regulate gene expression. However, the limited sensitivity of RNA-purification techniques has precluded the identification of genomic targets of cell-type specific lncRNAs. RNA-DamID is a powerful new approach to understand the mechanisms by which lncRNAs act in vivo. RNA-DamID is highly sensitive and accurate, and can resolve cell-type-specific chromatin binding patterns without cell isolation. The determinants of RNA-chromatin interactions can be identified with RNA-DamID by analyzing RNA and protein cofactor mutants. Here we describe how to implement RNA-DamID and the design considerations to take into account to accurately identify lncRNA-chromatin interactions in vivo. Key words RNA-DamID, lncRNAs, In vivo, Protein cofactor mutants

1

Introduction

1.1 Technologies for Understanding ChromatinAssociated RNAs

Abundant RNAs, distinct from mRNAs, which are chromatinassociated were first detected by biochemical fractionation of nuclei [1, 2]. The composition of chromatin-associated RNA varies by tissue-type and was hypothesized to control tissue-specific gene expression [3]. The discovery of chromatin-associated RNA led to a theory of RNA-based regulatory networks in eukaryotes [4– 6]. However, identifying the specific genomic loci bound by these RNAs was not possible. The development of fluorescence in situ hybridization (FISH) allowed the visualization of lncRNA molecules within the nucleus, revealing for example the binding of the dosage compensation RNAs, Xist, and roX1/2, to the X chromosome [7–9]. However, for lncRNAs whose targets were not restricted to a single chromosome, advances in RNA purification were required. A major breakthrough in the understanding of lncRNAchromatin interactions came with the development of chromatin

Ulf Andersson Vang Ørom (ed.), RNA-Chromatin Interactions: Methods and Protocols, Methods in Molecular Biology, vol. 2161, https://doi.org/10.1007/978-1-0716-0680-3_18, © Springer Science+Business Media, LLC, part of Springer Nature 2020

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isolation by RNA purification (ChIRP) and capture hybridization of RNA targets (CHART) [10, 11]. Both methods use biotinylated antisense oligonucleotides to purify chromatin-associated lncRNAs and their genomic targets. These techniques enabled the identification of the binding sites for the lncRNAs roX2, HOTAIR, and TERC. A third related approach, RNA affinity purification (RAP), uses long RNA (rather than DNA) oligonucleotides to identify the genomic occupancy of lncRNAs [12]. More recently, three new approaches, mapping RNA-genome interactions (MARGI), global RNA interactions with DNA sequencing (GRID-seq), and chromatin-associated RNA (ChARseq) sequencing have identified the genomic binding sites of all lncRNAs in cultured cells by ligating lncRNAs to associated genomic DNA [13–15]. While these approaches were highly effective in identifying the binding patterns of lncRNAs in cultured cells, their efficacy in vivo remains unclear. lncRNAs are expressed in highly spatially and temporally specific patterns [16–20]. Many lncRNAs are expressed in only a single cell type and are expressed, on average, at lower levels and in far more restricted patterns than protein-coding RNAs. RNA purification methods typically require millions of cells, making it difficult to study the mechanisms of RNAs that are so precisely expressed. Indeed, the bulk of studies examining the mechanisms of actions of lncRNAs focus on broadly or highly expressed lncRNAs [21]. RNA-DamID is a novel approach to decipher the mechanisms of action of cell-type-specific lncRNAs that does not rely on RNA-purification [22]. An lncRNA of interest is fused to several repeats of the MS2 stem loop. The tagged RNA is co-expressed with very low levels of an MS2 coat protein-Dam methylase fusion under the control of the GAL4 system (Fig. 1). The GAL4 system enables spatial and temporal restriction of the RNA-DamID signal. The MS2 coat protein binds to the MS2 stem loops with nanomolar affinity allowing the Dam methylase to methylate adenines in the sequence GATC close to sites of lncRNA-chromatin interaction (Fig. 1). These methylated sequences can be identified by digestion with methylation sensitive enzymes, adaptor ligation, and PCR amplification (Fig. 2). RNA-DamID has identified genome-wide, cell-type-specific, maps of RNA occupancy in vivo. It offers unparalleled sensitivity and was able to identify roX RNA binding sites from as few as ~30,000 neural stem cells. However, the maximum sensitivity of RNA-DamID may be considerably higher. DamID is capable of detecting chromatin interactions from single cells [23]. By combining RNA-DamID with mutant alleles, the determinants of RNA-binding can be identified. Targeted DamID has been implemented in a variety of model systems including mammalian cells [24–27], and it is anticipated that RNA-DamID will be adapted to other model organisms.

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Fig. 1 RNA-DamID schematic. An MS2-tagged lncRNA and MCP-Dam are expressed in the cell-type of interest. An upstream ORF titrates MCP-Dam levels to an extremely low level. The MCP-Dam fusion protein is recruited to sites of lncRNA-chromatin interactions. Dam methylates adenines in GATC motifs close to the sites of interaction. DamID identifies the methylated regions genome-wide

The key advantages of RNA-DamID are ultra-high sensitivity and cell-type-specificity without cell isolation. These features make RNA-DamID ideal for understanding the mechanisms of lncRNAs with restricted expression patterns in vivo. Experiments with the roX RNAs also suggested that RNA-DamID generates fewer false positives than ChIRP in vivo [22]. RNA-DamID involves the cell-type-specific expression of an MS2-tagged lncRNA and MCP-Dam fusion. DNA is extracted from the tissue and digested with the enzyme DpnI, which cleaves methylated GATC motifs. Adaptors are ligated to the methylated fragments, and residual unmethylated GATCs are cleaved by DpnII. Methylated fragments are then amplified using primers that bind to the adaptors. The amplified DNA is sonicated, the adaptors removed by AlwI digestion, and the DNA prepared for sequencing using the Illumina TruSeq protocol.

2 2.1

Materials DNA Extraction

1. DNA Micro kit. 2. Agarose (Molecular Biology Grade).

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Fig. 2 Major steps in the DamID workflow. DNA is extracted from tissues in which RNA-DamID constructs have been expressed. Genomic DNA is treated with the methyl-GATC-specific enzyme DpnI and adaptors are ligated. The methylated fragments are amplified by PCR and sequenced. The data are mapped to the genome and normalized to controls, identifying RNA-chromatin interactions

3. Ethidium bromide. 4. Absolute ethanol (Molecular Biology Grade). 5. 12.5 μg/μl RNAse A. 2.2

DpnI Digestion

1. DpnI enzyme. 2. DpnI buffer. 3. PCR Purification Kit.

Mapping RNA–Chromatin Interactions In Vivo with RNA-DamID

2.3

Adaptor Ligation

259

1. dsAdaptor (50 μM each of CTAATACGACTCACTATAGGGCAGCGTGGTCGCGGCCGAGGA and TCCTCGGCCG annealed by heating to 95
C and slowly cooling). 2. Adaptor ligation mix: 2 μl 10 NEB T4 Ligase Buffer, 0.8 μl dsAdaptor, 1.2 μl H2O.

2.4

DpnII Digestion

1. 10 DpnII Buffer. 2. DpnII Enzyme. 3. Magnetic beads. 4. 80% Ethanol in ddH2O.

2.5 PCR Amplification

1. MyTaq® Polymerase. 2. 5 MyTaq Master Mix. 3. DamID PCR primer (10 μM GGTCGCGGCCGAGGATC). 4. PCR Purification Kit.

2.6

Sonication

1. Sonicator and sonication tubes. 2. AlwI and Cutsmart® Buffer (10).

2.6.1 Sequencing and Analysis

3

1. Illumina TruSeq® Kit. 2. Computer or cluster with Unix and sufficient resources to process next generation sequencing data.

Methods

3.1 Designing Tagged RNA

The binding sites of either endogenously tagged or ectopically expressed chromatin-associated RNAs can be identified by RNA-DamID. For ectopically expressed tagged-RNAs, it is crucial that the RNA can function in trans. It should be experimentally confirmed that the addition of MS2 tags does not inhibit RNA function. When ectopically expressing MS2-tagged RNAs, it is recommended that the tag be located at the 50 end of the transcript to preclude premature transcriptional termination before the incorporation of a 30 tag. As the Dam-MCP fusion protein will be expressed at far lower levels than the tagged RNAs, the number of MS2 tags is unlikely to have a considerable impact. The binding sites of RNAs tagged with either three (roX2) or six (roX1) MS2 stem loops have been successfully mapped [22]. In all DamID experiments, it is critical to include a negative control. Dam preferentially methylates open chromatin and as such DamID data must be normalized to a Dam containing control [28, 29]. We recommend that the negative control be the untethered MS2 tags co-expressed with the Dam-MCP to control for background Dam-MCP methylation and any effects from MS2

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expression. RNA-DamID data normalized to this control or to Dam-alone has a high correlation, so Dam-alone is an alternative control. 3.2

DNA Extraction

1. Remove dissected tissue from storage in a 80
freezer and suspend in 180 μl of Buffer ATL. After the addition of buffer ATL care must be taken to minimize force that may shear DNA. Sheared DNA will be amplified by DamID, resulting in artefacts (see Note 1). Tissues rich in nucleases, e.g., digestive tissues, may require the addition of EDTA to prevent DNA degradation. 2. Add 20 μl of Proteinase K, gently mix by flicking, and heat at 56
C until tissue is completely digested. 3. After cooling to room temperature, add 20 μl RNase A (12.5 μg/μl) mix by gently flicking and incubate at room temperature for 2 min. 4. Mix 200 μl Buffer AL and 200 μl absolute ethanol in a tube by vortexing and add to digested tissue solution. 5. Mix by gentle inversion until homogenous. 6. Gently decant solution onto column. 7. Centrifuge at >6000  g for 1 min, discard flow-through and collection tube. 8. Add 500 μl of Buffer AW1 to the column in a fresh collection tube. 9. Centrifuge at >6000  g for 1 min, discard flow-through and collection tube. 10. Add 500 μl of Buffer AW2 to the column. 11. Centrifuge at >6000  g for 1 min, discard flow-through and collection tube. 12. Transfer column to a 1.5 ml tube and add 50 μl AE buffer to the center of the column. 13. Incubate for 1 min at room temperature. 14. Centrifuge at >6000  g for 1 min. 15. Run 1 μl of genomic DNA on a 0.8% ethidium bromidestained agarose gel. The DNA should appear as a single band higher than the ladder. Extensive smearing indicates degraded DNA and is associated with reduced data quality.

3.3

DpnI Digestion

1. Add 43.5 μl of eluate and 5 μl of Cutsmart® Buffer to a 1.5 ml tube. 2. Add 1.5 μl of DpnI to the DNA solution and mix very gently with a 1 ml pipette tip with the end cut off. 3. Incubate overnight at 37
C.

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4. Purify using a QIAquick PCR Purification Kit and elute DNA in 32 μl of distilled H2O. 3.4

Adaptor Ligation

1. Add 4 μl of adaptor ligation mix to 15 μl of DNA in a 0.2 ml tube. 2. Add 1 μl of T4 DNA ligase. 3. Incubate for 2 h at 16
C. 4. Inactivate ligase by heating to 65
C for 20 min.

3.5

DpnII Digestion

1. Add 4 μl of DpnII buffer (10), 15 μl of H2O to DNA solution. 2. Add 1 μl of DpnII and incubate at 37
C for 1 h. 3. Heat inactivate at 65
C for 20 min. 4. Add 60 μl of Ampure XP beads and incubate at room temperature for 10 min (see Note 2). 5. Place tubes on magnetic rack until beads separate and collect on the side (approximately 10 min). 6. Remove liquid, taking care to not disturb beads and wash twice with 190 μl of 80% ethanol. 7. Remove all ethanol (remove the majority with a P200 and then the remainder with a P20 pipette. 8. Allow beads to dry until they begin to appear cracked (approximately 5–10 min). 9. Resuspend beads in 38 μl resuspension buffer. 10. Incubate for 2 min. 11. Place on magnetic rack for 5 min or until clear, and then remove 36 μl in a new 0.2 ml tube.

3.6 PCR Amplification

1. Add 36 μl of DNA solution, 5 μl of MyTaq 5 master mix, 2.5 μl of DamID PCR primer (10 μM), and 1.5 μl of MyTaq. 2. Mix well and place in PCR machine. 3. Run the following program: 72
C for 10 min. 95
C for 30 s. 65
C for 5 min. 72
C for 15 min. Repeat 3


95 C for 30 s. 65
C for 1 min. 72
C for 10 min.

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Repeat 17


95 C for 30 s. 65
C for 1 min. 72
C for 2 min. 72
C for 5 min. 4
C hold. 4. Purify the DNA using a QIAquick PCR purification Kit and elute in 30 μl of ddH2O. 3.7

Sonication

1. Run 1 μl of DNA on a gel. The DNA should run as a smear. 2. Measure the DNA concentration. 3. Dilute 2 μg of DNA in 1 Cutsmart buffer in a sonication tube. 4. Sonicate to an average size of ~300 bp (see Note 3). 5. Add 1 μl of AlwI to cleave off DamID adaptors.

3.8 Sequencing and Analysis

1. Prepare libraries for sequencing using an Illumina TruSeq Kit. 2. Sequence to an average depth of at least five million reads (Drosophila) or 20–50 million reads (mammals). Single-end reads are sufficient for most purposes. 3. Map and normalize the DamID data according to the DamIDseq pipeline [30]. The DamID-seq pipeline normalises to Dam-alone and reduces background signal while retaining bound-signal. 4. We recommend that RNA-DamID data be represented as a log2 ratio over the negative control. 5. Determine sites of lncRNA-chromatin interaction using a peak finding algorithm, e.g., MACS2 [31].

4

Notes 1. Care must be taken to ensure that the DNA is of the highest possible integrity. Double stranded breaks induced by shearing provide substrates for blunt-ended adaptor ligation. This can result in the amplification of sheared genomic fragments that were not methylated by Dam. Solutions should be pipetted with a P1000 where possible and never vortexed. 2. Bead-purification removes the DpnII buffer, which is incompatible with some polymerases. We have found substantially increased yields from the DamID PCR following purification after DpnII digestion.

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3. The sonication time to reach an average size of 300 bp must be determined empirically according to the sonicator and tubes used. Each sonicator will give different results.

Acknowledgments A.H.B is funded by a Royal Society Darwin Trust Research Professorship and Wellcome Trust Senior Investigator Award 103792. A. H.B acknowledges core funding to The Gurdon Institute from the Wellcome Trust (092096) and CRUK (C6946/A14492). S.W.C. acknowledges support from a National Health and Medical Research Council (NHMRC) Early Career Fellowship (GNT1161832) and Mater Foundation. References 1. Warner JR, Soeiro R, Birnboim HC et al (1966) Rapidly labeled HeLa cell nuclear RNA. I. Identification by zone sedimentation of a heterogeneous fraction separate from ribosomal precursor RNA. J Mol Biol 19:349–361 2. Paul J, Duerksen JD (1975) Chromatinassociated RNA content of heterochromatin and euchromatin. Mol Cell Biochem 9:9–16 3. Mayfield JE, Bonner J (1971) Tissue differences in rat chromosomal RNA. Proc Natl Acad Sci U S A 68:2652–2655 4. Britten RJ, Davidson EH (1969) Gene regulation for higher cells: a theory. Science 165:349–357 5. Davidson EH, Klein WH, Britten RJ (1977) Sequence organization in animal DNA and a speculation on hnRNA as a coordinate regulatory transcript. Dev Biol 55:69–84 6. Mattick JS (1994) Introns: evolution and function. Curr Opin Genet Dev 4(6):823–831 7. Brown CJ, Hendrich BD, Rupert JL et al (1992) The human XIST gene: analysis of a 17 kb inactive X-specific RNA that contains conserved repeats and is highly localized within the nucleus. Cell 71:527–542 8. Brockdorff N, Ashworth A, Kay GF et al (1992) The product of the mouse Xist gene is a 15 kb inactive X-specific transcript containing no conserved ORF and located in the nucleus. Cell 71:515–526 9. Meller VH, Wu KH, Roman G et al (1997) roX1 RNA paints the X chromosome of male drosophila and is regulated by the dosage compensation system. Cell 88:445–457 10. Chu C, Qu K, Zhong FL et al (2011) Genomic maps of long noncoding RNA occupancy

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INDEX A

G

Antisense oligonucleotides ................................. 2, 18, 26, 30, 38, 46, 53–57, 256

Gapmers.....................................................................29–35 Gene expression ........................................... 6, 17, 37, 47, 54, 75, 116, 118, 143, 210, 230, 247, 255 Genome editing ................................................................ 2

B Biotinylated RNA.........................................................231, 235–237, 239, 241

C Chromatin ........................................1, 17–27, 30, 51–57, 75, 89–98, 115–118, 143, 144, 154, 158, 195, 200, 205, 247–253, 255, 256, 259 Chromatin-associated RNA (ChAR-seq).......................................115–141, 256 Chromatin immunoprecipitation (ChIP) ..................... 18, 57, 196, 204, 205, 210, 219 Chromatin oligo-affinity precipitation (ChOP) ..........................................................17–27 Cis-elements ......................................................... 143–159 CRISPR/Cas9.............................................................2, 52 CRISPR interference (CRISPRi) ................................ 2–4, 6, 11, 12

H High-throughput sequencing ............................. 162, 170 Histones...................................................... 2, 75–87, 116, 118, 158, 247, 248, 252 Hybridization-capture .................................................... 18

I ImmunoFISH........................................ 60, 63, 64, 66–73 Immunofluorescent............................................ 60, 70–72 Immunoprecipitation...........................18, 25, 76, 78, 79, 89, 161, 162, 167, 196, 197, 200, 204, 205, 210 In vivo crosslinking ...................................................76, 80

K Knockdown .................................................................3, 11

L

D Disuccinimidyl suberate.................................................. 76 DNA ............................................ 1, 2, 4, 5, 9, 17–19, 22, 23, 25–27, 30, 38, 40, 41, 43–45, 48, 52–57, 63, 82, 85, 116–118, 121, 124, 127, 130–132, 134–137, 139–141, 144, 146, 147, 149–153, 156–159, 170, 177, 180–183, 185–188, 195–206, 209–217, 220, 223–226, 229–232, 234–242, 247–249, 252, 253, 256, 257, 260–262 RNA immunoprecipitation (DRIP)....................... 213 Duchenne muscular dystrophy....................................... 53

E Electrophoretic mobility shift assay ............................. 230 3’end formation ........................................................55, 56

F FLAG-biotin-mediated cross-linking and immunoprecipitation (FbioCLIP-seq)............162, 163, 171

LIN28 ................................................................... 163, 171 LncRNAs .............................................. 1–4, 6, 10–13, 17, 18, 25–27, 29–35, 52, 53, 55, 116, 143, 229, 230, 255–257 Locked nucleic acids (LNAs)................................. 2, 4, 48 Long noncoding RNAs (lncRNAs) ...........................1, 17

M Mass spectrometry (MS)................................................ 84, 87, 89, 91, 96 Monte Carlo simulations .............................................176, 180, 189, 190

N Nascent RNA transcripts ................................... 51, 52, 54 Non-coding RNAs (ncRNAs) ...................................1–13, 17–27, 29, 30, 116, 118, 195, 247, 248 Non-denaturing bisulfite conversion ........................... 210 Nucleosomes ........................................................ 247–253

Ulf Andersson Vang Ørom (ed.), RNA-Chromatin Interactions: Methods and Protocols, Methods in Molecular Biology, vol. 2161, https://doi.org/10.1007/978-1-0716-0680-3, © Springer Science+Business Media, LLC, part of Springer Nature 2020

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AND

PROTOCOLS

O 2’OmePS ...................................................................53–57 2’-O-methyl phosphorothioate ...................................... 53

P Polyadenylation signal (pAS).......................................... 55 Protein-protein interactions ..........................75, 163, 167 Proteins................................... 1, 2, 4, 10, 17, 18, 20, 22, 25–27, 29, 37, 38, 46, 51, 52, 56, 59–73, 75–87, 89–98, 101–106, 108, 110–112, 118, 143, 158, 161–163, 165, 167, 198, 200, 205, 229–231, 247, 256, 259 Psoralen ......................................................................... 231 Pull-down .......................... 162, 231, 232, 235, 239–240

Q Quadruplexes ....................................................... 101–112

R Random mutagenesis.................................................... 157 REL-seq ................................................................ 143–159 R-loops .............................................. 195, 196, 209, 210, 212–214, 222–224, 226 RNA ................................................1–4, 6, 11, 17–20, 22, 25–27, 30–32, 34, 38, 39, 43, 48, 51–57, 59–61, 63, 64, 66, 70–72, 82, 89–98, 101–112, 115–141, 143–159, 161–163, 168–171, 175–206, 210, 212, 229–232, 234–236, 238–242, 247, 252, 253, 255, 256, 259 RNA-binding proteins (RBPs) ......................89, 161–163 RNA-DNA hybrid......................................................... 117 RNA-DNA triplexes............................................. 229, 230 RNA localization ...............................................29, 30, 52, 116, 143–159, 175–194

RNA-protein interactions ...................................... 37, 101 RNA-seq .................................................. 54, 59, 175–194 RNA-targeted therapy .................................................... 37

S Saturation transfer difference (STD) NMR ......................................................... 101–112 S9.6 Immunoprecipitation ........................................... 200 Single-molecule fluorescence in situ hybridization (smFISH) ............................ 60, 61, 63, 64, 71, 72 Single-Molecule Real Time sequencing (SMRT-seq) .............................................. 210, 212 Splice modulation .....................................................37, 46 Splice-switching oligonucleotides (SSOs) ...............................................37–39, 47, 48 Subcellular localizations.................................52, 143–159

T TERRA .......................................................................... 104 Transcriptional readthrough.....................................52, 57 Transcription termination......................... 51–53, 56, 210 Transcriptions............................................. 2–4, 6, 11, 17, 52–55, 57, 89, 116, 117, 148, 155–156, 162, 163, 169, 170, 176, 179, 180, 183, 186, 192, 193, 195, 196, 209, 213, 229, 230, 232, 237–239 Triple helix structures .......................................... 247–249 Triplex capture .............................................................231, 235, 236, 241–242 Triplex targeting sites.................................................... 248 Triplex-Forming Oligo (TFOs) ..................................248, 250, 252

W WDR43................................................................. 163, 171