RNA Abundance Analysis : Methods and Protocols [2nd ed.] 9781071607428, 9781071607435

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Methods in Molecular Biology 2170

Hailing Jin Isgouhi Kaloshian Editors

RNA Abundance Analysis Methods and Protocols Second Edition

METHODS

IN

MOLECULAR BIOLOGY

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

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

For over 35 years, biological scientists have come to rely on the research protocols and methodologies in the critically acclaimed Methods in Molecular Biology series. The series was the first to introduce the step-by-step protocols approach that has become the standard in all biomedical protocol publishing. Each protocol is provided in readily-reproducible stepbystep 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 Abundance Analysis Methods and Protocols Second Edition

Edited by

Hailing Jin Department of Plant Pathology and Microbiology, Center for Plant Cell Biology and Institute for Integrative Genome Biology, University of California, Riverside, CA, USA

Isgouhi Kaloshian Department of Nematology, University of California, Riverside, CA, USA

Editors Hailing Jin Department of Plant Pathology and Microbiology, Center for Plant Cell Biology and Institute for Integrative Genome Biology University of California Riverside, CA, USA

Isgouhi Kaloshian Department of Nematology University of California Riverside, CA, USA

ISSN 1064-3745 ISSN 1940-6029 (electronic) Methods in Molecular Biology ISBN 978-1-0716-0742-8 ISBN 978-1-0716-0743-5 (eBook) https://doi.org/10.1007/978-1-0716-0743-5 © Springer Science+Business Media, LLC, part of Springer Nature 2021 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 We are pleased to have this opportunity to edit the Second Edition of RNA Abundance Analysis. RNA abundance is one of the most important measurements for gene expression analysis in the field of molecular biology. Continuous progress in modern technology has empowered us to examine RNA expression more accurately and efficiently, with precision at the cellular and subcellular levels. A new collection of the rapid advances of methodology in RNA abundance analysis is important and timely. This book covers a wide range of techniques on RNA extraction, detection, quantification, visualization, and genome-wide profiling, from conventional methods to state-of-the-art high-throughput approaches. We include detailed techniques to examine mRNAs, small noncoding RNAs, protein-associated small RNAs, organelle RNAs, endosymbiont RNAs, and alternatively spliced RNA variants from various organisms. RNA editing and the computational data processing for genomewide datasets are also discussed. Collectively, these methods should provide helpful guidance to biologists in their gene expression and regulation studies. The beginning of many RNA studies is the isolation of RNAs. We have included methods for extracting RNAs from specific cells and tissues of plants, fungi, insect endosymbiont, and parasites (Chapters 3, 8, 13, and 14). Furthermore, we included detailed protocols on isolating RNAs from specific subcellular structures, such as chloroplasts and extracellular vesicles (Chapters 10 and 16). Isolating RNAs could be challenging if one wishes to address a process that is limited to a few cells within a tissue or organism, or a specific organelle or subcellular fraction of a cell. These chapters have provided excellent tools to achieve these goals. Once high-quality RNAs of specific cells, tissues, or subcellular structures have been extracted, the spatial and temporal expression patterns of an individual gene or the whole genome could be established. Therefore, Chapters 2, 5, 6, 7, 10, 11, 12, 13, and 14 present a set of diverse technologies to examine and analyze the expression of mRNAs and small RNAs. In particular, Chapters 2, 5, 6, 14, and 16 describe the application of highthroughput genome-wide next-generation sequencing approaches to study RNA-related parameters in organisms. While generating vast amounts of sequence data has become routine and increasingly economical, the bottleneck continues to be the computational analysis of the data. This edition therefore includes a chapter on bioinformatics methods to analyze high-throughput RNA and small RNA expression data collected by nextgeneration sequencing. RNAs function mostly through association with various proteins; the study of RNA-protein interaction is a key focus for understanding RNA regulation and gene expression. Chapters 4, 7, and 15 describe the methods to identify RNAs associated with specific protein or protein complexes and to understand the gene expression regulation mediated by RNA-protein interaction. In this edition, we have also included innovative emerging techniques, such as CRISPRCas-mediated RNA editing (Chapter 1) and titanium oxide nanofiber-mediated small RNA extraction (Chapter 8).

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Finally, we hope this new edition provides a comprehensive set of techniques and methods on isolating and analyzing mRNAs, small RNAs, and other RNA variants, which can assist you in your gene expression studies. Riverside, CA, USA

Hailing Jin Isgouhi Kaloshian

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

1 CRISPR-Cas RNA Targeting Using Transient Cas13a Expression in Nicotiana benthamiana . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Veerendra Sharma, Wenguang Zheng, Jun Huang, and David E. Cook 2 Strand-Specific RNA-Seq Applied to Malaria Samples . . . . . . . . . . . . . . . . . . . . . . . Xueqing Maggie Lu and Karine Le Roch 3 Laser Microdissection of Cells and Isolation of High-Quality RNA After Cryosectioning . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Marta Barcala, Carmen Fenoll, and Carolina Escobar 4 Detection of RNA in Ribonucleoprotein Complexes by Blue Native Northern Blotting . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . ¨ ßel, Steffen Ostendorp, Anna Ostendorp, and Julia Kehr Lena Kru 5 Quantitative Analysis of Plant miRNA Primary Transcripts . . . . . . . . . . . . . . . . . . . Jakub Dolata, Andrzej Zielezinski, Agata Stepien, Katarzyna Kruszka, Dawid Bielewicz, Andrzej Pacak, Artur Jarmolowski, Wojciech Karlowski, and Zofia Szweykowska-Kulinska 6 A Revised Adaptation of the Smart-Seq2 Protocol for Single-Nematode RNA-Seq . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Dennis Chang, Lorrayne Serra, Dihong Lu, Ali Mortazavi, and Adler Dillman 7 Analysis of RBP Regulation and Co-regulation of mRNA 30 UTR Regions in a Luciferase Reporter System . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Erin L. Sternburg and Fedor V. Karginov 8 Extraction of Small RNAs by Titanium Dioxide Nanofibers . . . . . . . . . . . . . . . . . . Luis A. Jimenez and Wenwan Zhong 9 Identification of MicroRNAs and Natural Antisense Transcript-Originated Endogenous siRNAs from Small-RNA Deep Sequencing Data . . . . . . . . . . . . . . . Weixiong Zhang, Xuefeng Zhou, Xiang Zhou, and Jing Xia 10 Purification and Analysis of Chloroplast RNAs in Arabidopsis . . . . . . . . . . . . . . . . Huan Wang and Hailing Jin 11 In Situ Detection of Mature miRNAs in Plants Using LNA-Modified DNA Probes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Xiaozhen Yao, Hai Huang, and Lin Xu 12 Northern Blotting Technique for Detection and Expression Analysis of mRNAs and Small RNAs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Ankur R. Bhardwaj, Ritu Pandey, Manu Agarwal, and Surekha Katiyar-Agarwal

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Contents

Isolation of Insect Bacteriocytes as a Platform for Transcriptomic Analyses . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Me´lanie Ribeiro Lopes, Pierre Simonet, Gabrielle Duport, Karen Gaget, Se´verine Balmand, Akiko Sugio, Jean-Christophe Simon, Nicolas Parisot, and Federica Calevro Small RNA Isolation and Library Construction for Expression Profiling of Small RNAs from Neurospora crassa and Fusarium oxysporum and Analysis of Small RNAs in Fusarium oxysporum-Infected Plant Root Tissue . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Shou-Qiang Ouyang, Gyungsoon Park, Hui-Min Ji, and Katherine A. Borkovich Studying RNA–Protein Interaction Using Riboproteomics. . . . . . . . . . . . . . . . . . . Sonali Chaturvedi and A. L. N. Rao Small RNA Extraction and Quantification of Isolated Fungal Cells from Plant Tissue by the Sequential Protoplastation. . . . . . . . . . . . . . . . . . . . . . . . . Qiang Cai and Hailing Jin

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

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Contributors MANU AGARWAL • Department of Botany, University of Delhi North Campus, Delhi, India SE´VERINE BALMAND • Univ Lyon, INSA-Lyon, INRAE, BF2i, UMR203, F-69621, Villeurbanne, France MARTA BARCALA • Facultad de Ciencias Ambientales y Bioquı´mica, Universidad de CastillaLa Mancha, Toledo, Spain; International Research Organization for Advanced Science and Technology (IROAST), Kumamoto University, Kumamoto, Japan ANKUR R. BHARDWAJ • Department of Botany, Ramjas College, University of Delhi North Campus, Delhi, India DAWID BIELEWICZ • Department of Gene Expression, Institute of Molecular Biology and Biotechnology, Faculty of Biology, Adam Mickiewicz University, Poznan´, Poznan´, Poland KATHERINE A. BORKOVICH • Department of Microbiology and Plant Pathology, Institute for Integrative Genome Biology, University of California, Riverside, CA, USA QIANG CAI • Department of Plant Pathology and Microbiology, Center for Plant Cell Biology and Institute for Integrative Genome Biology, University of California, Riverside, CA, USA FEDERICA CALEVRO • Univ Lyon, INSA-Lyon, INRAE, BF2i, UMR203, F-69621, Villeurbanne, France DENNIS CHANG • Department of Nematology, University of California, Riverside, CA, USA SONALI CHATURVEDI • Gladstone Institute of Virology and Immunology, Gladstone Institutes, San Francisco, CA, USA DAVID E. COOK • Department of Plant Pathology, Kansas State University, Manhattan, KS, USA ADLER DILLMAN • Department of Nematology, University of California, Riverside, CA, USA JAKUB DOLATA • Department of Gene Expression, Institute of Molecular Biology and Biotechnology, Faculty of Biology, Adam Mickiewicz University, Poznan´, Poznan´, Poland GABRIELLE DUPORT • Univ Lyon, INSA-Lyon, INRAE, BF2i, UMR203, F-69621, Villeurbanne, France CAROLINA ESCOBAR • Facultad de Ciencias Ambientales y Bioquı´mica, Universidad de Castilla-La Mancha, Toledo, Spain; International Research Organization for Advanced Science and Technology (IROAST), Kumamoto University, Kumamoto, Japan CARMEN FENOLL • Facultad de Ciencias Ambientales y Bioquı´mica, Universidad de Castilla-La Mancha, Toledo, Spain KAREN GAGET • Univ Lyon, INSA-Lyon, INRAE, BF2i, UMR203, F-69621, Villeurbanne, France HAI HUANG • National Laboratory of Plant Molecular Genetics, Shanghai Institute of Plant Physiology and Ecology, Shanghai Institutes for Biological Sciences, Chinese Academy of Sciences, Shanghai, China JUN HUANG • Department of Plant Pathology, Kansas State University, Manhattan, KS, USA ARTUR JARMOLOWSKI • Department of Gene Expression, Institute of Molecular Biology and Biotechnology, Faculty of Biology, Adam Mickiewicz University, Poznan´, Poznan´, Poland

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Contributors

HUI-MIN JI • College of Horticulture and Plant Protection, Yangzhou University, Yangzhou, China LUIS A. JIMENEZ • Program in Biomedical Sciences, University of California, Riverside, CA, USA HAILING JIN • Department of Plant Pathology and Microbiology, Center for Plant Cell Biology and Institute for Integrative Genome Biology, University of California, Riverside, CA, USA FEDOR V. KARGINOV • Department of Molecular, Cell and Systems Biology, Institute for Integrative Genome Biology, University of California, Riverside, CA, USA WOJCIECH KARLOWSKI • Department of Computational Biology, Institute of Molecular Biology and Biotechnology, Faculty of Biology, Adam Mickiewicz University, Poznan´, Poznan´, Poland SUREKHA KATIYAR-AGARWAL • Department of Plant Molecular Biology, University of Delhi South Campus, New Delhi, India JULIA KEHR • Molecular Plant Genetics, Universit€ at Hamburg, Institute of Plant Science and Microbiology, Hamburg, Germany LENA KRU¨ßEL • Molecular Plant Genetics, Universit€ at Hamburg, Institute of Plant Science and Microbiology, Hamburg, Germany KATARZYNA KRUSZKA • Department of Gene Expression, Institute of Molecular Biology and Biotechnology, Faculty of Biology, Adam Mickiewicz University, Poznan´, Poznan´, Poland KARINE LE ROCH • Department of Cell Biology and Neuroscience, Institute for Integrative Genome Biology, Center for Disease Vector Research, University of California, Riverside, CA, USA DIHONG LU • Department of Nematology, University of California, Riverside, CA, USA XUEQING MAGGIE LU • Department of Cell Biology and Neuroscience, Institute for Integrative Genome Biology, Center for Disease Vector Research, University of California, Riverside, CA, USA ALI MORTAZAVI • Department of Developmental and Cell Biology, Center for Complex Biological Systems, University of California, Irvine, CA, USA ANNA OSTENDORP • Molecular Plant Genetics, Universit€ a t Hamburg, Institute of Plant Science and Microbiology, Hamburg, Germany STEFFEN OSTENDORP • Molecular Plant Genetics, Universit€ a t Hamburg, Institute of Plant Science and Microbiology, Hamburg, Germany SHOU-QIANG OUYANG • College of Horticulture and Plant Protection, Yangzhou University, Yangzhou, China; Joint International Research Laboratory of Agriculture and AgriProduct Safety of Ministry of Education of China and Key Laboratory of Plant Functional Genomics of the Ministry of Education, Yangzhou University, Yangzhou, China ANDRZEJ PACAK • Department of Gene Expression, Institute of Molecular Biology and Biotechnology, Faculty of Biology, Adam Mickiewicz University, Poznan´, Poznan´, Poland RITU PANDEY • Department of Botany, SGTB Khalsa College, University of Delhi North Campus, Delhi, India NICOLAS PARISOT • Univ Lyon, INSA-Lyon, INRAE, BF2i, UMR203, F-69621, Villeurbanne, France GYUNGSOON PARK • Department of Electrical and Biological Physics, Plasma Bioscience Research Institute, Kwangwoon University, Seoul, Republic of Korea A. L. N. RAO • Department of Microbiology and Plant Pathology, University of California, Riverside, CA, USA

Contributors

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ME´LANIE RIBEIRO LOPES • Univ Lyon, INSA-Lyon, INRAE, BF2i, UMR203, F-69621, Villeurbanne, France LORRAYNE SERRA • Department of Developmental and Cell Biology, Center for Complex Biological Systems, University of California, Irvine, CA, USA VEERENDRA SHARMA • Department of Plant Pathology, Kansas State University, Manhattan, KS, USA JEAN-CHRISTOPHE SIMON • Agrocampus Ouest, Universite´ Rennes 1, INRAE, IGEPP, UMR 1349, BP 35327, Le Rheu, France PIERRE SIMONET • Univ Lyon, INSA-Lyon, INRAE, BF2i, UMR203, F-69621, Villeurbanne, France AGATA STEPIEN • Department of Gene Expression, Institute of Molecular Biology and Biotechnology, Faculty of Biology, Adam Mickiewicz University, Poznan´, Poznan´, Poland ERIN L. STERNBURG • Department of Molecular, Cell and Systems Biology, Institute for Integrative Genome Biology, University of California, Riverside, CA, USA AKIKO SUGIO • Agrocampus Ouest, Universite´ Rennes 1, INRAE, IGEPP, UMR 1349, BP 35327, Le Rheu, France ZOFIA SZWEYKOWSKA-KULINSKA • Department of Gene Expression, Institute of Molecular Biology and Biotechnology, Faculty of Biology, Adam Mickiewicz University, Poznan´, Poznan´, Poland HUAN WANG • Department of Plant Pathology and Microbiology, Center for Plant Cell Biology and Institute for Integrative Genome Biology, University of California, Riverside, CA, USA JING XIA • Department of Computer Science and Engineering, Washington University, St. Louis, MO, USA LIN XU • National Laboratory of Plant Molecular Genetics, Shanghai Institute of Plant Physiology and Ecology, Shanghai Institutes for Biological Sciences, Chinese Academy of Sciences, Shanghai, China XIAOZHEN YAO • National Laboratory of Plant Molecular Genetics, Shanghai Institute of Plant Physiology and Ecology, Shanghai Institutes for Biological Sciences, Chinese Academy of Sciences, Shanghai, China WEIXIONG ZHANG • Department of Computer Science and Engineering, Fudan University, Shanghai, China; Department of Computer Science and Engineering, Washington University, St. Louis, MO, USA; Department of Genetics, Washington University School of Medicine, St. Louis, MO, USA WENGUANG ZHENG • Department of Plant Pathology, Kansas State University, Manhattan, KS, USA WENWAN ZHONG • Department of Chemistry, University of California, Riverside, CA, USA XIANG ZHOU • Department of Computer Science and Engineering, Washington University, St. Louis, MO, USA XUEFENG ZHOU • Department of Computer Science and Engineering, Washington University, St. Louis, MO, USA ANDRZEJ ZIELEZINSKI • Department of Computational Biology, Institute of Molecular Biology and Biotechnology, Faculty of Biology, Adam Mickiewicz University, Poznan´, Poznan´, Poland

Chapter 1 CRISPR-Cas RNA Targeting Using Transient Cas13a Expression in Nicotiana benthamiana Veerendra Sharma, Wenguang Zheng, Jun Huang, and David E. Cook Abstract Application of the CRISPR-Cas prokaryotic immune system for single-stranded RNA targeting will have significant impacts on RNA analysis and engineering. The class 2 Type VI CRISPR-Cas13 system is an RNA-guided RNA-nuclease system capable of binding and cleaving target single-stranded RNA substrates in a sequence-specific manner. In addition to RNA interference, the Cas13a system has application from manipulating RNA modifications, to editing RNA sequence, to use as a nucleic acid detection tool. This protocol uses the Cas13a ortholog from Leptotrichia buccalis for transient expression in plant cells providing antiviral defense. We cover all the necessary information for cloning the Cas13 protein, crRNA guide cassette, performing transient Agrobacterium-mediated expression of the necessary Cas13a components and target RNA-virus, visualization of virus infection, and molecular quantification of viral accumulation using quantitative PCR. Key words CRISPR-Cas13, RNA targeting, mRNA interference, RNA editing, Transcriptome editing, Antiviral protection, Plant biotechnology, Plant virus

1

Introduction Understanding gene function, through the manipulation of DNA and subsequent experimental determination of phenotypic effects (i.e., functional genomics) remains a grand challenge across biological disciplines. Application of the clustered regularly interspaced short palindromic repeats (CRISPR) and CRISPR associated protein (Cas) prokaryotic immune system for eukaryotic DNA manipulation has ushered in a new approach for functional genomics and genome engineering [1–3]. A strength of using CRISPR-Cas systems for genome engineering is their general organism-agnostic function, plethora of homologs, ease of use, and their amenability to be redesigned to carry out novel functions [4]. The use of class 2 CRISPR-Cas systems, such as Cas9 and Cas12, for genome editing have been applied and previously

Hailing Jin and Isgouhi Kaloshian (eds.), RNA Abundance Analysis: Methods and Protocols, Methods in Molecular Biology, vol. 2170, https://doi.org/10.1007/978-1-0716-0743-5_1, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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detailed across a range of organisms and is not the focus of this methods chapter [5, 6]. The method in this chapter focuses on the use of class 2 type VI CRISPR-Cas systems referred to as Cas13, which target RNA as their substrate, delivering programmable single-stranded RNA interference [7, 8]. This approach opens new opportunities to manipulate and study gene function by targeting transcribed RNA. Also, while Cas13 has an inherent function as a RNA nuclease, research has shown it can also be engineered to carry-out novel functions which can aid in the study of RNA or address societal challenges [9, 10]. At this time, the RNA-targeting Cas13 systems have been subdivided into four families, termed Cas13a, Cas13b, Cas13c, and Cas13d [11–13]. While the Cas13 systems described to-date all require a crRNA and Cas13 effector protein, their sequence, structure, and mechanistic details can vary. Consequently, there is variation across the Cas13 systems for attributes such as target RNA binding affinity, guide processing and target RNA nuclease activity [10, 11]. There are also reports of additional helper proteins that are not required, but modulate target RNA degradation through various mechanisms [14, 15]. Significant mechanistic questions remain regarding Cas13a/b/c/d function in general and function within specific groups of organisms. This method will focus on Cas13a from Leptotrichia buccalis (Lbu) for transient expression in the plant Nicotiana benthamiana. We detail the use of Cas13a for targeted reduction of Turnip Mosaic Virus (TuMV), a single-stranded RNA virus of the largest plant infecting family, Poytviridae [16, 17]. Plant infecting viruses cause significant economic losses annually and threaten global food security. The method described here provides the plant with a new antiviral immune response, which has significant implications for improving crop production through biotechnology. More generally, the approach can be modified to study any cellular singlestranded RNA for a variety of experiments in planta.

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Materials

2.1 Synthetic DNA and Vectors

1. pGWB413 gateway cloning vector (Addgene plasmid ID: 74807). 2. Leptotrichia buccalis Cas13a DNA fragment including promoter and terminator (Integrated DNA Technologies) (Table 1 for sequence, see Notes 1–3). 3. NEBuilder HiFi DNA Assembly Master Mix (New England Biolabs). 4. pENTR/D-TOPO vector and cloning kit (Invitrogen).

Cas13a RNA-Targeting in Plants

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Table 1 Synthetic Cas13a cassette Sequence (50 ! 30 ) Sequence overlapping pGWB413, 50 of CaMV 35S promoter

Gtacaaagtggttgataacagcgggttaat

Lbu (Leptotrichia buccalis) Cas13a coding NCBI protein accession number WP_015770004, codon sequence optimized on IDT website HSP terminator

TATGAAGATGAAGATGAAATATTTGGTGTGTCAAA TA AAAAGCTAGCTTGTGTGCTTAAGTTTGTG TTTTTTTCT TGGCTTGTTGTGTTATGAATTTGTGGCTTTTTC TAATA TTAAATGAATGTAAGATCTCATTATAATGAA TAAACA AATGTTTCTATAATCCATTGTGAATGTTTTGTTGGA TC TCTTCGCATATAACTACTGTATGTGCTATGGTA TGGAC TATGGAATATGATTAAAGATAAG

Sequence overlapping pGWB413, 30 of HSP terminator

Ggcccgatcatattgtcgctcaggatcgtg

Table 2 Empty crRNA cassette and target mRNA guide oligos Name of oligos crRNA adaptor

HCPro guide-F HCPro guide-R

Sequence of oligos (5’ 3’) CACCtctagatGGAGTGATCAAAAGTCCCACATCGATCAGGTG ATATATAGCAGCTTAGTTTATATAATGATAGAGTCGACATAG CGATTgGATTTAGACCACCCCAAAAATGAAGGGGACTAAAA CAaGAGACCcagctGGTCTCgTTTTTTagcccggg AACACTGGGAAATCTTGTTGCGAAAGGACTTC AAAAGAAGTCCTTTCGCAACAAGATTTCCCAG

Note U6 promoter italic, Direct repeat shaded grey, BsaI sites bold

5. crRNA cloning adaptor (Integrated DNA Technologies) (Table 2 for sequence). 6. TuMV HCPro guide RNA oligos (Integrated DNA Technologies) (Table 3 for sequence). 7. 2
Annealing buffer: 20 mM Tris, 2 mM EDTA, 100 mM NaCl, pH 8.0. 8. TE buffer: 10 mM Tris, 1 mM EDTA, pH 8.0. 9. Gateway LR Clonase (Invitrogen). 10. Agarose (VWR). 11. Gel fragment extraction kit (Promega).

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Table 3 Oligos used to screen vectors for positive colonies and to perform qPCR Name of Oligo

Sequence of oligos (50 ! 30 )

Purpose

Cas13a-F

GCGAGGGTCGATTAGTGAAAT

Colony screening

Cas13a-R

CCAGGATGTCCGTTTCTGAATA

Colony screening

U6-F

GGAGTGATCAAAAGTCCCACATCG

Colony screening

BsaI-R

AAACGAGACCAGAACTAAGGGT

Colony screening

35S-R

TACGTCAGTGGAGATATCACATCA

Colony screening

TuMV_P1-F

TAGAGCGCAGCAACCAATTA

TuMV qPCR

TuMV_P1-R

CGAACCTCTTCTGCTTCGATTA

TuMV qPCR

EF1a-F

AGCTTTACCTCCCAAGTCATC

EF1a qPCR

EF1a-R

AGAACGCCTGTCAATCTTGG

EF1a qPCR

12. One Shot™ ccdB Survival™ 2 T1R Competent Cells (Thermo Fisher). 13. T4 DNA ligase (New England Biolabs). 14. PacI, PspOMI, KpnI, BsaI restriction enzymes (New England Biolabs). 15. Hotplate and water bath. 16. DNA oligos used for PCR identification during cloning (Integrated DNA Technologies) (Table 3 for sequence). 17. Luria Bertani (LB) medium: for LB broth, 0.5% yeast extract, 1% tryptone, 1% NaCl in deionized water. For LB agar, add 1.5% agar in LB broth. Autoclave at 121  C for 20 min to sterilize. 2.2 AgrobacteriumMediated Transient Transformation

1. Seeds of Nicotiana benthamiana plants. 2. Agrobacterium tumefaciens strain GV3101. 3. Turnip Mosaic Virus, TuMV infectious clone pCBTuMV-GFP (GenBank EF028235.1, [18]). 4. Infiltration buffer: 10 mM MgCl2, 10 mM MES buffer, pH 5.7 and 100 μM acetosyringone (prepared in DMSO). 5. 1.0 mL needleless syringes. 6. Spectrophotometer. 7. 50 mL conical tubes with screw top. 8. Table top centrifuge with rotor for 50 mL conical tubes. 9. Temperature controlled laboratory shaker.

Cas13a RNA-Targeting in Plants

2.3 Visualizing Virus Infection

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1. Hand held high-intensity UltraViolet lamp (Analytik Jena). 2. Nikon DSLR camera with stand. 3. Black cloth.

2.4 qPCR for Virus Quantification

1. TRIzol (Invitrogen). 2. Liquid Nitrogen. 3. Chloroform (Sigma-Aldrich). 4. Isopropanol (Fisher). 5. Ethanol. 6. Reinforced 2 mL tubes with screw tops. 7. 2.3 mm Zirconia/Silica beads (BioSpec). 8. Bead Ruptor Elite (Omni International). 9. NanoDrop ND1000 (Thermo Fisher). 10. Turbo DNA-free kit (Ambion). 11. 10 mM dNTP mix (New England Biolabs). 12. SuperScript II reverse transcriptase (Thermo Fisher). 13. RNaseOUT inhibitor (Thermo Fisher). 14. Random hexamer (Thermo Fisher). 15. 0.2 and 0.5 mL PCR tubes. 16. Thermocycler (MJ Research). 17. SYBR Select Master Mix for CFX (Applied Biosystems). 18. 96-well microplate. 19. Microplate sealing tape. 20. Centrifuge with rotor for 96-well plate. 21. CFX96 Touch Real-Time PCR Detection System (Bio-Rad).

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Methods

3.1 Cas13a Expression Vector

1. Synthesize the coding sequence for Cas13a from Leptotrichia buccalis (Lbu) following NCBI protein accession number WP_015770004.1 (see Notes 1–3). 2. Linearize pGWB413 vector by double digestion with PacI and PspOMI restriction enzymes (NEB) in a 20 μL reaction. To set up the reaction, add 2 μL of 10
CutSmart buffer (NEB), 5 U of PacI, 5 U of PspOMI, 800 ng of pGWB413 DNA, and use deionized water to make final volume of 20 μL. Incubate the reaction at 37  C for 2 h. 3. Run the digested products on 1% agarose gel, cut the agarose gel containing the vector fragment of about 10 kb and use a gel fragment extraction kit to purify the vector DNA.

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4. Combine the synthesized DNA fragments and linearized vector using NEBuilder HiFi DNA Assembly Master Mix. To prepare the assembly reaction system, add 0.1 pmol of linearized vector, 0.2 pmol of gene fragments, 10 μL of NEB HiFi DNA Assembly Master Mix, and deionized water to bring the final volume to 20 μL. Incubate the reaction at 50  C in a thermocycler for 60 min (Fig. 1a). 5. Take 10 μL of the assembly reaction, add to 50 μL of chemically competent E. coli ccdB survival cells. Incubate the reaction on ice for 30 min, heat shock at 42  C in a water bath for 90 s, and then cool promptly on ice. After 2 min on ice, add 250 μL of LB broth, incubate the cells at 37  C for 60 min with shaking at 200 rpm, and then coat the cells on LB agar plate with 75 μg/ mL spectinomycin and 50 μg/mL chloramphenicol. Incubate the plate overnight in 37  C incubator for the transformed cells to form single colonies. 6. Inoculate single colony to 5 mL of LB broth in a culture tube, shake the culture tube at 37  C for about 16 h. Select the positive colonies by PCR using primer set Cas13a-F and Cas13a-R and 1 μL of cell culture as template. The expected size of the amplicon is 255 bp (see Note 4). 7. Miniprep the plasmid DNA from the putative positive colonies, then perform enzyme digestion of the construct with KpnI and PspOMI at 37  C for 2 h. Run the digestion products on 1% agarose gel. The expected release from the construct is about 2150 bp (see Note 4). After verification, the pGWB413 vector harboring Lbu-Cas13a expression cassette will be used as destination vector for gateway cloning of crRNA. 3.2 Guide crRNA Cloning

1. Synthesize the empty crRNA cassette for Lbu-Cas13a (see Notes 5 and 6). 2. Clone the synthesized empty crRNA cassette into pENTR/DTOPO. Add 4 μL of crRNA adaptor DNA (about 100 ng), 1 μL of pENTR/D-TOPO vector, 1 μL of salt solution (available from the kit), incubate at room temperature for 30 min (Fig. 1b). 3. Clone into chemically competent E. coli following procedure described in Subheading 3.1, step 5. The selection antibiotic is 50 μg/mL kanamycin. 4. Screen E. coli colonies for the presence of the insert using PCR with the primer pair U6-F and BsaI-R. Positive colonies will yield a band of 143 bp. 5. Digest the positive pENTR/D-TOPO vector carrying the empty crRNA cassette using BsaI restriction enzyme at 37  C for 2 h. Run the digestion products on 1% agarose gel and clean

Cas13a RNA-Targeting in Plants

13

e

att R 2

Ca

att R 2

3 s1

R1 att

R1 att

Ca s

a

pGWB413

RB

RB

Gibson Assembly

f

LB

att L 2

ide gu

crR N

att L 2

RB

crRN A

e id

+

att L2

gu

L1 att

pENTR

Directional TOPO cloning

LB

L1 att

te set as

att L1

crRNA c

b

3 s1

A

Ca

LB

7

Gateway Cloning

Golden Gate Cloning

c

d

1 CACCtctagatggagTGATCAAAAGTCCCACATC 30 U6 Promoter

Topo-D

HCPro guide F AACACTGGGAAATCTTGTTGCGAAAGGACTTC

31 G ATC AG G TG ATATATAG C AG C T TAG T T TAT 60

HCPro guide R AAAAGAAGTCCTTTCGCAACAAGATTTCCCAG

A. thaliana U6 polymerase III promoter

61 ATAATGATAGAGTCGACATAGCGAT TgGAT 90 U6 Promoter

DR

91 TTAGACCACCCCAAAAATGAAGGGGACTAA 120 Lbu crRNA direct repeat BsaI

Annealed HCPro-crRNA ready to clone into BsaI digested crRNA cassette AACACTGGGAAATCTTGTTGCGAAAGGACT T C GACCCTT TAGAACAACGCT T T CCTGAAGAAAA

BsaI

121 A AC A a G AG ACC c t t a g t t c t G G TC TC gT T T DR

Boil sample Cool to RT

150

DNA removed guide crRNA target cloning site

151 T T Tagcccggg

161

Fig. 1 Schematic overview of Cas13a and associated crRNA cloning. (a) Plant codon optimized Lbu-Cas13a containing additional 30 bp overlapping DNA sequences was assembled with linearized pGWB413 by Gibson Assembly. (b) Synthesized crRNA cassette was inserted pENTR/D-TOPO vector by directional TOPO cloning. The Golden Gate cloning method was used for cloning the guide sequences into the pENTR/D-TOPO vector containing Empty-crRNA. (c) Sequence details for the empty Lbu-crRNA cassette are shown with annotation. The 50 end contains the sequence CACC (highlighted with a black to white gradient bar) to ensure directional TOPO cloning. The RNA polymerase III U6 promoter (indicated by a solid black bar beneath the sequence) is used to direct transcription of the crRNA. A guanine (g) nucleotide is added between the end of the U6 promoter and start of the Lbu-Cas13a crRNA direct repeat (DR) based on observations of a (g) requirement for PolIII promoters. Two BsaI sites (sequence shaded in green) after the Lbu-Cas13a direct repeat are used for cloning the guide target sequence into the empty-crRNA. The DNA region highlighted in grey is removed during

8

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up the linearized vector DNA from gel using gel fragment extraction kit. 6. Synthesize the oligos HCPro guide-F and HCPro guide-R needed for target mRNA binding (see Notes 7 and 8). 7. Anneal oligos obtained from step 6 in 1
annealing buffer with final concentration of 1 μM by boiling for 5 min and gradually cooling down to room temperature in 400 mL of water on a hotplate (Fig. 1d; see Note 9). 8. Ligate the annealed HCPro guide into the BsaI-digested empty crRNA cassette with T4 DNA ligase (NEB) at room temperature for 30 min, and transform the ligated products into chemically competent E. coli cells as described in Subheading 3.1, step 5. 9. Select positive colonies using PCR with the primers of U6-F and HCPro guide-R. Positive colonies are expected to generate a DNA band of size 149 bp. Isolate plasmid from PCR positive colonies containing crRNA + guide in pENTR/D-TOPO vector. 10. Clone the crRNA cassette contained in the pENTR/D-TOPO vector to the destination vector pGWB413 harboring Lbu-Cas13a by gateway LR reaction (Fig. 1). In a reaction tube, add 1 μL of pENTR plasmid DNA (about 150 ng), 1 μL of pGWB413 vector DNA (about 150 ng), 5 μL of TE buffer, 2 μL of 5
LR reaction buffer, and 1 μL of LR Clonase enzyme mix. Incubate the reaction at 25  C for 1 h in water bath, and then add 1 μL of proteinase K and incubate at 37  C for 10 min in water bath (see Note 10). 11. Transform 2 μL of the gateway reaction product into 50 μL of chemically competent E. coli cells and incubate the coated LB agar plate at 37  C overnight allowing the formation of single colonies. Use 75 μg/mL spectinomycin for selection. 12. Identify positive single colonies by PCR using primers U6-F and 35S-R. Positive colonies containing both Lbu-Cas13 and the crRNA + guide will produce a PCR DNA band of about 500 bp. Miniprep the plasmid DNA from positive colony. The resulting construct carries both Lbu-Cas13a and HCProcrRNA (Fig. 1f). ä Fig. 1 (continued) this cloning step and replaced by the target-specific guide sequence. A series of thymine nucleotides serve as a transcriptional termination sequence for PolIII (highlighted in red) (d) Representation of target guide oligo annealing. The two single-stranded DNA oligos are combined and heated in a water bath and then cooled to room temperature. Following annealing, the 50 end contains an AACA overhang, while the 30 contains an AAAA overhang to produce compatible sticky ends with BsaI digested Empty-guide. (e) Gateway LR reaction takes place between attL1and attL2 sites in HCPro-crRNA cassette and attR1 and attR2 sites in pGWB413 destination vector containing Lbu-Cas13a. (f) Final vector following gateway LR reaction, containing the HCPro-crRNA cassette and Lbu-Cas13a

Cas13a RNA-Targeting in Plants

3.3 AgrobacteriumMediated Transient Expression of Cas13a and TuMV-GFP in N. benthamiana

9

1. Transform the confirmed vector from Subheading 3.2, step 12, containing Lbu-Cas13a and HCPro-crRNA, into competent cells of A. tumefaciens strain GV3101. Transform by adding 2.0 μL of plasmid DNA to thawed A. tumefaciens cells and mix gently by tapping. 2. Incubate cells on ice for another 20 min, then transfer the tubes to liquid nitrogen for 1 min (see Note 11). 3. Transfer the tube(s) to 37  C and allow to thaw. Move thawed cells to ice and incubate for 5 min. 4. Add 0.5 mL of LB broth to each tube and incubate at 28  C, 220 rpm for 2–3 h. 5. Spin tubes at 6000 rpm (3,500
g) for 5 min to pellet cells and discard LB broth, leaving 100 μL in the tube. Suspend the cells in the remaining LB broth and spread the cells with the help of spreader on LB-agar medium plate containing selection. For Lbu-Cas13a vectors: 75 μg/mL spectinomycin, 30 μg/mL rifampicin, 30 μg/mL gentamycin; TuMV-GFP vector: 50 μg/mL kanamycin, 30 μg/mL rifampicin, 30 μg/mL gentamycin. 6. Incubate the plates at 28  C in incubator for 48 h, at which time single colonies should be present and visible (see Note 12). 7. In 50 mL tubes, inoculate a single colony of A. tumefaciens containing Lbu-Cas13a + HCPro-crRNA in one tube and another with Lbu-Cas13a + Empty-crRNA in 10 mL of LB broth with selective antibiotics as indicated in Subheading 3.3, step 5. Incubate overnight in a shaker at 28  C and 220 rpm (Fig. 2, step 1). 8. Measure the optical density of the A. tumefaciens cultures using the 600 nm setting of a spectrophotometer. Let grow until the OD600 is 0.8–1.2. 9. Spin down A. tumefaciens cultures at 4  C and 4000 rpm (~1,800
g) for 15 min, discard the supernatant and put tubes upside down on paper towels to drain the remaining LB broth. 10. Re-suspend the A. tumefaciens cells in infiltration buffer, adjusting the optical density (OD600) of bacterial cells to 1.0 (see Note 13). Incubate at room temperature for 2–3 h. 11. Cover the surface of a large laboratory tray with 2–4 sheets of newspaper and perform Agro-infiltration inside the large tray to contain the A. tumefaciens. 12. Using a 1.0 mL needleless syringe, infiltrate the A. tumefaciens suspension into the abaxial side of N. benthamiana leaves. Inject the A. tumefaciens suspension into the leaf surface

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1) Grow A. tumefaciens for 20 h

28oC 220 rpm 2) Infiltrate abaxial side of leaf with separate A. tumefaciens strains carrying different vectors Wait 72 h

3) Visualize viral infection and collect samples for molecular analysis of viral accumulation

Image GFP

Collect Samples

Fig. 2 General workflow for Agrobacterium-mediated expression of Cas13a and TuMV-GFP in N. benthamiana. (Step 1) requires A. tumefaciens be grown to the appropriate cell density to optimize virulence and transfer of vector DNA. (Step 2) involves the delivery of the A. tumefaciens into the plant leaf using a syringe and physical force. The syringe is held firmly against the abaxial side of the leaf, and using gently pressure, the A. tumefaciens is infiltrated into the intercellular compartments of the leaf. A dark, water soaked ring will be visible present corresponding to the region in which A. tumefaciens was successfully infiltrated. This region should be marked with a marker as it will not be visible during subsequent steps. For this protocol, A. tumefaciens strains carrying the Lbu-Cas13a vectors are infiltrated 48 h prior to infiltration of A. tumefaciens strains carrying TuMV-GFP. The two components are then allowed to accumulate for an additional 72 h before proceeding. In (Step 3), virus accumulation is approximated using a high-intensity UV lamp, which allows visualization of GFP, which is a surrogate for virus accumulation. Following visualization, the infiltrated tissue is collected for further analysis

while applying the counter pressure from the other side (see Notes 14 and 15). After agroinfiltration, mark the boundary of the infiltrated area with a permanent marker (see Note 16) (Fig. 2, step 2).

Cas13a RNA-Targeting in Plants

11

13. 48 h after infiltration of a Lbu-Cas13a vector, follow Subheading 3.3, steps 7–13 to infiltrate A. tumefaciens containing TuMV-GFP. A. tumefaciens carrying TuMV-GFP should be delivered at an OD600 of 0.3 (see Note 13). 3.4 Visualizing Virus Infection

To assess the effect of Lbu-Cas13a and crRNA expression on TuMV-GFP infection, the agroinfiltrated leaves are visualized using a hand-held UV lamp (see Note 17). Keep the plant in dark and illuminate the infiltrated leaf with UV light and capture the GFP fluorescence with a digital camera. 1. Hang a dark cloth along a wall or bench. 2. Position the plant so that the abaxial side of the infiltrated leaf can be photographed. 3. Uniformly illuminate the infiltrated leaf with the UV light. As TuMV is expressing GFP, bright green fluorescence can be seen with the naked eye in the areas expressing TuMV-GFP. The rest of the leaf will be red to purple due to chloroplast autofluorescence (Fig. 3a). The infiltrated area expressing Lbu-Cas13a + HCPro-crRNA should have significantly less GFP fluorescence than the area infiltrated the Lbu-Cas13a + Empty-crRNA vector (Fig. 3a, b).

a

b Leaf One

2

Leaf Two

1

2

1: Lbu-Cas13a + Empty-crRNA 2: Lbu-Cas13a + HCPro-crRNA

TuMV quantification normalized to host EF1a expression

1

1.00

0.75

0.50

0.25

0.00 Lbu-Cas13a + Empty-crRNA

Lbu-Cas13a + HCPro-crRNA

Fig. 3 TuMV accumulation is significantly reduced in the presence of Cas13a with a crRNA targeting the virus. (a) Photographs of N. benthamiana leaves visualized under UV light. Leaf One and Leaf Two are replicates showing the effect of targeting the TuMV-GFP virus with Lbu-Cas13a. The regions of agroinfiltration are outlined by a white dashed line. The two regions labeled with a 1 were infiltrated with the Lbu-Cas13a vector carrying an Empty-crRNA, while the regions marked with a 2 were infiltrated with the Lbu-Cas13a vector carrying the HCPro-crRNA targeting the TuMV genome. Both samples expressing the TuMV-targeting crRNA show less GFP fluorescence, indicating less virus accumulates in the samples. (b) Quantitative assessment of TuMV accumulation from leaf samples expressing Lbu-Cas13a and either the Empty-crRNA or the HCProcrRNA targeting TuMV. The expression values were normalized to N. benthamiana EF1a and TuMV levels in the Empty-crRNA sample was set to 1

12

Veerendra Sharma et al.

3.5 qPCR for Virus Quantification

1. Collect leaf tissue corresponding to the area where A. tumefaciens was infiltrated. The area marked at the time of infiltration serves as a guide. Add the tissue to a 2.0 mL screw top tube containing 4–6 2.3 mm beads. Immediately flashfreeze the samples in liquid nitrogen (see Note 18). Samples can be stored at 80  C until ready to extract RNA. 2. Remove the samples from liquid nitrogen and place in a chilled holder. Quickly unscrew the tops and add 1 mL of TRIzol to the frozen sample. Secure the cap back on the tube (see Note 19). 3. Place samples in the Bead Ruptor Elite, secure the lid and run the machine at speed 5, for 2 cycles of 30 s. Immerse the samples in liquid nitrogen between grinding cycles to ensure they remain frozen. 4. Spin tubes at 12,000
g for 10 min at 4  C to remove beads and tissue debris. Transfer supernatant to new 2.0 mL tubes. 5. Add 0.2 mL chloroform for each 1.0 mL of TRIzol added and mix vigorously for 15 s. Incubate samples for 5 min at RT (see Note 20). 6. Spin tubes at 12,000
g for 15 min at 4  C and carefully transfer the upper layer to a new tube (see Note 21). 7. Add 0.5 mL isopropanol to tubes, mix well and incubate at RT for 10 min. Spin tubes at 12,000
g for 30 min at 4  C. 8. Discard supernatant and wash the pellet with 75% ethanol. Spin at 12,000
g for 10 min and discard the supernatant. Remove remaining ethanol with the help of pipetting and dry the pellet at room temperature for 5–10 min. 9. Add 70 μL of RNase free water to the samples and incubate at 55–60  C for 10 min to dissolve the pellet. 10. Quantify RNA samples using NanoDrop ND1000. Additionally, check the integrity of the RNA samples by separating 1.0 μg of RNA on a 1.2% agarose gel (see Note 22). Store samples at 80  C until further analysis. 11. Use 1.0 μg of total RNA and treat with 1.0 μL Turbo DNase in 1
Turbo DNase buffer in 10 μL final volume and incubate at 37  C for 30 min. 12. Add 2.0 μL of DNase inactivation reagent and incubate at room temperature for 5 min with occasional mixing. 13. Spin tubes at 10,000
g for 5 min. Carefully transfer the supernatant ~10 μL to new tubes without disturbing inactivation beads. 14. Add 1.0 μL of random hexamers (200 ng/μL) to DNase treated RNA, mix well by pipetting and incubate at 65  C for 5 min, immediately place samples on ice.

Cas13a RNA-Targeting in Plants

13

15. Add 1.0 μL of 10 mM dNTPs, 4.0 μL of 5
First-Strand Buffer, 1.0 μL of RNaseOUT (40 U/μL), 2.0 μL of 0.1 M DTT mix well and incubate at room temperature for 2 min. Add 1.0 μL of SuperScript II Reverse transcriptase, mix well by pipetting and spin briefly. 16. Incubate samples at room temperature for 10 min and then incubate at 42  C for 50 min, followed by incubation at 70  C for 15 min to inactivate SuperScript II reverse transcriptase. Chill samples on ice and proceed to qPCR analysis. Alternatively, cDNA can be store at 20  C for weeks or 80  C for months. 17. Dilute an aliquot of cDNA three times with nuclease free water for qPCR analysis. 18. For qPCR analysis using three biological replicates and two qPCR technical replicates for each sample primer pair, calculate the number of reactions using following equation: Total number of reactions ¼

Total number of treatments
3biological replicates
2technical replicates number of reactions for a primer pair
number of primer pairs

19. Prepare the master mix for the total number of calculated reactions. A separate master mix is prepared for each primer pair (i.e., target gene and reference gene). Add and mix all the components except for cDNA as detailed in Table 4 (see Note 23). 20. Aliquot 38 μL of master mix in separate tubes for each cDNA sample and add 2.0 μL of cDNA template to the corresponding tubes, mix several times by pipetting, briefly spin and pipette 20 μL of each reaction mixture into two wells of microplate, which corresponds to two technical replicates. Complete the plate setup for all the samples (see Note 24). 21. Cover the microplate with microplate sealing tape and spin the plate briefly to ensure samples are at the bottom of the wells. Table 4 Components for qPCR Master Mix Components

Volume (μL)

qPCR master mix (2
)

10.0

cDNA

1.0

Forward primer (10 μM)

1.0

Reverse primer (10 μM)

1.0

Nuclease-free water

7.0

Final volume

20.0

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Veerendra Sharma et al.

Table 5 Three step amplification program for thermocycler during qPCR Number

Steps

Temperature 

Duration

1

Initial incubation

50 C

2 min

2

Enzyme activation

95  C

2 min

3 4 5

Denaturation Annealing Extension

95  C 52  C 72  C

15 s 15 s 45 s

(39 cyclesa)

a

At the end of each cycle, set the thermocycler camera to capture the fluorescent signal for each well

22. Insert the covered plate containing the reaction mixtures into the thermocycler and perform qPCR using the cycling conditions specified in Table 5 (see Note 24).

4

Notes 1. The coding sequence for Lbu-Cas13a was optimized on IDT website using Codon Optimization Tool (www.idtdna.com/ CodonOpt) to be expressed in plant system. 2. CaMV 35S promoter was used to drive the Lbu-Cas13a gene expression with HSP18 terminator downstream of the coding sequence. The choice of promoter for Cas13a should be considered depending on the users system and desired expression pattern. 3. The synthesized DNA fragment including CaMV 35S promoter, optimized coding sequence for Lbu-Cas13a and HSP18 terminator should additionally contain 30 base pairs of DNA sequence overlapping with the linearized pGWB413 vector. 4. The use of primers Cas13a-F and Cas13a-R and digestion by KpnI and PspOMI is based on the Lbu-Cas13a sequence used here. Different coding sequences require appropriate primer design and restriction enzymes. 5. crRNA adaptor includes U6 promoter, direct repeat specific to Lbu-Cas13a protein, BsaI cloning sites for insertion of guide/ spacer complementary to the gene targeting site, and TTTTT downstream of the cloning sites as transcription termination signal (Fig. 1c). For different Cas13a proteins, the direct repeat needs to be changed accordingly. 6. BsaI is one of many type IIS restriction enzymes, commonly used in Golden Gate cloning. Other type IIS restriction

Cas13a RNA-Targeting in Plants

15

enzymes, such as BaeI could be used but needs to be designed with the entire cloning process in mind to achieve compatibility between vectors. 7. For guide RNA, the length of spacer sequence and the targeting site of choice need to be optimized regarding specific Cas13a and the target gene. The PFS (protospacer flanking sequence) of the targeting site might be a consideration during the guide RNA designing based on the functional property of Cas13a [7, 14]. In this method, we used 28 bp region of HCPro with flanking T downstream of the targeting site. To synthesize the guide RNA, the top strand of the guide RNA needs to be reverse complementary to the targeting site. 8. We have observed significant target mRNA reduction compared to empty guide control when expressing some guide crRNA in the absence of Cas13. That is, some guide crRNA have been observed to produce Cas13-independent target mRNA interference. Care should be taken to include a negative control agroinfiltration expressing the guide crRNA without Cas13. 9. Larger volume of water in a beaker for boiling is acceptable. After boiling for 5 min, turn off the hotplate and leave the beaker on the hotplate. Let the water cool gradually to room temperature for 3 h or overnight. Alternatively, oligo annealing can also be performed on a thermocycler with 95  C for 5 min and then ramp down to 25  C with rate of 0.1  C/s. 10. This method uses gateway LR Clonase enzyme mix to perform the LR recombination reaction. If gateway LR Clonase II is used, skip the 5
reaction buffer as it is included in the LR Clonase II enzyme mix. 11. Do not touch or handle liquid nitrogen with your bare hand, it can burn your skin. Use forceps or protect your hands with thick gloves. 12. If you already have a glycerol stock of the desired A. tumefaciens, prepare a fresh streak. It is important to use A. tumefaciens from plates that are not older than 7 days. Older plates should be discarded and fresh A. tumefaciens should be re-streaked. This is necessary to maintain efficient plant infection and delivery of the transgene. 13. A. tumefaciens cultures should be grown to the log phase, which can be approximated to occur around OD600 of 0.8–1.2. Overgrown cells lose competency to deliver the gene of interest. In case of A. tumefaciens strain GV3101, 20 h of growth is optimal. To analyze the effect of Lbu-Cas13a mediated silencing of TuMV-GFP, the OD600 ratio of effector:target is 1.0:0.3. We have observed that infiltrating Lbu-Cas13a constructs before infiltrating TuMV-GFP

16

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significantly increases Cas13a-mediated reduction of TuMVGFP. Additionally, the ratio of effector (Lbu-Cas13a and crRNA) and target (TuMV-GFP) impacts the amount of target mRNA reduction observed. These details should be considered depending on the experiment. 14. During agroinfiltration, care should be taken to avoid damaging the leaves. It is common to have dead plant cells at the site where the syringe contacts the leaves, but too much damage can compromise the infiltration. 15. To avoid cross contamination between A. tumefaciens strains carrying different vectors during infiltration, it is important to disinfect one’s gloves with 70% ethanol or change gloves before handling different constructs. 16. It is important to mark the boundary of agro-infiltrated area during the infiltration of Lbu-Cas13a constructs. For the second infiltration with TuMV-GFP, try to overlap the infiltrated area as much as possible. Virus infiltration that extends beyond the region of Lbu-Cas13a infiltration will not have virus reduction. Care should be taken to mark these areas to aid in later tissue collection. 17. High-intensity UV light is hazardous to unprotected eyes. Always wear protective glasses while working with a UV light source. Capturing images of GFP fluorescence in a low light setting requires longer exposure times with the camera settings. Take care to equally shine the UV lamp across the leaf surface. 18. To obtain high quality intact RNA from leaf tissues, it is important to avoid excessive handling and damage to the tissue during sample collection. Collected leaf tissue should be immediately frozen in liquid nitrogen. During the grinding process, the tissue should be kept frozen to avoid RNA degradation. 19. This is considered “wet” grinding, in which the samples are ground in the presence of TRIzol. It is also possible to perform “dry” pulverization depending on the researchers needs and equipment. The critical point is to not let the samples thaw during grinding. Additionally, the bead beater can be placed in a cold room to minimize sample-heating and samples can be placed back into liquid nitrogen between rounds of bead beating. 20. TRIzol and chloroform are hazardous chemicals. Always work in a fume hood while handling TRIzol and chloroform and follow proper hazardous chemical disposal. 21. While transferring the upper phase containing RNA to new tubes it is important to avoid the interphase which contains unwanted molecules and chloroform.

Cas13a RNA-Targeting in Plants

17

22. A 260/280 ratio of 2.0 is an indicator of good quality RNA. The integrity of RNA samples should be checked on agarose gel before proceeding for cDNA synthesis. On a 1.5% agarose gel, the two bands corresponding to the largest size should be sharp and bright. These correspond to intact 28S and 18S ribosomal RNA. Additional bands can also be seen, but they should not be smeared. Smeared bands or weak 28S and 18S bands indicates the samples have undergone RNA degradation. 23. To account for pipetting error during qPCR, it is advisable to prepare master mix with two additional reactions added. Care should be taken to minimize pipetting variation and mixing variation. It is advisable to always mix the components the same number of times (i.e., 10) and to pipette using the same stopping point (i.e., the first stopping point, do not push pipette plunger all the way down). qPCR is sensitive to changes in handling and pipetting and minimizing variation between samples will help ensure robust results. 24. Always make a replica qPCR microplate layout on paper to keep track of your samples while setting up qPCR plate. 25. The qPCR profile for your target gene and housekeeping gene may vary depending upon the primer specifications and should be standardized before performing qPCR.

Acknowledgments Funding related to the development of this protocol is provided to DEC by the Defense Advanced Research Projects Agency (DARPA) through a Young Faculty Award (DP17AP00034). The content does not necessarily reflect the position or the policy of the Government and does not imply an official endorsement. References ˇ erma´k T, Baltes NJ, C ˇ egan R et al (2015) 1. C High-frequency, precise modification of the tomato genome. Genome Biol 16:232. https://doi.org/10.1186/s13059-015-07969 2. Xie K, Minkenberg B, Yang Y (2015) Boosting CRISPR/Cas9 multiplex editing capability with the endogenous tRNA-processing system. Proc Natl Acad Sci U S A 112:3570–3575. https://doi.org/10.1073/pnas.1420294112 3. Tang X, Lowder LG, Zhang T et al (2017) A CRISPR-Cpf1 system for efficient genome editing and transcriptional repression in plants. Nat Plants 3:17103. https://doi.org/10. 1038/nplants.2017.103

4. Qi LS, Larson MH, Gilbert LA et al (2013) Repurposing CRISPR as an RNA-guided platform for sequence-specific control of gene expression. Cell 152:1173–1183. https://doi. org/10.1016/j.cell.2013.02.022 5. Baltes NJ, Voytas DF (2015) Enabling plant synthetic biology through genome engineering. Trends Biotechnol 33:120–131. https:// doi.org/10.1016/j.tibtech.2014.11.008 6. Sadhu MJ, Bloom JS, Day L et al (2018) Highly parallel genome variant engineering with CRISPR-Cas9. Nat Genet 50:510–514. https://doi.org/10.1038/s41588-018-0087y

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7. Abudayyeh OO, Gootenberg JS, Konermann S et al (2016) C2c2 is a single-component programmable RNA-guided RNA-targeting CRISPR effector. Science 353:aaf5573. https://doi.org/10.1126/science.aaf5573 8. East-Seletsky A, O’Connell MR, Knight SC et al (2016) Two distinct RNase activities of CRISPR-C2c2 enable guide-RNA processing and RNA detection. Nature 538:270–273. https://doi.org/10.1038/nature19802 9. Rauch S, He C, Dickinson BC (2018) Targeted m6A reader proteins to study Epitranscriptomic regulation of single RNAs. J Am Chem Soc 140:11974–11981. https://doi.org/10. 1021/jacs.8b05012 10. Cox DBT, Gootenberg JS, Abudayyeh OO et al (2017) RNA editing with CRISPRCas13. Science 358:1019–1027. https://doi. org/10.1126/science.aaq0180 11. East-Seletsky A, O’Connell MR, Burstein D et al (2017) RNA targeting by functionally orthogonal type VI-A CRISPR-Cas enzymes. Mol Cell 66:373–383.e3. https://doi.org/10. 1016/j.molcel.2017.04.008 12. Konermann S, Lotfy P, Brideau NJ et al (2018) Transcriptome engineering with RNA-targeting type VI-D CRISPR effectors. Cell 173:665–676.e14. https://doi.org/10. 1016/j.cell.2018.02.033 13. O’Connell MR (2018) Molecular mechanisms of RNA targeting by Cas13-containing type VI

CRISPR-Cas systems. J Mol Biol 431 (1):66–68. https://doi.org/10.1016/j.jmb. 2018.06.029 14. Smargon AA, Cox DBT, Pyzocha NK et al (2017) Cas13b is a type VI-B CRISPR-associated RNA-guided RNase differentially regulated by accessory proteins Csx27 and Csx28. Mol Cell 65:618–630.e7 15. Yan WX, Chong S, Zhang H et al (2018) Cas13d is a compact RNA-targeting type VI CRISPR effector positively modulated by a WYL-domain-containing accessory protein. Mol Cell 70:327–339.e5. https://doi.org/ 10.1016/j.molcel.2018.02.028 16. Garcia-Ruiz H, Carbonell A, Hoyer JS et al (2015) Roles and programming of Arabidopsis ARGONAUTE proteins during turnip mosaic virus infection. PLoS Pathog 11:1–27. https:// doi.org/10.1371/journal.ppat.1004755 17. Aman R, Ali Z, Butt H et al (2018) RNA virus interference via CRISPR/Cas13a system in plants. Genome Biol 19:1. https://doi.org/ 10.1186/s13059-017-1381-1 18. Lellis AD, Kasschau KD, Whitham SA, Carrington JC (2002) Loss-of-susceptibility mutants of Arabidopsis thaliana reveal an essential role for elF(iso)4E during potyvirus infection. Curr Biol 12:1046–1051. https:// doi.org/10.1016/S0960-9822(02)00898-9

Chapter 2 Strand-Specific RNA-Seq Applied to Malaria Samples Xueqing Maggie Lu and Karine Le Roch Abstract Over the past few years only, next-generation sequencing technologies became accessible and many applications were rapidly derived, such as the development of RNA-seq, a technique that uses deep sequencing to profile whole transcriptomes. RNA-seq has the power to discover new transcripts and splicing variants, single nucleotide variations, fusion genes, and mRNA levels-based expression profiles. Preparing RNA-seq libraries can be delicate and usually obligates buying expensive kits that require large amounts of stating materials. The method presented here is flexible and cost-effective. Using this method, we prepared high-quality strand-specific RNA-seq libraries from RNA extracted from the human malaria parasite Plasmodium falciparum. The libraries are compatible with Illumina®’s sequencers Genome Analyzer and Hi-Seq. The method can however be easily adapted to other platforms. Key words Strand-specific RNA-seq, High-throughput sequencing, Malaria, Plasmodium falciparum, Splicing variant discovery, Transcript discovery

1

Introduction The advent of high throughput sequencing technologies marked the beginning of a new era for whole genome analysis. The cost for sequencing a genome dropped considerably over the past 5 years, a revolution for labs focusing on genome mining. Applications were rapidly derived, applying deep sequencing to various “omics” such as the development of RNA-seq to analyze whole transcriptomes. Where microarray-based techniques proved to be powerful tools in exploring gene expression profiles, RNA-seq has the power to establishing expression profiles in a more quantitative manner and to discover new transcripts and splicing variants, single nucleotide variations, and fusion genes at the single-base resolution. The dark side of the application is the considerable amount of complex computations that accompany RNA-seq. In addition, where preparing gDNA libraries is robust and affordable, with a wide range of reagent options on the market, preparing RNA-seq libraries is more expensive and can be more challenging.

Hailing Jin and Isgouhi Kaloshian (eds.), RNA Abundance Analysis: Methods and Protocols, Methods in Molecular Biology, vol. 2170, https://doi.org/10.1007/978-1-0716-0743-5_2, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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Here we present a method that has the double advantage to use reagents originally designed for genomic DNA library preparation and to ultimately provide strand-specific information that simplifies downstream analysis. The described method is used to prepare, in a flexible and cost-effective manner, high quality libraries from small amounts of RNA extracted from the human malaria parasite Plasmodium falciparum to be sequenced on Illumina®’s sequencers Genome Analyzer and Hi-Seq. It can however be adapted to a wide array of other organisms and platforms.

2

Materials All materials and reagents must be molecular biology grade and nuclease-free. All solutions must be freshly prepared before each experiment. Lab benches and pipettes must be clean. The regular use of cleaning solutions such as RNaseZap® (Ambion) is recommended. Nuclease-free barrier tips should be used at all times. Always wear gloves and change them often. After tissue homogenization, samples should always be kept on ice. Using non-stick (low retention) RNase-free tubes and tips can be beneficiary when working with low amounts of RNA. 1. Parasite cultures grown in complete RPMI medium at 5% hematocrit (see Note 1). 2. TRIzol® LS Reagent (Invitrogen™) pre-warmed at 37
C. 3. Chloroform. 4. Isopropanol pre-chilled on ice. 5. Nuclease-free non-DEPC water. 6. DNAse I RNAse-free (Ambion®). 7. Deionized formamide. 8. Formaldehyde 37%. 9. 10 MOPS EDTA buffer pH 7.0. 10. Glycerol 50%. 11. Bromophenol blue powder. 12. Ethidium bromide 20 mg/mL. 13. GenElute™ mRNA Miniprep Kit (Sigma-Aldrich). 14. 5 RNA storage solution (Ambion). 15. HPLC-purified random hexamers and Anchored OligodT(20). 16. SuperScript® VILO™ cDNA synthesis kit (Invitrogen™). 17. DNA Clean & Concentrator™ (Zymo Research). 18. 5 First Strand Buffer (Invitrogen™): 250 mM Tris–HCl (pH 8.3), 375 mM KCl, 15 mM MgCl2.

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19. 5 Second Strand Buffer (Invitrogen™): 100 mM Tris–HCl (pH 6.9), 450 mM KCl, 23 mM MgCl2, 0.75 mM β-NAD+, 50 mM (NH4)2SO4. 20. Set of dATP, dGTP, dCTP, and dUTP. 21. E. coli DNA Polymerase I 10 U/μL (Invitrogen™). 22. E. coli DNA Ligase 10 U/μL (Invitrogen™). 23. E. coli DNA RNase H 2 U/μL (Invitrogen™). 24. 0.1 M DTT. 25. dsDNA Shearase™ (Zymo Research). 26. Encore™ NGS Library System I (NuGEN®). 27. Same-day 70% ethanol in nuclease-free water. 28. USER™ Enzyme (New England Biolabs®). 29. 1 TE buffer pH 8.0.

3

Methods

3.1 Total RNA Extraction from Parasite Cultures

1. Spin down the cultures at 700  g for 5 min with brake level set at the minimum. Aspirate off the supernatant. 2. Add 5 volumes of pre-warmed TRIzol® LS (37
C) and mix thoroughly to dissolve all clumps (see Note 2). 3. Incubate at 37
C for 5 min to ensure the complete de-proteinization of nucleic acids. 4. Stopping point: the samples can be stored at 80
C until further processing. They must be thawed on ice before resuming the protocol. 5. Keep the samples on ice. For each 5 mL of TRIzol® LS that was used in step 2, add 1 mL of chloroform and vortex for 1 min. 6. Centrifuge at 12,000  g for 30 min at 4
C. 7. Carefully transfer the upper aqueous layer to a fresh tube (see Note 3) and add 0.8 volume of prechilled isopropanol to precipitate the RNA. Mix carefully by inverting. 8. Stopping point: the tubes can be stored at 20
C overnight until further processing. Their temperature must be equilibrated on ice for a few minutes before resuming the protocol (see Note 4). 9. Mix by inverting and centrifuge at 12,000  g for 30 min and 4
C. Carefully aspirate off the supernatant. 10. Allow the pellet to air-dry on ice for 5 min and add 30–100 μL of RNase-free non-DEPC treated water. 11. Heat tubes at 60
C for 10 min and then place on ice.

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DNase Treatment

1. To 100 μL of RNA solution add 11.3 μL of 10 DNase I Buffer and 2 μL (4 U) of DNase I (2 U/μL). 2. Incubate the tube at 37
C for 30 min. 3. Inactivate the DNAse at room temperature for 5 min in the presence of 1 mM EDTA. Transfer the tube on ice. 4. Stopping point: RNA solutions can be aliquoted and stored at 80
C. When needed, thaw tubes on ice. Avoid repeated freeze/thaw cycles.

3.3 Verification of the Quality and the Quantity of Total RNA

1. Quantify the concentration of the total RNA solution by UV spectrophotometry, such as a NanoDrop (Thermo Scientific). Typically, a clean solution of nucleic acid in nuclease-free water has an OD ~ 1.85. A ratio ranging from 1.8 to 2.2 is therefore recommended. 2. Check RNA integrity by agarose gel electrophoresis (see Notes 5 and 6). 3. If genomic DNA is visible on the gel repeat the DNase treatment (see Note 7). 4. Verify the absence of trace amounts of genomic DNA by 40 cycles of PCR on a chosen control gene using 50–500 ng of total RNA solution. Repeat the DNase treatment if necessary. 5. Stopping point: Store the total RNA solution at 80
C.

3.4 Purification of polyA+ mRNA from Total RNA

This protocol uses the reagents from the GenElute™ mRNA Miniprep Kit (Sigma-Aldrich). Before starting, equilibrate a heating block for microcentrifuge tubes at 70
C. Keep the elution solution at 70
C. If the beads were kept at 4
C let them sit on the bench top for at least 15 min. Cold beads reduce yields. 1. Thaw the total RNA sample on ice. The amount of starting material should be 150–500 μg of purified total RNA. The remaining steps are performed at room temperature unless specified otherwise. 2. Adjust volume of total RNA to 250 μL with RNase-free water. Add 250 μL of 2 binding solution and vortex briefly. 3. Add 15 μL of oligo(dT) polystyrene beads and vortex thoroughly. 4. Heat the mixture at 70
C for 3 min to denature the RNA and let it cool down for 10 min at room temperature. 5. Centrifuge 2 min at maximum speed (14,000–16,000  g) in a tabletop microcentrifuge. Carefully pipette off the supernatant without disturbing the bead pellet (see Note 8).

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6. Add 500 μL of wash solution mix by vortexing. Transfer the mixture to a GenElute spin filter/collection tube assembly. Failure to transfer all traces of mixtures will result in lower mRNA yields. 7. Centrifuge 1 min at maximum speed (14,000–16,000  g) in a tabletop microcentrifuge. Discard the flow-through and place the collection tube back on the GenElute spin filter. 8. Add 500 μL of Wash Solution onto the GenElute spin filter and centrifuge 2 min at maximum speed. Transfer the GenElute spin filter to a fresh nuclease-free microcentrifuge tube. 9. Add 50 μL of elution solution heated at 70
C onto the center of the GenElute spin filter and incubate 5 min at 70
C. Centrifuge 1 min at maximum speed. 10. Repeat step 10 for a second elution. 11. Check the mRNA quantity by UV spectrometry. Expect 1.5 to 2.5% of the starting amount total RNA, depending on the considered morphological stage of the parasite. 12. Stopping point: The mRNA solutions can be stored at 80
C. When needed, thaw tubes on ice. Avoid repeated freeze/thaw cycles. 3.5 Fragmentation of the polyA+ mRNAs

1. Reduce sample volume to 15–20 μL in a vacuum concentrator type SpeedVac® without heating. Do not let the sample dry. 2. Add 4 volumes of 5X RNA storage solution and incubate for 40 min at 98
C (see Note 9). 3. Reduce sample volume to 10 μL in a vacuum concentrator without heating. Do not let the sample dry. 4. Stopping point: The mRNA solutions can be stored at 80
C. When needed, thaw tubes on ice. Avoid repeated freeze/thaw cycles.

3.6 First Strand cDNA Synthesis

First strand cDNA is synthesized using the SuperScript® VILO™ cDNA synthesis kit (Invitrogen™). All reagents and buffers mentioned in this section refer to elements of the kit. Frozen items should be kept on ice after thawing. 1. In a thin-wall nuclease-free 0.2 mL PCR-grade tube, mix 3 μg of random hexamers and 1 μg of Anchored oligodT(20) to the fragmented mRNA in 14 μL final volume (see Note 10). 2. Incubate the tube in a pre-heated thermal cycler at 70
C for 10 min and quickly chill on ice for 5 min. Do not reduce this time. 3. On ice, add the following reagents to the tube from step 2: 4 μL of 5 VILO™ Reaction Mix, 2 μL of 10 SuperScript® Enzyme Mix (see Note 11). If you prepare multiple samples at

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the same time, make a master mix containing the 5 VILO™ Reaction Mix and the 10 SuperScript® Enzyme Mix and add 6 μL of it to each sample (see Note 12). 4. Gently mix the sample by flicking the bottom of the tube with fingertips. Spin, place on ice. 5. Incubate the sample in a thermal cycler using the following program: 25
C for 10 min, 42
C for 90 min, 85
C for 5 min, and hold at 4
C. 6. Remove promptly from the thermal cycler and place the tube on ice. 7. Purify first strand cDNA using the DNA Clean & Concentrator™ (Zymo Research): (a) Add 100 μL of DNA Binding Buffer to the reaction mixture and mix well by pipetting up and down. (b) Transfer to a Zymo-Spin™ Column/collection tube assembly and centrifuge 30 s at maximum speed (14,000–16,000  g) in a tabletop microcentrifuge. Discard the flow-through. (c) Add 200 μL of wash buffer (freshly prepared with absolute ethanol, see Note 13). Centrifuge 30 s at maximum speed. (d) Discard the flow-through and repeat c. for a second wash. (e) Transfer the Zymo-Spin™ Column to a fresh nucleasefree microcentrifuge tube. (f) Pipet 20 μL of nuclease-free water to the column matrix and let stand 1 min. Centrifuge 30 s at maximum speed to elute the nucleic acid. (g) Repeat step f. 8. Adjust sample volume to 47 μL with non-DEPC nuclease-free water (see Note 14). 3.7 Second Strand cDNA Synthesis

All reagents and buffers mentioned in this section should be made freshly. Frozen items should be kept on ice after thawing. 1. Prepare a dNTP mix containing dATP, dCTP, dGTP, and dUTP (instead of dTTP) each at 10 mM final concentration (see Note 15). 2. Chill all reagents on ice. 3. Set up the following reaction on ice and in the provided order:

Malaria Strand-Specific RNA-Seq

25

First strand cDNA

47 μL

5 First strand buffer

2 μL

100 mM DTT

1 μL

5 Second strand buffer

15 μL

10 mM dNTP (w/dUTP) mix

4 μL

E. coli DNA polymerase I 10 U/μL

4 μL

E. coli DNA ligase 10 U/μL

1 μL

E. coli RNase H 2 U/μL

1 μL

4. Mix gently by pipetting and incubate at 16
C for 2 h. 5. Chill the reaction on ice for at least 5 min. 6. Purify ds cDNA using the DNA Clean & Concentrator™ (Zymo Research): (a) Add 375 μL of DNA binding buffer to the reaction mixture and mix well by pipetting up and down. (b) Transfer to a Zymo-Spin™ Column/collection tube assembly and centrifuge 30 s at maximum speed (14,000–16,000  g) in a tabletop microcentrifuge. Discard the flow-through. (c) Add 200 μL of wash buffer (freshly prepared with absolute ethanol, see Note 13). Centrifuge 30 s at maximum speed. (d) Discard the flow-through and repeat c. for a second wash. (e) Transfer the Zymo-Spin™ column to a fresh nuclease-free microcentrifuge tube. (f) Pipet 6 μL of nuclease-free water to the column matrix and let stand for 1 min. Centrifuge 30 s at maximum speed to elute the nucleic acid. (g) Repeat step f. 7. Check the ds cDNA quantity by UV spectrometry and quality by visualization on a 1.2% agarose gel electrophoresis. A smear should be easily detected (see Note 16). 8. Stopping point: The sample is now ds cDNA and is relatively stable. It can be stored at 20
C. When needed, thaw tubes on ice. Avoid repeated freeze/thaw cycles. 3.8 ds cDNA Fragmentation

1. Mix 700 ng of ds cDNA with 11.5 μL of 3 dsDNA Shearase™ reaction buffer and 3.5 μL of dsDNA Shearase™ (Zymo Research). Reach a final volume of 35 μL final with nucleasefree water. 2. Incubate at 37
C for 40 min (see Note 17).

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Xueqing Maggie Lu and Karine Le Roch

3. Purify ds cDNA and inactivate the dsDNA Shearase™ by adding 175 μL of the DNA Clean & Concentrator™ DNA binding buffer (Zymo Research). 4. Mix well by pipetting up and down and transfer to a ZymoSpin™ column/collection tube assembly and centrifuge 30 s at maximum speed (14,000–16,000  g) in a tabletop microcentrifuge. Discard the flow-through. 5. Add 200 μL of wash buffer (freshly prepared with absolute ethanol, see Note 13). Centrifuge 30 s at maximum speed. 6. Discard the flow-through and repeat step 5 for a second wash. 7. Transfer the Zymo-Spin™ column to a fresh nuclease-free microcentrifuge tube. 8. Pipet 10 μL of nuclease-free water to the column matrix and let stand 1 min. Centrifuge 30 s at maximum speed to elute the nucleic acid. 9. Repeat step 8. 10. Check the size range and the concentration of the sample using microfluidic-based separation devices suitable for small amounts of starting materials, such as an Agilent 2100 Bioanalyzer (Agilent Technologies) or a LabChip® GX (Caliper Life Sciences) (see Note 18). Repeat the fragmentation procedure if necessary. 11. Stopping point: The sample can be stored at 20
C. When needed, thaw tubes on ice. Avoid repeated freeze/thaw cycles. 3.9 Library Preparation

3.9.1 End Repair

The protocol described here uses the NuGEN® Encore™ NGS Library System I, compatible with the Illumina® Genome Analyzer and Hi-Seq sequencing platforms, and all mentioned reagents refer to components of this kit (see Note 19). However, since the starting material is double stranded DNA, it can be easily adapted to any gDNA library preparation kit (including multiplexing) or set of reagents. The Agencourt® magnetic beads used for sample cleanup must be incubated at room temperature for at least 15 min before use. Cold beads reduce yields. Before each use, beads must be fully resuspended by inverting and tapping the tube. Thaw all necessary reagents, mix by vortexing, spin, and keep them on ice until use. Keep the nuclease-free water at room temperature. 1. Dilute 200 ng of fragmented ds cDNA to a volume of 7 μL with nuclease-free water in a 0.2 mL thin-wall nuclease-free PCR tube. Place on ice. 2. On ice, add 2.5 μL of End Repair Buffer Mix and 0.5 μL of End Repair Enzyme Mix to the sample and mix by pipetting up and down. If more than one sample is treated, prepare a master mix

Malaria Strand-Specific RNA-Seq

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of sufficient amounts of End Repair Buffer Mix and End Repair Enzyme Mix before adding 3 μL to each sample (see Note 12). 3. Place the tube in a pre-warmed thermal cycler (lid heated at 100
C) with the following program: 30 min at 25
C; 10 min at 70
C; hold at 4
C. 4. Remove the sample promptly from the thermal cycler, give a quick spin, and place on ice. 5. Resuspend Agencourt® RNAClean XP magnetic beads by inverting and tapping the tube on the bench top. Do not spin the tube. 6. Add 12 μL of the bead slurry to the sample and mix thoroughly by pipetting up and down. Incubate at room temperature for 10 min. 7. Transfer tubes to the magnetic separation device and let them stand for 5 min (see Note 20). 8. While still on the magnet, carefully pipet off 15 μL of liquid without disturbing the beads (see Note 21). Dispersion and loss of significant amounts of beads will reduce yields. 9. While still on the magnet, gently add 200 μL of freshly made 70% ethanol and let stand for 30 s (see Note 22). 10. While still on the magnet, remove 200 μL of the ethanol wash (see Note 23). 11. Repeat step 9. 12. While still on the magnet, remove all of the ethanol wash. Carefully inspect the tube for the absence of ethanol drops. 13. While still on the magnet, air-dry the beads for 5–10 min. Carefully inspect the tube to ensure the ethanol has entirely evaporated. 14. Remove from the magnet and add 12 μL of nuclease-free water to the dried beads. Resuspend carefully by pipetting up and down. 15. Transfer the tubes to the magnet and let them stand for 1 min. 16. While on the magnet, carefully remove 11 μL of the eluate without disturbing the beads and transfer to a fresh tube. 17. Repeat step 15 to minimize the carryover of beads into the next stage of the library preparation. 18. While on the magnet, carefully remove 10 μL of the eluate without disturbing the beads and transfer to a fresh nucleasefree thin-wall 0.2 mL PCR tube. Place on ice. 19. Proceed immediately to Subheading 3.9.2 (see Note 24).

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3.9.2 Ligation

1. On ice, add 1 μL of Adaptor Mix to the sample (see Note 25). Mix by pipetting thoroughly with the pipette set to 5 μL. 2. On ice, add 12.5 μL of Ligation Buffer Mix and 1.5 μL of Ligation Enzyme Mix to the sample (the Ligation Buffer Mix is very viscous and should be pipetted slowly). If more than one sample is treated, prepare a master mix of sufficient amounts of Ligation Buffer Mix and Ligation Enzyme Mix before adding 14 μL to each sample (see Note 12). 3. Carefully mix by pipetting slowly up and down without forming bubbles with the pipette set at 20 μL. Spin down the tube for 2 s. 4. Place the tube in a pre-warmed thermal cycler (lid not heated) with the following program: 10 min at 25
C; hold at 4
C. IMPORTANT : Use this incubation time to prepare the Amplification Master Mix to be used in the library amplification reaction (see Subheading 3.9.3, step 1). The adapterligated sample must not remain on ice more than 10 min from the end of the ligation reaction to the beginning of the amplification reaction. 5. Remove the sample promptly from the thermal cycler, give a quick spin, and place on ice. 6. Proceed immediately to Subheading 3.9.3.

3.9.3 Library Amplification

1. Prepare an Amplification Master Mix by sequentially mixing the following reagents: 64 μL of Amplification Buffer Mix, 3 μL of Amplification Primer Mix, 4 μL of DMSO (this mix should have been prepared during the incubation indicated at Subheading 3.9.2, step 4). Place tube on ice. If more than one sample is treated, adapt volumes to prepare a sufficient quantity of master mix. 2. On ice, add 3 μL of Amplification enzyme mix and 1 μL of USER™ enzyme to the Amplification Master Mix immediately before adding to the adapter-ligated sample (the USER™ enzyme will degrade the second strand of the ds cDNA prior amplification to achieve strand specificity, see Note 26). If more than one sample is treated, adapt volumes to prepare a sufficient quantity of master mix. 3. Mix well by pipetting slowly, avoiding bubbles, spin, and place on ice. 4. Add 73 μL of Amplification Master Mix to a clean 0.2 mL thinwall nuclease-free PCR tube. 5. Add 7 μL of adapter-ligated sample to the tube prepared at step 4. Mix well by pipetting slowly up and down at the 73 μL pipette setting, avoiding bubbles, spin, and place on ice. The remaining adapter-ligated sample can be discarded.

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6. Place the tube in a pre-warmed thermal cycler (lid heated at 100
C) with the following program: 5 min at 95
C; 2 min at 72
C; 5 cycles of (30 s at 94
C—30 s at 55
C—1 min at 72
C); 10 cycles of (30 s at 94
C—30 s at 63
C—1 min at 72
C); 5 min at 72
C; hold at 10
C. 7. Remove the sample promptly from the thermal cycler, give a quick spin, and place on ice. 8. Resuspend Agencourt® RNAClean XP magnetic beads by inverting and tapping the tube on the bench top. Do not spin the tube. 9. Add 80 μL of the bead slurry to the amplified library and mix thoroughly by pipetting up and down (see Note 27). Incubate at room temperature for 10 min. 10. Transfer tubes to the magnetic separation device and let them stand for 5 min (see Note 20). 11. While still on the magnet, carefully pipet off 140 μL of liquid without disturbing the beads (see Note 21). Dispersion and loss of significant amounts of beads will reduce yields. 12. While still on the magnet, gently add 200 μL of freshly made 70% ethanol and let stand for 30 s (see Note 22). 13. While still on the magnet, remove 200 μL of the ethanol wash (see Note 23). 14. Repeat steps 12 and 13 two more times for a total of three washes. 15. While still on the magnet, remove all of the ethanol wash. Carefully inspect the tube for the absence of ethanol drops. 16. While still on the magnet, air-dry the beads for at 10–15 min. Carefully inspect the tube to ensure the ethanol has entirely evaporated. 17. Remove from the magnet and add 33 μL of 1 TE to the dried beads. Resuspend carefully by pipetting up and down. 18. Transfer the tubes to the magnet and let stand for 2 min. 19. While on the magnet, carefully remove 30 μL of the eluate without disturbing the beads and transfer to a fresh tube. Place on ice. 20. Stopping point: The amplified libraries can be stored at 20
C. When needed, thaw tubes on ice. Avoid repeated freeze/thaw cycles. 3.9.4 Qualitative and Quantitative Evaluation of the Library

1. Analyze 3 μL of the library on a 1.6% agarose gel electrophoresis (see Note 28) and check the size and the purity of the library. Quantify by UV spectrophotometry.

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Notes 1. Typically, parasites are cultured in 25 mL total volume at 5% hematocrit until a parasitemia of 6–10% is reached. If a synchronization is performed (e.g., using sorbitol) make sure to let the parasites recover from the stress of the treatment, ideally wait for one cycle of invasion, before harvesting the RNAs. Waiting will minimize the background caused by stress-related variations. This RNA-seq protocol is typically prepared using four different cultures pooled together. 2. It is crucial to dissolve everything at this step for an optimal yield. 3. Do not transfer any of the lower phase to the next step. Phenol inhibits downstream enzymatic reactions including reverse transcription. 4. Tubes containing nucleic acids that were precipitated at 20
C or 80
C should always be allowed to equilibrate on ice before centrifugation. At these low temperatures the samples tend to become very viscous and the efficiency of centrifugation is lower. 5. Typically, 0.5–1 μg of total RNA should be loaded on a 1.2% agarose gel. Mix sample with 10 volumes of denaturing RNA loading buffer (for 1.5 mL stock loading buffer mix: 750 μL of deionized formamide, 240 μL of formaldehyde 37%, 150 μL of 10X MOPS EDTA buffer pH 7.0, 200 μL of 50% glycerol, 0.5 mg of bromophenol blue, and 10 μL of ethidium bromide 10 mg/mL) and heat for 5 min at 65
C. Ensure that all solutions and hardware, including electrophoresis tank and gel combs, are RNAse-free. The 28S and 18S rRNAs should appear as two clean bands around 5.3 and 2 kb, respectively. The upper band should be more intense. The presence of significant smearing or a lower intensity of the upper band indicates degradation of the extracted material. 6. If the purity of the RNA solution is questioned, e.g., presence of phenol, the samples can be further purified on RNeasy® (QIAGEN) cleanup columns according to the “RNA Cleanup” manufacturer’s protocol. All solutions must be fresh. 7. It is crucial to eliminate all contamination with genomic DNA in order to avoid competition in downstream reaction and inaccurate quantitative analysis of RNA levels or false discovery of alternative transcripts. 8. Any loss in beads will result in a loss of material. For maximum yield, about 50 μL of sample should remain in the tube after removing the supernatant.

Malaria Strand-Specific RNA-Seq

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9. The efficiency of this step is directly linked to the amount of starting material. If desired, the incubation time can be adjusted accordingly but should not exceed 60 min. 10. A combination of random primers and oligos dT should always be used in experiments dealing with Plasmodium falciparum’s AT-rich genome to maximize the reverse transcription of all possible transcripts regardless of their GC content. 11. The 5 VILO™ Reaction mix already contains random primers, MgCl2, and dNTPs. The 10 SuperScript® Enzyme Mix includes the SuperScript® III Reverse Transcriptase (reduced RNase H activity and high thermal stability for extended synthesis), the RNAseOUT™ Recombinant Ribonuclease Inhibitor, and a helper protein proprietary to Invitrogen™. 12. When dealing with multiple samples at the same time, the delay between the preparation of the first sample and the preparation of the last sample should be kept to a minimum to ensure uniformity. Do not prepare more than eight samples at a time. 13. As a general rule when using nucleic acid cleanup and purification reagents, buffers containing ethanol should always be as fresh as possible. Aging solutions can cause dramatic losses in material. 14. At that point, the samples can theoretically be frozen at 20
C until further processing. Empirical observations seem to indicate, however, that the performances are significantly increased when second strand cDNA is synthesized immediately after first strand. Therefore, we do not recommend the freezing of first strand cDNA. 15. The substitution of the dTTP by dUTP in the dNTP mix is critical in this protocol since it will allow using the USER™ (Uracil-Specific Excision Reagent) enzyme prior library amplification and achieving strand specificity. The USER™ enzyme will leave a nucleotide gap at the location of a uracil in the second strand of the cDNA. 16. Obtaining high-quality ds cDNA is an absolute prerequisite for a successful preparation of a sequencing library. We recommend not proceeding if the ds cDNA is not of satisfactory quality (the presence of a regular smear on the gel and an OD  1.8 is an example of satisfactory quality). 17. These reaction conditions have been optimized to obtain fragments ranging 150 bp to 300 bp in size. Increase incubation time for shorter fragments, decrease it for longer ones. 18. Small amounts of nucleic acids cannot be detected by classical agarose gel electrophoresis. In order to avoid wasting large amounts of samples we recommend using microfluidic-based devices that can quantify and display the size distribution of a

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few microliters of a sample concentrated in the picogram per microliter range. The Bioanalyzer DNA High Sensitivity Chip (Agilent Technologies) can resolve 3 μL of purified DNA at 5 μg/μL in TE for sizes ranging 50–7000 bp. The LabChip® GX can resolve bands as low as 5 bp and features a sensitivity of 0.1 ng/μL. 19. The NuGEN® Encore™ NGS Library System I uses magnetic beads (RNAClean® XP Purification Beads supplied in the kit) for the successive purification of the samples through the library preparation steps rather than silicate-based spin columns. Magnetic beads allows for minimized sample loss and reduction of input material for library preparation. A magnetic separation device, such as the Agencourt® SPRIStand, is thus necessary to perform the purification steps. When using the Agencourt® SPRIStand, 96-well plates or tube strips are preferred rather than single tubes for greater stability in the stand and better separation. 20. Reduction in the incubation time of the beads on the magnetic stands will result in reduced recovery of the samples. Similarly, the various incubation times have been optimized to obtain reproducible results in terms of nucleic acid yield and size range. They must be strictly observed. 21. While on the magnet, the beads will stay on the walls of the tube and form a ring. Use a small volume pipette tip to reach the bottom of the tube without touching the sides and gently aspirate the desired volume. 22. If multiple samples are treated simultaneously, monitor the time spent in reaching the last tube and deduct it from the 30 s. 23. Always use the smallest volume pipette tip that allows the removal of the desired volume within 2 to 3 withdrawals. Do not try to get everything in one-step and pipet slowly to prevent any bead loss. 24. NuGEN® developed proprietary adapter and primer sequences directly compatible with the Illumina® Genome Analyzer and Hi-Seq systems. Their use differ from the more common Illumina® ones mostly in the fact that the step for 30 -end A-tailing of the fragments that is usually carried on prior adapter ligation is absent in the NuGEN® protocol. This specificity significantly reduces the hands-on time of the protocol. In addition, NuGEN® adapters generate libraries free from adaptor dimers, unlike Illumina®’s adapters. 25. The adapters are partly complementary and provided partially annealed to each other. This condition is necessary for a successful ligation to the sample of interest. Make sure to always keep the tube of Adapter Mix on ice so that the adapter duplex does not denature.

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26. The USER™ (Uracil-Specific Excision Reagent) enzyme is added to the amplification mix. During a short denaturing step (5 min at 95
C) prior to the actual amplification cycles, the USER™ enzyme nicks the second strand of the ds cDNA at uracil locations. Only the first strand is amplified and strand specificity is achieved. 27. If multiple samples are processed at the same time, it may be useful to use a multi-channel pipette to ensure consistent incubation times. 28. A high percentage of agarose is necessary to resolve small libraries. Increasing the amount of agarose, however, significantly increases the detection threshold using intercalant agents such as ethidium bromide. Here, preparing a gel at 1.5–1.8% agarose is a good compromise between resolution and sensitivity. In addition, the use of low range agarose, such as the Certified Low Range Ultra Agarose (Bio-Rad), greatly improves the resolution of small bands without having to increase the agarose content.

Acknowledgements The authors thank Courtney Brady (NuGEN®), and Barbara Walter, John Weger, Rebecca Sun, and Glenn Hicks (Institute for Integrative Genome Biology, University of California Riverside) for their assistance in the library preparation and sequencing processes. References 1. Le Roch KG et al (2003) Discovery of gene function by expression profiling of the malaria parasite life cycle. Science 301:1503–1508 2. Bozdech Z et al (2003) The transcriptome of the intraerythrocytic developmental cycle of Plasmodium falciparum. PLoS Biol 1:E5 3. Otto TD et al (2010) New insights into the blood-stage transcriptome of Plasmodium

falciparum using RNA-Seq. Mol Microbiol 76:12–24 4. Sorber K, Dimon MT, DeRisi JL (2011) RNA-Seq analysis of splicing in Plasmodium falciparum uncovers new splice junctions, alternative splicing and splicing of antisense transcripts. Nucleic Acids Res 39:3820–3835

Chapter 3 Laser Microdissection of Cells and Isolation of High-Quality RNA After Cryosectioning Marta Barcala, Carmen Fenoll, and Carolina Escobar Abstract Laser capture microdissection (LCM) has become a powerful technique that allows analyzing gene expression in specific target cells from complex tissues. Widely used in animal research, still few studies on plants have been carried out. We have applied this technique to the plant–nematode interaction by isolating feeding cells (giant cells; GCs) immersed inside complex swelled root structures (galls) induced by root-knot nematodes. For this purpose, a protocol that combines good morphology preservation with RNA integrity maintenance was developed, and successfully applied to Arabidopsis and tomato galls. Specifically, early developing GCs at 3 and 7 days post-infection (dpi) were analyzed; RNA from LCM GCs was amplified and used successfully for microarray assays. Key words Laser-capture microdissection, RNA isolation, Cryosectioning, Arabidopsis, Tomato, Galls, Root-knot nematode, Giant cells

1

Introduction Laser-capture microdissection (LCM) is a technique that allows harvesting specific cells from complex tissues or populations for specific RNA, DNA, or protein isolation [1, 2] so that they can be used in downstream applications such as microarray hybridization, cDNA library construction, proteomic analysis, etc. So far several laser-capture equipments have been developed by different companies [3]. We performed LCM with the PixCell II system (Arcturus), which allows isolating cells using a low-power (infrared) laser and retaining them in a thermoplastic film; as the laser radiation is absorbed by the film instead of by the cell samples, this procedure should preserve the integrity of the captured material. The technique is applied to tissue sections that can be prepared from either frozen or paraffin-embedded biological samples. In general, better morphology can be observed in paraffin-embedded tissues fixed with non-coagulating fixatives, whereas the use of coagulating fixatives (such as ethanol: acetic acid) for frozen tissues

Hailing Jin and Isgouhi Kaloshian (eds.), RNA Abundance Analysis: Methods and Protocols, Methods in Molecular Biology, vol. 2170, https://doi.org/10.1007/978-1-0716-0743-5_3, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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renders higher RNA yield and quality [4]. Thus, a compromise between both good morphology and RNA preservation should be achieved. A few attempts to study specifically nematode feeding sites (NFS) have been carried out [5, 6]. The first report was based on micro-aspiration of the cytosolic content of tomato GCs [7], but this method could only be applied to large GCs at late differentiation stages. In contrast, LCM represented an advance as it permits precise NFS collection at any infection time as long as they can be identified in histological sections. The first reported NFS isolation by LCM was applied to syncytia induced by cyst nematodes [8]. We have developed a protocol to both preserve morphology and render high-quality RNA from GCs formed by root-knot nematodes at different developmental stages (3 and 7 days post-infection (dpi)) of either Arabidopsis or tomato. Briefly, this protocol consists of a mild fixation step with a non-crosslinking fixative, gall cryosectioning, GCs LCM, and eventually RNA extraction. The RNA obtained by this protocol has been successfully used for microarray analysis. Safety handling measures to avoid RNA degradation are strongly recommended, such as always wearing gloves, preparing all the solutions with diethyl pyrocarbonate (DEPC)-treated deionized water and using RNase-free plastic and glassware. To prepare 0.1% DEPC-treated deionized water: add 1 ml DEPC to 1 l of deionized water (see Note 1) and stir overnight at room temperature (RT) and autoclave it for 20 min at 121  C to destroy DEPC.

2 2.1

Materials Tissue Fixation

1. Ethanol-acetic acid (EAA) fixative solution: 3 parts of absolute ethanol (molecular biology grade) and one part of glacial acetic acid (3:1 v/v). Prepare it in a conical tube or a glass bottle by mixing the ethanol and acetic acid and keep it tightly closed on ice. Only freshly made fixative solution should be used. 2. Microcentrifuge tubes. 3. Surgical blade in a scalpel with handle. 4. Pointed tip tweezers.

2.2 Cryoprotective Solutions

1. 0.01 M phosphate buffered saline solution (PBS), pH 7.4: 0.138 M NaCl, 2.7 mM KCl. Dissolve a pouch in 1 l of DEPC-water (see Note 2) and store at RT. 2. 10% sucrose in 0.01 M PBS pH 7.4. Add 5 g of sucrose to PBS for a final volume of 50 ml. Dissolve completely and store at 4  C (see Note 3). 3. 15% sucrose in 0.01 M PBS pH 7.4. Add 7.5 g of sucrose to PBS for a final volume of 50 ml. Dissolve completely and store at 4  C.

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4. 34.3% sucrose in 0.01 M PBS pH 7.4. Add 17.25 g of sucrose to PBS for a final volume of 50 ml. Dissolve completely and store at 4  C. 5. 34.3% sucrose, 0.01% safranine-O dye, 0.01 M PBS pH 7.4. Prepare this solution by adding safranine-O from a 1000 stock to the previous solution. 6. 10% safranine-O (1000): weigh 100 mg of safranine-O and add it to 1 ml of 0.01 M PBS, pH 7.4. 7. Vacuum eppendorf concentrator 5301 (Eppendorf, Hamburg, Germany). 8. Orbital shaker. 2.3 Embedding and Cryosectioning

1. Embedding media: Tissue-TEK®, optimal cutting temperature grade (O.C.T) media (Sakura Finetek, AV Alphen aan den Rijn, The Netherlands). 2. Cryomoulds: mould disposable base of 7  7  5 mm (see Note 4). 3. 2-Isopentane (Methyl butane). 4. Liquid nitrogen. 5. Long forceps. 6. Poly-L-Lysine coated slides: Polysine® (BDH, Poole, UK). 7. Vertical glass staining jars. 8. Ethanol solutions: 70% and 95% Ethanol. 9. Xylene. 10. Desiccant (silica gel). 11. Cryostat with disposable blades and anti-roll device. We have developed the protocol using a Leica CM3050S cryostat (Leica Microsystems, Wetzlar, Germany).

2.4 Laser Capture Microdissection and RNA Extraction

1. PixCell II Laser Capture Microdissection system (Arcturus®, Life Technologies, California, USA). 2. CapSure® HS LCM caps (Arcturus). 3. ExtracSureTM Sample Extraction device (Arcturus). 4. Absolutely RNA Nanoprep kit (Stratagene, California, USA).

3 3.1

Methods Tissue Fixation

Unless otherwise specified, all the steps are carried out at 4  C by placing the microcentrifuge tubes on ice. 1. Clean the working surface and all metallic dissection instruments, such as the scalpel handle and the tweezers, with acetone.

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2. Prepare aliquots with 1.5 ml of EAA fixative in as many microcentrifuge tubes as needed (see Note 5), and let them cool down on ice. 3. Localize the galls and root pieces to dissect under a stereo microscope; open the plate containing the in vitro grown infected Arabidopsis plants and cover them with cool freshly made fixative EAA (see Note 6). Collect all the galls and control root pieces one by one, and transfer them into the fixative-filled tubes very quickly. Cut the samples carefully, leaving a small portion of root at both sides of the gall to facilitate sample handling with a pair of tweezers (i.e., when orienting the sample in the moulds). 4. To facilitate fixative infiltration apply vacuum to the samples for 15 min (see Note 7). Then gently swirl the tubes for 1 h (i.e., in a shaker at 70 rpm). 5. Replace the solution with fresh fixative; repeat the infiltration as in step 4 and swirl samples for 2 h (see Note 8). 6. Discard the fixative and add 10% sucrose in 0.01 M PBS pH 7.4 to each tube. Apply vacuum for 10 min and swirl the samples for 3 h as described in step 4. 7. Continue with the cryoprotective treatment by infiltrating samples successively in 15% sucrose in 0.01 M PBS pH 7.4, and 34.3% sucrose, 0.01% safranine-O, 0.01 M PBS pH 7.4 (see Note 9) as described in step 6. Swirl the samples for 3 h and overnight, respectively, as described in step 4. 8. Rinse the samples in 34.3% sucrose in 0.01 M PBS pH 7.4 to remove excess safranine-O dye immediately prior to embedding (see Note 10). 3.2

Embedding

1. Cool down isopentane almost to its freezing point by submersion in a liquid nitrogen bath (see Note 11) (Fig. 1a). 2. Label carefully the moulds. 3. Fill a mould with Tissue -Tek® O.C.T. compound, taking special care to avoid air bubbles. This prevents moulds from cracking during freezing and sectioning (see Note 12). 4. Carefully introduce a sample in the O.C.T. media and orient it with the tweezers. It is important to drain as much sucrose solution as possible from the sample (see Note 13). 5. Freeze the sample by immersing the entire mould in precooled isopentane for at least 20 s. Large samples will need longer immersion times. Long, wide open forceps are recommended as they facilitate handling the mould during submersion into the isopentane without disturbing sample orientation (Fig. 1a). O.C.T. will turn white upon freezing.

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Fig. 1 Some steps of the LCM process. (a) Custom-made setup for cooling isopentane in a liquid nitrogen bath. (b) Cryosections of Arabidopsis galls before (upper panels) and after (lower panel) dehydration. Giant cells are labeled by white asterisks. (c) LCM PixCell II instrument. (d) The LCM process, showing a CapSure® placed over a slide and the laser beam, an Arabidopsis root section after exposure to the laser beam, and the microdissected area captured onto a CapSure®

6. Transfer the mould to liquid nitrogen until you have finished freezing all the samples; then store the moulds at 80  C for later cryosectioning. 3.3 Cryosectioning and Laser Capture Microdissection

1. Let the cryostat cool down to 20  C (see Note 14). Then, place the samples inside and let them equilibrate to the cryostat temperature for at least 15 min. Also keep a staining jar filled with 70% ethanol inside the cryostat.

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2. Trim off the mould and cut it in 10 μm sections. During cryosectioning we check tissue morphology by picking a section in a slide (see next step) and mounting it with glycerol to avoid desiccation. The section can then be observed and photographed under a microscope (Fig. 1b). 3. When GCs are discerned within a section, label a slide appropriately with a pencil and pick up the section. For this purpose, drop a slide (stored at room temperature) carefully onto the cut section/s. We usually mount 3–4 sections per slide (see Note 15). Keep then the slide inside the cryostat until you finish collecting the 3–4 sections, and then transfer it to the staining jar filled with 70% ethanol. Store the staining jar at 20  C until sectioning is completed (see Note 16). 4. Prior to LCM, the O.C.T. compound has to be removed and sections dehydrated (Fig. 1b). To do this, perform sequential washes in staining jars as follows: 70% ethanol (30 s, RT), 0.01 M PBS pH 7.4 (30 s, RT), 70% ethanol (30 s, RT), 95% ethanol (overnight, at 20  C), and 100% ethanol (30 s, RT). Finally, dehydrate sections completely by washing slides twice in xylene (10 min each) (see Note 17). Let slides air-dry until xylene has completely evaporated, and store them at RT in a box with desiccant (such as silica gel). Sections should be used for LCM immediately. 5. Follow manufacturer’s instructions for laser capture microdissection (see Note 18; Fig. 1c). 6. Microdissect as many giant- or vascular control cells per cap as possible. This number varies depending on the tissue size, position in the slide, etc. We usually collect around 50 giant cells (3 dpi) from either Arabidopsis or tomato galls, and 100 vascular control cells on each cap (see Note 19; Fig. 1d). Once the cap is full of microdissected tissue, attach the ExtracSure Sample Extraction device to the cap and store it at RT until finishing with LCM (see Note 20). 3.4

RNA Extraction

1. RNA extraction is performed with the Absolutely RNA Nanoprep kit. Manufacturer’s instructions are followed with minor modifications (see Note 21). 2. Eluted RNA is stored at amplification.

4

80



C until subsequent

Notes 1. DEPC is highly toxic and volatile, and it must always be handled in a fume hood.

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2. Instead of purchasing PBS buffer, it can be also prepared as described in [9] and later it can be treated with DEPC. 3. Sucrose solutions become easily contaminated; therefore, we recommend preparing a reasonable volume and discard the remaining solutions once the procedure is completed. 4. Different base moulds can be used according to the size of the processed tissue; usually, the smaller the base mould the better, as large blocks tend to crack when being frozen. 5. In general, several samples can be fixed in each microcentrifuge tube, but this depends on the sample size, as enough fixative must be in contact and completely cover each sample. Usually, up to five 3-dpi galls were added per tube. 6. This step can be omitted when only a root piece has to be dissected from a plate. If several plants or root pieces have to be dissected from a single plate, it is advisable to cover the plate with approximately 5 ml of fixative to prevent root desiccation while the plate remains opened. Desiccation will collapse the tissue and destroy its morphology. 7. Depending on the tissue, vacuum treatment has to be empirically adjusted. Usually 10–15 min is sufficient to make tomato or Arabidopsis root tissue sink to the bottom of the tube. Avoid strong vacuum pumps as the tissue could be damaged, and bubbling air can eject samples from the tube. To avoid this, tubes can be wrapped with parafilm-like plastic with some holes in it. It is important to point out that, when not using the same vacuum device as described in this protocol, the incubation times required for best results could vary considerably depending on the pump strength used. 8. Replace the fixative very carefully without damaging or loosing the galls or the control roots. For that purpose use a Pasteur pipette or a thin plastic tip and keep samples always floating in solution; do not let samples be uncovered at any time. It is recommended to leave less than 20 μl (ideally 10 μl) of the previous solution to avoid desiccation. It is also important to visually localize samples throughout the process in order to avoid having them get stuck to the tube walls. 9. The concentration of sucrose as cryoprotectant may be adjusted depending on the characteristics of the cells to be preserved. Additionally, although the use of dyes might not be advisable [10], there are exceptions with tissues (e.g., very young Arabidopsis galls or roots) that become nearly transparent and can be hardly identified inside the tubes, especially in later steps such as while embedding and cryosectioning. 10. As safranine-O is a water-soluble dye, long incubation in 0.01 M PBS solution will fade sample staining.

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11. Work in a fume hood, as isopentane is very volatile. Fill half of a 100 ml beaker with isopentane and slowly transfer it to a Dewar flask containing liquid nitrogen. Leave the base of the beaker in contact with liquid nitrogen until solution becomes misty and thickens at the bottom. If a solid opaque white surface appears at the bottom (frozen isopentane), lift up the beaker out of the liquid nitrogen to warm it, and repeat the cooling process. 12. Keep the O.C.T. compound bottle upside down and squeeze it very gently when filling the cryomould to avoid bubble formation. If any bubble appears, remove it with a thin needle or a pipette tip. Some other protocols recommend adding a thin layer of O.C.T. to the mould, then orient the sample and finally fill the mould with O.C.T. In our hands, this was not adequate as we added several samples per mould (see Note 13), and the samples were easily disturbed. 13. Use very thin tip tweezers to take samples out of the tube and to orient them in the O.C.T. We usually place 2–3 galls or control roots per mould; this step is a bit tricky, mostly due to the bubbles that might be formed. Sucrose solution could be easily drained from the sample with a gentle touch onto a filter paper; however, this should be done with extreme care as it could over-dry the sample. Therefore, we omitted this step during Arabidopsis gall processing. 14. Before cryosectioning, be sure that the anti-roll plate and the disposable blades have been cleaned and cooled down, since otherwise the sample could melt. It is also advisable to keep spare disposable blades inside the cryostat, just in case additional ones are needed. 15. At least for some laser capture microdissectors, technical limitations require sections to be centered in the slide. Therefore, when several sections per slide are needed, ideally a ribbon of 3–4 sections (as those from paraffin sectioning) is required; this implies becoming skilled in cryosectioning. 16. We usually place two 70% ethanol staining jars inside the cryostat. If there is not enough room inside the cryostat, prepare in advance several staining jars with 70% ethanol and keep them in the freezer. 17. Xylene is hazardous and volatile; always handle it in a fume hood. 18. We used an Arcturus® PixCell II system for LCM. Laser beam power and length must be empirically established; for our tissue, it was typically set to 90–110 mW, 70 μs and a spot size of 7.5 μm. Recommended materials from the manufacturer were used, such as Arcturus CapSure® HS LCM caps for LCM, and the ExtracSure sample extraction device for RNA extraction.

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19. Sometimes non-desired tissue remains attached to the cap after microdissection. This can be easily eliminated by gently pressing the cap onto a post-it note without affecting RNA quality at all. 20. We processed caps immediately after LCM; it is not recommended to keep them at RT for more than 2 h, the longest time we tested with no detectable effects on RNA quality. 21. Caps were incubated in lysis buffer-β-mercaptoethanol mixture for 2–3 min at 60  C. DNase treatment was performed as indicated by the manufacturer. Finally, RNA was eluted twice with 20 μl elution buffer pre-warmed to 60  C. For RNA extraction from tomato microdissected GC, elution in a smaller volume (twice in 10 μl) has been successfully used.

Acknowledgements The authors thank K. Lindsey and J. Topping (University of Durham, UK) for the use of their equipment and for their advice during the development of the microdissection protocol. Work in the laboratory was supported by grants from the Ministry of Education (AGL2016-75287-R) and by the Castilla-La Mancha Government (SBPLY/17/180501/000287) to CE. M.B. was a recipient of a technologist contract from the University of CastillaLa Mancha and the Fondo Europeo de Desarrollo Regional (FEDER). References 1. Bonner RF et al (1997) Laser capture microdissection: molecular analysis of tissue. Science 278:1481–1483 2. Buck MR et al (1996) Laser capture microdissection. Science 274:998–1001 3. Bagnell C (2006) Laser capture microdissection. In: Coleman W, Tsongalis G (eds) Molecular diagnostics: for the clinical laboratorian. 2. Humana Press Inc, Totowa, NJ, US, pp 219–224 4. Portillo M et al (2009) Isolation of RNA from laser-capture-microdissected giant cells at early differentiation stages suitable for differential transcriptome analysis. Mol Plant Pathol 10:523–535 5. Escobar C, Sigal B, Mitchum MG (2011) Transcriptomic and proteomic analysis of the plant response to nematode infection. In: Jones J, Fenoll C, Gheysen G (eds) Genomics and molecular genetics of plant-nematode interactions. Springer science, Berlin/Heidelberg, pp 155–171

6. Shahzad MA et al. (2016) An improved procedure for isolation of high-quality RNA from nematode-infected Arabidopsis roots through laser capture microdissection. Plant Methods 12:25. 7. Wang Z, Potter R, Jones MGK (2001) A novel approach to extract and analyse cytoplasmic contents from individual giant cells in tomato roots induced by Meloidogyne javanica. Int J Nematol 11:219–225 8. Ithal N et al (2007) Developmental transcript profiling of cyst nematode feeding cells in soybean roots. Mol Plant-Microbe Interact 20:510–525 9. Sambrook J, Russell D (2001) Molecular cloning: a laboratory manual, vol 3, 3rd edn. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, New York 10. Ramsay K, Wang Z, Jones M (2004) Using laser capture microdissection to study gene expression in early stages of giant cells induced by root-knot nematodes. Mol Plant Pathol 5:587–592

Chapter 4 Detection of RNA in Ribonucleoprotein Complexes by Blue Native Northern Blotting Lena Kru¨ßel, Steffen Ostendorp, Anna Ostendorp, and Julia Kehr Abstract Northern blotting is a classical technique that allows the detection of specific nucleic acids using radioactive or non-radioactive probes. Normally, nucleic acids are denatured and separated by agarose or polyacrylamide gel electrophoresis and transferred and fixed to a membrane prior to detection. Here, we describe a method to analyze specific RNA in native ribonucleoprotein complexes using blue native PAGE with subsequent northern blotting, crosslinking of RNA onto a suitable membrane, and detection using non-radioactive probes. Key words Native northern blot, Non-radioactive nucleic acid detection, Ribonucleoprotein complex, RNA-binding proteins

1

Introduction Large RNA–protein interactions and ribonucleoprotein complexes are crucial for biosynthesis, gene regulation, and maintaining the structural integrity of the cell [1, 2]. To get further functional insights it is crucial to identify the proteins and RNAs these complexes are composed of. There are two basic approaches to analyze RNA–protein complexes, protein-centric, and RNA-centric [3]. Protein-centric approaches imply knowledge about the protein of interest and often a purified protein, whereas RNA-centric methods focus on binding of unknown proteins with specific RNAs [3]. These approaches include both native and fully denaturing methods [3]. Nevertheless it is still difficult to obtain RNA:protein interactions that occur in vivo. Our approach can be assigned to the RNA-centric methods. Normally northern blotting includes a denaturing polyacrylamide gel electrophoresis separating the RNA with subsequent transfer and fixation of RNA to a suitable membrane like nylon or nitrocellulose. Afterwards specific RNAs can be detected by either radioactively labelled probes or by non-radioactive DNA probes,

Hailing Jin and Isgouhi Kaloshian (eds.), RNA Abundance Analysis: Methods and Protocols, Methods in Molecular Biology, vol. 2170, https://doi.org/10.1007/978-1-0716-0743-5_4, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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e.g., digoxigenin or biotin-labeled. In contrast to that, our method combines the separation of large native protein complexes by blue native PAGE according to [4] and a subsequent non-radioactive northern blot approach to detect specific RNAs. Earlier reports already demonstrated that native RNA can be detected by northern blotting [5], but the technique has been rarely applied. Using our method allows to define protein complexes that comprise specific RNAs like non-coding RNAs or mRNAs. It can be combined with further investigation of these complexes by additional separation steps using electrophoresis and subsequent identification of the proteins by mass spectrometry (see also [6, 7]).

2

Materials Prepare all solutions using deionized RNase-free water and analytical grade RNase-free reagents. Prepare and store all reagents at room temperature unless indicated otherwise.

2.1 Blue Native Gel Electrophoresis

1. Novex® NativePAGE™ (Invitrogen, USA).

4–16%

Bis-Tris

Protein

Gel

2. Cathode buffer B (15 mM Bis-Tris pH 7.0, 50 mM Tricine, 0.02% (w/v) Coomassie blue G250). 3. Anode buffer (50 mM Bis-Tris pH 7.0). 4. Cathode buffer B/10 (15 mM Bis-Tris pH 7.0, 50 mM Tricine, 0.002% (w/v) Coomassie blue G250). 2.2 Blue Native Northern Blotting

1. Fastblot 43B semidry Blot (Biometra, Germany). 2. Positively charged nylon membrane (e.g., Amersham HybondN+, GE Healthcare Life Sciences, Sweden). 3. Filter paper (Whatman™ 3MM Chr, GE Healthcare Life Sciences, Germany). 4. UV-crosslinker (Stratalinker 2400, Agilent, USA). 5. ULTRAhyb© Ultrasensitive Hybridization Buffer (Ambion®, life technologies, Lithuania). 6. 30 biotinylated oligonucleotide probes (see Note 1). 7. Transfer buffer: 1 TBE buffer (89 mM Tris pH 7.6, 89 mM boric acid, 2 mM EDTA) (see Note 2). 8. Washing buffer: 2 SSC buffer, 0.1% SDS (300 mM NaCl, 30 mM sodium citrate pH 7.0) (see Note 3). 9. Pierce Chemiluminescent Nucleic Acid Detection Module Kit (ThermoFisher Scientific, Germany).

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10. Plastic container. 11. Hybridization oven. 12. CCD camera or X-ray film.

3

Methods Carry out all procedures at room temperature unless otherwise specified.

3.1 Blue Native Gel Electrophoresis

Carry out the electrophoresis steps at 4  C. Precool the buffers to 4  C. 1. Use a commercial Bis-Tris gradient gel (e.g., Novex NativePAGE 4–16% Bis-Tris, Invitrogen) (see Note 4). 2. Assemble the gel system (e.g., Mini-PROTEAN system, Bio-Rad) (see Note 5). 3. Fill the inner chamber with dark blue cathode buffer B to the half. Rinse the wells with buffer B and load 20–30 μg of the concentrated protein sample per well onto the gel by using e.g., a syringe for a better loading. Fill up the outer chamber with anode buffer (see Note 6). 4. Run the gel at 150 V for approximately 20–30 min, until the dye front is at 1/3 of the gel. 5. Then pause the run and remove the dark blue cathode buffer B. Fill up the inner chamber with light blue cathode buffer B/10 and continue the run. Stop the run when the blue dye front starts to run out of the gel (see Note 7).

3.2 Blue Native Northern Blotting

1. To prepare northern blotting cut the nylon membrane and 12 filter papers to the size of the gel. 2. Remove the gel plates, transfer the gel into a plastic container filled with transfer buffer, and equilibrate it for 10 min (see Note 8). 3. Wet membrane and filter papers in a plastic container containing transfer buffer. Assemble filter papers, nylon membrane and gel as shown in Fig. 1. Place six filter papers onto the semidry blotter. Place the membrane on the filter papers and put the gel on top. Mark the gel position and wells on the membrane with a pencil. Finally place six filter papers soaked with transfer buffer onto the gel. Remove bubbles carefully by rolling with e.g., a serological pipette after every layer. Start semidry blotting with 3.5 mA/cm2 for 1 h (see Note 9). 4. Rinse the membrane with DEPC-treated water and dry between two filter papers.

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Fig. 1 Blue native gel after electrophoresis. Arrows indicating Coomassie stained, visible protein complexes of concentrated phloem sap from Brassica napus. Box and scissor indicate a lane cut out for subsequent northern blotting

Fig. 2 Northern blot assembly for the semidry transfer of native RNA onto a nylon membrane

5. Immobilise RNA on the membrane by UV-crosslinking with 120,000 μJ/cm2 (autocrosslink program with Stratalinker 2400) (see Note 10). 6. To test different DNA probes, the membrane can be cut to use only one lane per probe. 7. For pre-hybridization, place one membrane piece into a 15-mL reaction tube (for larger membranes use larger reaction tubes or a plastic container) and add 6 mL of hybridization buffer. Pre-hybridize for 1 h at 68  C in a hybridization oven. 8. Prepare 1 mL hybridization buffer containing 4 μL of 30 -biotinylated DNA probe (100 nm) in a 1.5-mL reaction tube, mix well and add it to the membrane (see Note 11). 9. For hybridization, place the membrane back into the same hybridization oven. Set the temperature to 37  C and let it cool down over night. 10. Continue on the next day with chemiluminescent detection of the nucleic acid (Fig. 2). Place the membrane into a new plastic container and wash two times with 2 SSC, 0.1% SDS.

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11. Detection of biotin-labeled probes will be performed via the Chemiluminescent Nucleic Acid Detection Module (Thermo Fisher Scientific) following manufacturer’s instructions. The sensitive chemiluminescent reaction is performed by a streptavidin-conjugated horseradish peroxidase and luminol as substrate. 12. Imaging can be done on an X-ray film or by a CCD imaging instrument. 13. The membrane can be reused (see Note 10).

4

Notes 1. Probes can be biotinylated by an appropriate biotinylation kit (e.g., Pierce™ Biotin 30 End DNA Labeling Kit, Thermo Fisher Scientific). 2. Prepare 10 transfer buffer (890 mM Tris base, 890 mM boric acid, 20 mM EDTA-Na2). Weigh 53.9 g Tris base, 27.5 g boric acid, and 20 mL 0.5 M EDTA-Na2. Then fill up with DEPCtreated water to 400 mL. Adjust pH to 8.3 and fill up to 500 mL. Store at room temperature up to 6 months and dilute to 1 prior to use. 3. Prepare 20 SSC washing buffer (3 M sodium chloride, 300 mM sodium citrate). Weigh 175.3 g sodium chloride and 88.2 g sodium citrate and dissolve in 800 mL DEPC-treated water. Adjust the pH to 7 by adding HCl and fill up to 1 L with DEPC-treated water. Sterilization can be done by autoclaving. Prepare 10% SDS stock solution. SDS can cause irritations. Wear gloves and lab coat when working with SDS and SDS containing solutions. For 1 L 2 SSC + 0.1% SDS wash solution, add 100 mL 20 SSC and 10 mL 10% SDS solution to 500 mL DEPC-treated water. Fill up to 1 L with DEPCtreated water. 4. Alternatively you can use a self-casted blue native gradient gel, as described e.g., in [8]. 5. Test with water if the inner chamber is tight prior to adding buffer into the gel chambers. 6. The clear sample can be seen well in the dark blue buffer and no dye is required. 7. Fill the used cathode buffer B in a separate bottle. The buffer can be reused. After the run the gel can be stored at 4  C for at least 1 week. 8. Either the whole gel or only part of the gel can be transferred onto a membrane (Fig. 3), the residual part can be used for other experiments like protein identification via a 2nd (and 3rd) gel dimension with subsequent mass spectrometric analysis.

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Fig. 3 Result of a blue native northern blotting showing a blue native gel after gel electrophoresis in lane 1 and the developed nylon membrane in lane 2. Arrow indicates a signal for the tRNA GlyCCC

9. Bubbles will hinder RNA transfer. 10. After crosslinking, the membrane can be stored for at least 1 year at room temperature. 11. The membrane can be reused after stripping the membrane with boiling 0.1% SDS solution. The solution should be prepared with DEPC-treated water to avoid degradation of the RNA by RNases. The membrane can then be stored dry at room temperature between filter paper until reuse.

Acknowledgements This work was financially supported by a Career Integration Grant (CIG; PCIG14-GA-2013-63 0734) by the European Commission within the 7th framework program, the grant LFF-GK06 ‘DELIGRAH’ (Landesforschungsfo¨rderung Hamburg), and the DFG grant (DFG KE 856_6-1) awarded to JK. References 1. Glisovic T, Bachorik JL, Yong J et al (2008) RNA-binding proteins and post-transcriptional gene regulation. FEBS Lett 582:1977–1986 2. Nissen P (2000) The structural basis of ribosome activity in peptide bond synthesis. Science 289:920–930 3. Mchugh CA, Russell P, Guttman M (2014) Methods for comprehensive experimental

identification of RNA-protein interactions. Genome Biol 15:203 4. Schagger H, Cramer WA, von Jagow G (1994) Analysis of molecular masses and oligomeric states of protein complexes by blue native electrophoresis and isolation of membrane protein complexes by two-dimensional native electrophoresis. Anal Biochem 217:220–230

Blue Native Northern Blotting 5. Khandjian EW, Me´ric C (1986) A procedure for northern blot analysis of native RNA. Anal Biochem 159:227–232 6. Ostendorp A, Pahlow S, Kru¨ßel L et al (2017) Functional analysis of Brassica napus phloem protein and ribonucleoprotein complexes. New Phytol 214:1188–1197

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7. Pahlow S, Ostendorp A, Kru¨ßel L et al (2018) Phloem sap sampling from Brassica napus for 3D-PAGE of protein and ribonucleoprotein complexes. J Vis Exp 131:e57097 8. Fiala GJ, Schamel WW, Blumenthal B (2011) Blue native polyacrylamide gel electrophoresis (BN-PAGE) for analysis of multiprotein complexes from cellular lysates. J Vis Exp 48:e2164

Chapter 5 Quantitative Analysis of Plant miRNA Primary Transcripts Jakub Dolata, Andrzej Zielezinski, Agata Stepien, Katarzyna Kruszka, Dawid Bielewicz, Andrzej Pacak, Artur Jarmolowski, Wojciech Karlowski, and Zofia Szweykowska-Kulinska Abstract MicroRNAs control plant development and are key regulators of plant responses to biotic and abiotic stresses. Thus, their expression must be carefully controlled since both excess and deficiency of a given microRNA may be deleterious to plant cell. MicroRNA expression regulation can occur at several stages of their biogenesis pathway. One of the most important of these regulatory checkpoints is transcription efficiency. mirEX database is a tool for exploration and visualization of plant pri-miRNA expression profiles. It includes results obtained using high-throughput RT-qPCR platform designed to monitor pri-miRNA expression in different miRNA biogenesis mutants and developmental stages of Arabidopsis, barley, and Pellia plants. A step-by-step instruction for browsing the database and detailed protocol for highthroughput RT-qPCR experiments, including list of primers designed for the amplification of pri-miRNAs, are presented. Key words mirEX database, microRNA, pri-microRNA, RT-qPCR, Gene expression

1

Introduction MicroRNAs (miRNAs) are 20–24 nucleotides long noncoding, single-stranded RNA molecules that take part in a wide variety of physiological and cellular processes. To this day, the participation of plant miRNAs in leaf, root, and floral development, vegetative/ generative phase change, signal transduction, environmental, and biotic stress response has been experimentally proven [1–4]. To fulfill their role as a quick reaction force that allows rapid developmental changes and responses to environmental cues, their biogenesis must be adjusted to the current cell need and tightly controlled. In contrast to animals, the biogenesis of plant miRNAs occurs entirely in the cell nucleus (reviewed in [5]). MicroRNA genes (MIRs) are transcribed exclusively by RNA polymerase II into primary precursors (pri-miRNAs). Nascent pri-miRNAs are capped at the 50 end and polyadenylated at the 30 end, and contain

Hailing Jin and Isgouhi Kaloshian (eds.), RNA Abundance Analysis: Methods and Protocols, Methods in Molecular Biology, vol. 2170, https://doi.org/10.1007/978-1-0716-0743-5_5, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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characteristic hairpin-like secondary structures in which sequences of mature miRNAs are embedded [6]. These pri-miRNAs are then converted into shorter miRNA precursors, called pre-miRNAs, that consist only of miRNA-containing stem-loop structures which are then processed into miRNA/miRNA∗ duplexes with 2 nt overhang at the 30 -terminus of each duplex strand. The cleavage of pri- and pre-miRNAs is conducted always by enzymes that are members of the ribonuclease III family (mainly DICER-LIKE1 (DCL1)) [7]. However, for efficient and precise processing of plant pri-miRNAs by DCL1, two other proteins, HYPONASTIC LEAVES 1 (HYL1) and SERRATE (SE), are required. These three proteins, DCL1, HYL1, and SE, form the core of the plant miRNA biogenesis machinery. However, plenty of additional protein factors were recently identified to participate in pri-miRNA processing (reviewed in [8]). In the next step, the miRNA/ miRNA∗ duplex is exported to the cytoplasm and loaded into RNA-induced silencing complex (RISC), where miRNA∗ is removed and the mature miRNA fulfils a role of a guide leading to mRNA that can be simply cleaved or stopped in translation. In addition to the involvement of many different proteins in regulation of miRNA biogenesis, the complex structure of miRNA genes may result in an elaborate pattern of pri-miRNA processing. The majority of MIRs in plants represent independent transcription units (67%), and the remaining genes encode miRNAs located within the introns of protein-coding or noncoding RNA genes [9, 10]. miRNAs can be found in exons (exonic miRNAs) and introns (intronic miRNAs) and their biogenesis may involve regulatory events such as alternative splicing, alternative polyA site selection, and/or multiple transcription start sites (reviewed in [8]). Indeed, the significance of crosstalk between the microprocessor, spliceosome, and polyadenylation machinery in miRNA production in Arabidopsis has been reported in several recent studies [11–16]. Additionally, the interplay between posttranscriptional processing events might by developmentally specific [17]. In order to understand the mechanisms responsible for plant growth and response to stress, it is of thus utmost importance to quantify precisely the expression level of miRNA genes at different stages of miRNA biogenesis in studied conditions. The pri-miRNA level is the element that allows to monitor transcriptional and posttranscriptional regulation of miRNA biogenesis. There are several methods to evaluate transcript levels: Northern blot, RT-qPCR, microarrays, and RNA sequencing. While Northern blot is a low-throughput technique, the other three allow to analyze more than one transcript at a time. DNA microarray technology is a well-established and powerful tool, that is mainly used in biomedical science for diagnostics [18]. However, it utilizes probes of known cDNA sequences and thus does not allow to identify novel transcripts and their variants. In contrast,

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RNA-seq enables whole-transcriptome analyses including alternative splice variants and due to the decreased running costs it has become the gold standard for gene expression quantification [19]. However, the RNA-seq field still encounters challenges in terms of data processing and analyses, since depending on the processing workflow, slightly different sets of differentially expressed genes may be obtained [20]. Additionally, the levels of pri-miRNAs are rather low in the cell and not all of them can be easily detected by RNA-seq. For instance, from 329 Arabidopsis pre-miRNAs predicted and deposited in miRBase database (Release 22) Lepe-Soltero and colleagues have been recently able to detect only 133 primary precursors in se-1, the mutant of SE in which the processing of pri-miRNAs is impaired and thus their levels are upregulated [10, 21]. Alternatively, the RT-qPCR technique is highly quantitative and sensitive, that together with possibility of performing over 300 reactions in a single run makes it a highthroughput approach in quantifying transcript levels [22]. Nevertheless, there is one major potential difficulty that arises when using this powerful technique—the requirement of normalization based on reference genes. Several groups found that many housekeeping genes considered to be stably expressed show variation in distinct plant tissues and species and thus it is highly recommended to systematically validate reference genes based on the actual experimental conditions under investigation [23–26]. This chapter is devoted to the mirEX platform that was developed to be a comprehensive starting point for a comparative investigation of miRNA gene expression [9, 27]. We will present how to prepare RNA and cDNA as well as how much of cDNA is required to run this high-throughput qPCR platform using primers amplifying all pri-miRNAs in the same temperature profile. We will also explain how to explore the mirEX database to get knowledge about miRNA expression and its regulation in Arabidopsis and barley.

2

Materials The protocol is optimized for Arabidopsis thaliana (Col-0), its mutants (e.g., dcl1-7, see Note 1) [28] and barley (spring cultivar Rolap) plants [29].

2.1 Plant Growth and Harvesting (Arabidopsis thaliana)

1. Arabidopsis thaliana (L.) Heynh., Columbia-0 seeds. 2. Seed sterilization solution: 5.25% sodium hypochlorite, 3% hydrochloric acid, MilliQ water up to 100 mL. 3. ½ MS solid medium (1 L): 2.2% (w/v) Murashige & Skoog medium concentrate, 0.5% (w/v) sucrose, 0.8% (w/v) plant culture agar, MilliQ water up to 1000 mL. Adjust pH 5.5–5.6 using 0.1 M KOH before adding agar. Autoclave (121  C, 1 atm., 20 min).

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4. Jiffy-7 42 mm pots soil (Jiffy International AS, Norway). 5. VEFI Trays 36  22 cm (800130). 6. SANYO (MLR-350H) growth chamber. 2.2 Plant Growth and Harvesting (Hordeum vulgare)

1. Hordeum vulgare cv. Rolap kernels. 2. Sterilizing agent T75DS/WS (Organika-Azot, Jaworzno). 3. 5 L pots containing field soil (pH 6.0) mixed with sand (7:2). 4. Multinutrient fertilizer—supplement soil before kernel sowing. For 5 L of soil (1 pot—5.5 kg of soil) prepare 200 mL of fertilize containing: NH4NO3—2.11 g, KH2PO4—2.51 g, K2SO4—0.61 g, MgSO4·7H2O—3.78 g, 10 mM H3BO3— 4.5 mL, 10 mM Cu2SO4—0.7 mL, 10 mM MnSO4·H2O— 0.33 mL, 10 mM FeCl3—10.33 mL. 5. Nitrogen fertilizer (used for tillering plants)—for 1 pot solution containing 2.11 g of NH4NO3. 6. Conviron environmental chamber (Conviron, Winnipeg, Manitoba, Canada).

2.3 RNA Isolation and cDNA Preparation

1. RNA isolation reagent: 0.8 M guanidine thiocyanate, 0.4 M ammonium thiocyanate, 0.1 M sodium acetate pH 5.0, 5% (v/v) glycerol, 38% (v/v) saturated acidic phenol, 5 mM EDTA, 0.5% (w/v) sodium lauroylsarcosine, and diethylpyrocarbonate-treated water (0.1% DEPC). 2. Direct-zol™ RNA MiniPrep (Zymo Research). 3. RNA loading buffer (2): 10 mM Tris HCl pH 7.5, 2.5 mM EDTA, 90% (v/v) formamide, 0.01% (w/v) xylencyanol, 0.01% (w/v) bromophenol blue. Autoclave (121  C, 1 atm., 20 min). 4. TURBO DNA-free™ Kit (Thermo Fisher Scientific). 5. Oligo-dT(18) primer (Thermo Fisher Scientific). 6. dNTPs’ Mix (10 mM each). 7. RNasin® Plus RNase Inhibitor (Promega). 8. SuperScript III™ Reverse Transcriptase (Thermo Fisher Scientific). 9. Power SYBR™ Green PCR Master Mix (Thermo Fisher Scientific). 10. Primers for cDNA quality check (0.5 μM each), see Table 1. 11. PCR machine ProFlex PCR System (Applied Biosystems). 12. QuantStudio 7 Flex Real-Time PCR System (Applied Biosystems).

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Table 1 Primers for cDNA quality check (0.5 μM each) Gene ID

Primer sequence 50 –30

Notes

GAPDH

At1g13440

Fw: TCTCGATCTCAATTTCGCAAAA Rev: CGAAACCGTTGATTCCGATTC

50 region of the cDNA

GAPDH

At1g13440

Fw: TTGGTGACAACAGGTCAAGCA Rev: AAACTTGTCGCTCAATGCAATC

30 region of the cDNA

ARF1

AJ508228.2

Fw: CACGACAGCCATCTTCATCT Rev: GCGTGCTCCGCCTGATCT

50 region of the cDNA

ARF1

AJ508228.2

Fw: CGTATGTGACAGGCAGAAG Rev: TGTAAAACCACGGCACAGAA

30 region of the cDNA

Gene Arabidopsis

Barley

2.4 HighThroughput qPCR

1. Power SYBR™ Green PCR Master Mix (Thermo Fisher Scientific). 2. mirEX pri-miRNA primers (http://www.combio.pl/mirex2. download/primers/), primers for quality control genes (see Table 1). 3. QuantStudio 7 Flex Real-Time PCR System (Applied Biosystems).

2.5 Control of qPCR Quality

1. LinRegPCR software.

2.6 Calculations and Data Analysis

1. QuantStudio Real-Time PCR software (Applied Biosystems).

2.7 Web-Based Protocols

3

2. Microsoft Excel.

2. Microsoft Office Excel. The only requirement needed for exploring mirEX is a computer with Internet connection and a standard web browser (e.g., Mozilla Firefox, Safari, Google Chrome, Internet Explorer).

Methods

3.1 Plant Growth and Harvesting (Arabidopsis thaliana)

We recommend to perform experiment in three independent biological replicates for each selected developmental stage [30] or plant organ. In the case of wild-type and various mutants, plants should be grown side by side. Each biological replicate should be sown at least 1 week after the previous one. For studies on young seedlings (1.02 or 1.03 stage) we recommend to grow plants in vitro, on a ½ MS solid medium. To study pri-miRNA expression in further stages of Arabidopsis development or plant organs, we recommend to grow plants in soil.

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1. For in vitro culture, sterilize A. thaliana seeds (approximately 100 μL of each genotype). Incubate seeds for 6 h in fumes of hydrochloric acid solution (see Subheading 2.1, item 2). Open tubes (2 mL) containing seeds, place together with a solutioncontaining beaker, in an exicator and tightly covered with a grease-sealed lid. Sow A. thaliana seeds immediately after sterilization. Do not store sterilized seeds. 2. Sow sterilized seeds on plates with a ½ MS solid medium (see Subheading 2.1.3) under sterile conditions. Close the plates using parafilm. 3. For “in soil culture,” sow seeds directly in soaked with water Jiffy-7 42 mm pots soil placed on trays. 4. Place plates/trays in the dark at 4  C for 48 h. Then, move the plates/trays into a SANYO growth chamber with a 16 h day (130–150 μM m2/s) and a day/night temperature of 22  C with 70% relative humidity. 5. For particular stage of development [30] collect plant tissue after the indicated number of days in Table 2. 6. For “in soil culture” water plants every 2 days with 0.2–0.5 L of tap water. Avoid soil desiccation as well as overwatering. 7. For each stage or plant organ collect tissue into aluminum foil envelopes and immediately freeze in liquid nitrogen. Store material at 80  C. 8. Transfer plant tissue into a mortar and grind using liquid nitrogen and pestle until you see a layer of fine dust. Store homogenized material at 80  C or use immediately.

Table 2 Arabidopsis developmental stages Stage

Approx. days of growth

Culture

Tissue

1.02

10

In vitro

Whole plant

1.03

14

In soil

Whole plant

1.05

14

In vitro

Whole plant

1.09

21

In soil

Aerial part

3.50

25

In soil

Leaves

5.10

35

In soil

Leaves

6.00

42

In soil

Leaves, stem

6.50

53

In soil

Leaves, stem, flowers, siliques

pri-microRNA Expression

3.2 Growth and Harvesting (Hordeum vulgare)

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We recommend to perform the experiment in three independent biological replicates. Each biological replicate should be sown at least 1 week after the previous one. 1. Mix barley kernels with T75DS/WS powder and sow directly in soil. 2. Grow barley plants in 5 L pots in a walk-in Conviron environmental chamber with a 16 h day/8 h night photoperiod, 800 μmol light conditions, and 22  C day/15  C night temperature. Maintain optimal growth conditions of 70% SWC (soil water content). 3. Collect barley plants in five major growth points according to Zadoks et al. cereal development decimal code [31]: 1-weekold plants (code 11, one leaf developed), 2-week-old plants (code 13, third leaf developed), 3-week-old plants (code 20–21, beginning of tillering), 6-week-old plants (code 32–36, stem elongation), and 68-day-old plants (code 75–77, kernels in milk ripeness). 4. Collect whole plants (roots and shoots) into aluminum foil envelopes and immediately freeze in liquid nitrogen. Store material at 80  C. 5. Transfer plant tissue into a mortar and grind using liquid nitrogen and pestle until you see a layer of fine dust. Store homogenized material at 80  C or use immediately.

3.3 RNA Isolation and cDNA Preparation

1. Into a cooled 2.0 mL RNase-free tubes (use liquid nitrogen for cooling) weight up to 100 mg of ground plant tissue. The given amount of plant tissue will suffice for cDNA preparation for running the whole mirEX platform for one biological replicate. 2. Add 1.0 mL of RNA isolation reagent (see Subheading 2.3, item 1) and homogenize mixture by vortexing thoroughly for 10–20 s (see Note 2). 3. Centrifuge samples at 14,000  g for 10 min at room temperature. Next, transfer the supernatant carefully into a new 2.0 mL RNase-free tubes. Do not disturb the pellet. 4. Repeat centrifugation of your supernatant again until you will not see any pellet. Usually, up to 1–2 more centrifugations are required. 5. Following last centrifugation, add 1.0 volume of chilled 95–100% ethanol to the supernatant. 6. Apply up to 800 μL of the mixture from point 6 to Zymo-Spin IIC Column placed in Collection Tube (provided) and centrifuge at 14,000  g for 1 min at room temperature. Discard flow-through and centrifuge residual volume.

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7. Transfer the column into a new collection tube (provided). 8. Add 400 μL of Direct-zol RNA PreWash directly to the column and centrifuge. Discard the flow-through and repeat this step. 9. Add 700 μL RNA Wash Buffer to the column and centrifuge for 2 min to ensure complete removal of the wash buffer. Transfer the column carefully into a new 1.5 mL RNase-free tube (not provided). 10. To elute RNA, add 50 μL of DNase/RNase-free water directly to the column matrix, incubate for 2 min at room temperature and centrifuge. Quantify each sample using Nanodrop Spectrophotometer. 11. For checking RNA quality, denature 0.5 μg of RNA sample in 1 RNA loading buffer for 5 min at 70  C and place RNA sample on ice for 2 min. Run samples on a 1.2% agarose gel for 30 min at 10 V per cm of your gel length. You can store RNA sample in 80  C freezer or proceed directly to the next step (see Note 3). 12. To remove DNA contamination from RNA sample use TURBO DNA-free™ Kit. After RNA quality control, take 30–50 μg of RNA and incubate with 0.1 volume 10 TURBO DNase Buffer and 1 μL of TURBO DNase (2 Units/μL) for 1 h at 37  C. A typical reaction volume is 50 μL. Next, add 0.1 volume freshly agitated DNase inactivation reagent, incubate 5 min at room temperature, vortex gently every 30 s. Centrifuge at 10,000  g for 1.5 min at 4  C and transfer supernatant containing RNA to a fresh RNase-free tube (do not disturb or aspirate the pellet). Determine the RNA concentration and quality as in the point 10 and 11, respectively (see Note 4). You can store RNA sample in 80  C freezer or proceed directly to the next step (see Note 3). At least 30 μg of RNA should be obtained when following this protocol. 13. Thaw the reagents (oligo-dT(18) primer, dNTPs’ mix, 5 RT Buffer, 0.1 M DTT) and total RNA samples on ice. 14. Mix and briefly spin each component and combine the following in a 0.2 mL RNase-free PCR tube (Table 3). 15. Prepare five identical reactions for each Arabidopsis RNA sample and four identical reactions for each barley RNA sample (see Note 5). 16. Incubate for 5 min at 65  C and put reaction on ice for 2 min. 17. Collect the content of the tube by brief centrifugation and add the rest of the RT mixture prepared as seen in Table 4. 18. Mix by pipetting gently up and down and incubate for 1 h at 50  C (Fig. 1).

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Table 3 Reverse Transcription (RT) reaction composition per one reaction (part I)

Component

Amount for single reaction (20 μL)

Total RNA

4.00 μga

10 mM dNTPs’ mix

1.00 μL

Oligo-dT(18) primer

1.00 μL

RNase-free water

Up to 13.00 μL∗

Partial volume

13.00 μL

a

Add appropriate volume according to RNA sample concentration Add appropriate volume according to RNA sample volume

∗

Table 4 Reverse Transcription reaction composition (part II)

Component

Volume for single reaction (20 μL)

5 RT buffer

4.00 μL

0.1 M DTT

1.00 μL

SuperScript III reverse transcriptase 200 U/μL

1.00 μL

RNase inhibitor 20 U/μL

1.00 μL

Partial volume

7.00 μL

Fig. 1 Temperature profile of reverse transcription reaction. The temperature of incubation and time for each step are indicated above and below the profile line respectively. The 1 symbol indicates the steps in which the thermocycler is ready to immediately proceeds to required incubation temperature

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19. Inactivate the reaction by heating at 70  C for 15 min. 20. Collect the content of the tube by brief centrifugation and dilute four times with MilliQ water (to 80 μL final volume). 21. Combine diluted cDNA obtained from the same RNA sample into 1.5 mL tube (for Arabidopsis 5  80 μL ¼ 400 μL, for barley 4  80 μL ¼ 320 μL). 22. After cDNA synthesis proceed directly to qPCR or store cDNA at 20  C. 23. To test the cDNA quality by RT-qPCR perform independently quantitative analysis of glyceraldehyde-3-phosphate dehydrogenase mRNA 50 and 30 fragments (GAPDH ; At1g13440) level for Arabidopsis cDNA samples and 50 and 30 fragments level of ADP-ribosylation factor 1-like protein mRNA (ARF1, AJ508228.2) for barley cDNA samples, respectively. Use separate primers for the amplification of 50 and 30 GAPDH mRNA fragments. Set up the reaction Master Mix as seen in Table 5. 24. Start the RT-qPCR reaction program (Fig. 2).

Table 5 Real-time PCR reaction mixture for testing cDNA quality Component

Volume for single reaction (10 μL)

cDNA

1.00 μL

Power SYBR™ green PCR master mix

5.00 μL

Primer mix (0.5 μM each)

4.00 μL

Total volume

10.00 μL

Fig. 2 Real-time PCR profile. The temperature of incubation and time for each step are indicated above and below the profile line, respectively. Camera sign indicates for fluorescence data acquisition after each cycle

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Fig. 3 Preparation of diluted primers for real-time PCR. The volume and concentration of primers for each step of dilution are indicated

25. Calculate the Ct difference for 50 and 30 GAPDH or ARF1 mRNA fragments. It should be lower than 1, see Note 6 [23]. 3.4 HighThroughput qPCR

Real-time PCRs were performed using the QuantStudio 7 Flex Real-Time PCR System (Applied Biosystems) and SYBR Green to monitor dsDNA synthesis. 1. Prepare 96-well plate with 5 μM (F+R) primers (see Note 7 and Fig. 3 below). (a) Pipette 10 μL of 100 μM Forward (F) and 10 μL of 100 μM Reverse (R) primers for one particular gene into one well. Add water to final volume of 200 μL. (b) Repeat this step for all pairs of primers. (c) For a reference gene, pipette the same primer mix to three separate wells. 2. Prepare 96-well plate with 0.5 μM (F+R) primers. (a) Pipette 20 μL of 5 μM primer mix to a new plate. (b) Add water to final volume of 200 μL. (c) For a reference gene, pipette the same primer mix to three separate wells. Two-step dilution procedure (Subheadings 3.4, steps 1 and 2) assures more precise primer dosage in the experiment.

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Fig. 4 Experimental plates layout for running high-throughput qPCR platform for Arabidopsis pri-miRNAs. The position of the primers on reaction plate is indicated Table 6 Master Mix composition Component

Volume for single reaction (6 μL)

cDNA

1.00 μL

Power SYBR™ green PCR master mix

5.00 μL

3. Prepare 384-well reaction plates. Pipette 4 μL of 0.5 μM primer mix to a new 384-well plate (Fig. 4). 4. Set up the reaction Master Mix as seen in Table 6.

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5. Pipette 6 μL of Master Mix to each well with 4 μL (0.5 μM) primer mix on 384-well plate. 6. Start the RT-qPCR reaction program (the same as in Subheading 3.3, step 24 and Fig. 2). 3.5 Control of qPCR Quality

Export from the real-time PCR apparatus fluorescence data per cycle. Use this data in LingRegPCR software to calculate PCR efficiency for each reaction. For further analysis keep Ct values where coefficient of determination (R2) value is higher than 0.9 (see Note 8) [32].

3.6 Calculations and Data Analysis

Determine the relative expression level/fold change of pri-miRNAs in/between your probes (e.g., A. thaliana wt plants from two developmental stages and A. thaliana wt plants and its dcl1-7 mutant) (see Note 9). How to calculate relative expression level of pri-miRNAs? 1. Calculate the average Ct values for the reference genes for wt samples. 2. Substract mean Ct values for a reference gene from Ct values obtained for pri-miRNA (ΔCt). Perform subtractions separately for all reference genes used. 3. Calculate the relative expression level (REL) using 2ΔCt equation and log10 for REL. 4. Repeat steps 1–3 for all biological replicates. 5. An example of Subheadings 3.6, steps 1–4 calculations are given below. Example: Calculations for pri-miR319a/b/c REL in Arabidopsis thaliana (two developmental stages (1.02; 1.03), three biological replicates, actin gene as a reference gene), Table 7. How to calculate fold change in pri-miRNA expression between two samples? 6. Calculate the average Ct values for the reference genes for wt and mutant samples. 7. Substract mean Ct values for a reference gene from Ct values obtained for pri-miRNA (ΔCt). Perform subtractions separately for all reference genes used. Calculate mean ΔCt values from biological replicates. 8. Subtract mutantΔCt value from wt ΔCt value (ΔΔCt). 9. Use 2ΔΔCt to calculate the fold change. 10. Repeat steps 6–8 for all biological replicates and reference genes. 11. An example of Subheading 3.6, steps 6–10 calculations are given below.

319a 319b 319c

1.03

25.342 22.198 22.818

Ct

Ct

R2 Ct

2nd 1st biol biol

ΔCt

2nd 3rd biol 1st biol biol

avg Ct (reference gene actin)

1.000 27.171 0.999 26.702 12.860 13.248 11.369 12.482 13.923 1.000 26.531 0.999 24.608 9.337 13.283 1.000 25.242 0.996 23.816 9.958 11.993

R2

3rd biol

1.000 27.810 0.997 32.096 1.000 30.663 13.572 18.111 17.789 14.238 13.986 1.000 26.774 0.991 30.285 1.000 29.332 13.203 12.174 0.621 13.898∗ 0.999 28.210 0.998 28.902 – 10.099

0.999 1.000 0.999

R2

2nd biol

avg ΔCt

12.873 13.699 11.542 12.306 11.112 10.606

15.332 13.912 13.238 11.953 12.447 11.466

3rd biol

0.0000751887 0.0001974306 0.0006417210

0.0000648518 0.0002522230 0.0003534876

relative expression level 2ΔCt

4.12 3.70 3.19

4.19 3.60 3.45

Log10 (REL)

Please notice that the cross through Ct value of pri-miR319c for the 1st biological replicate, due to law R2 value from LinRegPCR program, was omitted from further analysis.

∗

319a 319b 319c

primiRNA

1.02

developmental stage

1st biol

Table 7 An example of RT-qPCR results data analysis (developmental stages)

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Table 8 An example of RT-qPCR results data analysis (mutant vs. control plants) ΔCt wt pri1st miRNA biol

ΔCt dcl1–7 2nd biol

3rd biol

1st mean biol

2nd biol

3rd biol

mean

ΔΔCt

Fold change (2ΔΔCt)

319a

15.445 13.169 11.94 13.52 12.546 12.821 13.538 12.968 0.549

1.463

319b

12.508 10.238 10.27 11.00

9.494 1.511

2.849

319c

15.475 15.674 14.64 15.26 10.995 11.865 11.012 11.291 3.971 15.681

9.814

9.094

9.574

Example: Calculations for pri-miR319a/b/c in Arabidopsis thaliana (comparison between wt and dcl1-7 mutant, 3.50 developmental stage, actin gene as a reference gene), Table 8. 12. Apply Student’s t-test for statistical analyses. Use 2ΔCt (relative expression level) for this purpose (see Note 10). 3.7 mirEX: An OnLine Expression Atlas of Plant pri-miRNAs

mirEX (http://www.combio.pl/mirex) is a web application dedicated to comparative data mining of pri-miRNA expression information in three plants (A. thaliana, H. vulgare, P. endiviifolia (see Note 11). The methods presented in this chapter describe how to use the mirEX web interface to browse pri-miRNA expression levels through various developmental plant stages (see Subheading 3.7.1), comparing expression profiles in miRNA biogenesis mutants (see Subheading 3.7.2), or performing multi-species comparisons of pri-miRNA expression levels (see Subheading 3.7.3), and take the user through the information that is available for any individual pri-miRNA member or family (see Subheading 3.7.4).

3.7.1 Investigating primiRNAs Expression Across Various Developmental Stages

In this exercise, we will investigate the expression profile of pri-miRNA319 gene family across all tissues and developmental stages in A. thaliana. 1. Go to http://www.combio.pl/mirex. 2. Click “Browse” from the main menu located at the top of the page. 3. The graphical query builder allows for the selection of plant species of user’s interest as well as tissues and developmental stages (see Note 12). Select tissues from all the developmental stages in A. thaliana either by selecting every green square near the plant drawing or by clicking a “Select all” link located on the top-right side of the query builder. 4. Click the “next” button. 5. User can now select one or more of pri-miRNA genes of interest. This can be done in three ways: (1) by typing pri-miRNA names (e.g., ath-miR319a) in the input text box,

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Fig. 5 Expression profile of pri-miRNA319 family across 13 tissues/developmental stages selected in Arabidopsis thaliana. The chart is customizable by setting different gene as reference, showing the expression of mature miRNA in addition to pri-miRNA, zooming in/out by dragging out the rectangle in the chart

(2) by selecting individual or groups of pri-miRNAs from the tree-like expandable menu, or (3) by mixing both input methods. Choose the input method that suits you best and select all the ath-miR319 family members (i.e., ath-miR319a, ath-miR319b, and ath-miR319c). 6. Click the “next” button to go to the results. The resulting page of the ath-miR319 expression profile provides three types of visualized analyses, including interactive line graph (Fig. 5), searchable and sortable table and heatmap visualization. The latter two are described later in Subheadings 3.7.2 and 3.7.3, respectively. By default, the line graph shows expression levels as logarithm of REL of pri-miRNA transcript in respect to the reference actin gene. However, user can customize the graph by changing a reference gene expression to other than actin mRNA or by showing expression levels of mature miRNA. 7. In the default graph line, you can observe an extensive difference in expression levels among pri-miRNA319 members in rosette leaves at the flowering stage (6.50-rosette_leaf-soil). Use the select box “Reference gene” and set a different gene as a reference (e.g., elongation factor 1-alpha) and see how the line graph changes.

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8. Use select box “Molecule type” and switch the expression profile to mature miRNA. The accumulation of mature molecules is presented as RPM (reads per million) counts normalized to all miRNAs identified in the sample. You can observe increased accumulation of mature miR319 molecules in the inflorescence. 9. Display primary and mature miRNA on the same graph by selecting “primary + mature microRNA” option from the Molecule select box. The differences in transcript and mature miRNA accumulation can be the result of pri-miRNA transcription and maturation efficiency and/or miRNA stability within the RISC complex [3, 33]. 3.7.2 Comparing primiRNA Expression Levels in microRNA Biogenesis Mutants

mirEX provides measurements of pri-miRNA expression levels in four Arabidopsis mutants that represent the key components of the microRNA biogenesis pathway in the plants. In this exercise, we will investigate differences in expression levels of pri-miRNAs in dicer-like 1 mutant (dcl1-7) compared to control wild-type plant. 1. Click “Browse” from the main menu. 2. Click the “dcl” blue button, located at the bottom of 3.50 Arabidopsis stage. 3. The resulting page has four separate panels. First panel provides description of dcl1-7 mutant including effects on miRNA biogenesis and phenotype as well as list of relevant references. Second and third panels—“Mutant Chart” and “Mutant Table”—provide information about expression levels of 296 pri-miRNAs in mutant and control plants (the level of pri-miRNAs derived from three MIR genes was not detectable in the dcl1-7 mutant (MIR832, MIR845b, MIR5026)). Fourth panel shows pictures of dcl1-7 knock-downed plant compared to its control. Click on the video camera located at the top of the “Mutant Chart” panel and watch a short video showing how to customize the chart. 4. Expression values are listed in the searchable and sortable table (Fig. 6). Click the “Fold change” column to sort the table by increasing fold-change values (from 0.000567529 for MIR849 to 593.27 for MIR395d). Red and green arrows show the fold change values below 1 or above 1, respectively. You can see pri-miRNA names that have similar expression levels in mutant and control (e.g., MIR849, MIR776, MIR778). Click the “Fold change” column again. The fold change of pri-miRNA 395d is approximately 593, meaning that the level of this transcript in mutant is 593 higher than in the control wildtype plants. Use the “Search” text field located at the top of the table to find out which member of the MIR319 gene family has the greatest expression level in the mutant plant.

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Fig. 6 Expression profile of pri-miRNAs in the dcl1-7 mutant of Arabidopsis 3.50 stage (rosette in half of the final size) compared to wild-type control plant. Select box “Molecule” allows for displaying expression measurements (RPM) for mature miRNA molecules. The table provides search-based text filtering, column sorting, and pagination

5. In the select box “Molecule” set the mature miRNA molecule. Are there any miRNAs with high expression in mutant? What is the expression level of miR319a? 3.7.3 Inter-Species Comparison of pri-miRNA Expression Levels

In addition to intra-species comparisons of pri-miRNA/miRNA expression levels, mirEX also offers a heat map-based display of multiple miRNA entries across multi-tissue and multi-species comparisons. 1. Click “Browse” from the main menu. 2. Select all tissues from A. thaliana and H. vulgare and click the “next” button. 3. Using the expendable tree menu select the following pri-miRNAs: ath-miR159a, hvu-miR159a, all members of mir164, ath-miR167a, hvu-miR167a, all members of miR319 and miR399. 4. Click “next” to go to the resulting page (Fig. 7). 5. The level of gene expression in the heat map is color-coded: blue and red cells indicate low and high expression levels, respectively, and grey cells indicate low quality data (i.e., correlation coefficient of expression values is below 0.95).

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Fig. 7 Heat map-based display of comparison of pri-miRNAs level from Arabidopsis and barley in different developmental stages

6. Hover the mouse pointer over the MIR319b name to see a tooltip providing the list of selected tissues ordered by decreasing fold-change values. You can see that the molecule has the highest expression in stem from Arabidopsis. Click MIR319b to dynamically sort the matrix according to that tissue order. 7. Click the “ath-6.00-stem-soil” tissue name to sort pri-miRNA according to decreasing expression levels in that tissue. 8. Under the “Clustering” option, click “Dynamic” to find groups of pri-miRNAs with similar expression levels. In addition to the heap map you can also generate a dendrogram by clicking the “+Dendrogram” button. 3.7.4 Accessing Information Regarding Individual pri-miRNA

The pri-miRNA entry page is the central source of information regarding any given pri-miRNA/miRNA member as well as its family. Among the basic data characterizing each miRNA molecule (e.g., hairpin pre-miRNA structure, Northern hybridizations, or external database links), in this exercise we will cover the following features: (1) a graphical presentation of the expression levels in all of the tested growth stages and tissues in the form of pictographs,

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(2) expression profiles in mutant plants (where applicable), (3) mature miRNA NGS-based data with multiple sequence alignment, (4) the exon-intron structure of the pri-miRNA transcripts with options to highlight and download all of the presented features, and (5) automatic retrieval of the most recent articles from PubMed. Although pri-miRNA entry is accessible by clicking a miRNA gene name from most pages, the fastest and easiest way to navigate to specific pri-miRNA record is by using the search feature located on the right side of the main menu across every page. 1. Clicking on “Search” will open the simple search box where you can enter your search term. Type “319” and click “athMIR319b” from the auto-suggestion list. 2. The pictographic view maps the expression levels of ath-MIR319b into different parts and stages of Arabidopsis. For example, you can observe the highest expression level (as red color intensity) in stem and inflorescence. Switch the expression pictographs to mature miRNA to see the expression levels of mature miRNA. Which part of the plant has the highest abundance in miR319b? You can also switch the pictographic view between the two other MIR319 family members (MIR319a, MIR319c). 3. Scroll down the page to the “Mutant profile” panel. The column chart shows the expression levels of pri-miR319b across all seven miRNA biogenesis mutants. Which mutant does affect the expression of MIR319b the most? You can also switch the expression profile between other members of the MIR319 family using the “Family member” select box. 4. Scroll down to the “Pri-miRNA NGS reads coverage” panel showing the coverage pattern of sRNA fragments on the pri-miR319b transcript sequence (from 0 to 873 nt). The structural elements of gene (e.g., exon, pre-miRNA, mature miRNA) are highlighted as color-coded backgrounds. Mouseover the grey peak in the chart—you can see exact RPM values for each nucleotide position in pri-miRNA319b sequence. Use “Tissue” select box and switch the expression profile to 5.10rosette_leaf-soil. You can observe a clear change between the processing efficiency of miR319b/miR319b∗. 5. The next panel “NGS reads data table” shows the abundance of each short read (RPM) related to pri-miRNA319b across all Arabidopsis tissues. Use the “Show/hide table columns” to hide all tissue columns other than “6.50-inflorescence-soil” (leave columns from sequence to annotation). Sort the table according to the decreasing RPM in the “6.50” column. You can see the most abundant sequences in this tissue with the exact location in regard to pri-miRNA319b.

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6. The next panel “Pre-miRNA structure” maps the RPM values onto pre-miRNA 319b secondary structure. Switch the view to the “6.50-inflorescence-soil” tissue. You can see the mature miRNA highlighted in red on the structure. 7. The panel “miRNA gene structure” shows schematically all the gene features (e.g., exon, pre-miRNA, mature miRNA) with their positions in the full-length pri-miRNA sequence. For example, hovering your mouse pointer over the purple rectangle corresponding to pre-miRNA will highlight its start and end positions in pri-miRNA and clicking on the feature will highlight the corresponding sequence. In addition to the gene feature, mirEX also shows primers used to assess pri-miRNA expression level by RT–qPCR (see Note 13). 8. The last panel “Most recent articles from PubMed” shows a list of five most recent papers from PubMed concerning ath-MIR319b in title and/or abstract. Clicking on the “See all 5 articles in PubMed” link will direct you to the list of articles in PubMed. 3.7.5 Quiz Questions

If you get this far, try to answer the following questions: (a) What pri-miRNA has the highest expression level in seeds from Arabidopsis? (b) The expression of which member of the pri-miRNA172 family seems to have the most stable expression across all Arabidopsis tissues. (c) Display the expression levels of all pri-miRNA in all P. endiviifolia tissues. What pri-miRNA seems to be the most variable in terms of expression level? Check your answers in Note 14 (see Note 14).

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Notes 1. Arabidopsis dcl1-7 mutant carry point mutation (P415S) in the RNA-helicase domain (exon 3). The seeds obtained from heterozygous plants have to be sown plants but for the experiment, only homozygous plants should be selected. 2. To remove plant contaminants, polyphenolics and polysaccharides, it is recommended to add 70 μL Plant RNA Isolation Aid reagent (Invitrogen, Thermo Fisher Scientific) to the lysis buffer. Then vortex the tube and incubate in room temperature by 5 min, gently rotate the tube during this time. Proceed to the next step. 3. It is recommended to carry out the entire procedure in 1 day to avoid RNA freeze–thaw cycles.

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4. We highly recommend to check the DNA digestion efficiency by traditional PCR-based amplification using primers for 200–500 bp genomic fragment i.e., promoter region. Set the reaction program to at least 40–45 cycles. If you notice any band on the gel after electrophoresis, repeat the DNAse I treatment as described in the Methods Subheading 3.3, step 12. 5. Five reverse transcription reactions will allow to run mirEX platform for all Arabidopsis pri-miRNAs (299 pri-miRNAs), while four RT reactions will be sufficient for pri-miRNA detection from barley (140 pri-miRNAs). 6. The primer sequences for GAPDH cDNA are designed based on the exon-exon junction and they do not recognize splice isoforms in splicing mutants. Thus, it is recommended to test splicing efficiency of these pre-mRNAs in differently treated plants as well as in the case of mutant plants. 7. We recommend to use multichannel pipettes for preparation of primers plates. 8. LinRegPCR is a program for the analysis of qPCR data performed with SYBR™ Green or similar fluorescent dyes. The program after determination of a baseline of fluorescence and baseline subtraction creates a Window-of-Linearity for PCR efficiency calculation. We recommend to use the latest available software version [32]. 9. We recommend to input your data into Microsoft Excel (separate sheet for one reference gene) and conduct calculation for ΔCt, ΔΔCt, 2ΔCt, 2ΔΔCt using this software. 10. If you will use raw Ct values variance between the data from given two samples will be falsely underrepresented and this will affect statistical analysis [34]. 11. The data available in the mirEX cover the expression profiles for pri-miRNAs measured by RT-qPCR, and contain information about mature miRNA levels extracted from Next-Generation Sequencing (NGS) as well as Northern blot hybridization experiments. The data set of Arabidopsis thaliana miRNA sequences includes 299 pri-miRNAs representing 194 families. The expression data for microRNA primary transcripts from barley H. vulgare and liverwort P. endiviifolia contain 140 (57 families) and 22 (17 families) pri-miRNA sequences, respectively. 12. You can switch among three plants by using the grey-colored tabs located at the top of the graphical query builder. Each plant’s developmental stage is shown as a pictogram located in separate panel. Hovering the mouse cursor over the information icon provides short description of a given stage of the plant. Green query icons are present for every stage and tissue.

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13. Primers for all pri-miRNAs (299 in A. thaliana, 140 in H. vulgare, 22 in P. endiviifolia) used in mirEX are available for download at http://www.combio.pl/mirex2.download/ primers/. 14. Answers: (a) pr-miRNA5642a; (b) pri-miRNA172e; (c). pri-miRNA408b.

Acknowledgements The project is supported by the National Science Centre, Poland (NCN): UMO-2017/25/B/NZ1/00603, UMO-2016/23/D/ NZ1/00152, UMO-2016/21/B/NZ9/00550, UMO-2016/ 23/B/NZ9/00857, 2013/10/A/NZ1/00557, UMO-2016/ 23/B/NZ9/00862, by the Foundation for Polish Science (grant START 2017 to Agata S) and by KNOW RNA Research Centre in Poznan´ (No. 01/KNOW2/2014). References 1. Chen X (2009) Small RNAs and their roles in plant development. Annu Rev Cell Dev Biol 25:21–44. https://doi.org/10.1146/ annurev.cellbio.042308.113417 2. Sunkar R, Li YF, Jagadeeswaran G (2012) Functions of microRNAs in plant stress responses. Trends Plant Sci 17:196–203. https://doi.org/10.1016/j.tplants.2012.01. 010 3. Barciszewska-Pacak M, Milanowska K, Knop K, Bielewicz D, Nuc P, Plewka P, Pacak AM, Vazquez F, Karlowski W, Jarmolowski A, Szweykowska-Kulinska Z (2015) Arabidopsis microRNA expression regulation in a wide range of abiotic stress responses. Front Plant Sci 6:410. https://doi.org/10.3389/fpls. 2015.00410 4. Smoczynska A, Szweykowska-Kulinska Z (2017) MicroRNA-mediated regulation of flower development in grasses. Acta Biochim Pol 63:687–692. https://doi.org/10.18388/ abp.2016_1358 5. Yu Y, Jia T, Chen X (2017) The ‘how’ and ‘where’ of plant microRNAs. New Phytol 216:1002–1017. https://doi.org/10.1111/ nph.14834 6. Xie Z, Allen E, Fahlgren N, Calamar A, Givan SA, Carrington JC (2005) Expression of Arabidopsis MIRNA genes. Plant Physiol 138:2145–2154. https://doi.org/10.1104/ pp.105.062943 7. Park W, Li J, Song R, Messing J, Chen X (2002) CARPEL FACTORY, a dicer

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Chapter 6 A Revised Adaptation of the Smart-Seq2 Protocol for Single-Nematode RNA-Seq Dennis Chang, Lorrayne Serra, Dihong Lu, Ali Mortazavi, and Adler Dillman Abstract The advancement of transcriptomic studies in plant parasitic nematodes will greatly benefit from the development of single-nematode RNA-seq methods. Since many plant parasitic nematodes are obligate parasites, it is often difficult to efficiently obtain sufficient amounts of nematodes for transcriptomic studies. Here we have adapted SMART-Seq2 for single-nematode RNA-seq requiring only an individual nematode for a sample replicate. This protocol provides a detailed step-by-step procedure of the RNA-seq workflow starting from lysis of the nematode to quantification of transcripts using a user-friendly online platform. Key words RNA-seq, Nematode, Smart-seq2

1

Introduction Studying the transcriptomic changes in organisms is becoming much more widespread and seemingly the standard as the molecular window to view the biology of organisms. RNA-seq methods and technology have been continually developed and improved. Current RNA-seq workflows follow a few basic steps: (1) lysis of cells to access the RNA and often purification of the RNA, (2) cDNA synthesis using reverse transcriptase, typically a modified M-MLV (Moloney-Murine Leukemia Virus reverse transcriptase [1], (3) amplification of the cDNA by PCR, and (4) sequencing of the DNA. One of the more popular methods is known as the SMART-Seq (Switching Mechanism at the 50 end of the RNA transcript) [2, 3] with an updated iteration called SMART-Seq2 [4, 5]. M-MLV reverse transcriptase is known to add additional cytosines to the 30 end of the synthesized cDNA after transcribing the template RNA and SMART-Seq targets these additional

Dennis Chang and Lorrayne Serra contributed equally to this work. Hailing Jin and Isgouhi Kaloshian (eds.), RNA Abundance Analysis: Methods and Protocols, Methods in Molecular Biology, vol. 2170, https://doi.org/10.1007/978-1-0716-0743-5_6, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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nucleotides using TSO (Template Switching Oligonucleotides) that allow the reverse transcriptase to switch from the RNA template to the TSO template. This allows addition of specific sequences to the 30 end of cDNA resulting in simple primer design and subsequent PCR amplification with increased coverage of the 50 end of RNA transcripts [2, 6], i.e., better generation of fulllength transcripts. SMART-Seq2 has been optimized for single cell or low RNA input samples by modifications including (but not limited to) the addition Betaine and MgCl2 to increase reverse transcriptase processivity, LNA (locked nucleic acid) [7] TSO to increase thermal stability and annealing to the additional cytosines at the 30 end of the synthesized cDNA, and use of reagents that result in overall increased transcript coverage and cDNA yield [8]. Low input and single cell RNA-seq kits such as SMART-Seq v4 (Takara Bio USA, Inc) and NEBnext Single cell/Low input (New England Biolabs, Inc) are just two examples of commercially available kits that utilize low input RNA optimized SMART-Seq and they demonstrate the value that scientists find in studying low input RNA systems. As single cell and low input RNA-seq technology and methodology continues to advance nematologists can adapt and improve these methods, particularly for single-nematode RNA-seq (Fig. 1). Developing and advancing single-nematode RNA-seq methods would benefit the field of nematology (applicable to science in general) in three major ways: (1) it allows transcriptomic study of limited quantities of nematode samples, especially those from natural or agricultural environments; (2) it could obtain higher resolution transcriptomic data of developing nematodes by sampling individuals of different stages without the need to synchronize a population; and (3) it may better address research questions related to population/sample heterogeneity since individual nematodes are analyzed separately rather than being pooled. RNA input from free-living nematode species such as C. elegans is typically not a limiting factor because they can generally be reared and studied in relatively large numbers. Parasitic nematodes and difficult to culture non-model species, on the other hand, are often limited in the number of nematodes obtainable and therefore limited in the RNA starting material. Plant parasitic nematodes (PPNs) are obligate biotrophs and need to be cultured on plant hosts which presents the first hurdle for obtaining sufficient amounts of nematodes. RNA-seq studies of plant parasitic nematodes still generally require a relatively high degree of processing to isolate PPNs from plants and generally entails blending of plant tissues (particularly root tissues) and filtering through sieves/meshes (typically in series) [9–13]. These methods are currently the most efficient ways to obtain sufficient numbers of PPNs (or PPN eggs) to yield enough RNA material for traditional RNA-seq. Single-nematode RNA-seq requires minimal numbers of individuals and presents the

Fig. 1 SMART-Seq2 principles and workflow. (a) Cutting the nematode with a syringe needle and lysis of the tissue with lysis buffer and proteinase K to release mRNA. (b) The first cDNA strand is synthesized by binding of the Oligo-dT30-VN primer to the poly-A tail of mRNA. The reverse transcriptase will use the mRNA as the template until it reaches the 50 end of the mRNA where it will add additional cytosines to the new cDNA strand. The LNA-modified TSO (Locked nucleic acid, Template switching oligonucleotide) will then anneal to the overhanging cytosines and the reverse transcriptase will switch from mRNA to the new TSO as the template for reverse transcription. (c) cDNA is amplified by PCR. (d) Amplified DNA getting fragmented and tagmented by transposomes carrying the tag sequences (transposomes not shown). (e) Indexes are added to the DNA fragments via PCR to prepare the library for sequencing

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opportunity to significantly reduce the time and effort put into isolating nematodes from plant hosts. Single-nematode RNA-seq would also benefit studies of rare or difficult to access PPNs which could be due to various factors such as low population density, limited host range, or seasonal variation. A major aspect often studied in PPNs (and parasites in general) is gene expression throughout the different life stages. These life stages fulfill specific purposes for the nematodes and encompass different morphological and physiological changes [14, 15]. A single infested plant is expected to contain a population of PPNs with mixed life stages. So, extra care must be taken to assess and isolate the nematodes based on life stages. Processing a relatively large amount of nematodes can take more time and comes with the increased risk of contaminating nematodes of different life stages. Single-nematode RNA-seq requires very few individuals thereby reducing the risk of contamination during processing. Additionally, the higher resolution of transcriptomic data obtained from sequencing individual nematodes leads to more efficient discovery of important gene expression differences/similarities between the life stages. Even in studies where the life stage is not considered, other specific factors such as morphological, physiological, or behavioral changes of individuals can be more efficiently assessed and correlated with gene expression profiles of individual nematodes. Heterogenous populations of some PPN species can be studied by isolating nematodes from specific plant tissues or parts. In the case of migratory endoparasitic PPNs, which feed while traveling through host tissue causing damage along the way, many species exhibit different life stages that can often be associated with specific parts of the plant [16]. Since this undoubtedly result in low amounts of nematodes, traditional RNA-seq would require multiple plants to be processed taking up more time and effort. Singlenematode RNA-seq not only saves the time and effort but it can facilitate analysis of the heterogeneity of a population from a single host, therefore limiting the impact of host variance. Another potential benefit of single-nematode RNA-seq is in its application to the issue of how PPNs react to different plant hosts. Some PPNs have very narrow host ranges such as Ditylenchus africanus which primarily targets peanuts while Ditylenchus dipsaci is known to infect more than 500 different plant species [17]. PPNs that target multiple plant species could be expected to utilize different strategies of infection and regulate gene expression in a host-specific manner; however, this topic is still underexplored [18]. Understanding whether a species of PPN uses only a specific set of genes to infect plants or whether different genes are utilized

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based on the host could lead to improved targeting of essential infection genes controlling PPNs. In our own need to circumvent the issue of low RNA input due to the limited number of parasitic nematodes from in vivo insect infections, we have adapted SMART-Seq2 protocols [5, 19] into a streamlined protocol for single-nematode RNA-seq [20]. While the protocol was originally adapted for insect-parasitic nematodes [21] and their embryos [22], it could easily be adapted to other nematodes. The protocol in this book chapter is an updated adaptation of the aforementioned protocol [20] covering nematode lysis, reverse transcription of RNA, amplification of cDNA, DNA library labeling/preparation for sequencing, and a basic description of how to use Galaxy [23] to quantify transcript abundance from raw RNA-seq data with the Salmon program [24]. The user may choose any program/software they are familiar with to quantify transcript abundance; however, for researchers without any prior bioinformatic analysis experience we especially recommend Galaxy. It is a free web-based data analysis platform and its user-friendly interface is designed to make bioinformatic analysis more accessible to researchers without extensive experience in specialized software or programming. Salmon is a quick and accurate program for quantifying RNA transcript abundance and is available on the Galaxy platform. Note: Salmon quantification requires a reference transcriptome so use of Salmon is only applicable to organisms with an assembled transcriptome (either de novo assembly or predicted from a sequenced genome).

2 2.1

Materials Consumables

1. 0.2 ml thin-walled PCR tubes, nuclease free. 2. Syringe needles (needles should be less than 100 in length and any gauge between 25 and 31 G). 3. Pipette tips, nuclease free. 4. 1.5 ml microfuge tubes, nuclease free. 5. Nitrile or Latex gloves. 6. 70% ethanol in a spray bottle. 7. RNase decontaminating solution for wiping down surfaces and tools.

2.2

Reagents

1. Molecular grade water, nuclease free. 2. Lysis buffer stock [25]. 20 μl of 1 M Tris–HCl pH 8.0. 20 μl of 100% Triton X-100 (see Note 1). 200 μl of 10% Tween 20.

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2 μl of 0.5 M EDTA. 1.628 ml nuclease-free water. Total 1.871 ml. 3. Proteinase K (QUIAGEN, #19131). 4. RNasin ribonuclease inhibitor (Promega, #N2611). 5. dTNP (10 mM), (Thermo Fisher Scientific, #R0192). 6. Superscript II reverse transcriptase (Thermo Fisher Scientific, #18064014). 7. Betaine (BioUltra 99%, Sigma-Aldrich, #61962). 8. Magnesium chloride (Sigma-Aldrich, #M8266). 9. Phusion® High-Fidelity PCR Master Mix with HF Buffer (New England Biolabs, #M0531L). 10. Agencourt Ampure XP beads (Beckman Coulter, #A63881). 11. 80% ethanol in nuclease-free water. 12. Elution Buffer of choice (10 mM Tris–Cl pH 8.5). 13. Nextera DNA Flex Library Prep Kit (Illumina, Inc. #20018704). 2.3

Oligonucleotides

1. Olidgo-dT30VN primer (ordered from idtdna.com). 50 -AAGCAGTGGTATCAACGCAGAGTACT30VN0 3. Oligonucleotide primer for annealing to the poly(A)-tail of mRNAs. V can be either A, C, or G, N can be any base. The primers should be solubilized in TE buffer to 100 μM. Store in aliquots at 20  C for 6 months. 2. LNA-modified TSO (Locked Nucleic Acid-modified Template Switching Oligonucleotide) (ordered from exiqon.com). 50 -AAGCAGTGGTATCAACGCAGAGTACATrGrG +G-30 . The two rG (riboguanosines) and +G (LNA-modified guanosine) help to facilitate template switching. Store in TE buffer at 100 μM aliquots at 80  C for up to 6 months. Minimize repeated free-thaw cycles. 3. IS PCR primers (ordered from idtdna.com). 50 -AAGCAGTGGTATCAACGCAGAGT-30 . Solubilize in TE buffer and store in 100 μM aliquots at 20  C for up to 6 months. 4. Sequencing index primers [26]. See Table 1. Every sample will receive the Ad1_noMX primer while each sample receives a different Ad2.# primer. Primers should be stored in TE buffer in 25 μM aliquots at 20  C.

50 -CAAGCAGAAGACGGCATACGAGATTCGCCTTAGTCTCGTGGGCTCGGAGATGT-30 50 -CAAGCAGAAGACGGCATACGAGATCTAGTACGGTCTCGTGGGCTCGGAGATGT-30 50 -CAAGCAGAAGACGGCATACGAGATTTCTGCCTGTCTCGTGGGCTCGGAGATGT-30 50 -CAAGCAGAAGACGGCATACGAGATGCTCAGGAGTCTCGTGGGCTCGGAGATGT-30 50 -CAAGCAGAAGACGGCATACGAGATAGGAGTCCGTCTCGTGGGCTCGGAGATGT-30 50 -CAAGCAGAAGACGGCATACGAGATCATGCCTAGTCTCGTGGGCTCGGAGATGT-30 50 -CAAGCAGAAGACGGCATACGAGATGTAGAGAGGTCTCGTGGGCTCGGAGATGT-30 50 -CAAGCAGAAGACGGCATACGAGATCCTCTCTGGTCTCGTGGGCTCGGAGATGT-30 50 -CAAGCAGAAGACGGCATACGAGATAGCGTAGCGTCTCGTGGGCTCGGAGATGT-30 50 -CAAGCAGAAGACGGCATACGAGATCAGCCTCGGTCTCGTGGGCTCGGAGATGT-30 50 -CAAGCAGAAGACGGCATACGAGATTGCCTCTTGTCTCGTGGGCTCGGAGATGT-30 50 -CAAGCAGAAGACGGCATACGAGATTCCTCTACGTCTCGTGGGCTCGGAGATGT-30 50 -CAAGCAGAAGACGGCATACGAGATATCACGACGTCTCGTGGGCTCGGAGATGT-30 50 -CAAGCAGAAGACGGCATACGAGATACAGTGGTGTCTCGTGGGCTCGGAGATGT-30 50 -CAAGCAGAAGACGGCATACGAGATCAGATCCAGTCTCGTGGGCTCGGAGATGT-30 50 -CAAGCAGAAGACGGCATACGAGATACAAACGGGTCTCGTGGGCTCGGAGATGT-30 50 -CAAGCAGAAGACGGCATACGAGATACCCAGCAGTCTCGTGGGCTCGGAGATGT-30 50 -CAAGCAGAAGACGGCATACGAGATAACCCCTCGTCTCGTGGGCTCGGAGATGT-30 50 -CAAGCAGAAGACGGCATACGAGATCCCAACCTGTCTCGTGGGCTCGGAGATGT-30

Ad2.1_TAAGGCGA

Ad2.2_CGTACTAG

Ad2.3_AGGCAGAA

Ad2.4_TCCTGAGC

Ad2.5_GGACTCCT

Ad2.6_TAGGCATG

Ad2.7_CTCTCTAC

Ad2.8_CAGAGAGG

Ad2.9_GCTACGCT

Ad2.10_CGAGGCTG

Ad2.11_AAGAGGCA

Ad2.12_GTAGAGGA

Ad2.13_GTCGTGAT

Ad2.14_ACCACTGT

Ad2.15_TGGATCTG

Ad2.16_CCGTTTGT

Ad2.17_TGCTGGGT

Ad2.18_GAGGGGTT

Ad2.19_AGGTTGGG

(continued)

50 -AATGATACGGCGACCACCGAGATCTACACTCGTCGGCAGCGTCAGATGTG-30

Ad1_noMX

Table 1 Indexing primers

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50 -CAAGCAGAAGACGGCATACGAGATGAAACCCAGTCTCGTGGGCTCGGAGATGT-30 50 -CAAGCAGAAGACGGCATACGAGATTGTGACCAGTCTCGTGGGCTCGGAGATGT-30 50 -CAAGCAGAAGACGGCATACGAGATAGGGTCAAGTCTCGTGGGCTCGGAGATGT-30 50 -CAAGCAGAAGACGGCATACGAGATAGGAGTGGGTCTCGTGGGCTCGGAGATGT-30

Ad2.21_TGGGTTTC

Ad2.22_TGGTCACA

Ad2.23_TTGACCCT

Ad2.24_CCACTCCT

Primers to index label tagmented DNA fragments for library sequencing

50 -CAAGCAGAAGACGGCATACGAGATCACCACACGTCTCGTGGGCTCGGAGATGT-30

Ad2.20_GTGTGGTG

Table 1 (continued)

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Equipment

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1. Microcentrifuge. 2. Dissecting microscope. 3. Thermocycler. 4. Magnetic separation stand (for 1.5 ml centrifuge tubes). 5. Heat block. 6. Access to a DNA fluorometric quantification device (i.e., Qubit fluorometer, Thermo Fisher Scientific Inc.) and its associated reagents. 7. Access to use or submit samples to a BioAnalyzer (Agilent Technologies, Inc). 8. Access to use or submit samples to a Next-Gen nucleic acid sequencer.

3

Methods Clean all surface areas, pipettes, and equipment with 70% ethanol followed by an RNase decontaminant such as RNase away or RNaseZap. RNA can be extremely sensitive to degradation so after lysis of the nematode, do not let the mRNA samples simply sit on ice for more than an hour. Work hastily and always keep samples on ice. Be sure to change gloves frequently (see Note 2).

3.1 Isolation and Lysis of Nematodes

1. Prepare incomplete lysis buffer. 46.8 μl of lysis buffer stock. 3.2 μl of 20 mg/ml. 50 μl Total volume. 2. Prepare complete lysis buffer. 18 μl of incomplete lysis buffer. 2 μl of RNase inhibitor. 20 μl Total volume. 3. Add 2 μl of complete lysis buffer to the bottom of the 0.2 ml PCR tube(s). Briefly centrifuge to ensure the buffer is at the bottom of the tube and place the tubes on ice. 4. Wash the nematode three times in nuclease-free water. 5. Gently transfer the nematode in 2 μl of nuclease-free water to the wall of the PCR tubes. 6. Use the syringe needle to cut the nematode into 3–4 pieces while observing under a dissecting microscope (Fig. 2). Use a new needle for every nematode. 7. Quickly spin the nematode contents down into the lysis buffer at the bottom of tube and place it on ice.

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Fig. 2 Isolation and cutting of individual nematodes. (a) An example set up of a dissecting microscope station to cut nematodes. (b) Transferring of a single nematode to the wall of a PCR tube. (c) A nematode in 2 μl of nuclease-free water before cutting. (d) A nematode is cut with the tip of 29 G syringe needle. (Figure reproduced from Serra et al. [20])

8. Incubate the samples in the thermocycler to digest nematode tissues with proteinase K at 65  C for 10 min and inactivate the enzyme at 85  C for 1 min. Step

Temp.

Time

1

65  C

10 min

2

85  C

1 min

3



4 C

Continuous

9. Promptly remove the samples, briefly centrifuge, and place them back on ice (see Note 3).

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1. Add 1 μl of the oligo-dT VN primer (10 μM) and 1 μl of dNTP (10 mM) to the samples, mix by pipetting, briefly centrifuge, and place on ice. 2. Incubate the samples in a thermocycler at 72  C for 3 min and promptly place them back on ice. 3. Prepare the reverse transcription master mix.

Reagent

Volume per sample (μl)

Final concentration

Betaine (5 M)

2

1M

DTT (100 mM)

0.5

5 mM

MgCl2 (1 M)

0.06

6 mM

TSO (100 μM)

0.1

1 μM

Superscript II first-strand buffer (5)

2

1

SuperScript II reverse transcriptase (200 U/μl)

0.5

100 U

RNasin ribonuclease inhibitor (40 U/μl)

0.25

10 U

Nuclease-free water

0.29

–

Total volume

5.7

–

4. Add 5.7 μl of the reverse transcription master mix to each sample (now 10 μl total), mix by pipetting, briefly centrifuge, and place back on ice. 5. Run the first-strand synthesis reaction in the thermocycler with the program below: Step

Temp. 

Time

1

42 C

90 min

2

50  C 42  C

2 min 2 min

Go to step 2

14 70  C 

4 C

15 min Continuous

This is a good stopping point and the samples can be stored at 20  C. 3.3 PCR Amplification

1. Place/thaw the first-strand samples on ice and prepare the PCR amplification master mix following the recipe below:

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Reagent KAPA HiFi HotStart ReadyMix (2)

Volume per sample (μl)

Final Concentration

12.5

1

IS PCR primer (10 μM)

0.25

0.1 μM

Nuclease-free water

2.25

–

Total volume

15

2. Add 15 μl of the PCR amplification master mix to each firststrand synthesis sample (now 25 μl in total). 3. Run the samples with the PCR program below. Step 1 2

Temp. 

Time

98 C 98  C

3 min 20 s

67  C 72  C

15 s 6 min

Go to step 2

17

20

72  C

5 min

21

4 C

Continuous

This is a good stopping point and the samples can be stored at 20  C. 3.4 Cleanup of PCR Amplicons

1. Thaw the PCR samples and allow them to come to room temperature, approx. 10 min. 2. Vortex the Ampure XP beads to thoroughly mix the beads in solution. 3. Aliquot 26 μl of the beads into 1.5 ml microfuge tubes and allow them to warm up to room temperature (8 min). 4. Add the PCR sample (26 μl) to the beads at a 1:1 ratio. Mix thoroughly by pipetting up and down ten times. Do NOT discard the old PCR tubes yet. 5. Incubate the sample at room temperature for 8 min. 6. After incubation, place the samples on the magnetic separation stand for 5 min. Ensure the solution becomes clear. 7. Be careful not to disturb the magnetized bead pellet and keeping the tube on the magnetic stand, transfer the supernatant to its previously associated PCR tube (see Note 4).

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8. Quickly add 200 μl of 80% ethanol to the bead pellet and incubate for 30 s. (Do not mix). 9. Carefully remove the ethanol wash without disturbing the bead pellet. 10. Repeat steps 8 and 9 one more time. 11. Allow the beads to air-dry on the magnetic rack for ~5 min or until the first signs of minor cracks. Observe under a microscope frequently for minor cracks (see Note 5). 12. Quickly add 17.5 μl of EB to the bead pellets for each sample, pipette to mix, and place on a nonmagnetic tube rack (see Note 6). 13. Incubate the samples at room temperature for 2 min. 14. Place the samples on the magnetic stand for 2–3 min until the beads pellet and the solution becomes clear. 15. Collect 15 μl of the supernatant and place it on ice. If there are any visible remnants of the brown beads in the collected supernatant, transfer it back into the bead tube and repeat step 13. 3.5 Amplified cDNA Concentration and Quality Check (Not Detailed)

The user should choose an appropriate method of DNA quantification. Fluorometric quantification methods are more accurate then UV-based (Nanodrop) methods and we recommend using the Qubit Fluorometer and its associated reagents. The quality of the amplified DNA should be checked with the BioAnalyzer (Agilent Technologies). Figure 3 shows examples of BioAnalzyer profiles of a high-quality DNA sample (Fig. 3a) and a low-quality DNA sample (Fig. 3b). The distribution of the DNA lengths (correlating to mRNA transcript lengths) should show a strong group of peaks toward the 500–10,000 bp range for samples with minimal degradation (Fig. 3a). Profiles without peaks or many smaller fragmented peaks indicate mRNA transcript degradation and should not be used (Fig. 3b) (see Note 7).

Fig. 3 Example bioanalyzer profiles of amplified DNA quality. (a) Profile of a DNA sample made from RNA with minimal degradation. The distribution of the large peaks is around 1000 bp and peaks around 70 bp are not present, indicating no PCR primer contamination in the DNA. (b) Profile of a low-quality DNA sample made from RNA that has been significantly degraded showing fragmented and small peaks. (Figure reproduced from Serra et al. [20])

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3.6 Tagmentation of Amplified DNA

The user can use their own Tagmentation kit of choice so long as the proper protocol for tagmentation and post-tagmentation cleanup is followed. The protocol below details the use of the Nextera DNA Flex Library Prep kit (Illumina, Inc) and is adapted from its associated protocol. 1. Bring BLT and TB1 to room temperature (~8 min). 2. Set one heat block to 55  C and a second heat block to 37  C. 3. Transfer 20 ng of DNA samples into 1.5 ml tubes and bring the volume to 8 μl with nuclease-free water. 4. Vortex the BLT and TB1 for at least 10 s to thoroughly mix solutions. In a new PCR tube prepare the tagmentation master mix by combining 5.2 μl of BLT and 4.8 μl of TB1 per sample. Vortex the mix one more time. 5. Add 10 μl of the tagmentation master mix into the DNA and pipette up and down at least ten times. 6. Place sample for 15 min in the 55  C heating block to initiate tagmentation. Remove samples immediately after 15 min. 7. Bring TSB and TWB to room temperature (~8 min). If TSB contains visible precipitates, it can be heated at 37  C for 10 min followed with some vortexing. 8. Add 5 μl of TSB to each tagmented sample and slowly mix by pipetting up and down ten times. 9. Place sample for 15 min in the 37  C heating block to stop tagmentation. Remove samples immediately after 15 min. 10. Place samples on the magnetic rack for 2–3 min until solution is clear. 11. Carefully remove and discard the supernatant. 12. Remove the samples from the magnetic rack and wash the beads with 50 μl of TWB. Pipette up and down slowly to thoroughly mix the beads in solution. 13. Place the samples back on the magnetic rack and discard the solution. Repeat steps 9–13 a second time. 14. After removal of the TWB from the second wash, add 50 μl of TWB and pipette slowly to mix the beads. 15. Keep sample in a magnetic stand without removing TWB until Subheading 3.7, step 4.

3.7 Indexing Tagmented DNA by PCR Amplification

1. Thaw EPM on ice and invert to mix. Then briefly centrifuge and place it back on ice. 2. Thaw index primers at room temperature, mix by flicking or pipetting, briefly centrifuge, then place them back on ice. 3. Prepare the Indexing PCR master mix by mixing 5 μl EPM with 5 μl of nuclease-free water and 2.5 μl of Ad1_nMX. Vortex and briefly centrifuge the master mix.

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4. Keeping the tubes on the magnetic rack carefully remove and discard the 50 μl of TWB supernatant of the first sample. 5. Remove the first sample from the magnetic rack and resuspend beads in 12.5 μl of the Indexing PCR master mix. Mix by pipetting up and down ten times. Transfer the mix with beads to a PCR tube. Repeat this for each sample. 6. Add 2.5 μl of the Ad2.# to each sample (each sample receives a different Ad2.# primer, see Table 1). 7. Mix thoroughly and briefly centrifuge. 8. Run the indexing PCR with the following program: Step

Temp. 

Time

1

68 C

3 min

2 3

98  C 98  C 62  C 68  C

3 min 45 s 30 s 2 min

Go to step 3

9 

4

68 C

1 min

5

10  C

Continuous

9. After PCR, centrifuge the samples at 280  g for 1 min. ∗This is a good stopping point and the samples can be stored at 2–8  C up to 3 days 3.8 DNA Library Clean Up

1. Bring Sample Purification Beads (SPB) and Resuspension Buffer (RSB) to room temperature (~8 min). Vortex SPB frequently while using it, otherwise, the beads will settle in the bottom of the tube and samples won’t be cleaned properly. 2. Transfer samples from PCR tubes to 1.5 ml microfuge tubes. 3. Place tubes in a magnetic stand and transfer 15 μl of supernatant to a new 1.5 ml microfuge tube. 4. Add 20 μl of nuclease-free water and 22.5 μl of SPB to microfuge tube containing supernatant and pipette mixture ten times. 5. Incubate samples at room temperature for 5 min. 6. While samples are incubating, vortex SPB and add 8 μl to a new 1.5 ml microfuge tube. 7. Place samples from step 5 on a magnetic stand and wait a 2–3 min for solution to turn clear. 8. Transfer all the supernatant to the 1.5 ml microfuge tube prepared in step 6, mix thoroughly.

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9. Incubate sample at room temperature for 5 min. 10. Place sample from step 9 in a magnetic stand and wait for solution to turn clear. 11. Without disturbing the beads, remove and discard the supernatant. 12. Quickly add 200 μl of 80% ethanol to the bead pellet (still on the magnetic stand) and incubate for 30 s (Do not mix). 13. Carefully remove the 80% ethanol wash without disturbing the bead pellet. 14. Repeat steps 12 and 13 two more times. 15. Allow beads to air-dry on the magnetic stand for 5 min. 16. Remove samples from magnetic stand and resuspend beads in 17.5 μl of RSB. 17. Incubate at room temperature for 2 min on a tube rack. 18. Place tubes back in the magnetic stand and wait until solution is clear. 19. Transfer 15 μl of supernatant containing the cDNA library to a new 1.5 μl microfuge tube. 3.9 DNA Library Concentration and Quality Check

The DNA library concentration should be measured as previously done, and the quality should be checked by BioAnalyzer. Figure 4 shows examples of a fully tagmented library (Fig. 4a) and partially tagmented library (Fig. 4b). Most of the DNA that has been tagmented (along with fragmentation) should be around 200 bp long and that should be indicated by a central peak around 200 bp in the profile. A DNA library that has many partially or un-tagmented DNA will result in varying sizes of DNA and is demonstrated by a broad peak or sometimes multiple smaller peaks across from 150 to 1000 bp.

3.10 Sequencing of the DNA Library (Not Detailed)

For RNA-seq data, sequence coverage/depth for each sample should be at least 1  107 reads to reliably detect one transcript per million (TPM). Sequencing should be performed as pairedend, 43 bp reads (see Note 8). Sequencing with single-end reads is a viable alternative that is typically less expensive and quicker, however paired-end sequencing allows for better alignment of reads to the reference genome resulting in a higher quality data set.

3.11 Use Galaxy to Generate Relative mRNA Abundance Data from Raw Sequencing Data

Before beginning: Log onto usegalaxy.org and create an account (Fig. 5). Download your reference transcriptome in the fasta/fastq file format. Download your RNA-seq reads (raw sequencing files) in the fasta/fastq file format. If sequencing was performed as paired-end reads, you should have a “read 1” file and a “read 2” file for each sample.

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Fig. 4 Example BioAnalyzer profiles of DNA library quality. (a) Example profile of a tagmented DNA library with a central peak around 200 bp. (b) Example profile of DNA library with deficient fragmentation and tagmentation resulting in a broad distribution of fragmented peaks from 150–1000 bp. (Figure reproduced from Serra et al. [20])

1. On the Galaxy web-page at the left side under the “Tools” menu, click on “Get Data.” 2. In the new window, click “Upload File.” 3. Click “Choose local file” and choose your Transcriptome file, Read1 file, and Read2 file (the order does not matter). 4. Once your files are queued, you can change the file “Type” box to their appropriate file type or leave the selection at “Autodetect.” 5. Click “Start” and the files will begin to upload. The files will turn green when they are uploaded. 6. Close the upload window and you will see the files under the “History” column on right side of the web page. 7. On the left side under the “Tools” column, scroll down and click “RNA-seq,” then scroll down and click “Salmon.” 8. In the middle of the page under the menu option “Select a reference transcriptome. . .,” click on the entry box and set it to “Use one from the history.” 9. Under “Select reference transcriptome,” click the entry box and select your transcriptome file. 10. Under “Is this library mate-paired?”, set the entry box to “Paired-end” (or single-end instead, if that was part of the sequencing protocol). 11. Under “Mate pair 1,” select your RNA-seq read 1 file. 12. Under “Mate pair 2,” select your RNA-seq read 2 file. 13. The other data menu options such as “Relative orientation of reads within a pair” and “Specify the strandedness of the reads” should be set according to your sequencing and library preparation protocol.

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Fig. 5 Galaxy web-interface displaying RNA-seq analysis. Upload the transcriptome and FASTQ RNA-seq files under “Get data.” Then under “RNA-seq” run Salmon to quantify gene expression. The results can be downloaded from the “History” panel

14. The options further below are parameters for running Salmon and can be left as their default options or adjusted as needed. Most of the parameters are supplemented with a description of their functions and effect on quantification. More detailed descriptions can be found at https://salmon.readthedocs.io/

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en/latest/ and guidance can be found at https://combine-lab. github.io/salmon/faq/ 15. Important parameters such as “Perform sequence-specific bias correction” and “Perform fragment GC bias correction” should typically be set to “Yes.” 16. Once all parameters are set, click “Execute.” The analysis will be displayed in yellow under the “History” panel to the right of the web page and will turn green when completed. 17. After the analysis is completed, click on the file name and select the small floppy disk/save icon to download the file as a tab-delimited text file. The information in the file should be organized into columns: Transcript ID/Name, Transcript Length, Transcripts per Million (TPM), and Estimated number of reads. The TPM information can then be used for further analysis such as differential expression analysis.

4

Notes 1. The final concentration of Triton X-100 in the lysis buffer is 1%, however for nematode embryos we have also used a final concentration of 0.3% Triton X-100. 2. This protocol was optimized for handling a few samples at a time using individual PCR tubes and 1.5 ml microfuge tubes however, the protocol can be re-optimized for a higher volume of samples using multi-well plates (Thermofisher Scientific, #AB-0859) with magnetic plates (Thermofisher Scientific, # AM10027). 3. DNase can be used prior to reverse transcription (Subheading 3.1, step 9) to remove the genomic DNA. However, it is not required and we have not used DNase in our preparations. 4. Save the supernatant in case of insufficient DNA binding to the beads and repeating incubation of the supernatant with the beads if needed. 5. Do not over dry the beads as this will reduce DNA elution. The beads can sometimes dry in under 5 min, so we recommend frequently checking the bead pellet of each sample under the microscope and placing them back on the magnetic rack. At the very first sign of minor cracking, the bead pellet is dry enough. Other protocols do not recommend drying to the point of cracking however we found this degree of drying did not affect our DNA yield. 6. Pipette the EB solution up and down on the bead pellet until it breaks and dissolves. To minimize over drying of the beads we

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recommend quick initial breaking of the pellet with the EB solution for each sample before returning to thoroughly mix and dissolve the beads for each sample. Ensure the solution is homogenously brown. 7. In our hands, 8/10 of the DNA samples typically pass the BioAnalyzer quality check. 8. Regarding sequencing, samples with less than 1  106 reads are poor quality sequences and should not be used.

Acknowledgments This work is supported in part by NIH NIAID R21 AI142121 to A. Dillman. References 1. Gerard GF, Fox DK, Nathan M, Dalessio JM (1997) Reverse transcriptase—the use of cloned moloney murine leukemia virus reverse transcriptase to synthesize DNA from RNA. Mol Biotechnol 8:61–77. https://doi.org/10. 1007/bf02762340 2. Zhu YY, Machleder EM, Chenchik A, Li R, Siebert PD (2001) Reverse transcriptase template switching: a SMART (TM) approach for full-length cDNA library construction. BioTechniques 30:892–897. https://doi.org/10. 2144/01304pf02 3. Chenchik A, Diachenko L, Moqadam F, Tarabykin V, Lukyanov S, Siebert PD (1996) Full-length cDNA cloning and determination of mRNA 50 and 30 ends by amplification of adaptor-ligated cDNA. BioTechniques 21:526–534 4. Picelli S, Bjorklund AK, Faridani OR, Sagasser S, Winberg G, Sandberg R (2013) Smart-seq2 for sensitive full-length transcriptome profiling in single cells. Nat Methods 10:1096–1098. https://doi.org/10.1038/ nmeth.2639 5. Picelli S, Faridani OR, Bjorklund AK, Winberg G, Sagasser S, Sandberg R (2014) Full-length RNA-seq from single cells using Smart-seq2. Nat Protoc 9:171–181. https:// doi.org/10.1038/nprot.2014.006 6. Kulpa D, Topping R, Telesnitsky A (1997) Determination of the site of first strand transfer during Moloney murine leukemia virus reverse transcription and identification of strand transfer-associated reverse transcriptase errors. EMBO J 16:856–865. https://doi.org/10. 1093/emboj/16.4.856

7. Petersen M, Wengel J (2003) LNA: a versatile tool for therapeutics and genomics. Trends Biotechnol 21:74–81. https://doi.org/10. 1016/s0167-7799(02)00038-0 8. Picelli S (2017) Single-cell RNA-sequencing: the future of genome biology is now. RNA Biol 14:637–650. https://doi.org/10.1080/ 15476286.2016.1201618 9. Gardner M, Dhroso A, Johnson N, Davis EL, Baum TJ, Korkin D, Mitchum MG (2018) Novel global effector mining from the transcriptome of early life stages of the soybean cyst nematode Heterodera glycines. Sci Rep 8:15. https://doi.org/10.1038/s41598-01820536-5 10. Kumar M, Gantasala NP, Roychowdhury T, Thakur PK, Banakar P, Shukla RN, Jones MGK, Rao U (2014) De Novo transcriptome sequencing and analysis of the cereal cyst nematode, Heterodera avenae. PLoS One 9:16. https://doi.org/10.1371/journal.pone. 0096311 11. Cotton JA, Lilley CJ, Jones LM, Kikuchi T, Reid AJ, Thorpe P, Tsai IJ, Beasley H, Blok V, Cock PJA, Eves-van den Akker S, Holroyd N, Hunt M, Mantelin S, Naghra H, Pain A, Palomares-Rius JE, Zarowiecki M, Berriman M, Jones JT, Urwin PE (2014) The genome and life-stage specific transcriptomes of Globodera pallida elucidate key aspects of plant parasitism by a cyst nematode. Genome Biol 15:17. https://doi.org/10.1186/gb2014-15-3-r43 12. Eves-van den Akker S, Lilley CJ, Danchin EGJ, Rancurel C, Cock PJA, Urwin PE, Jones JT (2014) The transcriptome of Nacobbus

Smart-Seq2 for a Single Nematode aberrans reveals insights into the evolution of sedentary endoparasitism in plant-parasitic nematodes. Genome Biol Evol 6:2181–2194. https://doi.org/10.1093/gbe/evu171 13. Choi I, Subramanian P, Shim D, Oh BJ, Hahn BS (2017) RNA-Seq of plant-parasitic nematode Meloidogyne incognita at various stages of its development. Front Genet 8:3. https://doi. org/10.3389/fgene.2017.00190 14. Perry RN, Moens M (2011) Survival of parasitic nematodes outside the host. In: Perry RN, Wharton DA (eds) Molecular and physiological basis of nematode survival. Cabi Publishing-C a B Int, Wallingford, pp 1–27 15. Grencis R, Harnett W (2011) Survival of animal-parasitic nematodes inside the animal host. In: Perry RN, Wharton DA (eds) Molecular and physiological basis of nematode survival. Cabi Publishing-C a B Int, Wallingford, pp 66–85 16. Jones JT, Haegeman A, Danchin EGJ, Gaur HS, Helder J, Jones MGK, Kikuchi T, Manzanilla-Lopez R, Palomares-Rius JE, Wesemael WML, Perry RN (2013) Top 10 plant-parasitic nematodes in molecular plant pathology. Mol Plant Pathol 14:946–961. https://doi.org/10.1111/mpp. 12057 17. Moens M, Perry RN (2009) Migratory plant endoparasitic nematodes: a group rich in contrasts and divergence. Annu Rev Phytopathol 47:313–332. https://doi.org/10.1146/ annurev-phyto-080508-081846 18. Bell CA, Lilley CJ, McCarthy J, Atkinson HJ, Urwin PE (2019) Plant-parasitic nematodes respond to root exudate signals with hostspecific gene expression patterns. PLoS Pathog 15:19. https://doi.org/10.1371/journal. ppat.1007503 19. Trombetta JJ, Gennert D, Lu D, Satija R, Shalek AK, Regev A (2014) Preparation of singlecell RNA-Seq libraries for next generation sequencing. Curr Protoc Mol Biol 107:4.22.21–4.22.17. https://doi.org/10. 1002/0471142727.mb0422s107

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20. Serra L, Chang D, Macchietto M, Williams K, Murad R, Lu D, Dillman AR, Mortazavi A (2018) Adapting the Smart-seq2 protocol for robust single worm RNA-seq. Bio Protoc 8: e2729. https://doi.org/10.21769/ BioProtoc.2729 21. Lu DH, Macchietto M, Chang D, Barros MM, Baldwin J, Mortazavi A, Dillman AR (2017) Activated entomopathogenic nematode infective juveniles release lethal venom proteins. PLoS Pathog 13:31. https://doi.org/10. 1371/journal.ppat.1006302 22. Macchietto M, Angdembey D, Heidarpour N, Serra L, Rodriguez B, El-Ali N, Mortazavi A (2017) Comparative transcriptomics of Steinernema and Caenorhabditis single embryos reveals orthologous gene expression convergence during late embryogenesis. Genome Biol Evol 9:2681–2696. https://doi.org/10. 1093/gbe/evx195 23. Afgan E, Baker D, Batut B, van den Beek M, Bouvier D, Cech M, Chilton J, Clements D, Coraor N, Gruning BA, Guerler A, HillmanJackson J, Hiltemann S, Jalili V, Rasche H, Soranzo N, Goecks J, Taylor J, Nekrutenko A, Blankenberg D (2018) The galaxy platform for accessible, reproducible and collaborative biomedical analyses: 2018 update. Nucleic Acids Res 46:W537–W544. https://doi.org/10.1093/nar/gky379 24. Patro R, Duggal G, Love MI, Irizarry RA, Kingsford C (2016) Salmon provides accurate, fast, and bias-aware transcript expression estimates using dual-phase inference. bioRxiv:021592. https://doi.org/10.1101/ 021592 25. Shaham S (2006) Methods in cell biology. WormBook. https://doi.org/10.1895/ wormbook.1.49.1 26. Buenrostro JD, Wu B, Chang HY, Greenleaf WJ (2015) ATAC-seq: a method for assaying chromatin accessibility genome-wide. Curr Protoc Mol Biol 109:21.29.21–21.29.29. https://doi.org/10.1002/0471142727. mb2129s109

Chapter 7 Analysis of RBP Regulation and Co-regulation of mRNA 30 UTR Regions in a Luciferase Reporter System Erin L. Sternburg and Fedor V. Karginov Abstract The luciferase reporter assay is a widely used tool to study the cis and trans factors controlling regulation of gene expression. In this assay, regulatory elements can be fused to the luciferase gene, and as a result effect protein output by changing rates of transcription, rates of translation, or mRNA stability. This protocol focuses on probing the function of RNA-binding proteins (RBPs) through their interactions with the 30 untranslated region (UTR), thus examining gene expression regulation on the mRNA level. Assessment of 30 UTR sequence requirements, as well as single and co-regulatory roles of RBPs in regulation of mRNAs will be discussed. Key words RNA-binding proteins (RBP), 30 untranslated region (UTR), Reporter assays, Gene regulation

1

Introduction RNA-binding proteins (RBPs) regulate many aspects of cellular function through their interactions with mRNA transcripts, often through sequence-specific binding to 30 untranslated regions (UTRs), which can modulate mRNA stability and translation [1, 2]. RBPs can recognize binding motifs directly, or through association with small RNA partners [3], and can recognize primary and secondary sequence elements [4]. Luciferase assays provide a convenient, inexpensive, and quantitative method of measuring 30 UTR regulatory effects by controlling reporter gene expression with the regulatory region of interest [5–8]. The luciferase reporter is transiently expressed from a plasmid, which can be easily modified to test sites of interest. Once expressed, overall protein expression can be measured in a cell lysate by observing chemical luminescence produced by the luciferase protein in the presence of substrate. The luciferase assay consists of cloning a 30 UTR region of interest into the 30 UTR of a luciferase reporter gene. If the region confers regulatory activity when the reporter is introduced into

Hailing Jin and Isgouhi Kaloshian (eds.), RNA Abundance Analysis: Methods and Protocols, Methods in Molecular Biology, vol. 2170, https://doi.org/10.1007/978-1-0716-0743-5_7, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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cells, the overall luciferase expression will be affected. Often, a control co-expression of an orthogonal luciferase protein is used to normalize for transfection and other technical differences (dual luciferase assays). Activity of the luciferase protein under wildtype conditions can then be compared to conditions where alterations in 30 UTR sequence or protein availability have been made. Similarly, when probing the function of the miRNA machinery, manipulation of miRNA abundance/availability can also be performed. 30 UTR site selection can be designed to include full-length 30 UTRs or short regulatory regions of interest. For the latter, concatenation of several copies in the reporter can be used to increase the sensitivity of the assay. In this protocol, variations in luciferase assay design are discussed. Detailed instructions are provided for the generation and cloning of short concatenated regulatory regions into the 30 UTR of the Renilla luciferase gene located in the psiCheck2 dual luciferase plasmid. Use of wildtype and mutant sequences in combination with protein/miRNA level manipulation is used to determine regulation by single RBPs, as well as in the detection of co-regulatory interactions.

2

Materials

2.1 Generation of psiCheck2-BsmBI Plasmid

1. top and bottom strand oligonucleotides (top strand: TCGACACCAGAGACGGACGTCTCTGTTT, bottom strand: GGCCAAACAGAGACGTCCGTCTCTGGTG). 2. PsiCheck2 plasmid (Promega). 3. NotI restriction enzyme (New England Biolabs). 4. XhoI restriction enzyme (New England Biolabs). 5. Calf Intestinal Alkaline Phosphatase (New England Biolabs). 6. T4 Polynucleotide kinase. 7. ATP, 10 mM. 8. T4 ligase. 9. Competent E. coli cells. 10. LB media.

2.2 Monomer Generation

1. Synthetic oligonucleotides corresponding to your 30 UTR region of interest. 2. Primers with the following overhang sequences to introduce BsmBI sites and unique overhangs to each monomer: Monomer 1 forward primer: GCGTCTCTCACC NNNNNNNNNN. Monomer 2 forward ANNNNNNNNNN.

primer:

GCGTCTCTAAC

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Monomer 3 forward NNNNNNNNNN.

primer:

GCGTCTCTGGCT

Monomer 4 forward NNNNNNNNNN.

primer:

GCGTCTCTTCAG

Monomer 1 reverse NNNNNNNNNN.

primer:

GCGTCTCTTGTT

Monomer 2 reverse NNNNNNNNNN.

primer:

GCGTCTCTAGCC

Monomer 3 reverse NNNNNNNNNN.

primer:

GCGTCTCTCTGA

Monomer 4 reverse NNNNNNNNNN.

primer:

GCGTCTCTAAAC

Where NNNNNNNNNN corresponds to the 10 nucleotides flanking each end of your insert region. 3. KOD Hot Start DNA Polymerase. 2.3 Golden Gate Assembly into psiCheck2-BsmBI Plasmid

1. psiCheck2-BsmBI plasmid. 2. T7 ligase. 3. Tango buffer, 10 (NEB). 4. ATP, 10 mM. 5. DTT, 10 mM. 6. BsmBI. 7. PlasmidSafe exonuclease (recommended).

2.4 Dual Luciferase Assay

1. HEK293 cells, cultured at 37  C and 5% CO2, or other transfectable cells grown under recommended conditions. 2. TransIT-LT1 transfection reagent (Mirus). 3. Opti-MEM Reduced serum media. 4. Dual luciferase assay kit (Promega). 5. White 96-well plates. 6. Luminometer.

3

Methods

3.1 Site Selection for Testing

A few strategies can be employed when choosing 30 UTR sites or regions to test in a luciferase assay. If the study’s aim is to determine the regulatory role of a specific RNA-binding protein, its target sites can be identified using one or a combination of the following criteria:

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1. Crosslinking-Immunoprecipitation followed by sequencing (CLIP-seq) datasets can be used to determine sites of direct binding for a given RNA-binding protein. This is particularly useful in identifying the target genes and binding locations of RBPs with undefined or loosely defined sequence specificities. 2. If the RNA-binding protein of interest has a known sequence specificity, bioinformatic motif searches can identify potential target sites within 30 UTR regions. This criterion is also useful for defining targets of the miRNA machinery, since complementarity and conservation of the “seed” region is a reasonable predictor of regulation. Since occupancy of an RBP at a known motif can be affected by many cellular factors, this method is best used as an additional filter together with other criteria. 3. RNA-sequencing data and/or microarray data can be used to identify differentially expressed genes under conditions where levels of your RNA-binding protein of interest have been manipulated. Note that changes in gene expression can represent secondary effects of protein knockout, so this method is best used as an additional filter. It is also important to consider whether a full-length 30 UTR or a smaller segment of the 30 UTR will be incorporated. Using a fulllength 30 UTR will test a regulatory region or element in a more native context, since the recruitment of additional factors will be maintained. Using a full-length 30 UTR is also beneficial when the exact binding site of a protein within a 30 UTR is not known. However, using a full-length 30 UTR increases the likelihood that other elements may mask the tested regulatory site. Additionally, challenges can arise when cloning particularly long 30 UTRs (greater than 3 kb). Selection of a smaller region of a 30 UTR can be useful in dissecting the regulation of a single element, or of a small subset of elements. This design requires that the RBP binding site is known. When using small regions, it is also possible to concatenate multiple copies of the sequence, which can enhance the regulatory signal. This is particularly useful for regulatory regions that do not lead to large changes in downstream protein synthesis. In Subheadings 3.4–3.6, a protocol is outlined to clone a 4 concatenated sequence into the dual luciferase plasmid, psiCheck2 (see Note 1). 3.2 Mutant Site Design

Comparing luciferase reporter activity between constructs and/or conditions can be used to determine the strength of regulation of a 30 UTR or region of interest. Comparisons can include manipulation of protein levels, co-regulatory factor levels, or manipulation of the 30 UTR sequence. When comparing a native sequence to a manipulated sequence, the following strategies can be used:

Luciferase Reporters for 3’UTR Analysis

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Fig. 1 Schematic of luciferase reporter assay design. psiCheck2 plasmid with a wildtype site (green) is transfected in parallel with a mutant site plasmid (red) into HEK293 cells. In this example, the site has a stabilizing effect, since mutation of the site leads to a decrease in reporter signal as compared to the wildtype construct

1. To determine the regulatory effect of an RNA-binding protein binding to a specific site, precise mutations can be introduced within the region that the RBP is known/predicted to bind. This design requires that information about RBP binding is available, either through a predicted motif/miRNA seed or through CLIP-sequencing data. If only CLIP-binding data exists, mutation of the entire CLIP peak region by sequence shuffling can abolish any sequence specificity while maintaining overall nucleotide composition. 2. To determine the overall effect of the 30 UTR on gene expression, full 30 UTR constructs can be compared to an empty vector control, where the luciferase gene has a minimal 30 UTR. 3. To identify regions of a 30 UTR where regulation is important, full 30 UTR constructs can be compared to constructs where large sections of the 30 UTR have been deleted. This can be done in a tiled fashion to profile the length of the UTR. Mutant or empty vector constructs will be transfected in parallel to the wildtype construct to determine the regulatory effects. Comparing the wildtype construct to the mutant construct can determine whether the region is stabilizing, repressive, or has no effect (Fig. 1). If the region is stabilizing, then luciferase reporter activity should decrease for constructs where the region has been mutated or removed (wildtype to mutant ratio is >1). If the region is repressive, then the level of luciferase should increase under conditions where the UTR/region has been mutated or removed (wildtype to mutant ratio is 1) or repressive effect (