Methods and Protocols

Methods in Molecular Biology 1296

Mathieu Rederstorff Editor

Small Non-Coding RNAs Methods and Protocols

METHODS

IN

MOLECULAR BIOLOGY

Series Editor John M. Walker School of Life and Medical Sciences University of Hertfordshire Hatfield, Hertfordshire, AL10 9AB, UK

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Small Non-Coding RNAs Methods and Protocols

Edited by

Mathieu Rederstorff Université de Lorraine, Biopôle, CNRS UMR 7365, IMoPA, Vandoeuvre-lès-Nancy, France

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Editor Mathieu Rederstorff Université de Lorraine, Biopôle, CNRS UMR 7365, IMoPA Vandoeuvre-lès-Nancy, France

ISSN 1064-3745 ISSN 1940-6029 (electronic) Methods in Molecular Biology ISBN 978-1-4939-2546-9 ISBN 978-1-4939-2547-6 (eBook) DOI 10.1007/978-1-4939-2547-6 Library of Congress Control Number: 2015934152 Springer New York Heidelberg Dordrecht London © Springer Science+Business Media New York 2015 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, express or implied, with respect to the material contained herein or for any errors or omissions that may have been made. Printed on acid-free paper Humana Press is a brand of Springer Springer Science+Business Media LLC New York is part of Springer Science+Business Media (www.springer.com)

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Preface The reality of pervasive transcription, or the fact that most if not the whole human genome is being actively transcribed, is still debated. However, while only about 1.2 % of the human genome encode for protein coding genes, it is well admitted now that a vast majority of transcripts corresponds to non-protein coding transcripts, the so-called non-coding RNAs (ncRNAs). While more and more (very) long non-coding RNAs (lincRNAs) are being identified, with sizes up to several kilobases, most non-coding RNAs known to date are rather small, with sizes ranging from 20 or less to 200 nucleotides. The discovery of the fascinating class of microRNAs (miRNAs) about a decade ago as well as the availabilities of genome sequences and progresses in next-generation sequencing techniques dramatically boosted the attention of researchers in the field. We now know that the many classes of small non-coding RNAs are involved in all biological pathways, such as RNA processing or modification, gene expression regulation at the transcriptional or posttranscriptional levels, translation, or even protein secretion. However, our actual knowledge is only the tip of the iceberg. Many questions are yet to be answered, especially regarding the implications, direct or indirect, of small non-coding RNAs with numerous disorders, suggesting more and more their possible and powerful usage as diagnostic markers and/or therapeutic tools or targets. Owing to their small sizes, tridimensional structures, low abundances, or differential expression levels, small non-coding RNAs require customized dedicated protocols for their identification and study, compared, for instance, to messenger RNAs (mRNAs). Small Non-coding RNAs: Methods and Protocols is a laboratory protocols book dedicated to biochemists or cellular/molecular biologists, already working in the field of RNA biology or willing to start studying small non-coding RNAs in their projects. It describes basic as well as more sophisticated, state-of-the-art methods to tackle all aspects of small non-coding RNAs biology, from their identification or biogenesis to their use in therapeutics. This survey of technologies will be of valuable help to all those willing to contribute deciphering the numerous functions of small non-coding RNAs. I thank all the authors and editors for their outstanding contributions to this volume. Vandoeuvre-lès-Nancy, France

Mathieu Rederstorff

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

PART I

INTRODUCTION TO SMALL NON-CODING RNAS

1 Small Non-Coding RNAs: A Quick Look in the Rearview Mirror . . . . . . . . . . Guillaume Clerget, Yoann Abel, and Mathieu Rederstorff 2 Alcoholic Precipitation of Small Non-Coding RNAs . . . . . . . . . . . . . . . . . . . . Guillaume Clerget, Valérie Bourguignon-Igel, and Mathieu Rederstorff 3 Quantification and Quality Control of a Small Non-Coding RNA Preparation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Virginie Marchand and Christiane Branlant 4 Impact of RNA Isolation Protocols on RNA Detection by Northern Blotting . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Katrin Damm, Simone Bach, Katrin M.H. Müller, Gabriele Klug, Olga Y. Burenina, Elena A. Kubareva, Arnold Grünweller, and Roland K. Hartmann

PART II

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VISUALIZATION AND ANALYSIS OF SMALL NON-CODING RNAS

5 Improved Northern Blot Detection of Small RNAs Using EDC Crosslinking and DNA/LNA Probes. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Katrin Damm, Simone Bach, Katrin M.H. Müller, Gabriele Klug, Olga Y. Burenina, Elena A. Kubareva, Arnold Grünweller, and Roland K. Hartmann 6 Direct Cloning of Double-Stranded RNAs . . . . . . . . . . . . . . . . . . . . . . . . . . . Manli Shen, Marina Falaleeva, Natalia Korotkova, and Stefan Stamm 7 Detection and Labeling of Small Non-Coding RNAs by Splinted Ligation. . . . Gabrielle Bourgeois, Florian Chardon, Anne-Sophie Tillault, and Magali Blaud 8 Fluorescence In Situ Hybridization of Small Non-Coding RNAs . . . . . . . . . . . Valentin Vautrot, Christelle Aigueperse, Christiane Branlant, and Isabelle Behm-Ansmant 9 RT-qPCR-Based Quantification of Small Non-Coding RNAs . . . . . . . . . . . . . Fjoralba Zeka, Pieter Mestdagh, and Jo Vandesompele 10 Stem-Loop RT-PCR Based Quantification of Small Non-Coding RNAs . . . . . Véronique Salone and Mathieu Rederstorff 11 miR-RACE: An Effective Approach to Accurately Determine the Sequence of Computationally Identified miRNAs . . . . . . . . . . . . . . . . . . . Chen Wang and Jinggui Fang

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12 Probing Small Non-Coding RNAs Structures . . . . . . . . . . . . . . . . . . . . . . . . . Jean-Vincent Philippe, Lilia Ayadi, Christiane Branlant, and Isabelle Behm-Ansmant

PART III

HIGH-THROUGHPUT APPROACHES TO STUDY NON-CODING RNAS

13 cDNA Library Generation for the Analysis of Small RNAs by High-Throughput Sequencing . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Jennifer Gebetsberger, Roger Fricker, and Norbert Polacek 14 CLIP-Seq to Discover Transcriptome-Wide Imprinting of RNA Binding Proteins in Living Cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Jérôme Saulière and Hervé Le Hir 15 Microarray Analysis of Small Non-Coding RNAs. . . . . . . . . . . . . . . . . . . . . . . Michael Karbiener and Marcel Scheideler

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SMALL NON-CODING RNAS APPLICATIONS

16 RLM-RACE, PPM-RACE, and qRT-PCR: An Integrated Strategy to Accurately Validate miRNA Target Genes . . . . . . . . . . . . . . . . . . . . . . . . . . Chen Wang and Jinggui Fang 17 Dual Luciferase Gene Reporter Assays to Study miRNA Function . . . . . . . . . . Thomas Clément, Véronique Salone, and Mathieu Rederstorff 18 Gene Expression Knockdown by Transfection of siRNAs into Mammalian Cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Yoann Abel and Mathieu Rederstorff 19 Efficient and Selective Knockdown of Small Non-Coding RNAs . . . . . . . . . . . Xue-Hai Liang, Wen Shen, and Stanley T. Crooke 20 Cell-SELEX: In Vitro Selection of Synthetic Small Specific Ligands . . . . . . . . . Helena Dickinson, Melanie Lukasser, Günter Mayer, and Alexander Hüttenhofer 21 Small Non-Coding RNAs and Aptamers in Diagnostics and Therapeutics . . . . Marissa Leonard, Yijuan Zhang, and Xiaoting Zhang Index . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Contributors YOANN ABEL • CNRS UMR 7365 IMoPA, Université de Lorraine, Vandoeuvre-lès-Nancy, France CHRISTELLE AIGUEPERSE • CNRS UMR 7365 IMoPA, Université de Lorraine, Vandoeuvre-lès-Nancy, France LILIA AYADI • CNRS UMR 7365 IMoPA, Université de Lorraine, Vandoeuvre-lès-Nancy, France SIMONE BACH • Institut für Pharmazeutische Chemie, Philipps-Universität Marburg, Marburg, Germany ISABELLE BEHM-ANSMANT • CNRS UMR 7365 IMoPA, Université de Lorraine, Vandoeuvre-lès-Nancy, France MAGALI BLAUD • Laboratoire de Cristallographie et RMN Biologiques, CNRS UMR 8015, Université Paris Descartes, Paris, France GABRIELLE BOURGEOIS • Laboratoire de Biochimie, CNRS UMR 7654, Ecole Polytechnique, Palaiseau, France VALERIE BOURGUIGNON-IGEL • CNRS UMR 7365 IMoPA, Université de Lorraine, Vandoeuvre-lès-Nancy, France CHRISTIANE BRANLANT • CNRS UMR 7365 IMoPA, Université de Lorraine, Vandoeuvre-lès-Nancy, France; Centre Hospitalier Universitaire de Nancy, Vandoeuvre-lès-Nancy, France OLGA Y. BURENINA • Chemistry Department and A.N. Belozersky Institute of Physico-Chemical Biology, M.V. Lomonosov Moscow State University, Moscow, Russia FLORIAN CHARDON • Laboratoire de Cristallographie et RMN Biologiques, CNRS UMR 8015, Université Paris Descartes, Paris, France THOMAS CLÉMENT • CNRS UMR 7365 IMoPA, Université de Lorraine, Vandoeuvre-lès-Nancy, France GUILLAUME CLERGET • CNRS UMR 7365 IMoPA, Université de Lorraine, Vandoeuvre-lès-Nancy, France STANLEY T. CROOKE • Department of Core Antisense Research, ISIS Pharmaceuticals Inc., Carlsbad, CA, USA KATRIN DAMM • Institut für Pharmazeutische Chemie, Philipps-Universität Marburg, Marburg, Germany HELENA DICKINSON • Center of Chemistry & Biomedicine, Innsbruck Medical University, Innsbruck, Austria MARINA FALALEEVA • University of Kentucky, Lexington, KY, USA JINGGUI FANG • College of Horticulture, Nanjing Agricultural University, Nanjing, China ROGER FRICKER • Department of Chemistry and Biochemistry, Graduate School for Cellular and Biomedical Sciences, University of Bern, Bern, Switzerland JENNIFER GEBETSBERGER • Department of Chemistry and Biochemistry, Graduate School for Cellular and Biomedical Sciences, University of Bern, Bern, Switzerland ARNOLD GRÜNWELLER • Institut für Pharmazeutische Chemie, Philipps-Universität Marburg, Marburg, Germany

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ROLAND K. HARTMANN • Institut für Pharmazeutische Chemie, Philipps-Universität Marburg, Marburg, Germany HERVÉ LE HIR • Ecole Normale Supérieure, Institut de Biologie de l’ENS (IBENS), Inserm U1024, and CNRS UMR 8197, Paris, France ALEXANDER HÜTTENHOFER: • Center of Chemistry & Biomedicine, Innsbruck Medical University, Innsbruck, Austria MICHAEL KARBIENER • RNA Biology Group, Institute of Molecular Biotechnology, Graz University of Technology, Graz, Austria GABRIELE KLUG • Institut für Mikrobiologie und Molekularbiologie, Justus-Liebig-Universität Gießen, Gießen, Germany NATALIA KOROTKOVA • University of Kentucky, Lexington, KY, USA ELENA A. KUBAREVA • Chemistry Department and A.N. Belozersky Institute of Physico-Chemical Biology, M.V. Lomonosov Moscow State University, Moscow, Russia MARISSA LEONARD • Department of Cancer Biology, Graduate Program in Cancer and Cell Biology, Vontz Center for Molecular Studies, University of Cincinnati College of Medicine, Cincinnati, OH, USA XUE-HAI LIANG • Department of Core Antisense Research, ISIS Pharmaceuticals Inc., Carlsbad, CA, USA MELANIE LUKASSER • Center of Chemistry & Biomedicine, Innsbruck Medical University, Innsbruck, Austria VIRGINIE MARCHAND • Centre Hospitalier Universitaire de Nancy, Nancy, France; CNRS UMR 7365 IMoPA, Université de Lorraine, Vandoeuvre-lès-Nancy, France GÜNTER MAYER • Life & Medical Sciences Institute (LIMES), Chemical Biology, University of Bonn, Bonn, Germany PIETER MESTDAGH • Center for Medical Genetics, Ghent University, Ghent, Belgium KATRIN M.H. MÜLLER • Institut für Mikrobiologie und Molekularbiologie, Justus-Liebig-Universität Gießen, Gießen, Germany JEAN-VINCENT PHILIPPE • CNRS UMR 7365 IMoP, Université de Lorraine, Vandoeuvre-lès-Nancy, France NORBERT POLACEK • Department of Chemistry and Biochemistry, University of Bern, Bern, Switzerland MATHIEU REDERSTORFF • Université de Lorraine, Biopôle, CNRS UMR 7365, IMoPA, Vandoeuvre-lès-Nancy, France VÉRONIQUE SALONE • CNRS UMR 7365 IMoPA, Université de Lorraine, Vandoeuvre-lès-Nancy, France JÉRÔME SAULIÈRE • Ecole Normale Supérieure, Institut de Biologie de l’ENS (IBENS), Inserm U1024, and CNRS UMR 8197, Paris, France MARCEL SCHEIDELER • Institute for Diabetes and Cancer (IDC), Helmholtz Zentrum München, Neuherberg, Germany; German Center for Diabetes Research (DZD), Neuherberg, Germany MANLI SHEN • University of Kentucky, Lexington, KY, USA WEN SHEN • Department of Core Antisense Research, ISIS Pharmaceuticals Inc., Carlsbad, CA, USA STEFAN STAMM • University of Kentucky, Lexington, KY, USA ANNE-SOPHIE TILLAULT • Department of Chemistry and Biochemistry, Alberta RNA Research and Training Institute, University of Lethbridge, Lethbridge, AB, Canada JO VANDESOMPELE • Center for Medical Genetics, Ghent University, Ghent, Belgium

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Contributors

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VALENTIN VAUTROT • CNRS UMR 7365 IMoPA, Université de Lorraine, Vandoeuvre-lès-Nancy, France CHEN WANG • College of Horticulture, Nanjing Agricultural University, Nanjing, China FJORALBA ZEKA • Center for Medical Genetics, Ghent University, Ghent, Belgium XIAOTING ZHANG • Department of Cancer Biology, Vontz Center for Molecular Studies, Cincinnati Cancer Center, University of Cincinnati College of Medicine, Cincinnati, OH, USA YIJUAN ZHANG • Department of Cancer Biology, University of Cincinnati College of Medicine, Cincinnati, OH, USA

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Part I Introduction to Small Non-Coding RNAs

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Chapter 1 Small Non-Coding RNAs: A Quick Look in the Rearview Mirror Guillaume Clerget, Yoann Abel, and Mathieu Rederstorff Abstract The revolution of miRNA discovery, in the early 2000s, shed a new light in the exciting field of small non-coding RNAs. Since then, and owing to outstanding breakthroughs in RNomic techniques, novel small non-coding RNA families have been regularly discovered, e.g., piRNAs, tiRNAs, and many others. In this review, we provide a very succinct historical and functional overview on most prominent small non-coding RNA families. Key words miRNA, snoRNA, piRNA, tRNA, rRNA, tiRNA, snRNA

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Introduction

1.1 Dawn of Small Non-Coding RNAs

It has been more than 60 years that the first soluble cytoplasmic small non-coding RNA (sncRNA), a yeast alanine transfer RNA (tRNA), was identified and its primary structure determined [1, 2]. Together with ribosomal RNAs (rRNA), the nucleic acid moieties of the ribosomes, tRNAs play a central role in protein synthesis, as the adaptor molecules providing the correct amino acid in response to a specific codon on the mRNA [3]. Only 20 years later were the first nuclear RNAs uncovered: uridine-rich RNAs, or U-RNAs, were observed to be very abundant, nucleus or sometimes even nucleolus specific, and highly conserved [4, 5]; small nuclear RNAs (snRNA) U1, U2, U4, U5, and U6 were later shown to be involved in splicing, in association with several proteins within a ribonucleoprotein particle (RNP) called spliceosome [6]. Base pairing among snRNAs and between snRNAs and pre-messenger RNA (pre-mRNA) regions account for the proper conformation of both spliceosome and premRNA, eventually leading to release of the introns and mature mRNAs [7]. Interestingly, most small nucleolar RNAs (snoRNAs) are processed from the released introns [8]. Within snoRNPs,

Author contributed equally with all other contributors. Mathieu Rederstorff (ed.), Small Non-Coding RNAs: Methods and Protocols, Methods in Molecular Biology, vol. 1296, DOI 10.1007/978-1-4939-2547-6_1, © Springer Science+Business Media New York 2015

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snoRNAs guide posttranscriptional RNA modifications of other ncRNAs, e.g., rRNAs and snRNAs: 2′-O-methylations for C/D box snoRNAs and pseudouridylation for H/ACA box snoRNAs. Some snoRNAs, such as U3 or U8, are instead involved in early cleavages of pre-rRNA during ribosome biogenesis in the nucleolus [9]. In the late 1990s, pioneering studies aimed at identifying novel non-coding RNAs using experimental approaches combining library generation and sequencing identified numerous novel snoRNAs in various models including human, using material that was usually discarded: small total RNA in the size range of 50–500 nucleotides [10]. At that time, why would one have looked for smaller functional RNAs? It would both have been nonsense and a waste of time. Nonsense? Actually no… antisense! 1.2 miRNAs: The Revolution!

In the early 1990s, the lin-14 gene was shown to be involved in temporal development in the worm C. elegans. The lin-4 gene product, most probably a polypeptide, was shown to negatively regulate lin-14 expression by directly acting on the 3′ untranslated region of its pre-mRNA [11]. In 1993, both Ambros and Ruvkun labs discovered that the lin-4 gene product actually was a small non-coding RNA, of 22 nucleotides in length in its mature form, targeting the 3′ UTR of the lin-14 pre-mRNA several times, owing to imperfect antisense base pairing [12, 13]. Unfortunately enough, apparently, the lin-4 small RNA neither had additional counterparts in worm nor was it found to be conserved in other species. However, in 2000, Pasquinelli and coworkers identified that let-7, another 22-nucleotide-long small temporal RNA (stRNA) involved in C. elegans developmental timing, was conserved in human and drosophila, anticipating the likely discovery of many other small regulatory RNAs [14]. Revolution came less than 1 year later, when Tuschl, Bartel, and Ambros labs reported, in the same issue of Science, the observation of an abundant class of novel small regulatory RNAs in worm, human, and drosophila, termed since microRNAs (miRNAs) [15–17]. Even certain viruses were next shown to encode their own miRNAs [18, 19]. It was rapidly observed that the miRNA maturation and assembly machinery were the same as the one implicated in the formation of short interfering RNAs (siRNAs), involved in the RNA interference (RNAi) pathway described in worm a couple of years earlier by the 2006 Nobel Prize awardees Andrew Fire and Craig Mello [20, 21] and even earlier in plants [22]. miRNA/siRNA maturation, from a double-stranded precursor, is now well understood [23] but still open to surprises; recently it was discovered that miRNAs could be processed from other types of ncRNAs, such as snoRNAs [24, 25]. In human, siRNAs, within the RNA-induced silencing complex (RISC), mediate perfectly complementary mRNA target cleavages

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ncRNAs Overview

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[26, 27]. On the other hand, miRNAs regulate gene expression by imperfectly base pairing to the 3′ UTR of the targeted mRNA, repressing its translation, which eventually leads to the mRNA decay by different possible mechanisms [28, 29]. miRNAs can act as regulators of epigenetic modifications as well, via DNA or histone modifications, therefore controlling gene expression at the transcriptional level [30]. 1.3 Small Non-Coding RNAs Return!

Another sncRNA family, the Piwi-interacting RNAs (piRNA), discovered about half a decade after miRNAs, also regulates transcription [31]. piRNAs are 27–29-nucleotide-long RNAs. They prevent transposon dissemination in germinal cell lines, owing to an original “ping-pong” mechanism [32, 33]. The smallest eukaryotic ncRNAs known to date, the transcription initiation RNAs (tiRNAs), sized 17–18 nucleotides, are generated upon stalling or backtracking of RNA polymerase II (RNAPII) near the transcription start sites (TSSs) and might be involved in transcription initiation regulation as well [34, 35]. TSSs appear to be the source of bunches of small and long non-coding RNAs (lncRNAs) [36], such as transcription start site-associated RNAs (TSSa-RNAs) and promoter-associated RNAs (PASRs) in animals [37] or promoter upstream transcripts (PROMPTs) [38] and cryptic/Xrn1-sensitive unstable transcripts (CUTs/XUTs) in yeast [39–41]. While lncRNAs are generally involved in epigenetic and chromatin modifications [42–44], the functionality of short transcripts is debated. They might be involved in maintaining chromatin into a transcriptionally active state or in maintaining RNAPII available near TSSs [45].

1.4 Small Non-Coding RNA: Not the End

Most if not all the human genome appears to be actively transcribed [46–48]. Nevertheless, most transcripts rather belong to the TUF (transcripts of unknown function) rather than to the ncRNA family. Progresses in transcriptomics/RNomics enabled the identification of thousands of short and long transcripts [49–55]. Many of them were attributed a function but it is only the top of the iceberg. Whether every TUF is indeed functional is debated, and experimental efforts will be mandatory to answer this question. There are no doubts that novel ncRNAs and novel ncRNA families will still be discovered in the (next) future.

Acknowledgment This work was supported by the Centre National pour la Recherche Scientifique, the Université de Lorraine, the Région Lorraine and La Ligue contre le Cancer.

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Guillaume Clerget et al. yeast. Nature 457(7232):1033–1037. doi:10.1038/nature07728, nature07728 [pii] Mercer TR, Dinger ME, Mattick JS (2009) Long non-coding RNAs: insights into functions. Nat Rev Genet 10(3):155–159. doi:10.1038/nrg2521, nrg2521 [pii] Ponting CP, Oliver PL, Reik W (2009) Evolution and functions of long noncoding RNAs. Cell 136(4):629–641. doi:10.1016/j. cell.2009.02.006, S0092-8674(09)00142-1 [pii] Yazgan O, Krebs JE (2007) Noncoding but nonexpendable: transcriptional regulation by large noncoding RNA in eukaryotes. Biochem Cell Biol 85(4):484–496. doi:10.1139/O07061, o07-061 [pii] Aalto AP, Pasquinelli AE (2012) Small noncoding RNAs mount a silent revolution in gene expression. Curr Opin Cell Biol 24(3):333– 340. doi:10.1016/j.ceb.2012.03.006, S09550674(12)00037-3 [pii] Birney E, Stamatoyannopoulos JA, Dutta A, Guigo R, Gingeras TR, Margulies EH, Weng Z, Snyder M, Dermitzakis ET, Thurman RE, Kuehn MS, Taylor CM, Neph S, Koch CM, Asthana S, Malhotra A, Adzhubei I, Greenbaum JA, Andrews RM, Flicek P, Boyle PJ, Cao H, Carter NP, Clelland GK, Davis S, Day N, Dhami P, Dillon SC, Dorschner MO, Fiegler H, Giresi PG, Goldy J, Hawrylycz M, Haydock A, Humbert R, James KD, Johnson BE, Johnson EM, Frum TT, Rosenzweig ER, Karnani N, Lee K, Lefebvre GC, Navas PA, Neri F, Parker SC, Sabo PJ, Sandstrom R, Shafer A, Vetrie D, Weaver M, Wilcox S, Yu M, Collins FS, Dekker J, Lieb JD, Tullius TD, Crawford GE, Sunyaev S, Noble WS, Dunham I, Denoeud F, Reymond A, Kapranov P, Rozowsky J, Zheng D, Castelo R, Frankish A, Harrow J, Ghosh S, Sandelin A, Hofacker IL, Baertsch R, Keefe D, Dike S, Cheng J, Hirsch HA, Sekinger EA, Lagarde J, Abril JF, Shahab A, Flamm C, Fried C, Hackermuller J, Hertel J, Lindemeyer M, Missal K, Tanzer A, Washietl S, Korbel J, Emanuelsson O, Pedersen JS, Holroyd N, Taylor R, Swarbreck D, Matthews N, Dickson MC, Thomas DJ, Weirauch MT, Gilbert J, Drenkow J, Bell I, Zhao X, Srinivasan KG, Sung WK, Ooi HS, Chiu KP, Foissac S, Alioto T, Brent M, Pachter L, Tress ML, Valencia A, Choo SW, Choo CY, Ucla C, Manzano C, Wyss C, Cheung E, Clark TG, Brown JB, Ganesh M, Patel S, Tammana H, Chrast J, Henrichsen CN, Kai C, Kawai J, Nagalakshmi U, Wu J, Lian Z, Lian J, Newburger P, Zhang X, Bickel P, Mattick JS, Carninci P, Hayashizaki Y, Weissman S, Hubbard T, Myers RM, Rogers J, Stadler PF,

Lowe TM, Wei CL, Ruan Y, Struhl K, Gerstein M, Antonarakis SE, Fu Y, Green ED, Karaoz U, Siepel A, Taylor J, Liefer LA, Wetterstrand KA, Good PJ, Feingold EA, Guyer MS, Cooper GM, Asimenos G, Dewey CN, Hou M, Nikolaev S, Montoya-Burgos JI, Loytynoja A, Whelan S, Pardi F, Massingham T, Huang H, Zhang NR, Holmes I, Mullikin JC, UretaVidal A, Paten B, Seringhaus M, Church D, Rosenbloom K, Kent WJ, Stone EA, Batzoglou S, Goldman N, Hardison RC, Haussler D, Miller W, Sidow A, Trinklein ND, Zhang ZD, Barrera L, Stuart R, King DC, Ameur A, Enroth S, Bieda MC, Kim J, Bhinge AA, Jiang N, Liu J, Yao F, Vega VB, Lee CW, Ng P, Yang A, Moqtaderi Z, Zhu Z, Xu X, Squazzo S, Oberley MJ, Inman D, Singer MA, Richmond TA, Munn KJ, Rada-Iglesias A, Wallerman O, Komorowski J, Fowler JC, Couttet P, Bruce AW, Dovey OM, Ellis PD, Langford CF, Nix DA, Euskirchen G, Hartman S, Urban AE, Kraus P, Van Calcar S, Heintzman N, Kim TH, Wang K, Qu C, Hon G, Luna R, Glass CK, Rosenfeld MG, Aldred SF, Cooper SJ, Halees A, Lin JM, Shulha HP, Xu M, Haidar JN, Yu Y, Iyer VR, Green RD, Wadelius C, Farnham PJ, Ren B, Harte RA, Hinrichs AS, Trumbower H, Clawson H, Hillman-Jackson J, Zweig AS, Smith K, Thakkapallayil A, Barber G, Kuhn RM, Karolchik D, Armengol L, Bird CP, de Bakker PI, Kern AD, Lopez-Bigas N, Martin JD, Stranger BE, Woodroffe A, Davydov E, Dimas A, Eyras E, Hallgrimsdottir IB, Huppert J, Zody MC, Abecasis GR, Estivill X, Bouffard GG, Guan X, Hansen NF, Idol JR, Maduro VV, Maskeri B, McDowell JC, Park M, Thomas PJ, Young AC, Blakesley RW, Muzny DM, Sodergren E, Wheeler DA, Worley KC, Jiang H, Weinstock GM, Gibbs RA, Graves T, Fulton R, Mardis ER, Wilson RK, Clamp M, Cuff J, Gnerre S, Jaffe DB, Chang JL, Lindblad-Toh K, Lander ES, Koriabine M, Nefedov M, Osoegawa K, Yoshinaga Y, Zhu B, De Jong PJ (2007) Identification and analysis of functional elements in 1% of the human genome by the ENCODE pilot project. Nature 447(7146):799–816. doi:10.1038/nature05874 47. Clark MB, Amaral PP, Schlesinger FJ, Dinger ME, Taft RJ, Rinn JL, Ponting CP, Stadler PF, Morris KV, Morillon A, Rozowsky JS, Gerstein MB, Wahlestedt C, Hayashizaki Y, Carninci P, Gingeras TR, Mattick JS (2011) The reality of pervasive transcription. PLoS Biol 9(7):e1000625. doi:10.1371/journal. pbio.1000625, discussion e1001102, 10-PLBI-PS-8040R3 [pii] 48. Clark MB, Choudhary A, Smith MA, Taft RJ, Mattick JS (2013) The dark matter rises: the expanding world of regulatory RNAs. Essays

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Biochem 54:1–16. doi:10.1042/bse0540001, bse0540001 [pii] Jochl C, Rederstorff M, Hertel J, Stadler PF, Hofacker IL, Schrettl M, Haas H, Huttenhofer A (2008) Small ncRNA transcriptome analysis from Aspergillus fumigatus suggests a novel mechanism for regulation of protein synthesis. Nucleic Acids Res 36(8):2677–2689. doi:10.1093/nar/gkn123, gkn123 [pii] Metzker ML (2010) Sequencing technologies – the next generation. Nat Rev Genet 11(1):31– 46. doi:10.1038/nrg2626, nrg2626 [pii] Rederstorff M (2012) Generation of cDNA libraries from RNP-derived regulatory noncoding RNAs. Methods Mol Biol 925:211– 218. doi:10.1007/978-1-62703-011-3_14 Rederstorff M, Bernhart SH, Tanzer A, Zywicki M, Perfler K, Lukasser M, Hofacker IL, Huttenhofer A (2010) RNPomics: defining the ncRNA transcriptome by cDNA library generation from ribonucleo-protein particles.

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Nucleic Acids Res 38(10):e113. doi:10.1093/ nar/gkq057, gkq057 [pii] 53. Rederstorff M, Huttenhofer A (2011) cDNA library generation from ribonucleoprotein particles. Nat Protoc 6(2):166–174. doi:10.1038/ nprot.2010.186, nprot.2010.186 [pii] 54. Shen M, Eyras E, Wu J, Khanna A, Josiah S, Rederstorff M, Zhang MQ, Stamm S (2011) Direct cloning of double-stranded RNAs from RNase protection analysis reveals processing patterns of C/D box snoRNAs and provides evidence for widespread antisense transcript expression. Nucleic Acids Res 39(22):9720– 9730. doi:10.1093/nar/gkr684, gkr684 [pii] 55. Skreka K, Schafferer S, Nat IR, Zywicki M, Salti A, Apostolova G, Griehl M, Rederstorff M, Dechant G, Huttenhofer A (2012) Identification of differentially expressed non-coding RNAs in embryonic stem cell neural differentiation. Nucleic Acids Res 40(13):6001–6015. doi:10.1093/nar/gks311, gks311 [pii]

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Chapter 2 Alcoholic Precipitation of Small Non-Coding RNAs Guillaume Clerget, Valérie Bourguignon-Igel, and Mathieu Rederstorff Abstract Alcoholic precipitation is a critical step to recover RNA of high purity. This chapter describes the principles of alcoholic precipitation as well as a standard, basic protocol with key advices to observe, but numerous variations on the theme are discussed. Indeed, several important parameters, such as the choice of salt, alcohol, or carrier, have to be considered to improve the efficiency of precipitation and the yield of RNA recovery. Key words RNA, Precipitation, Alcohol, Salt, Carrier

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Introduction Alcoholic precipitation is used to recover, purify, or concentrate nucleic acids from a total extract. It relies on nucleic acids’ differential solubility in solution upon addition of alcohol and salts. Nucleic acids are soluble in water because both nucleic acids and water are polar molecules that can therefore interact together [1]. The negatively charged phosphate groups (PO3−) of nucleic acids electrostatically interact with water. Hence, water molecules form a solvation or hydration shell around each nucleic acid molecule, isolating them from each other and subsequently dissolving them in solution. Alcohol and salt enable to disrupt this hydration shell. In water, salts used for nucleic acid precipitation completely dissociate into their constituting anions and cations. Cations neutralize the negative charges of nucleic acids (PO3− groups) while alcohol, which has a much lower dielectric constant than water, increases the electrostatic interaction force between the phosphate groups and the cations. Once neutralized, nucleic acids become less hydrophilic and thus precipitate out of solution. Additionally, alcohol decreases the repulsive forces between the inter-helical phosphates so that nucleic acid molecules aggregate. Depending on the sample type or the subsequent experiments to be performed, several parameters such as the type of salt

Mathieu Rederstorff (ed.), Small Non-Coding RNAs: Methods and Protocols, Methods in Molecular Biology, vol. 1296, DOI 10.1007/978-1-4939-2547-6_2, © Springer Science+Business Media New York 2015

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or alcohol to be used, the addition of a carrier, the precipitation temperature, or the centrifugation speed have to be considered to improve RNA quality and recovery yield and are discussed (see Notes 1–5).

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Materials 1. 100 % ethanol. 2. 70 % ethanol. 3. 100 % isopropanol. 4. 3 M sodium acetate, pH 5.2. 5. 5 M ammonium acetate. 6. 8 M lithium chloride. 7. 2 M sodium chloride. 8. Yeast tRNA (10–20 μg/ml). 9. Salmon sperm DNA (10–20 μg/ml). 10. Glycogen (50–150 μg/ml). 11. GlycoBlue (50–150 μg/ml) (Ambion). 12. Linear polyacrylamide (10–20 μg/ml). 13. 1 M MgCl2. 14. RNase-free, DEPC-treated water. 15. 1× TE buffer, pH 8.0: 10 mM Tris–HCl, pH 8.0, 1 mM EDTA, pH 8.0.

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Methods To avoid RNase contaminations when preparing and handling RNAs, always use appropriate precautions such as RNase-free water (see Note 6), gloves, and RNase-free tubes and tips.

3.1 Ethanol Precipitation

1. Adjust sample volume to a minimum of 100 μl (see Note 7). 2. Add sodium acetate to the sample to a final concentration of 0.3 M (about 1/10 of volume) (see Notes 1, 3 and 8). 3. Add 2.5–3 volumes of ice-cold 100 % ethanol (see Note 2). 4. Mix thoroughly by inverting the tube or pipetting up and down (see Note 9). 5. Incubate on ice for 15 min to 1 h (see Note 4). 6. Centrifuge at 12,000 × g for 30 min at 4 °C (see Note 5). 7. Carefully discard the supernatant without disturbing the pellet.

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8. Wash the RNA pellet with 0.5 ml of 70 % ethanol and mix gently. 9. Centrifuge at 12,000 × g for 15 min at 4 °C. 10. Carefully discard the supernatant without disturbing the pellet (see Note 10). 11. Air-dry the RNA pellet for 5–10 min at room temperature (see Note 11). 12. Dissolve the pellet in RNase-free water (see Note 12).

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Notes 1. Different salts can be used during alcoholic precipitation, the choice of which depending on the subsequent applications. Indeed, each salt features specific properties providing different advantages and/or drawbacks [1, 2]. Sodium acetate (0.3 M final concentration, pH 5.2) is the most widely used salt for alcoholic precipitation [1, 2]. However, if the sample contains SDS, sodium chloride (0.2 M final concentration) could be alternatively chosen. In this case, SDS would remain soluble in ethanol [1, 2], which would allow its separation and removal from the RNA. Ammonium acetate (2.5 M final concentration) is another possible choice. It allows to greatly reduce coprecipitation of both dNTPs and oligosaccharides. However, ammonium acetate should not be used if a phosphorylation reaction is planned after precipitation since ammonium ions inhibit T4 polynucleotide kinase [1, 2]. Finally, a last broadly used salt for RNA precipitation is lithium chloride (0.8 M final concentration). It is very soluble in alcohol and therefore coprecipitates with RNA to a smaller extent than others salts [1, 2]. Moreover, DNA, proteins, and carbohydrates do not efficiently precipitate with lithium chloride, which therefore leads to a greater purity of RNAs precipitated with this salt [3]. Unfortunately, possible loss of small RNAs has been reported (≈100 nt or less, e.g., miRNAs, tRNAs, snand snoRNAs, or 5S/5.8S rRNAs). Anyway, as small and large RNAs feature different solubility in high ionic strength solutions, small RNAs remaining soluble while larger ones do not, this property enables to selectively purify small RNAs with high concentrations of LiCl (up to 2.5 M) even without alcohol [1], as small RNAs will remain in solution. Finally, lithium chloride should not be employed if subsequent reverse transcription or in vitro translation reactions are planned, as the chloride ions at this concentration inhibit RNA-dependent DNA polymerase as well as initiation of protein synthesis in most cell-free systems [1].

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2. Alternatively, isopropanol can replace ethanol as a precipitation alcohol. However, since isopropanol is less polar than ethanol, RNAs are even less soluble in isopropanol than they are in ethanol. Therefore, for a similar precipitation efficiency, a lower volume of isopropanol is necessary [1, 2]. Isopropanol can therefore replace ethanol in the case of large sample volume, using about one volume of isopropanol per volume of sample. However, as salts are also less soluble in isopropanol, they coprecipitate and contaminate RNA preparation to a higher extent [2]. Therefore, washing steps after precipitation with isopropanol are very important. Finally, isopropanol is less volatile than ethanol and will require more time for the RNA pellet to dry [2]. 3. If you expect RNA quantity to be very low or if RNAs are very small in size (8 nt). However, glycogen may prevent reverse transcription in the case of very large templates, as well as nucleic acid interactions with proteins [7] in a concentrationdependent manner [8]. Finally, linear polyacrylamide is another DNA/RNA-free inert molecule. As glycogen, it neither

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interferes with enzymatic reactions such as phosphorylation by polynucleotide kinase, ligation, cDNA synthesis, in vitro transcription, and digestion by endonucleases nor with electrophoresis [9]. Linear polyacrylamide does not inhibit nucleic acid interactions with proteins [10]; however, it does not trigger coprecipitation of very small RNA (≤20 bases) [7]. 4. The incubation step for the alcoholic precipitation of RNAs is generally performed at very low temperature (−20 °C or −80 °C) in most protocols. However, although possible RNases might be less active at lower temperatures, this appears to be rather counterproductive [1, 2]. Indeed, the lower the temperature, the higher the viscosity, which decreases RNA movements and thus aggregation and precipitation. Additionally, the dielectric constant increases when the temperature diminishes; therefore precipitation efficiency decreases with the temperature. Finally, salts’ solubility decreases with the temperature as well, which leads to stronger coprecipitation of salts at lower temperatures [11]. Therefore, incubation at room temperature or on ice is sufficient and recommended, provided you work in RNase-free conditions. If you retrieve a low amount of RNA or if you do not see the pellet, you can increase the incubation time or add a carrier if not done already. An average time of about 15 min is generally sufficient. 5. If the amount of RNA is low, if you do not see the pellet, or if you work with very small RNAs (100 nt), whereas TRIzol methods are the methods of choice to enrich tiny RNAs (~14 nt).

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Materials Prepare all solutions with autoclaved ddH2O and analytical grade reagents for molecular biology use. Prepare reagents at room temperature; store them at room temperature (short-term storage) or as aliquots at −20 °C (long-term storage), if not stated otherwise. The procedures described herein have to be performed under biosafety S1 conditions.

2.1 Bacterial Cell Cultures

1. Warm air incubation shaker. 2. Water bath shaker. 3. Dewar flask with handle for liquid nitrogen. 4. Centrifuges for 1.5, 15, and 50-ml tubes. 5. LB medium: 10 g/l peptone, 5 g/l yeast extract, 10 g/l NaCl. Adjust to pH 7.5 with NaOH and autoclave. 6. The following bacterial strains were used in this protocol: Bacillus subtilis PY79 (wt) and B. subtilis PY79 ΔbsrA (6S-1 RNA deletion strain). Glycerol stocks stored at −80 °C were used to streak out bacteria on appropriate agar plates, and single colonies were picked to inoculate fresh liquid LB media.

2.2

RNA Isolation

1. Extraction buffer: 10 mM NaOAc, 150 mM sucrose, adjusted to pH 4.8 with acetic acid. Sterilize by filtration. 2. Lysozyme solution: 20 mg/ml in 1× TE buffer. Sterilize by filtration. 3. 20 % (w/v) SDS solution. 4. Acidic phenol (Roth). Store at 4–10 °C. 5. Chloroform. 6. TRIzol® Reagent (Ambion). 7. Phase Lock Gel™ (5Prime).

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8. 3 M sodium acetate, pH 5.0. Store at −20 °C. 9. 100 % ethanol. Store at −20 °C. 10. Isopropanol. 11. 75 % ethanol (100 % ethanol diluted with ddH2O (v/v)). 2.3 Control of Total RNA Quality

1. 2× denaturing loading buffer: 0.02 % (w/v) bromophenol blue, 0.02 % (w/v) xylene cyanol blue, 8 M urea, 50 % (v/v) formamide, 2× TBE. 2. Denaturing polyacrylamide gel: 1× TBE, polyacrylamide/ bisacrylamide (24:1), 8 M urea. For polymerization, add 0.1 % (w/v) of 10 % ammonium persulfate (APS) and 0.1 % (v/v) of (N,N,N′,N′-tetramethylethylenediamine) (TEMED). 3. Ethidium bromide nucleic acid staining solution: 0.5 μg/ml ethidium bromide, 1× TBE. 4. UV transilluminator.

3

Methods Carry out all procedures at room temperature unless otherwise specified. Total RNA was extracted from three pellets of B. subtilis wt and one pellet of ΔbsrA cells for each extraction method (four samples per method).

3.1 Bacterial Cell Cultures

1. Grow B. subtilis wt and ΔbsrA bacteria in 20 ml LB medium at 37 °C in an incubation shaker at 200 rpm overnight. 2. Inoculate 300 ml of fresh medium with an overnight culture to an OD600 of 0.05. 3. Let the cells grow to stationary phase for about 28 h in a water bath shaker. Control growth by regularly measuring the OD600. 4. Induce outgrowth by diluting 40 ml stationary culture (OD600 = 4 to 4.5) with 160 ml fresh prewarmed (37 °C) LB medium and incubate under shaking (200 rpm). 5. After 3 min, harvest 40 ml of outgrowth culture for each extraction in a 50-ml tube. Pellet the cells by centrifugation at 8,200 × g for 10 min at 4 °C (approx. 0.43 g wet cell pellet). 6. Quick-freeze cells in liquid nitrogen and store pellets at −80 °C.

3.2

RNA Isolation

3.2.1 Method 1: Extracting RNA Three Times with Hot Phenol

1. Resuspend harvested B. subtilis cells from Subheading 3.1 in 1.6 ml extraction buffer on ice, and split the cell suspension equally into two 2-ml tubes. The following steps refer to 800 μl samples. 2. Add 75 μl lysozyme solution per reaction tube and incubate for 10 min.

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3. Add 40 μl of 20 % SDS solution and vortex thoroughly (see Note 1). 4. For complete lysis of bacteria, put the tubes into a 65 °C water bath for 90 s. 5. Add 800 μl of acidic phenol preheated to 65 °C and vortex thoroughly for 15 s. 6. Invert the tube several times and vortex again for 15 s until the sample is homogenous. 7. Incubate for 4 min at 65 °C in the water bath. 8. Dip the tubes into liquid nitrogen for at least 30 s. 9. Quickly thaw the sample in the 65 °C water bath. 10. Vortex for 5 s and centrifuge at 15,700 × g for 10 min to separate the aqueous and organic phases. 11. Carefully pipette 3× 192 μl aliquots of the aqueous phase into a single new tube. 12. Repeat steps 5–10 and pipette 3× 160 μl of the aqueous phase into a single new tube. 13. Repeat steps 5–10 and finally pipette 2× 171 μl of the aqueous phase into a single new tube (see Note 2). 14. Add 800 μl of chloroform and vortex thoroughly for 20 s. Centrifuge at 15,700 × g for 5 min. 15. Carefully remove 2× 128 μl of the aqueous phase without any withdrawal of interphase and organic phase. 16. For RNA precipitation, add 25.6 μl of precooled 3 M sodium acetate, pH 5.0 (0.1 vol), and 640 μl ethanol (2.5 vol). 17. Briefly vortex and incubate for 10–20 min at −80 °C or at least for 2 h at −20 °C. 18. Centrifuge at 15,500 × g for 30 min at 4 °C. 19. Remove and discard the supernatant (see Note 3). 20. Wash the pellet with 200 μl of 75 % ethanol by carefully adding the ethanol solution until the entire pellet is submerged. 21. Let stand for 5 min at room temperature. Remove the supernatant and air-dry the pellet for about 5–10 min (see Note 4). 22. Dissolve the RNA pellet in 20 μl ddH2O and store at −20 °C. Store at −80 °C for longer periods (see Notes 5–8; Figs. 1 and 2). 3.2.2 Method 2: Extracting RNA Once with Hot and Once with Cold Phenol

1. Resuspend bacterial cells from Subheading 3.1 in 4 ml extraction buffer at 4 °C. 2. Add 360 μl lysozyme solution to lyse bacterial cells and incubate for 10 min.

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a

b

Fig. 1 Detection of RNAs (115 nt and 190 nt) by Northern blot analysis using 10 % denaturing PAA gels, followed by membrane transfer and immobilization by UV crosslinking. For each extraction method, three independent RNA preparations (a, b, and c) were processed to assess signal fluctuations between individual RNA preparations. The experiment revealed that extraction of intermediate-sized RNAs is more efficient with phenol methods 1 and 2 than with the TRIzol methods 3 and 4. (a) The X-ray film was exposed for 2 min. 6S-1 RNA is detectable with all four extraction methods, although signals were stronger for RNAs prepared according to methods 1 and 2. However, the 5S rRNA loading control was hardly visible after 2 min when using the TRIzol methods 3 and 4. (b) After 10 min of film exposure, 6S-1 RNA and 5S rRNA signals became also visible for the TRIzol RNA preparations. However, for method 4 compared to method 3, the 5S rRNA controls showed reduced intensities and stronger fluctuations between individual samples. The signal above 6S-1 RNA is the 201-nt long 5′-precursor transcript of 6S-1 RNA [2]

3. Add 200 μl of 20 % SDS solution and vortex thoroughly (see Note 1). 4. Add 4 ml of acidic phenol preheated to 65 °C. 5. Vortex thoroughly and incubate for 5 min at 65 °C in a water bath. 6. Incubate for 5 min at 4 °C on ice. 7. Centrifuge at 8,200 × g for 30 min at 4 °C. 8. Transfer as much aqueous phase as possible into a new tube without any withdrawal of interphase and organic phase. 9. Add 4 ml of 4–10 °C cold acidic phenol. 10. Vortex thoroughly for 30 s and centrifuge at 8,200 × g for 30 min at 4 °C.

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Fig. 2 Northern blot detection of tiny RNAs (~14 nt) after separation on 10 % native PAA gels (for details, see Chapter 5 in this issue). In general, ~14-mers were enriched in RNA preparations using TRIzol as extraction reagent. (a) RNA fixation with EDC crosslinking. Both TRIzol methods gave rise to prominent signals specific to ~14-meric pRNA transcripts synthesized in vivo by B. subtilis σA-RNAP using 6S-1 RNA as the template. Specificity was inferred from the loss of signal in RNA prepared from the 6S-1 RNA knockout strain (ΔbsrA). Method 2 resulted in a fainter and more diffuse signal, whereas only a very faint signal was seen with RNA prepared by method 1. M, marker: 0.25 ng of a synthetic 6S-1 RNA-specific, 14 nt long pRNA (p146S-1, 5′-pGUU CGG UCA AAA CU-3′). Note that p146S-1 used marker carried a 5′-monophosphate terminus, and in vivo synthesized pRNAs detected in the other lanes were primary transcripts with 5′-triphosphate ends. (b) RNA fixation with UV crosslinking. The same trends as in panel A were observed, but the signals had a somewhat reduced intensity and less distinct appearance

11. Transfer as much aqueous phase as possible into a new tube. 12. Add 4 ml of chloroform and vortex thoroughly for 30 s. 13. Centrifuge at 8,200 × g for 30 min at 4 °C. 14. Remove the aqueous phase by pipetting it into a new 50-ml tube without any withdrawal of organic phase material. 15. Add 0.1 volume of precooled 3 M sodium acetate (pH 5.0) and 2.5 volume of 100 % ethanol. 16. Vortex and keep for 10–20 min at −80 °C or at least for 2 h at −20 °C. 17. Centrifuge at 8,200 × g for 30 min at 4 °C and discard the supernatant (see Note 3). 18. Wash with 2 ml of 75 % ethanol by carefully adding the ethanol solution until the entire pellet is submerged.

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19. Let stand for 5 min at room temperature. Remove the ethanol solution and air-dry the RNA pellet for about 10–15 min (see Note 4). 20. Dissolve the RNA pellet in 40 μl of ddH2O and store the RNA at −20 °C or −80 °C for longer periods (see Notes 5–9; Figs. 1 and 2). 3.2.3 Method 3: RNA Extraction Using TRIzol®

1. Add 1 ml TRIzol reagent to the bacterial cell pellets from Subheading 3.1 and resuspend the cells by pipetting up and down to accelerate lysis (see Note 10). 2. Incubate for 5 min at 4 °C on ice and transfer 1 ml of lysate to a new 2-ml tube. 3. Add 200 μl of chloroform per 1 ml of TRIzol® used for lysis and vortex the sample thoroughly. Chill on ice for 15 min. 4. Centrifuge the samples at 15,700 × g for 15 min to separate the phases (see Note 11). 5. Carefully pipette about 600 μl of aqueous phase into a new tube (see Note 12). 6. To precipitate the total RNA, add 600 μl of 100 % isopropanol to the aqueous phase, mix, and keep the sample at least for 1 h at −20 °C. 7. Centrifuge at 15,500 × g for 15 min at 4 °C. 8. Remove and discard the supernatant (see Note 13). 9. Add 200 μl of precooled 75 % ethanol, vortex gently, and centrifuge at 15,500 × g for 10 min at 4 °C. 10. Discard the supernatant, air-dry the pellet for 5–10 min, and dissolve the RNA pellet in 20 μl of ddH2O (see Note 4). 11. If not used immediately, store the RNA at −20 °C or −80 °C for extended periods (see Notes 5, 6, 8, 14, and 15) (Figs. 1 and 2).

3.2.4 Method 4: RNA Extraction Using a Combination of TRIzol® and Phase Lock Gel

1. Centrifuge the 15-ml tubes containing the Phase Lock Gel for 5 min at 8,200 × g to pellet the gel. 2. Add 1 ml of TRIzol® to the bacterial cell pellet from Subheading 3.1 and resuspend the cells by pipetting up and down several times to accelerate lysis (see Note 10). 3. Incubate for 5 min at 4 °C on ice and pipette 1 ml of the lysate onto the Phase Lock Gel matrix. 4. Invert the tube several times by hand. 5. Add 200 μl of chloroform per 1 ml of TRIzol® used for lysis and invert thoroughly several times. Incubate for 15 min at 4 °C on ice.

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6. Centrifuge the sample at 8,200 × g for 15 min to separate the mixture into three phases (see Note 16). 7. Pipette the upper aqueous phase into a new 2-ml tube and proceed with isopropanol precipitation as in step 6 of Subheading 3.2.3 (see Notes 6, 8, 14, and 15) (Figs. 1 and 2). 3.3 Control of Total RNA Quality

1. Add one volume of 2× denaturing loading buffer to 6 μg of total RNA. 2. Incubate the mixture at 98 °C for 3 min and put on ice immediately. 3. Load samples onto a 5 % denaturing PAA gel. 4. Run the gel (15 cm wide, 20 cm long, 1 mm thick) in 1× TBE buffer at 20 mA until the xylene cyanol dye has reached the second half of the gel. 5. Stain the gel for 5 min with ethidium bromide. 6. Visualize the stained RNA under UV light using a transilluminator (Fig. 3). 7. See Note 8 for general remarks.

Fig. 3 Ethidium bromide staining of total bacterial RNA separated on a 5 % denaturing PAA gel. Large RNAs, such as 23S and 16S rRNA, but also 5S rRNAs, are more abundant in preparations using the phenol methods, whereas smaller RNAs (roughly tRNAs and shorter RNAs) are enriched with TRIzol-based techniques. In all four samples, the total RNA is considered largely intact as distinct bands and little smearing are observed (see Note 8). However, some degradation of 23S rRNA may have occurred in method 1 and 2 preparations, as 16S rRNA bands were somewhat more intense than those for 23S rRNA in the corresponding lanes

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Notes 1. Add SDS solution at room temperature as SDS precipitates on ice. 2. The amount of aqueous phase decreases after each step, whereas the volume of hot phenol remains constant. 3. After precipitation, a white RNA pellet should be clearly visible at the wall of the tube. 4. Drying for longer periods may impair RNA dissolubility. 5. We advise to shock-freeze RNA extracts in liquid nitrogen before storage at −20 °C or −80 °C. 6. We get about tenfold higher amounts of total RNA with the hot phenol method 1 compared to the TRIzol methods, as inferred from UV spectroscopy. Note that the phenol methods 1 and 2 include a heating step to 65 °C. Therefore, these methods should be avoided when analysis of natively folded RNA is intended. 7. The total RNA yields (as inferred from UV spectroscopy) obtained with method 1 are about 2–3 times higher than those obtained with method 2. Furthermore, we observed that the signal intensity of the loading control (5S rRNA) is more uniform/reproducible with method 1 than with method 2. 8. After purification, the quality of the total cellular RNA needs to be analyzed by electrophoresis under denaturing conditions. We routinely use 5 % PAA gels containing 8 M urea. Alternatively, denaturing 1.2 % agarose gels containing formaldehyde [3] may be used to primarily assess the quality of longer RNAs. Gels can be stained with ethidium bromide or SYBR Gold. Bacterial 16S and 23S rRNAs and eukaryotic 18S and 28S rRNAs are the main gel bands (provided the RNA is not substantially degraded), with the larger rRNA species having around twice the intensity of the smaller rRNA. Distinct rRNA bands with little “smearing” in between or below the two rRNA species are a hallmark of integrity of RNA preparations. 9. If the RNA solution is too viscous, increase the volume with ddH2O. 10. TRIzol contains the chaotropic reagent guanidinium isothiocyanate which inactivates cellular nucleases. This efficiently protects the RNA from degradation during the isolation process. 11. In case of insufficient phase separation, repeat vortexing and centrifuge for another 10 min. 12. Approximately 1/4 of the aqueous phase remains in the TRIzol extraction tube.

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13. The RNA pellet often has a transparent, gel-like appearance after centrifugation. 14. A comparison of TRIzol methods with or without Phase Lock Gel revealed that method 3 extraction is well reproducible as inferred from 5S rRNA and 6S-1 RNA signals in the three parallel RNA samples (Fig. 1). In contrast, more fluctuations and reduced signal intensities are apparent when involving the Phase Lock Gel designed to facilitate phase separation (Fig. 1). 15. This method is applicable to extraction of tiny RNAs (Fig. 2). 16. The Phase Lock Gel forms a barrier between the aqueous and organic phases.

Acknowledgment This work was supported by the Deutsche Forschungsgemeinschaft (GK 1384) to R.K.H. and G.K. and the Russian Foundation for Basic Research (14-04-91336) to O.Y.B. and E.A.K. References 1. Beckmann BM, Grünweller A, Weber MH, Hartmann RK (2010) Northern blot detection of endogenous small RNAs (approximately 14 nt) in bacterial total RNA extracts. Nucleic Acids Res 38(14):e147. doi:10.1093/nar/ gkq437, gkq437 [pii] 2. Beckmann BM, Burenina OY, Hoch PG, Kubareva EA, Sharma CM, Hartmann RK (2011) In vivo

and in vitro analysis of 6S RNA-templated short transcripts in Bacillus subtilis. RNA Biol 8(5):839– 849. doi:10.4161/rna.8.5.16151, 16151 [pii] 3. Kroczek RA, Siebert E (1990) Optimization of northern analysis by vacuum-blotting, RNAtransfer visualization, and ultraviolet fixation. Anal Biochem 184(1):90–95, doi: 0003-2697 (90)90017-4 [pii]

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Part II Visualization and Analysis of Small Non-Coding RNAs

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Chapter 5 Improved Northern Blot Detection of Small RNAs Using EDC Crosslinking and DNA/LNA Probes Katrin Damm, Simone Bach, Katrin M.H. Müller, Gabriele Klug, Olga Y. Burenina, Elena A. Kubareva, Arnold Grünweller, and Roland K. Hartmann Abstract Successful detection of very small RNAs (tiny RNA, ~14 nt in length) by Northern blotting is dependent on improved Northern blot protocols that combine chemical crosslinking of RNA with 1-ethyl-3-(3dimethylaminopropyl)-carbodiimide (EDC) to positively charged membranes, the use of native polyacrylamide gels, and the development of highly sensitive and specific probes modified with locked nucleic acids (LNA). In this protocol, we show that Northern blot detection of tiny RNAs with 5′-digoxigenin-labeled DNA/LNA mixmer probes is a highly sensitive and specific method and, in our hands, more sensitive than using a corresponding DNA/LNA mixmer probe with a 5′-32P-end label. Key words Northern blot, EDC crosslinking, UV crosslinking, Digoxigenin, LNA, Native PAGE

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Introduction Recent advances in high-throughput sequencing methods and bioinformatics have led to the identification of new classes of small regulatory RNAs (sRNA) in pro- and eukaryotes. However, expression patterns and the proof of functionality of such sRNAs need to be confirmed with established standard techniques including RT-qPCR, microarrays, genetic methods, and, of course, Northern blotting. The detection of very small RNAs (miRNAs and shorter ones, termed “tiny RNAs” in the following) poses a challenge, as it requires highly sensitive and specific methods to confirm their identity. This prompted the development of a specialized RT-qPCR technique, the stem-loop primer method, to quantify individual miRNAs [1]. Nevertheless, Northern blotting remains indispensable for a direct visualization of cellular RNAs and their length variants ranging from primary transcripts over processing intermediates to mature forms. A major problem for

Mathieu Rederstorff (ed.), Small Non-Coding RNAs: Methods and Protocols, Methods in Molecular Biology, vol. 1296, DOI 10.1007/978-1-4939-2547-6_5, © Springer Science+Business Media New York 2015

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the detection of tiny RNAs by Northern blotting is their limited interaction surface, thus exacerbating the problem of sterically hindering probe accessibility upon covalent immobilization on hybridization membranes. Improved protocols have already been described to overcome several limitations of conventional Northern blot procedures [2, 3]. However, these advancements have turned out to be insufficient for the detection of tiny RNAs [4, 5]. We have recently described an approach combining native polyacrylamide gels, RNA immobilization on nylon membranes by 1-ethyl-3-(3-dimethylaminopropyl)-carbodiimide (EDC) crosslinking, and nonradioactive 5′-digoxigenin (DIG)-endlabeled oligonucleotide probes containing locked nucleic acid (LNA) modifications for the detection of tiny RNAs with high sensitivity and specificity [4, 5]. In this protocol, we initially describe denaturing PAGE for the separation of sRNAs, such as 5S rRNA and 6S-1 RNA, and native PAGE for the separation of tiny RNAs. Next, we describe the RNA transfer to nylon membranes, followed by RNA immobilization by UV (RNAs >50 nt) or EDC (RNAs