Plant Innate Immunity: Methods and Protocols [1st ed.] 978-1-4939-9457-1;978-1-4939-9458-8

Table of contents :
Front Matter ....Pages i-xiii
Using Bioinformatics and Molecular Biology to Streamline Construction of Effector Libraries for Phytopathogenic Pseudomonas syringae Strains (Jay Jayaraman, Morgan K. Halane, Sera Choi, Honour C. McCann, Kee Hoon Sohn)....Pages 1-12
Immunoprecipitation Under Non-Denaturing or Denaturing Conditions of Lysine-Acetylated Proteins Expressed in Planta (Maxime Escouboué, Laurent Deslandes)....Pages 13-21
Protein Acetylation in Pathogen Virulence and Host Defense: In Vitro Detection of Protein Acetylation by Radiolabeled Acetyl Coenzyme A (Karl J. Schreiber, Jennifer D. Lewis)....Pages 23-32
A Novel Assay Based on Confocal Microscopy to Test for Pathogen Silencing Suppressor Functions (Mustafa Adhab, James E. Schoelz)....Pages 33-42
Quantification of Extracellular ATP in Plant Suspension Cell Cultures (Sowmya R. Ramachandran, Sonika Kumar, Kiwamu Tanaka)....Pages 43-54
Measuring Pectin Properties to Track Cell Wall Alterations During Plant–Pathogen Interactions (Gerit Bethke, Jane Glazebrook)....Pages 55-60
Method to Study Dynamics of Membrane-Bound Plant Transcription Factors During Biotic Interactions in Tomato (Supriyo Chowdhury, Payel Bhattacharjee, Shrabani Basak, Shreya Chowdhury, Pallob Kundu)....Pages 61-68
Preparation of Plant Material for Analysis of Protein–Nucleic Acid Interactions by FRET-FLIM (Maxime Escouboué, Laurent Camborde, Alain Jauneau, Elodie Gaulin, Laurent Deslandes)....Pages 69-77
Analysis of DNA Methylation Profile in Plants by Chop-PCR (Pratiti Dasgupta, Shubho Chaudhuri)....Pages 79-90
Change in Nucleosome Dynamics During Stress Responses in Plants (Amit Paul, Shubho Chaudhuri)....Pages 91-100
Measuring Cell Ploidy Level in Arabidopsis thaliana by Flow Cytometry (Leiyun Yang, Zhixue Wang, Jian Hua)....Pages 101-106
A Method for Investigating the Pseudomonas syringae-Arabidopsis thaliana Pathosystem Under Various Light Environments (Daniel L. Leuchtman, Anthony D. Shumate, Walter Gassmann, Emmanuel Liscum)....Pages 107-113
Isolation and Characterization of Plant Metabolite Signals that Induce Type III Secretion by the Plant Pathogen Pseudomonas syringae (Conner J. Rogan, Jeffrey C. Anderson)....Pages 115-126
Quantification of Cauline Leaf Abscission in Response to Plant Pathogens (O. Rahul Patharkar)....Pages 127-139
Methods for Replicating Leaf Vibrations Induced by Insect Herbivores (Sabrina C. J. Michael, Heidi A. Appel, Reginald B. Cocroft)....Pages 141-157
Infection Assay for Xanthomonas campestris pv. campestris in Arabidopsis thaliana Mimicking Natural Entry via Hydathodes (Marieke van Hulten, Sayantani Chatterjee, Harrold A. van den Burg)....Pages 159-185
The Apple Fruitlet Model System for Fire Blight Disease (Sara M. Klee, Judith P. Sinn, Timothy W. McNellis)....Pages 187-198
Generating Transgenic Arabidopsis Plants for Functional Analysis of Pathogen Effectors and Corresponding R Proteins (Sharon Pike, Walter Gassmann, Jianbin Su)....Pages 199-206
Identification of Novel Pararetroviral Promoters for Designing Efficient Plant Gene Expression Systems (Ankita Shrestha, Ahamed Khan, Nrisingha Dey)....Pages 207-222
Biochemical and Molecular Characterization of Novel Pararetroviral Promoters in Plants (Ahamed Khan, Ankita Shrestha, Nrisingha Dey)....Pages 223-236
Sampling and Handling of Soil to Identify Microorganisms with Impacts on Plant Growth (Robert J. Kremer)....Pages 237-246
A New Strategy for the Selection of Epiphytic and Endophytic Bacteria for Enhanced Plant Performance (Eduardo Balsanelli, Vânia Carla Pankievicz, Valter Antonio Baura, Fábio de Oliveira Pedrosa, Emanuel Maltempi de Souza)....Pages 247-256
Back Matter ....Pages 257-260

Citation preview

Methods in Molecular Biology 1991

Walter Gassmann Editor

Plant Innate Immunity Methods and Protocols

METHODS

IN

MOLECULAR BIOLOGY

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

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

Plant Innate Immunity Methods and Protocols

Edited by

Walter Gassmann Division of Plant Sciences, Christopher S. Bond Life Sciences Center, and Interdisciplinary Plant Group, University of Missouri, Columbia, MO, USA

Editor Walter Gassmann Division of Plant Sciences, Christopher S. Bond Life Sciences Center, and Interdisciplinary Plant Group University of Missouri Columbia, MO, USA

ISSN 1064-3745 ISSN 1940-6029 (electronic) Methods in Molecular Biology ISBN 978-1-4939-9457-1 ISBN 978-1-4939-9458-8 (eBook) https://doi.org/10.1007/978-1-4939-9458-8 © Springer Science+Business Media, LLC, part of Springer Nature 2019 This work is subject to copyright. All rights are reserved by the Publisher, whether the whole or part of the material is concerned, specifically the rights of translation, reprinting, reuse of illustrations, recitation, broadcasting, reproduction on microfilms or in any other physical way, and transmission or information storage and retrieval, electronic adaptation, computer software, or by similar or dissimilar methodology now known or hereafter developed. The use of general descriptive names, registered names, trademarks, service marks, etc. in this publication does not imply, even in the absence of a specific statement, that such names are exempt from the relevant protective laws and regulations and therefore free for general use. The publisher, the authors, and the editors are safe to assume that the advice and information in this book are believed to be true and accurate at the date of publication. Neither the publisher nor the authors or the editors give a warranty, express or implied, with respect to the material contained herein or for any errors or omissions that may have been made. The publisher remains neutral with regard to jurisdictional claims in published maps and institutional affiliations. This Humana Press imprint is published by the registered company Springer Science+Business Media, LLC, part of Springer Nature. The registered company address is: 233 Spring Street, New York, NY 10013, U.S.A.

Preface The last decades have witnessed tremendous growth in our understanding of plant innate immune mechanisms. Building on solid foundations in plant genetics and molecular biology, the field has obtained an increasingly detailed description of processes with which plants recognize and respond to general classes of microbes and pests and to specific pathogendeployed molecules [1–5]. Much of this work utilized the power of model systems and interactions. What can get overlooked in these successes is that each model system provides both generally applicable, but also interaction-specific, information. At the same time, increasingly sophisticated techniques at all levels of biological inquiry provide fascinating insights into multifaceted aspects of plant-microbe interactions that ultimately are likely to touch on all fields of plant biology [6, 7]. Starting with the interaction of two genes [8], we now describe the manipulation of immune, hormonal, and developmental signaling pathways involving posttranslational protein modifications, transcription factors and chromatin states, organelles, secretion and protein trafficking by pathogens, and plant mechanisms to monitor these essential processes. This presents both an opportunity and a challenge when editing a book on methods used in the field of plant immunity. We start this book with chapters describing methods that characterize the basic framework of plant-microbe interactions. Whether a pathogen is successful in colonizing a given host is determined in large part by the pathogen’s tool kit of effectors that function to shut off immune responses and reprogram the host cell [1, 3, 9]. Chapter 1 deals with identification using bioinformatics of the complete effector suite of a new bacterial pathogen, with the causal agent of kiwi bacterial canker, Pseudomonas syringae pv. actinidiae, as an example. The next two chapters describe methods to ascertain a particular biochemical function of effectors, protein acetylation. Among the increasing ways that proteins are posttranslationally modified, acetylation is of growing interest in plant-pathogen interactions [10]. Chapter 2 focuses on identifying host targets of acetylating effectors by immunoprecipitation, while Chapter 3 describes an in vitro method to verify that a given protein is acetylated by a given effector. While the term "effector" is not commonly associated with viral pathogens, they too employ proteins to manipulate the host cell [11]. Chapter 4 addresses the shortcomings of current assays for identifying viral silencing suppressor proteins by introducing a silencing suppression assay with greatly increased sensitivity. The pathogen tool kit is only one side of the molecular dialog between pathogens and their hosts. Equally important in determining the final outcome are plant responses. While research used to focus mainly on the genetic determinants of resistance and susceptibility, there has been a renewed emphasis on physiological and cell biological plant responses that can now take advantage of well-defined plant-microbe interaction systems. Chapter 5 describes a method for quantifying ATP release from plant cells, a newly characterized plant response [12], while Chapter 6 deals with cell wall alterations that can occur either as a resistance mechanism or as a consequence of disease [13]. Moving from the cell surface into the interior, Chapter 7 describes a method to work with a class of membrane-tethered transcription factors that become activated when processed and released from the membrane [14]. Chapters 8–10 describe methods that take a closer look at parameters that ultimately have an impact on gene regulation [15], including protein–DNA interactions (Chapter 8), DNA methylation (Chapter 9), and chromatin states (Chapter 10). To round out this part of

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the book, Chapter 11 looks at the measurement of ploidy levels in plant nuclei, since endoreduplication has emerged as an important factor in host-microbe interactions [16]. As a field, the study of molecular plant-microbe interactions has made rapid progress by adopting currently well-characterized model interactions under narrow environmental conditions. With a wealth of current knowledge from these systems, it is possible to expand our understanding by branching out to additional parameters in plant-pathogen systems. Chapter 12 introduces a method to investigate plant pathogen responses under well-defined spectral light conditions [17], while Chapter 13 focuses on characterizing plant metabolites recently found to induce responses in bacterial leaf pathogens [18]. The next set of chapters explores plant responses that are not traditional mainstays of pathogen or pest studies, namely, leaf abscission [19], as a way to shed pathogens (Chapter 14) and leaf vibrations (Chapter 15) as a signal for attack by chewing insects [20]. Subsequent chapters describe methods for less well-studied pathogen systems, the Xanthomonas campestris pv. campestris-Arabidopsis interaction (Chapter 16), and the Erwinia amylovora-apple interaction (Chapter 17). While these pathogens have been studied for decades, the methods described in this book will hopefully contribute to the accelerated research into molecular mechanisms of disease and resistance [21, 22]. We round out this volume of Methods in Molecular Biology with several chapters describing basic technologies and tools in plant immunity research and how pathogens in turn can be used for biotechnological applications. A basic approach in plant molecular biology is the generation of transgenic plants. However, expression of deleterious genes such as pathogen effectors or plant resistance genes can lead to pleiotropic phenotypes that interfere with experimental analyses even when using inducible promoters. Chapter 18 describes a key approach to circumvent these problems through the facile generation of large numbers of transgenic lines from which appropriate transgenic events with the desired properties can be isolated. Chapters 19 and 20 describe a suite of methods for the development of promoters with novel properties using pararetroviral pathogens as raw material [23]. Finally, it is well-known that only a very small proportion of microbes is deleterious, and we have begun to appreciate that multicellular eukaryotic organisms depend on a beneficial microbiome to thrive [24]. The last two chapters therefore conclude this book by describing soil sampling techniques for isolating natural assemblies of microbes from well-defined sites (Chapter 21) and approaches to isolate beneficial plant-growth-promoting bacteria from the phytosphere (Chapter 22). In conclusion, with this volume, we endeavor to present the reader with useful methods that will allow an expansion of experimental inquiry to methods touching on new biological processes that perhaps are not traditionally associated with studies of plant immunity, and to non-model systems. We hope that practitioners in this ever-expanding field will feel inspired and that the chapters in this book will allow them to embark on new directions in their research. Columbia, MO, USA

Walter Gassmann

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References 1. Xin XF, He SY (2013) Pseudomonas syringae pv. tomato DC3000: a model pathogen for probing disease susceptibility and hormone signaling in plants. Annu Rev Phytopathol 51:473–98 2. Cook DE, Mesarich CH, Thomma BP (2015) Understanding plant immunity as a surveillance system to detect invasion. Annu Rev Phytopathol 53:541–63 3. Cui H, Tsuda K, Parker JE (2015) Effector-triggered immunity: from pathogen perception to robust defense. Annu Rev Plant Biol 66:487–511 4. Niks RE, Qi X, Marcel TC (2015) Quantitative resistance to biotrophic filamentous plant pathogens: concepts, misconceptions, and mechanisms. Annu Rev Phytopathol 53:445–70 5. Zipfel C, Oldroyd GE (2017) Plant signalling in symbiosis and immunity. Nature 543:328–36 6. Ning Y, Liu W, Wang GL (2017) Balancing immunity and yield in crop plants. Trends Plant Sci 22:1069–79 7. Su J, Spears BJ, Kim SH et al (2018) Constant vigilance: plant functions guarded by resistance proteins. Plant J 93:637–50 8. Flor HH (1971) Current status of the gene-for-gene concept. Annu Rev Phytopathol 9:275–96 9. Gassmann W, Bhattacharjee S (2012) Effector-triggered immunity signaling: from gene-for-gene pathways to protein-protein interaction networks. Mol Plant-Microbe Interact 25:862–8 10. Lewis JD, Lee A, Ma W et al (2011) The YopJ superfamily in plant-associated bacteria. Mol Plant Pathol 12:928–37 11. Leisner SM, Schoelz JE (2018) Joining the crowd: integrating plant virus proteins into the larger world of pathogen effectors. Annu Rev Phytopathol 56:89–110 12. Choi J, Tanaka K, Cao Y et al (2014) Identification of a plant receptor for extracellular ATP. Science 343:290–4 13. Bethke G, Thao A, Xiong G et al (2016) Pectin biosynthesis is critical for cell wall integrity and immunity in Arabidopsis thaliana. Plant Cell 28:537–56 14. Bhattacharjee P, Das R, Mandal A et al (2017) Functional characterization of tomato membranebound NAC transcription factors. Plant Mol Biol 93:511–32 15. Garner CM, Kim SH, Spears BJ et al (2016) Express yourself: transcriptional regulation of plant innate immunity. Semin Cell Dev Biol 56:150–62 16. Bao Z, Hua J (2015) Linking the cell cycle with innate immunity in Arabidopsis. Molecular Plant 8:980–2 17. Ballare´ CL (2014) Light regulation of plant defense. Annu Rev Plant Biol 65:335–63 18. Anderson JC, Wan Y, Kim YM et al (2014) Decreased abundance of type III secretion system-inducing signals in Arabidopsis mkp1 enhances resistance against Pseudomonas syringae. Proc Natl Acad Sci USA 111:6846–51 19. Patharkar OR, Gassmann W, Walker JC (2017) Leaf shedding as an anti-bacterial defense in Arabidopsis cauline leaves. PLoS Genet 13:e1007132 20. Appel HM, Cocroft RB (2014) Plants respond to leaf vibrations caused by insect herbivore chewing. Oecologia 175:1257–66 21. Vicente JG, Holub EB (2013) Xanthomonas campestris pv. campestris (cause of black rot of crucifers) in the genomic era is still a worldwide threat to brassica crops. Mol Plant Pathol 14:2–18 22. Klee SM, Mostafa I, Chen S et al (2018) An Erwinia amylovora yjeK mutant exhibits reduced virulence, increased chemical sensitivity and numerous environmentally dependent proteomic alterations. Mol Plant Pathol 19:1667–78 23. Khan A, Shrestha A, Bhuyan K et al (2018) Structural characterization of a novel full-length transcript promoter from Horseradish Latent Virus (HRLV) and its transcriptional regulation by multiple stress responsive transcription factors. Plant Mol Biol 96:179–96 24. Finkel OM, Castrillo G, Herrera Paredes S et al (2017) Understanding and exploiting plant beneficial microbes. Curr Opin Plant Biol 38:155–63

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

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1 Using Bioinformatics and Molecular Biology to Streamline Construction of Effector Libraries for Phytopathogenic Pseudomonas syringae Strains . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1 Jay Jayaraman, Morgan K. Halane, Sera Choi, Honour C. McCann, and Kee Hoon Sohn 2 Immunoprecipitation Under Non-Denaturing or Denaturing Conditions of Lysine-Acetylated Proteins Expressed in Planta . . . . . . . . . . . . . . . 13 Maxime Escouboue´ and Laurent Deslandes 3 Protein Acetylation in Pathogen Virulence and Host Defense: In Vitro Detection of Protein Acetylation by Radiolabeled Acetyl Coenzyme A . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 23 Karl J. Schreiber and Jennifer D. Lewis 4 A Novel Assay Based on Confocal Microscopy to Test for Pathogen Silencing Suppressor Functions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 33 Mustafa Adhab and James E. Schoelz 5 Quantification of Extracellular ATP in Plant Suspension Cell Cultures. . . . . . . . . 43 Sowmya R. Ramachandran, Sonika Kumar, and Kiwamu Tanaka 6 Measuring Pectin Properties to Track Cell Wall Alterations During Plant–Pathogen Interactions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 55 Gerit Bethke and Jane Glazebrook 7 Method to Study Dynamics of Membrane-Bound Plant Transcription Factors During Biotic Interactions in Tomato . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 61 Supriyo Chowdhury, Payel Bhattacharjee, Shrabani Basak, Shreya Chowdhury, and Pallob Kundu 8 Preparation of Plant Material for Analysis of Protein–Nucleic Acid Interactions by FRET-FLIM . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 69 Maxime Escouboue´, Laurent Camborde, Alain Jauneau, Elodie Gaulin, and Laurent Deslandes 9 Analysis of DNA Methylation Profile in Plants by Chop-PCR . . . . . . . . . . . . . . . . 79 Pratiti Dasgupta and Shubho Chaudhuri 10 Change in Nucleosome Dynamics During Stress Responses in Plants. . . . . . . . . . 91 Amit Paul and Shubho Chaudhuri 11 Measuring Cell Ploidy Level in Arabidopsis thaliana by Flow Cytometry . . . . . . 101 Leiyun Yang, Zhixue Wang, and Jian Hua 12 A Method for Investigating the Pseudomonas syringae-Arabidopsis thaliana Pathosystem Under Various Light Environments . . . . . . . . . . . . . . . . . . . 107 Daniel L. Leuchtman, Anthony D. Shumate, Walter Gassmann, and Emmanuel Liscum

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Isolation and Characterization of Plant Metabolite Signals that Induce Type III Secretion by the Plant Pathogen Pseudomonas syringae . . . . . . . Conner J. Rogan and Jeffrey C. Anderson Quantification of Cauline Leaf Abscission in Response to Plant Pathogens . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . O. Rahul Patharkar Methods for Replicating Leaf Vibrations Induced by Insect Herbivores . . . . . . . Sabrina C. J. Michael, Heidi A. Appel, and Reginald B. Cocroft Infection Assay for Xanthomonas campestris pv. campestris in Arabidopsis thaliana Mimicking Natural Entry via Hydathodes . . . . . . . . . . . . Marieke van Hulten, Sayantani Chatterjee, and Harrold A. van den Burg The Apple Fruitlet Model System for Fire Blight Disease . . . . . . . . . . . . . . . . . . . . Sara M. Klee, Judith P. Sinn, and Timothy W. McNellis Generating Transgenic Arabidopsis Plants for Functional Analysis of Pathogen Effectors and Corresponding R Proteins . . . . . . . . . . . . . . . . . . . . . . . Sharon Pike, Walter Gassmann, and Jianbin Su Identification of Novel Pararetroviral Promoters for Designing Efficient Plant Gene Expression Systems . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Ankita Shrestha, Ahamed Khan, and Nrisingha Dey Biochemical and Molecular Characterization of Novel Pararetroviral Promoters in Plants. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Ahamed Khan, Ankita Shrestha, and Nrisingha Dey Sampling and Handling of Soil to Identify Microorganisms with Impacts on Plant Growth . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Robert J. Kremer A New Strategy for the Selection of Epiphytic and Endophytic Bacteria for Enhanced Plant Performance . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Eduardo Balsanelli, Vaˆnia Carla Pankievicz, Valter Antonio Baura, Fa´bio de Oliveira Pedrosa, and Emanuel Maltempi de Souza

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

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17 18

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20

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127 141

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Contributors MUSTAFA ADHAB  Division of Plant Sciences, University of Missouri-Columbia, Columbia, MO, USA; Plant Protection Department, University of Baghdad, Baghdad, Iraq JEFFREY C. ANDERSON  Department of Botany and Plant Pathology, Oregon State University, Corvallis, OR, USA HEIDI A. APPEL  Department of Biology, University of Toledo, Toledo, OH, USA EDUARDO BALSANELLI  Department of Biochemistry and Molecular Biology, Federal University of Parana´, Curitiba, PR, Brazil SHRABANI BASAK  Division of Plant Biology, Bose Institute, Kolkata, West Bengal, India VALTER ANTONIO BAURA  Department of Biochemistry and Molecular Biology, Federal University of Parana´, Curitiba, PR, Brazil GERIT BETHKE  Department of Agronomy and Plant Genetics, University of Minnesota, Saint Paul, MN, USA PAYEL BHATTACHARJEE  Division of Plant Biology, Bose Institute, Kolkata, West Bengal, India LAURENT CAMBORDE  LIPM, Universite´ de Toulouse, INRA, CNRS, UPS, CastanetTolosan, France SAYANTANI CHATTERJEE  Molecular Plant Pathology, Swammerdam Institute for Life Sciences (SILS), University of Amsterdam, Amsterdam, The Netherlands SHUBHO CHAUDHURI  Division of Plant Biology, Bose Institute, Kolkata, India SERA CHOI  Department of Life Sciences, Pohang University of Science and Technology, Pohang, Republic of Korea SHREYA CHOWDHURY  Division of Plant Biology, Bose Institute, Kolkata, West Bengal, India SUPRIYO CHOWDHURY  Division of Plant Biology, Bose Institute, Kolkata, West Bengal, India REGINALD B. COCROFT  Division of Biological Sciences, University of Missouri, Columbia, MO, USA PRATITI DASGUPTA  Division of Plant Biology, Bose Institute, Kolkata, West Bengal, India FA´BIO DE OLIVEIRA PEDROSA  Department of Biochemistry and Molecular Biology, Federal University of Parana´, Curitiba, PR, Brazil EMANUEL MALTEMPI DE SOUZA  Department of Biochemistry and Molecular Biology, Federal University of Parana´, Curitiba, PR, Brazil LAURENT DESLANDES  LIPM, Universite´ de Toulouse, INRA, CNRS, UPS, Castanet-Tolosan, France NRISINGHA DEY  Division of Plant and Microbial Biotechnology, Institute of Life Sciences, Government of India, Bhubaneswar, Odisha, India MAXIME ESCOUBOUE´  LIPM, Universite´ de Toulouse, INRA, CNRS, UPS, Castanet-Tolosan, France WALTER GASSMANN  Division of Plant Sciences, Christopher S. Bond Life Sciences Center, and Interdisciplinary Plant Group, University of Missouri, Columbia, MO, USA ELODIE GAULIN  LIPM, Universite´ de Toulouse, INRA, CNRS, UPS, Castanet-Tolosan, France JANE GLAZEBROOK  Department of Plant and Microbial Biology, University of Minnesota, Saint Paul, MN, USA

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Contributors

MORGAN K. HALANE  Department of Life Sciences, Pohang University of Science and Technology, Pohang, Republic of Korea JIAN HUA  Plant Biology Section, School of Integrative Plant Science, Cornell University, Ithaca, NY, USA ALAIN JAUNEAU  LIPM, Universite´ de Toulouse, INRA, CNRS, UPS, Castanet-Tolosan, France JAY JAYARAMAN  Department of Life Sciences, Pohang University of Science and Technology, Pohang, Republic of Korea; The New Zealand Institute for Plant & Food Research Ltd., Auckland, New Zealand AHAMED KHAN  Division of Plant and Microbial Biotechnology, Institute of Life Sciences, Government of India, Bhubaneswar, Odisha, India SARA M. KLEE  Department of Plant Pathology & Environmental Microbiology, The Pennsylvania State University, University Park, PA, USA ROBERT J. KREMER  Division of Plant Sciences, University of Missouri, Columbia, MO, USA SONIKA KUMAR  Department of Plant Pathology, Washington State University, Pullman, WA, USA PALLOB KUNDU  Division of Plant Biology, Bose Institute, Kolkata, West Bengal, India DANIEL L. LEUCHTMAN  Bayer Crop Science, Chesterfield, MO, USA JENNIFER D. LEWIS  Department of Plant and Microbial Biology, University of California— Berkeley, Berkeley, CA, USA; Plant Gene Expression Center, United States Department of Agriculture, Albany, CA, USA EMMANUEL LISCUM  Division of Biological Sciences, Christopher S. Bond Life Sciences Center, and Interdisciplinary Plant Group, University of Missouri, Columbia, MO, USA HONOUR C. MCCANN  New Zealand Institute for Advanced Study, Massey University, Auckland, New Zealand TIMOTHY W. MCNELLIS  Department of Plant Pathology & Environmental Microbiology, The Pennsylvania State University, University Park, PA, USA SABRINA C. J. MICHAEL  Division of Biological Sciences, University of Missouri, Columbia, MO, USA VAˆNIA CARLA PANKIEVICZ  Department of Biochemistry and Molecular Biology, Federal University of Parana´, Curitiba, PR, Brazil O. RAHUL PATHARKAR  Division of Biological Sciences and Interdisciplinary Plant Group, University of Missouri, Columbia, MO, USA AMIT PAUL  Division of Plant Biology, Bose Institute, Kolkata, West Bengal, India SHARON PIKE  Division of Plant Sciences, Christopher S. Bond Life Sciences Center, and Interdisciplinary Plant Group, University of Missouri, Columbia, MO, USA SOWMYA R. RAMACHANDRAN  Department of Plant Pathology, Washington State University, Pullman, WA, USA CONNER J. ROGAN  Department of Botany and Plant Pathology, Oregon State University, Corvallis, OR, USA JAMES E. SCHOELZ  Division of Plant Sciences, University of Missouri-Columbia, Columbia, MO, USA KARL J. SCHREIBER  Department of Plant and Microbial Biology, University of California— Berkeley, Berkeley, CA, USA ANKITA SHRESTHA  Division of Plant and Microbial Biotechnology, Institute of Life Sciences, Government of India, Bhubaneswar, Odisha, India ANTHONY D. SHUMATE  Oregon Health and Science University, Portland, OR, USA

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xiii

JUDITH P. SINN  Department of Plant Pathology & Environmental Microbiology, The Pennsylvania State University, University Park, PA, USA KEE HOON SOHN  Department of Life Sciences, Pohang University of Science and Technology, Pohang, Republic of Korea; School of Interdisciplinary Bioscience and Bioengineering, Pohang University of Science and Technology, Pohang, Republic of Korea JIANBIN SU  Division of Plant Sciences, Christopher S. Bond Life Sciences Center, and Interdisciplinary Plant Group, University of Missouri, Columbia, MO, USA KIWAMU TANAKA  Department of Plant Pathology, Washington State University, Pullman, WA, USA MARIEKE VAN HULTEN  Molecular Plant Pathology, Swammerdam Institute for Life Sciences (SILS), University of Amsterdam, Amsterdam, The Netherlands HARROLD A. VAN DEN BURG  Molecular Plant Pathology, Swammerdam Institute for Life Sciences (SILS), University of Amsterdam, Amsterdam, The Netherlands ZHIXUE WANG  Plant Biology Section, School of Integrative Plant Science, Cornell University, Ithaca, NY, USA; State Key Laboratory of Rice Biology, Institute of Nuclear Agricultural Sciences, Zhejiang University, Hangzhou, China LEIYUN YANG  Plant Biology Section, School of Integrative Plant Science, Cornell University, Ithaca, NY, USA

Chapter 1 Using Bioinformatics and Molecular Biology to Streamline Construction of Effector Libraries for Phytopathogenic Pseudomonas syringae Strains Jay Jayaraman, Morgan K. Halane, Sera Choi, Honour C. McCann, and Kee Hoon Sohn Abstract The war between plants and their pathogens is endless, with plant resistance genes offering protection against pathogens until the pathogen evolves a way to overcome this resistance. Given how quickly new pathogen strains can arise and defeat plant defenses, it is critical to more rapidly identify and examine the specific genomic characteristics new virulent strains have gained which give them the upper hand. An indispensable tool is bioinformatics. Genome sequencing has advanced rapidly in the last decade, and labs are frequently uploading high-quality genomes of various organisms, including plant pathogenic bacteria such as Pseudomonas syringae. Pseudomonas syringae strains inject several effector proteins into host cells which often overcome host defenses. Probing online genomes provides a way to quickly and accurately predict effector repertoires of Pseudomonas, enabling the cloning of complete effector libraries of newly emerged strains. Here, we describe detailed protocols to rapidly clone bioinformatically predicted P. syringae effectors for various screening applications. Key words Genome sequencing, Arabidopsis, Pseudomonas syringae, Effectors, Plant disease resistance, Golden Gate cloning

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Introduction Plants are constantly under attack by pathogens; however, most plants are resistant to most pathogens. Plant disease resistance is governed mainly by plant resistance genes which encode resistance proteins [1]. These proteins can, directly or indirectly, recognize secreted pathogen effectors, leading to a rapid immune response. However, as plants have evolved resistance genes for protection, rapidly-evolving pathogens can also acquire new ways to overcome plant defenses via effectors. This is typically achieved through the

Jay Jayaraman and Morgan K. Halane are co-first authors. Walter Gassmann (ed.), Plant Innate Immunity: Methods and Protocols, Methods in Molecular Biology, vol. 1991, https://doi.org/10.1007/978-1-4939-9458-8_1, © Springer Science+Business Media, LLC, part of Springer Nature 2019

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procurement of new effectors via horizontal gene transfer, mutation of current effector genes in order to maintain virulence but evade host recognition, or jettisoning the recognized effector entirely. Whole genome sequencing has advanced quickly in recent years, making it a key tool in protecting plants from new virulent bacterial pathogens. By using bioinformatics to predict effector libraries from new virulent strains, it is possible to clone complete effector libraries from those strains and test them, quickly moving from in silico prediction to in planta experimentation. The genome of Pseudomonas syringae pv. actinidiae, causal agent of the bacterial canker disease in kiwifruit, was recently sequenced [2]. We used this advance to examine effectors secreted from a newly emerged Pseudomonas syringae pv. actinidiae strain, V13, a causal agent of the bacterial canker disease in kiwifruit which has wreaked havoc on the industry in diverse regions such as Italy, Chile, and New Zealand [3, 4]. Here, we describe a method in which a bioinformatic approach is used to predict the full type-III secreted effector repertoire of a highly virulent and destructive Pseudomonas syringae strain, and an effector library is constructed to examine these distinct effectors in plants.

2

Materials

2.1 Plasmid DNA Construction Using Golden Gate Cloning

1. Deionized sterile water (dH2O). 2. Genomic DNA Purification Kit (e.g., Wizard Kit from Promega). 3. BsaI restriction enzyme. 4. SmaI restriction enzyme. 5. Enzyme buffer(s). 6. High-fidelity polymerase (e.g., Phusion® from Thermo). 7. High-fidelity polymerase buffer. 8. Deoxynucleotide triphosphates (dNTPs). 9. T4 DNA ligase. 10. T4 DNA ligase buffer. 11. pICH41021 entry vector. 12. A DNA gel extraction kit (e.g., Zymoclean Gel DNA recovery kit from Zymo Research). 13. DNA Clean & Concentrator kit or Sepharose 4B. 14. Plasmid miniprep kit (e.g., ZymoPURE from Zymo Research). 15. Site-directed mutagenesis kit (e.g., QuickChange II from Agilent). 16. Yeast transformation kit (e.g., Frozen-EZ Yeast Transformation II kit from Zymo Research).

Bioinformatic Prediction and Library Construction of Pseudomonas. . .

3

17. Bovine serum albumin (BSA). 18. Dimethyl sulfoxide (DMSO). 2.2 Plasmid Mobilization

1. Sterile dH2O. 2. MicroPulser™ Electroporator (Biorad). 3. Low salt LB media (per liter): 10 g tryptone, 5 g yeast extract, 5 g NaCl, 1 g D-glucose, and 10 g agar for solid media. 4. N,N-Dimethylformamide (DMF). 5. Isopropyl-β-D-thiogalactoside (IPTG, 0.1 M solution in dH2O). 6. 5-Bromo-4-chloro-3-indolyl β-D-galactopyranoside (X-gal, 20 mg/mL solution in DMF). 7. King’s B media: 20 g bacto peptone, 1.5 g K2HPO4, 10 mL glycerol, and 15 g agar (for solid media).

3

Methods As an example for this protocol, sequences used were from a newly emerged Pseudomonas syringae pv. actinidiae (Psa) strain Psa V13 (ICMP18884) as well as from a nonpathogenic strain isolated from the same host plants, Pseudomonas syringae pv. actinidifoliorum (Pfm) strain Pfm LV5 (ICMP18803) previously also classified as Psa. Their sequenced genomes are publicly available from NCBI and at pseudomonas-syringae.org. A large selection of recommended Golden Gate modules for cloning and Level 1 vectors for a wide variety of applications are published and available [5].

3.1 Identification of Bioinformatically Predicted Effectors

1. Obtain effector protein sequences from the pseudomonassyringae.org database and use them as query sequences in a tBLASTn search of the bacterium of choice, in our case Psa V13, retaining hits with e-values 100 ng/ μL), 1.5 μL 10 T4 DNA ligase buffer (concentration may vary depending on manufacturer), 0.5 μL SmaI restriction enzyme (see Note 3), 0.5 μL T4 DNA ligase, and sterile dH2O for 20 μL total reaction volume. Incubate the SmaI/ ligase reaction mix for 2 h at room temperature. 8. Use a DNA Clean & Concentrator kit or Sepharose to clean the SmaI/ligase reaction mix. Elute in a 7 μL volume (see Note 5). 9. Transform 5–10 μL of the cleaned-up ligation reaction mix by mixing with a 40 μL aliquot of cold electrocompetent Escherichia coli cells in 0.1 cm width electroporation cuvettes. Pulse (1.8 kV) on the electroporator. Add 0.5 mL LB liquid medium to cells, recover at 37  C with shaking, and plate the cells on agar-solidified LB medium plates with 100 μg/mL ampicillin. Spread them with 10 μL 0.1 M IPTG and 40 μL 20 mg/mL X-gal. Incubate the plates at 37  C for 18–20 h, and select positive white colonies for further screening. 10. Inoculate white colonies in LB liquid medium with 100 μg/ mL ampicillin and miniprep pICH41021 plasmid carrying the inserted module using the plasmid miniprep kit. Elute in a 50 μL volume. Screen for inserted modules in plasmids by PCR or flanking BsaI restriction enzyme digest and agarose gel electrophoresis. 11. Sequence each module cloned in the pICH41021 vector before proceeding. Check each sequence by alignment to the ä Fig. 1 (continued) 4 bp with the Level 1 destination vector sequence, and last reverse primer (R4), which overlaps 4 bp with the C-terminal tag sequence, each forward primer overlaps with the next reverse primer for 4 bp native to GeneX. Each of the 4 bp overlaps is unique, allowing assembly of each of the modules in the correct order. Following PCR of each of the modules, gel extraction of PCR products, and four SmaI digest/ blunt-end ligation reactions of each of the PCR products into the Level 0 shuttle vector pICH41021, constructs are transformed into E. coli. Cloned constructs extracted from E. coli are then isolated and sequenced to confirm that there are no errors in cloning and that the flanking BsaI sites introduced in the primers are present and lead to expected 4 bp overlap sequences. Indicated in GeneX sequence (orange arrows) are two internal BsaI restriction sites (one in module 1 and the other in module 2) that will then be removed by site-directed mutagenesis and vectors re-sequenced to confirm BsaI site removal without frame-shift mutations. (b) Golden Gate assembly with the Level 1 destination vector of choice, each of the four modules of GeneX, and a selected epitope tag is conducted and cloned into E. coli. As an example, the vector indicated in (b) is the binary vector for Agrobacterium-mediated transient expression, pICH86988. By Golden Gate assembly, GeneX is reconstituted from its four modules into pICH86988 by replacing the lacZ sequence, within the right (RB) and left (LB) border sequences under the 35S-Ω promoter. Authentic assembly is confirmed by restriction digest and sequencing. These steps (a) and (b) can be repeated for each of the genes to be cloned generating a module library for each Pseudomonas strain and a selection of different Level 1 libraries for screening by different methods

Bioinformatic Prediction and Library Construction of Pseudomonas. . .

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original sequence of the effector, and look for correct positioning of BsaI sites and respective overhangs expected from the primer sequence design. 12. If desired, conduct site-direct mutagenesis at this point to remove any internal BsaI sites using a kit such as the Agilent QuickChange II Site-Directed Mutagenesis kit. After transformation of E. coli following mutagenesis, re-sequence the plasmid(s) to confirm their integrity and to visualize expected mutation(s). 3.3 Using Golden Gate Assembly to Build Multipurpose Effector Libraries

1. Assemble modules into a Golden Gate-compatible destination vector of interest (Fig. 1). Suitable Golden Gate-compatible vectors are listed in Table 1 with suggested screening methods/applications. 2. Set up Golden Gate assemblies with equimolar amounts of each module and, if required, C-terminal tag in 0.2 mL PCR tubes: X μL of each module and Golden Gate-compatible vector (and tags if required) in equimolar amounts (~50 ng each), 2 μL 10 T4 DNA ligase buffer, 2 μL 10 BSA, 1 μL T4 DNA ligase, 1 μL BsaI restriction enzyme, and sterile dH2O up to 20 μL. 3. Run the Golden Gate reaction on a thermocycler under the following conditions: 25 cycles of digestion for 3 min at 37  C, ligation for 4 min at 16  C; denature ligase for 5 min at 50  C; denature BsaI for 5 min at 80  C; and hold at 20  C. 4. Use a DNA Clean & Concentrator kit or Sepharose to clean the SmaI/ligase reaction mix. Elute in a 7 μL volume (see Note 5). 5. Transform 5 μL of the cleaned-up Golden Gate reaction mix by mixing with a 40 μL aliquot of cold electrocompetent Escherichia coli cells (see Note 6) in 0.1 cm width electroporation cuvettes, pulsing on the electroporator, recovering and plating on agar-solidified LB medium plates with appropriate antibiotic as in Subheading 3.2, step 9. 6. Inoculate white colonies in LB liquid medium with appropriate antibiotic and miniprep Golden Gate assembled plasmids with inserted modules (and tag if used) using the plasmid miniprep kit. Elute in a 50 μL volume. Screen for inserted modules in plasmids by PCR or restriction enzyme digest followed by agarose gel electrophoresis to confirm authentic assembly. 7. Assess effectors for the development of a hypersensitive response in plants by Pseudomonas type-III secretion system delivery [10] or analyze effectors using other standard recommended procedures (see Notes 8–10).

C-terminal

C-terminal

Yeast GAL1 promoter

Yeast GAL1 promoter

pLexA-GG

pB42AD-GG

TRP1 (Tryptophan prototrophy); AmpicillinR

Saccharomyces cerevisiae RFY206; Mat (α)

[11]

References

[13] Yeast prey vector carrying N-terminal fused LexA DNA-binding domain for GAL promoter

[13]

[12] Binary vector for Agrobacterium transient/ stable infiltration or floral dips

[12] Binary vector for Agrobacterium transient/ stable infiltration or floral dips

Broad host-range vector with avrRps4 promoter driven expression

Description

Saccharomyces cerevisiae Yeast bait vector carrying N-terminal fused EGY48 þ pSH18-34 activation domain for (lacZ reporter þ GAL promoter URA3, Uracil prototrophy); Mat (a)

A. tumefaciens

KanamycinR

N- or C-terminal

None native, assembled with selected promoter module

pICH86966

HIS3 (Histidine prototrophy); AmpicillinR

A. tumefaciens

KanamycinR

CaMV 35S promoter þ TMV C-terminal Ω leader

pICH86988

P. fluorescens Pf0–1 (T3S), or P. syringae

Host

Gentamicin

R

C-terminal

Selection marker

avrRps4 promoter (hrp box)

Epitope tag compatible

pBBR1MCS5B:: avrRps4pro

Level 1 vector Promoter element

Table 1 Golden Gate-compatible vectors recommended for screening effectors

8 Jay Jayaraman et al.

Bioinformatic Prediction and Library Construction of Pseudomonas. . .

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9

Notes 1. Bioinformatic methods for the identification of candidate effectors often use different BLAST algorithms to identify candidate effector sequences, followed by filtering with various minimum identity and coverage thresholds [3, 14, 15]. Most automated annotation methods do not correctly identify the position of effector start codons, so caution must be exercised if using the predicted proteome as a BLAST subject database. 2. Psa V13 candidate effector sequences were manually inspected to determine the position of the start codon by comparison with homologous effectors and the position of an upstream hrp box [2, 3]. 3. If even a single SmaI site is present in the module to be bluntend cloned into pICH41021, the cloned products will be truncated. To avoid this, a second restriction enzyme, Eco53kI, can be used to blunt-end clone into pICH41021 with the same reaction conditions as for SmaI. However, since SmaI enzymatic function is high at room temperature (25  C) while that for Eco53kI is 37  C, if using the latter enzyme, the digestion/ligation reaction should be run in a thermocycler under the Golden Gate assembly program (see Subheading 3.3, step 3). 4. If both Eco53kI and SmaI restriction sites are present in the module to be cloned, a TA cloning strategy can be used instead. Following the module PCR amplification and agarose gel extraction, the product can be incubated with Taq DNA polymerase briefly to add 30 adenine residues to the product: 6 μL purified PCR product, 1 μL dATP (10 mM), 5 μL Taq polymerase PCR buffer with MgCl2 (10), 0.2 μL Taq DNA polymerase (New England Biolabs), and 37.8 μL sterile dH2O. The reaction is incubated for 20 min at 72  C and then ligated into a TA cloning-compatible vector, e.g., pCR8 (Thermo) or pGEM-T (Promega) kits. 5. Alternatively, any reaction mix can be cleaned up (desalted) in a lab-made Sepharose column. Briefly, add 120 μL Sepharose 4B to a 0.6 mL microfuge tube pierced with a fine needle at the base of the tube and then placed in a holding 2 mL microcentrifuge tube. Centrifuge the Sepharose for 2 min at 600  g to set the column; then transfer the 0.6 mL tube with the Sepharose column to a new collection tube; load the sample to be desalted onto the column; and re-centrifuge for 2 min at 600  g to collect the sample. 6. Electrocompetent Escherichia coli (or Agrobacterium tumefaciens) cells can be prepared and stored at
80  C for extended periods for later use, after thawing on ice:

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(a) Inoculate a colony from a fresh (grown overnight) plate of E. coli strain to be made electrocompetent into 10 mL of liquid LB medium in a 50 mL sterile centrifuge tube and incubate for up to 20 h at 37  C with shaking at 200 rpm. (b) In a 2 L flask containing 500 mL of LB, add 5 mL of the overnight culture. (c) Shake at 37  C and 200 rpm until the cultures reach an OD600 of up to 0.7. Place the culture on ice immediately for 20 min. Precool the centrifuge and 1.5 L of sterile 10% glycerol to 4  C prior to harvesting cells. (d) Cells must now be kept ice cold at all times. Pour 250 mL culture each into two chilled 500 mL centrifuge bottles (Nalgene). (e) Centrifuge at 5000  g for 15 min. Pour off the supernatant (aspirate the residual media) and add 250 mL of 10% glycerol to each of the centrifuge bottles and completely suspend the cells by firmly swirling the glycerol solution, avoiding vigorous shaking. (f) Re-centrifuge at 5000  g for 15 min. Pour off the supernatant, and completely suspend the cells in 200 mL 10% glycerol. (g) Re-centrifuge at 5000  g for 15 min. Pour off the supernatant, and completely suspend the cells in 100 mL 10% glycerol, pooling both tubes into one. (h) Re-centrifuge at 5000  g for 15 min. Pour off the supernatant, and completely suspend the cells in 2 mL 10% glycerol; pipette gently to resuspend the cells. (Agrobacterium-competent cells can be mixed in a much larger volume—10 mL.) (i) Aliquot 40 μL volumes of the competent cells into pre-chilled sterile microcentrifuge tubes and transfer them to a
80  C freezer for long-term storage. 7. Alternatively, electroporation of Pseudomonas syringae is also a viable option. To prepare electrocompetent Pseudomonas syringae: (a) Inoculate a colony of the Pseudomonas strain of interest from a fresh (grown overnight) agar-solidified King’s B plate into 5 mL of liquid LB medium and grow overnight at 28  C. (b) Centrifuge cells at 5000  g for 3 min. Discard the supernatant, and resuspend the cells in 1 mL of 300 mM sucrose. (c) Re-centrifuge at 5000  g for 3 min. Discard the supernatant and re-centrifuge cells to remove all traces of supernatant. Resuspend cells in 50 μL of 300 mM sucrose.

Bioinformatic Prediction and Library Construction of Pseudomonas. . .

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(d) Add 1 μL of plasmid and pulse with the electroporator set at 2 kΩ. Add 500 μL LB to cells; recover at 28  C for 90 min with shaking (200 rpm). (e) Streak 150 uL of transformed cells onto LB plates containing appropriate antibiotic(s), and store the plates at 28  C until transformed bacterial colonies grow visibly (2–3 days). 8. You may wish to mobilize the assembled broad host-range vector library into a Pseudomonas strain of choice via triparental mating: (a) Grow overnight LB liquid medium cultures of (donor) E. coli containing your broad host-range vector with effector of interest, a helper E. coli strain (e.g., HB101 carrying the vector pRK2013), and the recipient Pseudomonas strain. (b) Mix suspensions of Pseudomonas strain, donor E. coli, and helper E. coli in a 6:1:1 ratio (final volume of 800 μL), centrifuge at 5000  g for 3 min, and resuspend in 100 μL. (c) Spot several 20 μL aliquots of cell mixture on agarsolidified LB medium plates and incubate at 28  C for 6–8 h. (d) Streak from spotted aliquots on agar-solidified King’s B medium plates with appropriate antibiotic and incubate for 2 days at 28  C for colony transformants to appear. 9. Effector-triggered cell death in Nicotiana plants, as well as in planta protein expression and subcellular localization, can be achieved by transient expression using Agrobacterium tumefaciens [16]. An A. tumefaciens strain of choice can be transformed through electroporation to carry the appropriate binary vector (pICH86988 or pICH86966) (see Note 7). Stable expression lines in Arabidopsis thaliana plants can also be made using the same library in A. tumefaciens by floral dip [17]. 10. Protein-protein interaction can be investigated for effectors by yeast two-hybrid assays [18]. Yeast can be transformed with Golden Gate assembled vectors using the Zymo Frozen-EZ Yeast Transformation II kit.

Acknowledgments This work was carried out with the support of the Basic Science Research Program through the National Research Foundation of Korea (NRF) funded by the Ministry of Education (NRF-2016R1D1A1B03934707), Republic of Korea.

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References 1. Jones JD, Dangl JL (2006) The plant immune system. Nature 444(7117):323–329. https:// doi.org/10.1038/nature05286 2. Templeton MD, Warren BA, Andersen MT et al (2015) Complete DNA sequence of pseudomonas syringae pv. actinidiae, the causal agent of Kiwifruit canker disease. Genome Announc 3(5):e01054. https://doi.org/10. 1128/genomeA.01054-15 3. McCann HC, Rikkerink EH, Bertels F et al (2013) Genomic analysis of the Kiwifruit pathogen Pseudomonas syringae pv. actinidiae provides insight into the origins of an emergent plant disease. PLoS Pathog 9(7):e1003503. https://doi.org/10.1371/journal.ppat. 1003503 4. Choi S, Jayaraman J, Segonzac C et al (2017) Pseudomonas syringae pv. actinidiae type III effectors localized at multiple cellular compartments activate or suppress innate immune responses in nicotiana benthamiana. Front Plant Sci 8:2157. https://doi.org/10.3389/ fpls.2017.02157 5. Engler C, Youles M, Gruetzner R et al (2014) A golden gate modular cloning toolbox for plants. ACS Synth Biol 3(11):839–843. https://doi.org/10.1021/sb4001504 6. Altschul SF, Gish W, Miller W et al (1990) Basic local alignment search tool. J Mol Biol 215(3):403–410. https://doi.org/10.1016/ S0022-2836(05)80360-2 7. Lindeberg M, Stavrinides J, Chang JH et al (2005) Proposed guidelines for a unified nomenclature and phylogenetic analysis of type III Hop effector proteins in the plant pathogen Pseudomonas syringae. Mol PlantMicrobe Interact 18(4):275–282. https://doi. org/10.1094/MPMI-18-0275 8. Stamatakis A (2014) RAxML version 8: a tool for phylogenetic analysis and post-analysis of large phylogenies. Bioinformatics 30 (9):1312–1313. https://doi.org/10.1093/ bioinformatics/btu033 9. Engler C, Kandzia R, Marillonnet S (2008) A one pot, one step, precision cloning method with high throughput capability. PLoS One 3 (11):e3647. https://doi.org/10.1371/jour nal.pone.0003647 10. Thomas WJ, Thireault CA, Kimbrel JA et al (2009) Recombineering and stable integration of the Pseudomonas syringae pv. syringae

61 hrp/hrc cluster into the genome of the soil bacterium Pseudomonas fluorescens Pf0-1. Plant J 60(5):919–928. https://doi.org/10. 1111/j.1365-313X.2009.03998.x 11. Jayaraman J, Choi S., Prokchorchik M, Choi DS, Spiandore A, Rikkerink EH, Templeton MD, Segonzac C, Sohn KH (2017) A bacterial acetyltransferase triggers immunity in Arabidopsis thaliana independent of hypersensitive response. Sci Rep 7:3557. https://doi.org/ 10.1111/j.1365-313X.2009.03998.x 12. Weber E, Engler C, Gruetzner R, Werner S, Marillonnet S (2011) A modular cloning system for standardized assembly of multigene constructs. PLoS ONE 6(2):e16765. https:// doi.org/10.1371/journal.pone.0016765 13. Segonzac C, Newman TE, Choi S, Jayaraman J, Choi DS, Jung, GY, Cho H, Lee YK, Sohn KH (2017) A conserved EAR motif is required for avirulence and stability of the Ralstonia solanacearum effector PopP2 in planta. Front Plant Sci 8. https://doi.org/10.3389/fpls. 2017.01330 14. Baltrus DA, Nishimura MT, Romanchuk A et al (2011) Dynamic evolution of pathogenicity revealed by sequencing and comparative genomics of 19 Pseudomonas syringae isolates. PLoS Pathog 7(7):e1002132. https://doi. org/10.1371/journal.ppat.1002132 15. Hulin MT, Armitage AD, Vicente JG et al (2018) Comparative genomics of Pseudomonas syringae reveals convergent gene gain and loss associated with specialization onto cherry (Prunus avium). New Phytol 219 (2):672–696. https://doi.org/10.1111/nph. 15182 16. Vaghchhipawala Z, Rojas CM, Senthil-Kumar M et al (2011) Agroinoculation and agroinfiltration: simple tools for complex gene function analyses. Methods Mol Biol 678:65–76. https://doi.org/10.1007/978-1-60761-6825_6 17. Clough SJ, Bent AF (1998) Floral dip: a simplified method for Agrobacterium-mediated transformation of Arabidopsis thaliana. Plant J 16(6):735–743 18. Fields S, Song O (1989) A novel genetic system to detect protein-protein interactions. Nature 340(6230):245–246. https://doi.org/10. 1038/340245a0

Chapter 2 Immunoprecipitation Under Non-Denaturing or Denaturing Conditions of Lysine-Acetylated Proteins Expressed in Planta Maxime Escouboue´ and Laurent Deslandes Abstract Protein lysine acetylation is a highly conserved posttranslational modification that plays key roles in many biological processes such as the regulation of gene expression, chromatin dynamics, and metabolic pathways. Recent studies revealed that various pathogens use lysine acetylation to interfere with host immune responses. Identification of lysine-acetylated host proteins resulting from virulence activities of pathogen effectors is therefore essential for understanding their biological functions. Here we provide a method for immunoprecipitating lysine-acetylated proteins transiently expressed in planta under non-denaturing or denaturing conditions and detecting them by immunoblotting. To illustrate this rapid and simple procedure, immunoprecipitation of the lysine-acetylated WRKY domain of the RRS1-R immune receptor, a substrate of the Ralstonia solanacearum PopP2 effector, is presented as a typical example. Key words Lysine acetylation, Effector, Agrobacterium-mediated transient expression, Nicotiana benthamiana

1

Introduction Posttranslational modifications (PTMs) of proteins such as phosphorylation, ubiquitination, methylation, and acetylation play a pivotal role in many biological processes [1, 2]. Acetylation of lysine residues represents one of the most universal PTMs and is conserved across prokaryotes and eukaryotes [3]. This modification is catalyzed by lysine acetyltransferases and consists of transferring an acetyl group (C2H3O) from acetyl coenzyme A to a specific residue. Two kinds of acetylations can be distinguished. The Nt-acetylation catalyzed by Nt-acetyltransferases (NATs) is irreversible and occurs on the N-terminal amino group of a polypeptide. By contrast, Nε-acetylation catalyzed by lysine (K) acetyltransferases (KATs) is reversible and occurs on the ε-amino group of specific lysine residues of substrate proteins [4]. As a consequence of the neutralization of the positive charge of lysine residues, lysine acetylation can

Walter Gassmann (ed.), Plant Innate Immunity: Methods and Protocols, Methods in Molecular Biology, vol. 1991, https://doi.org/10.1007/978-1-4939-9458-8_2, © Springer Science+Business Media, LLC, part of Springer Nature 2019

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affect biological functions such as protein activities, protein localizations, and protein-protein and protein-DNA interactions [5]. First discovered on histone tails [6] and involved in the regulation of chromatin structure and gene expression, lysine acetylation also occurs on a large number of nonhistone proteins in different subcellular compartments. In plant-pathogen interactions, effector-mediated lysine acetylation of host immune-related components represents a potent virulence strategy used by pathogens to inactivate immune complexes and suppress defense signaling [7, 8]. This is particularly well illustrated with the YopJ effector family that exists in many plant and animal pathogens [9]. YopJ effectors can affect the function and/or the stability of their targets by acetylating specific lysine, serine, and/or threonine residues. Modifications of these residues lead to many perturbations including inactivation of protein functions, inhibition of protein-protein and protein-DNA interactions, and interference with other PTMs [10, 11]. YopJ effectors typically contain a conserved catalytic cysteine (Cys) residue involved in acetyltransferase activity [12]. The founding member, the Yersinia pestis effector YopJ, acetylates critical residues of various mitogen-activated protein kinase kinases to block host immunity [13]. Recently, the mode of action of the YopJ member PopP2 from the soilborne bacterium Ralstonia solanacearum has been elucidated. PopP2 was shown to target many defensive WRKY transcription factors (TFs) by acetylating a key lysine residue located in the conserved heptad of their WRKY DNA-binding domains. PopP2-triggered acetylation of WRKY TFs inhibits their DNA-binding activity and blocks their transactivating functions needed for defense gene expression in basal resistance [14]. PopP2 also acetylates the same lysine residue located within the C-terminal WRKY domain of the Arabidopsis RRS1-R immune receptor that binds DNA. This disrupts RRS1-RDNA association and activates RPS4-dependent immunity [14]. Targeted assays using radioactive labeling or immunoblotting with anti-acetylated lysine antibodies are traditionally used for studying acetylation of candidate substrates. With these methods, the most significant challenge is detection of low-abundance acetylated proteins. To enhance the sensitivity of detection in immunoblotting, pre-enrichment of lysine-acetylated proteins can be useful. Here we describe a simple and rapid method for immunoprecipitating lysine-acetylated candidate proteins transiently expressed in N. benthamiana cells. In some cases, lysine-acetylated residues of native proteins are not accessible to antibodies that only recognize denaturated proteins. Therefore, we present two immunoprecipitation procedures performed under either non-denaturing or denaturing conditions, respectively. Lysine acetylation of the WRKY domain of RRS1-R by the bacterial acetyltransferase PopP2 [14] is presented as a typical example. The principles of these two protocols are applicable to other lysine-acetylated candidate proteins.

Detection of Lysine‐Acetylated Proteins

2

15

Materials Unless otherwise noted, deionized water and basic molecular biology reagents and equipment are used to prepare the specified solutions.

2.1 AgrobacteriumMediated Transient Expression in N. benthamiana Leaves

1. Three- to four-week-old N. benthamiana plants grown in a greenhouse under natural daylight conditions. 2. Agrobacterium tumefaciens strain transformed with a binary vector allowing constitutive expression of your protein of interest. As a typical example, we constitutively overexpressed the C-terminal WRKY domain of RRS1-R tagged with a triple FLAG epitope (35S-WRKY-R-3FLAG) [14]. Active PopP2 and the catalytically inactive mutant PopP2-C321A were fused to a triple HA epitope (35S-PopP2-3HA and 35S-PopP2-C321A-3HA). Introduce binary vectors by electroporation in Agrobacterium tumefaciens (see Note 1). 3. YEB Agrobacterium growth medium: 5 g/L beef extract, 1 g/ L yeast extract, 5 g/L peptone, 5 g/L sucrose, and 0.5 g/L MgCl2. Combine the reagents and shake until complete dissolution. Adjust the final volume of the solution to 1 L with H2O. Sterilize by autoclaving. 4. Antibiotics used for Agrobacterium tumefaciens: gentamicin 15 μg/mL and carbenicillin 25 μg/mL. 5. Agromix resuspension buffer: 10 mM 2-(N-morpholino)ethanesulfonic acid (MES)-KOH pH 5.6, 10 mM MgCl2, 150 mM acetosyringone (from a 0.5 M stock solution of 30 ,50 -dimethoxy40 -hydroxyacetophenone dissolved in DMSO, aliquoted (50 μL) and stored at 20  C).

2.2 Extraction and Immunoprecipitation of Lysine-Acetylated Proteins from Plant Tissues

1. 1 M Tris–HCl pH 7.5, autoclaved. 2. 3 M NaCl, autoclaved. 3. 0.5 M ethylenediaminetetraacetic acid (EDTA)/NaOH, pH 8.0, autoclaved. 4. 1 M dithiothreitol (DTT), sterilized by filtration, aliquoted (100 μL), and stored at 20  C. 5. 10% Triton™ X-100 diluted in water. 6. Protease inhibitor cocktail for plant cell extracts  100 (SIGMA). 7. 1 M sodium butyrate dissolved in water, sterilized by filtration, aliquoted, and stored at 20  C (see Note 2). 8. 20% sodium dodecyl sulfate (SDS) dissolved in water and stored (up to 1 year) at room temperature. 9. Bromophenol blue.

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10. Non-denaturing IP buffer: 50 mM Tris–HCl pH 7.5, 150 mM NaCl, 5 mM EDTA pH 8.0, 2 mM DTT, 0.2% Triton™ X-100, 1% (v/v) protease inhibitor cocktail, and 10 mM sodium butyrate. 11. Denaturing lysis buffer: 50 mM Tris–HCl pH 7.5, 5 mM EDTA pH 8.0, 10 mM DTT, 1% SDS, 1% (v/v) protease inhibitor cocktail, and 10 mM sodium butyrate. Keep at room temperature to avoid SDS precipitation. 12. 4 Laemmli denaturation buffer: 0.25 M Tris–HCl pH 7.5, 40% glycerol (v/v), 8% SDS, 0.4 M DTT, and 0.04% bromophenol blue. Aliquot and store at 20  C. 13. Protein A agarose (SIGMA). 14. Anti-acetylated lysine antibody (e.g., cell signaling, #9441) (see Note 3). 15. 15 mL sterile plastic centrifuge tubes. 16. Coring tool (7 mm diameter). 2.3 Immunoblot Detection

1. Standard equipment for SDS-PAGE and semidry transfer. 2. Tris-buffered saline containing 0.1% Tween 20 (TBS-T): 10 mM Tris–HCl pH 7.5, 150 mM NaCl, and 0.1% (v/v) Tween® 20. 3. Hybond™-ECL™ nitrocellulose membrane (0.45 μm) (e.g., GE Healthcare). 4. Skim milk powder. 5. Blocking TBS-T buffer: TBS-T buffer with 1% skim milk powder. 6. Anti-HA-Peroxidase and anti-FLAG M2-Peroxidase antibody from mouse (Sigma). 7. Chemiluminescent immunoblot blot detection reagents (e.g., Clarity Western ECL Blotting Substrates, Bio-Rad). 8. Chemiluminescence imager (e.g., ChemiDoc™ MP Imaging System Bio-Rad).

3

Methods We present the immunoprecipitation under non-denaturing and denaturing conditions of the WRKY domain of the RRS1-R immune (WRKY-R-3FLAG) that was shown to be acetylated on a specific lysine residue with active PopP2 acetyltransferase (PopP23HA) but not with catalytically inactive PopP2 C321A mutant (PopP2-C321A-3HA) [14] (see Fig. 1).

Detection of Lysine‐Acetylated Proteins

17

Fig. 1 Immunoprecipitation under non-denaturing and denaturing conditions of Lys-acetylated WRKY-R co-expressed with PopP2. The WRKY domain of RRS1-R (WRKY-R-3FLAG) was transiently expressed with 3HA-tagged PopP2 or C321A in N. benthamiana leaves. Protein extracts were immunoblotted with anti-HA (α-HA) and anti-FLAG antibodies (α-FLAG) (CE, input). Lysine-acetylated WRKY-R-FLAG was immunoprecipitated with anti-acetylated lysine antibody (IP AcK) under non-denaturing (left panel) or denaturing conditions (right panel). Immunoprecipitated proteins were analyzed on immunoblots with α-FLAG antibodies for detection of acetylated WRKY-R (Ac WRKY-R-3FLAG). Protein loading was monitored by Ponceau staining. The pre-stained protein ladder is visible 4th lane from the left 3.1 AgrobacteriumMediated Transient Expression in N. benthamiana Leaves

1. Inoculate one single colony of Agrobacterium tumefaciens containing the 35S:WRKY-R-3FLAG, 35S-PopP2-3HA, and 35S-PopP2-C321A-3HA constructs (pAM-PAT based vectors [14]) in 10 mL of YEB with appropriate antibiotics (gentamicin 15 μg/mL and carbenicillin 25 μg/mL). Incubate overnight at 28  C, shaking at 140 rpm. 2. Centrifuge the Agrobacterium cells at 4500  g for 10 min; then resuspend the pellet in 2 mL of Agromix resuspension buffer. 3. Measure the optical density (OD) at 600 nm. For singleprotein expression, adjust the OD600 to 0.25. 4. For co-expression of two proteins, combine 0.25 OD of each Agrobacterium strain, final OD600 ¼ 0.5. 5. In some cases, the co-expression of 35S:p19 (0.25 OD600), a suppressor of RNA silencing, can promote the accumulation of the GFP fusion proteins (see Note 4). 6. Incubate the suspension at room temperature for at least 60 min before infiltration. 7. Infiltrate the bacterial suspension into the underside of a N. benthamiana leaf using a 1 mL syringe without a needle.

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3.2 Immunoprecipitation of Lysine-Acetylated Proteins Under Nondenaturing Conditions

1. 36–48 h after infiltration, harvest four 7 mm leaf discs with a coring tool or equivalent for each protein combination.

3.2.1 Protein Extraction

4. Transfer supernatants carefully to clean chilled microcentrifuge tubes, and keep on ice.

2. Grind leaf material in liquid nitrogen, and thaw in 1 mL of non-denaturing IP buffer. Incubate on ice for 5 min. 3. Centrifuge the mixture at 9000  g for 5 min at 4  C.

5. To prepare an aliquot for an input control, transfer 50 μL of the extract into a 1.5 mL microcentrifuge tube, and add 50 μL of 4 Laemmli denaturation buffer. Mix by flicking the tube. 6. Denature the input control samples in 2 Laemmli denaturation buffer at 95  C for 3 min. These samples correspond to the input for IP under non-denaturing conditions and are ready for immunoblotting as described in Subheading 3.4, step 1. 3.2.2 Immunoprecipitation

1. To the protein extract kept on ice (Subheading 3.2.1, step 4), add 20 μL 50% slurry of protein A agarose pre-equilibrated with non-denaturing IP buffer. 2. Incubate for 20 min at 4 (preclearing step).



C with gentle shaking

3. Centrifuge at 800  g for 2 min at 4  C. 4. Transfer the supernatant to a clean microcentrifuge tube, and discard the agarose beads. 5. Add 1 μL of acetylated lysine antibody. Incubate for 2 h at 4  C with gentle shaking. 6. Add 20 μL of a 50% slurry of pre-equilibrated protein A agarose in non-denaturing IP buffer. 7. Incubate for 1–2 h at 4  C with gentle shaking. 8. Centrifuge the tube at 800  g for 1 min at 4  C. Discard the supernatant. 9. Wash the resin with 1 mL of non-denaturing IP buffer (3  10 min at 4  C with gentle shaking). 10. Centrifuge the tube at 800  g for 1 min at 4  C. Discard the supernatant. 11. Resuspend the resin in 40 μL of 2 Laemmli denaturation buffer. 12. Heat samples to 95  C for 3 min to denature. These samples correspond to the acetylated lysine IP and are ready for immunoblotting as described in Subheading 3.4, step 1.

Detection of Lysine‐Acetylated Proteins

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3.3 Immunoprecipitation of Lysine-Acetylated Proteins Under Denaturing Conditions

1. 36–48 h after infiltration, harvest four 7 mm leaf discs with a coring tool, or cut equivalent material, for each protein combination.

3.3.1 Protein Extraction

3. Heat samples to 95  C for 3 min to denature.

2. Grind the leaf material in liquid nitrogen, and thaw in 400 μL of denaturing lysis buffer. 4. Centrifuge the mixture at 16,000  g for 2 min at room temperature. 5. Transfer 50 μL of the extract into a 1.5 mL microcentrifuge tube, and add 50 μL of 4 Laemmli denaturation buffer. Mix by flicking the tube. Denatured samples corresponding to the input for IP under denaturing conditions are ready for immunoblotting as described in Subheading 3.4, step 1.

3.3.2 Immunoprecipitation

1. Transfer the remaining denatured protein extract (~350 μL) into 15 mL sterile plastic centrifuge tubes containing 20 volumes (~7 mL) of non-denaturing IP buffer. Add 0.1 volume (~700 μL) of 10% Triton™ X-100 (see Note 5). 2. Add 20 μL of a 50% slurry of pre-equilibrated protein A agarose with non-denaturing IP buffer into each 15 mL tube. 3. Incubate for 20 min at 4 (preclearing step).



C with gentle shaking

4. Centrifuge at 800  g for 2 min at 4  C. 5. Transfer the supernatant to a clean 15 mL centrifuge tube. 6. Add 1 μL of anti-acetylated lysine antibody. Incubate for 2 h at 4  C with gentle shaking. 7. Add 20 μL of a 50% slurry of pre-equilibrated protein A agarose with non-denaturing IP buffer. 8. Incubate for 1–2 h at 4  C with gentle shaking. 9. Centrifuge the tube at 800  g for 2 min at 4  C. Carefully discard the supernatant. 10. Add 1 mL of non-denaturing IP buffer to resuspend the resin, and transfer into a clean 1.5 mL microcentrifuge tube. 11. Centrifuge the tube at 800  g for 1 min at 4  C. Discard the supernatant. 12. Wash the resin three times with 1 mL of non-denaturing IP buffer (3  10 min at 4  C with gentle shaking). 13. Resuspend the resin in 40 μL of 2 Laemmli denaturation buffer. 14. Heat samples to 95  C for 3 min. Denatured samples, corresponding to IP under denaturing conditions, are ready for immunoblotting as described in Subheading 3.4, step 1.

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3.4 Immunoblot Detection

1. Separate extracted samples (10 μL) by electrophoresis on a 10% SDS-polyacrylamide gel, and transfer to a nitrocellulose membrane. 2. Block the membrane with 2.5% (w/v) skim milk in TBS-T for 1 h at room temperature. 3. Incubate with HRP-conjugated antibodies at the desired dilution (e.g., 1:5000 dilution for both anti-HA- and anti-FLAG HRP-conjugated) in 0.5% (w/v) skim milk in TBS-T for 1 h at room temperature. 4. Wash three times with TBS-T (3  10 min) at room temperature. 5. Use chemiluminescent immunoblot blot detection reagents and a chemiluminescence imager to detect immunoreactive signals. See Fig. 1 for an example of immunoblotting showing lysine-acetylated WRKY-R-3FLAG upon co-expression with active PopP2-3HA.

4

Notes 1. Agrobacterium tumefaciens C58-derived strains such as GV3103 can be used [15]. 2. Na butyrate is a water soluble histone deacetylase inhibitor. This compound is used to prevent deacetylation activities during extraction and immunoprecipitation. 3. Anti-acetylated lysine antibody (cell signaling) detects proteins posttranslationally modified by acetylation on the ε-epsilonamine groups of lysine residues. The antibody recognizes acetylated lysine in a wide range of sequence contexts. 4. Co-expression with the p19 protein of tomato bushy stunt virus, a suppressor of RNA silencing, prevents possible RNA silencing and might promote increased transient expression of transgenes. 5. To prevent the SDS from denaturing the immunoprecipitating antibody, the protein samples must be diluted at least tenfold in SDS-free IP buffer to decrease the concentration of SDS to 0.1% (w/v) or less. An excess of 1% Triton™ X-100 in the IP s SDS.

Acknowledgments This work was performed at the Laboratory of Plant-Microbe Interactions, part of the French Laboratory of Excellence “TULIP” (ANR-10-LABX-41; ANR-11-IDEX-0002-02). L.D. was supported by a grant from the Agence Nationale de la Recherche (RADAR, ANR-15-CE20-0016-01).

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References 1. Hartl M, Finkemeier I (2012) Plant mitochondrial retrograde signaling: post-translational modifications enter the stage. Front Plant Sci 3:253 2. Johnova´ P, Skala´k J, Saiz-Ferna´ndez I et al (2016) Plant responses to ambient temperature fluctuations and water-limiting conditions: a proteome-wide perspective. Biochim Biophys Acta 1864(8):916–931 3. Mischerikow N, Heck AJ (2011) Targeted large-scale analysis of protein acetylation. Proteomics 11(4):571–589 4. Drazic A, Myklebust LM, Ree R et al (2016) The world of protein acetylation. Biochim Biophys Acta 1864(10):1372–1401 5. Yang XJ, Seto E (2008) Lysine acetylation: codified crosstalk with other posttranslational modifications. Mol Cell 31(4):449–461 6. Allfrey VG, Faulkner R, Mirsky AE (1964) Acetylation and methylation of histones and their possible role in the regulation of RNA synthesis. Proc Natl Acad Sci U S A 51:786–794 7. Lee J, Manning AJ, Wolfgeher D et al (2015) Acetylation of an NB-LRR plant immuneeffector complex suppresses immunity. Cell Rep 13(8):1670–1682 8. Ma KW, Ma W (2016) YopJ family effectors promote bacterial infection through a unique acetyltransferase activity. Microbiol Mol Biol Rev 80(4):1011–1027

9. Lewis JD, Lee A, Ma W et al (2011) The YopJ superfamily in plant-associated bacteria. Mol Plant Pathol 12(9):928–937 10. Meinzer U, Barreau F, Esmiol-Welterlin S et al (2012) Yersinia pseudotuberculosis effector YopJ subverts the Nod2/RICK/TAK1 pathway and activates caspase-1 to induce intestinal barrier dysfunction. Cell Host Microbe 11 (4):337–351 11. Jiang S, Yao J, Ma KW et al (2013) Bacterial effector activates jasmonate signaling by directly targeting JAZ transcriptional repressors. PLoS Pathog 9(10):e1003715 12. Falgarone G, Blanchard HS et al (1999) Coordinate involvement of invasin and Yop proteins in a Yersinia pseudotuberculosis-specific class I-restricted cytotoxic T cell-mediated response. J Immunol 162(5):2875–2883 13. Mukherjee S, Keitany G, Li Y et al (2006) Yersinia YopJ acetylates and inhibits kinase activation by blocking phosphorylation. Science 312(5777):1211–1214 14. Le Roux C, Huet G, Jauneau A et al (2015) A receptor pair with an integrated decoy converts pathogen disabling of transcription factors to immunity. Cell 161(5):1074–1088 15. Luo ZQ, Clemente TE, Farrand SK (2001) Construction of a derivative of Agrobacterium tumefaciens C58 that does not mutate to tetracycline resistance. Mol Plant-Microbe Interact 14(1):98–103

Chapter 3 Protein Acetylation in Pathogen Virulence and Host Defense: In Vitro Detection of Protein Acetylation by Radiolabeled Acetyl Coenzyme A Karl J. Schreiber and Jennifer D. Lewis Abstract Protein acetylation has emerged as a common modification that modulates multiple aspects of protein function, including localization, stability, and protein-protein interactions. It is increasingly evident that protein acetylation significantly impacts the outcome of host-microbe interactions. In order to characterize novel putative acetyltransferase enzymes and their substrates, we describe a simple protocol for the detection of acetyltransferase activity in vitro. Purified proteins are incubated with 14C-acetyl CoA and separated electrophoretically, and acetylated proteins are detected by phosphorimaging or autoradiography. Key words Protein acetylation, Radiolabeling, 14C-acetyl CoA, In vitro assay, Pathogen

1

Introduction Protein acetylation involves the transfer of an acetyl group (CH3CO) from acetyl coenzyme A (CoA) to a specific protein residue, as catalyzed by acetyltransferase enzymes [1]. These reactions can occur co- or posttranslationally and result in the covalent attachment of an acetyl group at either the α-amino group at the N-terminus of a protein or at lysine, serine, threonine, or histidine residues within a protein [2, 3]. N-terminal acetylation modulates protein function by influencing characteristics such as subcellular localization [4–6], protein-protein interactions [7, 8], protein folding [9], and protein stability [10–12]. The earliest identified and most widely studied example of acetylation at internal residues involves lysine residues of histone proteins [13]. This modification significantly influences chromatin accessibility [14], possibly through the recruitment of acetyllysine-binding chromatin remodeling enzymes [15] as well as altered histone-histone interactions resulting from acetylation at specific sites [16]. Additional known targets of internal residue acetylation include cytoskeletal proteins

Walter Gassmann (ed.), Plant Innate Immunity: Methods and Protocols, Methods in Molecular Biology, vol. 1991, https://doi.org/10.1007/978-1-4939-9458-8_3, © Springer Science+Business Media, LLC, part of Springer Nature 2019

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(actin and tubulin), transcription factors, chaperones, metabolic enzymes, and various regulatory proteins [17–20]. Given the range of physiological processes regulated by acetylation, it is perhaps not surprising that protein acetylation comprises an important biochemical interface for host-microbe interactions. Following pathogen infection, transcriptional changes in histone (de)acetyltransferase genes lead to shifts in global histone acetylation patterns that facilitate the coordinated expression of immunity-related genes [21, 22]. To counteract this response, many pathogens have evolved virulence-promoting molecules, termed effectors, that are secreted into host cells and act as suppressors of host immunity. For example, the fungal pathogen Cochliobolus carbonum produces the effector molecule HC-toxin, whose activity as an inhibitor of histone deacetyltransferases is required for virulence on maize [23, 24]. The effector protein PsAvh23 from the oomycete pathogen Phytophthora sojae disrupts the assembly of a histone acetyltransferase protein complex to modulate plant immunity [25]. In animal hosts, the Legionella pneumophila effector RomA methylates histones at specific residues in order to block their subsequent acetylation and thus suppress immunity-related changes in gene expression [26]. Finally, the effector SseF from Salmonella enterica interacts with and enhances the activity of the acetyltransferase TIP60, which is thought to alter patterns of histone acetylation and thus promote pathogen replication in host cells [27]. Direct acetyltransferase activity has been demonstrated for other effectors, which act on numerous nonhistone substrates in host cells. The Xanthomonas euvesicatoria effector AvrBsT acetylates the microtubule-associated protein ACIP1, which plays a significant yet currently undefined role in plant defense [28]. A more well-understood class of effector targets comprises nucleotide-binding leucine-rich repeat (NLR) proteins, which have evolved in plants to recognize pathogen effectors and elicit a strong immune response known as effector-triggered immunity (ETI) [29]. The effector HopZ3 from Pseudomonas syringae acetylates the NLR protein RPM1, an associated protein (RIN4), and the effector AvrB, all of which serve to suppress the recognition of AvrB by RPM1 [3]. Another effector from Pseudomonas syringae, HopZ1a, acetylates tubulin, causing destabilization of microtubules, disruption of the plant secretory pathway, and overall suppression of cell wall-based defense mechanisms [30]. HopZ1amediated acetylation of jasmonate ZIM domain transcriptional repressors also enhances pathogen virulence by activating jasmonic acid-induced signaling which suppresses salicylic acid-induced immunity against P. syringae [31]. The enzymatic activity of HopZ1a can, however, be recognized in plants when HopZ1a acetylates the pseudokinase ZED1 which, in complex with the protein ZAR1, results in the activation of ETI [32]. A similar dual

In Vitro Detection of Protein Acetylation

25

role in virulence/resistance is illustrated by the Ralstonia solanacearum effector PopP2, which promotes pathogen virulence by acetylating WRKY transcription factors as well as the WRKY domain of the NLR protein RRS1 [33–35]. In the presence of the NLR RPS4, RRS1 acetylation induces ETI. Effectors with acetyltransferase activity have also been identified in animal pathogens, including Vibrio parahaemolyticus (VopA), Salmonella enterica (AvrA), and various Yersinia species (YopJ/YopP), all of which target mitogen-activated protein kinase signaling pathways in order to suppress inflammatory host immune responses [2, 36–40]. Clearly, acetylation events significantly influence the outcome of host-microbe interactions. Methods for the detection of protein acetylation are generally dictated by the intended scope of analysis. Mass spectrometry enables global acetylation profiling [41, 42], which can be aided by prior anti-acetyllysine antibody-mediated immunopurification to enrich for proteins acetylated at lysines [20]. Specific acetylation sites on purified proteins or peptides can also be identified by mass spectrometry [3, 43] or nuclear magnetic resonance [44]. While these approaches yield detailed datasets, they require significant inputs of labor and equipment. When more qualitative results are desired, such as the determination of whether or not a protein of interest possesses acetyltransferase activity or acts as an acetyltransferase substrate, then more straightforward analyses are appropriate. Here, we describe a simple protocol for the detection of protein acetylation in vitro, based on the incorporation of radiolabeled acetyl CoA.

2

Materials 1. Purified proteins of interest (minimum concentration: 0.3 μg/ μL) in a solution buffered to pH 7–8 (see Note 1). 2. 5 acetylation reaction buffer (prepared fresh): 250 mM 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid (HEPES) pH 8.0 with KOH, 50% glycerol, 5 mM dithiotreitol, 5 μM phytate (optional) (see Note 2). 3.

14

C-acetyl CoA (see Note 3).

4. Heat block or incubator set to 30  C. 5. SDS polyacrylamide gel (continuous or discontinuous) made with an acrylamide concentration that will provide adequate resolution of the proteins of interest. 6. 10 SDS-PAGE loading buffer: 0.6 M Tris-HCl pH 6.8, 20% sodium dodecyl sulfate (SDS), 20% glycerol, 1 mM dithiothreitol, 2.9 M β-mercaptoethanol, 0.05% bromophenol blue.

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7. Coomassie stain: 40% methanol, 10% acetic acid, 0.1% Coomassie Brilliant Blue R-250. 8. Coomassie destain: 40% methanol, 10% acetic acid. 9. SDS-PAGE gel-fixing solution: 50% methanol, 10% acetic acid. 10. Amplify™ Fluorographic Reagent (GE Healthcare) (see Note 4). 11. Gel dryer. 12. Whatman 3 mm paper and plastic wrap. 13. X-ray film cassette with a phosphor screen. 14. Phosphorimager. 15. Platform shaker.

3

Methods

3.1 In Vitro Acetylation Assay

1. For each reaction, combine the following in a 20 μL volume: ~2 μg of acetyltransferase enzyme, ~2 μg of test substrate protein, 1 acetylation reaction buffer, 2 μL of 14C-acetyl CoA (see Note 2). 2. Incubate the reaction at 30  C for 1 h. 3. Add 2 μL of 10 SDS-PAGE loading buffer, and load the entire reaction onto an SDS-PAGE gel (see Note 5). 4. To verify input protein levels, load a separate SDS-PAGE gel with the same reactions as described in step 1 but only containing the proteins diluted in an appropriate buffer (i.e., lacking 14 C-acetyl CoA and acetylation reaction buffer). Run this gel in a separate electrophoresis apparatus to avoid 14C contamination, and visualize proteins by Coomassie staining (see Subheading 3.2). 5. For the gel containing the 14C-labeled samples, when adequate electrophoretic separation is achieved (see Note 6), remove the gel from the electrophoresis apparatus, and fix the proteins by soaking the gel in SDS-PAGE gel-fixing solution for 30 min at room temperature. 6. Transfer the gel to Amplify™ Fluorographic Reagent, and soak for 15 min at room temperature. 7. Dry the gel in a gel dryer, e.g., 2 h at 80  C (see Notes 7 and 8). 8. Wrap the gel in plastic wrap, and place it in a cassette with a phosphor screen (see Note 8). 9. Incubate the cassette at 20  C for an empirically determined length of time (see Note 9). 10. Survey the work area for radioactive contamination, and decontaminate as required (see Note 10).

In Vitro Detection of Protein Acetylation

27

Fig. 1 Demonstration of acetyltransferase activity for the Pseudomonas syringae effector HopZ1a, acting on the substrate ZED1. (Upper) Purified GST-HopZ1a or the catalytic mutant GST-HopZ1aC216A (70 kDa) and 6xHis-ZED1 (41 kDa) were incubated with phytate in the presence of 14C-acetyl CoA. The amount of 6xHis-ZED1 in the reactions ranged from 2.5 to 4.5 μg. Samples were separated on a 9% SDS polyacrylamide gel, and 14C incorporation was visualized by phosphorimaging. *, GST-HopZ1a or GST-HopZ1aC216A; **, His-ZED1. (Lower) Coomassie-stained gel of proteins. Reproduced from [32] with permission from the National Academy of Sciences

11. Visualize the phosphor screen using a phosphorimager. If necessary, clear the phosphor screen, and incubate the gel with the phosphor screen for more time (see Note 11). 12. For semiquantitative comparisons of samples, analyze digital images using the phosphorimager software (see Note 12). An example of a typical phosphorimage is shown in Fig. 1. 3.2 Coomassie Staining

1. Place the gel in a small tray, and add enough Coomassie stain to cover the gel. Incubate at room temperature with gentle shaking overnight (see Notes 13 and 14). 2. Decant the stain into a bottle for future reuse (see Note 14), and then add Coomassie destain solution to the gel. Incubate at room temperature with gentle shaking for several hours until background staining is sufficiently reduced. To accelerate destaining, include rolled up Kimwipes in the destain solution. 3. Place the gel in a sheet protector, and scan it in a standard flatbed scanner for a permanent record of the data.

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Notes 1. There are numerous potential avenues for the preparation of purified proteins for the acetyltransferase assay. Variables include the choice of protein expression vector, expression host, and expression conditions, all of which must be empirically optimized for maximal yield of soluble protein. As the recombinant proteins are separated by size, it is also important to consider the relative masses of the proteins of interest with their affinity tags. The conditions for protein purification will depend on the affinity tag included in the chosen expression vector and may also require empirical refinement to obtain protein samples with high degrees of purity [45, 46]. If the affinity tag is relatively large (e.g., glutathione S-transferase or maltose-binding protein) and is not cleaved from the protein of interest prior to the assay, it is important to include a control reaction with the affinity tag alone to evaluate whether this polypeptide influences the acetyltransferase reaction in some way. In addition, acetyltransferase activity is governed by a conserved catalytic triad (Cys-Asp-His, and occasionally Cys-Glu-His) that is similar to papain-like cysteine proteases [47–49]. A Cys-to-Ala mutant of a putative acetyltransferase will provide an important negative control to verify that acetyltransferase activity is responsible for any signals detected in the phosphorimage. 2. Several pathogen effectors are activated in the presence of phytate (inositol hexakisphosphate) as a mechanism for ensuring that these acetyltransferases are only activated once inside a eukaryotic host [30, 50]. Acetyltransferases of eukaryotic origin may not require phytate as a cofactor. If needed for this assay, a 10 mM stock of phytate can be filter-sterilized and stored as aliquots at 20  C. Serial dilutions of this stock should be used to achieve the final micromolar concentration. 3.

14

C-acetyl CoA is primarily supplied by PerkinElmer, Inc. in units of 50 μCi with a specific activity in the range of 40–60 mCi/mmol. According to the manufacturer, optimal storage conditions for 14C-acetyl CoA are 20  C, pH 5.0, under nitrogen. Note that this compound readily hydrolyzes at pH >8.5 and in strongly acidic solutions. To preserve chemical stability, the 14C-acetyl CoA sample should be thawed slowly at 4  C and aliquoted to avoid repeated freeze-thaw cycles. 3Hacetyl CoA can also be used in this assay. Commercially available stocks of 3H-acetyl CoA tend to have a higher specific activity (1–10 Ci/mmol) than 14C-acetyl CoA, but 14C has a higher energy and higher efficiency in liquid scintillation counting than 3H. These and other factors, such as the

In Vitro Detection of Protein Acetylation

29

materials permitted under one’s institutional radioactivity license, should be considered in selecting an appropriate radioisotope [51]. Use of radioactivity typically requires training by the institution to ensure that materials are handled in an appropriate manner. 4. Amplify™ Fluorographic Reagent is used to improve the detection efficiency of weak beta particle emitters such as 14C and thus reduce the exposure time required to observe a signal on a phosphor screen. 5. It is preferable that the loading dye and sample are not boiled, as boiling may volatilize the 14C. 6. To minimize contact with 14C, it is best to stop the gel when the dye front is approximately 0.5 cm from the bottom of the gel. The bottom part of the gel containing the dye front should be cut off and disposed of as solid radioactive waste, since it contains unincorporated 14C-acetyl CoA. 7. The gel should be dried on a piece of Whatman paper and covered with a piece of plastic wrap to avoid contaminating the gel dryer with radioactivity. While the wet gel can be placed on multiple pieces of Whatman paper to soak up any liquid, use of more than one piece of Whatman in the gel dryer should be avoided, as the gel may not dry. Alternatively, the gel may be air-dried in cellophane for ~1 week. However, the gel is more likely to crack using this method. 8. For a detailed discussion of autoradiography and phosphorimaging, please see reference [52]. 9. Depending on the signal intensity, exposure times will vary from days to weeks. 10.

14

C contamination is not detectable using a Geiger-Mu¨ller counter, so the workspace should be monitored by wipe surveys combined with liquid scintillation counting. Follow local regulations for the disposal of waste materials and liquids containing 14C.

11. Alternatively, X-ray film and an X-ray film developer can be used in place of the phosphorimager [52]. 12. If X-ray film is used, digital images of films can be analyzed using programs such as ImageJ to calculate relative signal intensities among different samples. 13. The gel tray should be covered with plastic wrap to minimize release of the pungent fumes generated by both the staining and destaining solutions. 14. For a more rapid protocol, the gel can be heated in a microwave for 10–20 s at both the staining and destaining steps and then incubated in the respective solutions for 15–20 min [53]. The staining solution can be reused multiple times.

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Acknowledgments We thank Dr. Mae¨l Baudin, Dr. Yuan Chen, and Jana Hassan for constructive feedback on the manuscript. Research on plant immunity in the Lewis laboratory was supported by the United States Department of Agriculture-Agricultural Research Service 203021000-046-00D (J.D.L), and NSF IOS-1557661 (J.D.L.). References 1. Drazic A, Myklebust LM, Ree R et al (2016) The world of protein acetylation. Biochim Biophys Acta Proteins Proteomics 1864:1372–1401 2. Mukherjee S, Keitany G, Li Y et al (2006) Yersinia YopJ acetylates and inhibits kinase activation by blocking phosphorylation. Science 312:1211–1214 3. Lee J, Manning AJ, Wolfgeher D et al (2015) Acetylation of an NB-LRR plant immuneeffector complex suppresses immunity. Cell Rep 13:1–13 4. Behnia R, Panic B, Whyte JRC et al (2004) Targeting of the Arf-like GTPase Arl3p to the Golgi requires N-terminal acetylation and the membrane protein Sys1p. Nat Cell Biol 6:405–413 5. Setty SRG, Strochlic TI, Tong AHY et al (2004) Golgi targeting of Arf-like GTPase Arl3p requires its Nα-acetylation and the integral membrane protein Sys1p. Nat Cell Biol 6:414–419 6. Murthi A, Hopper AK (2005) Genome-wide screen for inner nuclear membrane protein targeting in Saccharomyces cerevisiae: roles for N-acetylation and an integral membrane protein. Genetics 170:1553–1560 7. Scott DC, Monda JK, Bennett EJ et al (2011) N-terminal acetylation acts as an avidity enhancer within an interconnected multiprotein complex. Science 334:674–678 8. Monda JK, Scott DC, Miller DJ et al (2013) Structural conservation of distinctive N-terminal acetylation-dependent interactions across a family of mammalian NEDD8 ligation enzymes. Structure 21:42–53 9. Holmes WM, Mannakee BK, Gutenkunst RN et al (2014) Loss of N-terminal acetylation suppresses a prion phenotype by modulating global protein folding. Nat Commun 5:4383 10. Kuo HP, Lee DF, Chen CT et al (2010) ARD1 stabilization of TSC2 suppresses tumorigenesis through the mTOR signaling pathway. Sci Signal 3:ra9

11. Varshavsky A (2011) The N-end rule pathway and regulation by proteolysis. Protein Sci 20:1298–1345 12. Xu F, Huang Y, Li L et al (2015) Two N-terminal acetyltransferases antagonistically regulate the stability of a Nod-like receptor in Arabidopsis. Plant Cell 27:1547–1562 13. Allfrey VG, Faulkner R, Mirsky AE (1964) Acetylation and methylation of histones and their possible role in the regulation of RNA synthesis. Proc Natl Acad Sci U S A 315:786–794 14. Eberharter A, Becker PB (2002) Histone acetylation: a switch between repressive and permissive chromatin. EMBO Rep 3:224–229 15. Swygert SG, Peterson CL (2014) Chromatin dynamics: interplay between remodeling enzymes and histone modifications. Biochim Biophys Acta Gene Regul Mech 1839:728–736 16. Ye J, Ai X, Eugeni EE et al (2005) Histone H4 lysine 91 acetylation: a core domain modification associated with chromatin assembly. Mol Cell 18:123–130 17. Glozak MA, Sengupta N, Zhang X et al (2005) Acetylation and deacetylation of non-histone proteins. Gene 363:15–23 18. Kim SC, Sprung R, Chen Y et al (2006) Substrate and functional diversity of lysine acetylation revealed by a proteomics survey. Mol Cell 23:607–618 19. Spange S, Wagner T, Heinzel T et al (2009) Acetylation of non-histone proteins modulates cellular signalling at multiple levels. Int J Biochem Cell Biol 41:185–198 20. Choudhary C, Kumar C, Gnad F et al (2009) Lysine acetylation targets protein complexes and co-regulated major cellular functions. Science 325:834–840 21. Xu J, Xu H, Liu Y et al (2015) Genome-wide identification of sweet orange (Citrus sinensis) histone modification gene families and their expression analysis during the fruit development and fruit-blue mold infection process. Front Plant Sci 6:1–16

In Vitro Detection of Protein Acetylation 22. DeFraia CT, Wang Y, Yao J et al (2013) Elongator subunit 3 positively regulates plant immunity through its histone acetyltransferase and radical S-adenosylmethionine domains. BMC Plant Biol 13:102 23. Brosch G, Ransom R, Lechner T et al (1995) Inhibition of maize histone deacetylases by HC toxin, the host-selective toxin of Cochliobolus carbonum. Plant Cell 7:1941–1950 24. Ransom RF, Walton JD (1997) Histone hyperacetylation in maize in response to treatment with HC-toxin or infection by the filamentous fungus Cochliobolus carbonum. Plant Physiol 115:1021–1027 25. Kong L, Qiu X, Kang J et al (2017) A Phytophthora effector manipulates host histone acetylation and reprograms defense gene expression to promote infection. Curr Biol 27:981–991 26. Rolando M, Sanulli S, Rusniok C et al (2013) Legionella pneumophila effector RomA uniquely modifies host chromatin to repress gene expression and promote intracellular bacterial replication. Cell Host Microbe 13:395–405 27. Wang X, Li D, Qu D et al (2010) Involvement of TIP60 acetyltransferase in intracellular Salmonella replication. BMC Microbiol 10:228 28. Cheong MS, Kirik A, Kim JG et al (2014) AvrBsT acetylates Arabidopsis ACIP1, a protein that associates with microtubules and is required for immunity. PLoS Pathog 10: e1003952 29. Schreiber KJ, Baudin M, Hassan JA et al (2016) Die another day: Molecular mechanisms of effector-triggered immunity elicited by type III secreted effector proteins. Sem Cell Dev Biol 56:124–133 30. Lee AHY, Hurley B, Felsensteiner C et al (2012) A bacterial acetyltransferase destroys plant microtubule networks and blocks secretion. PLoS Pathog 8:e1002523 31. Jiang S, Yao J, Ma KW et al (2013) Bacterial effector activates jasmonate signaling by directly targeting JAZ transcriptional repressors. PLoS Pathog 9:e1003715 32. Lewis JD, Lee AHY, Hassan JA et al (2013) The Arabidopsis ZED1 pseudokinase is required for ZAR1-mediated immunity induced by the Pseudomonas syringae type III effector HopZ1a. Proc Natl Acad Sci U S A 110:18722–18727 33. Tasset C, Bernoux M, Jauneau A et al (2010) Autoacetylation of the Ralstonia solanacearum effector PopP2 targets a lysine residue essential for RRS1-R-mediated immunity in Arabidopsis. PLoS Pathog 6:e1001202

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34. Le Roux C, Huet G, Jauneau A et al (2015) A receptor pair with an integrated decoy converts pathogen disabling of transcription factors to immunity. Cell 161:1074–1088 35. Sarris PF, Duxbury Z, Huh SU et al (2015) A plant immune receptor detects pathogen effectors that target WRKY transcription factors. Cell 161:1089–1100 36. Trosky JE, Mukherjee S, Burdette DL et al (2004) Inhibition of MAPK signaling pathways by VopA from Vibrio parahaemolyticus. J Biol Chem 279:51953–51957 37. Trosky JE, Li Y, Mukherjee S et al (2007) VopA inhibits ATP binding by acetylating the catalytic loop of MAPK kinases. J Biol Chem 282:34299–34305 38. Jones RM, Wu H, Wentworth C et al (2008) Salmonella AvrA coordinates suppression of host immune and apoptotic defenses via JNK pathway blockade. Cell Host Microbe 3:233–244 39. Orth K, Xu Z, Mudgett MB et al (2000) Disruption of signaling by Yersinia effector YopJ, a ubiquitin-like protein protease. Science 290:1594–1597 40. Paquette N, Conlon J, Sweet C et al (2012) Serine/threonine acetylation of TGFβ-activated kinase (TAK1) by Yersinia pestis YopJ inhibits innate immune signaling. Proc Natl Acad Sci U S A 109:12710–12715 41. Li Y, Silva JC, Skinner ME et al (2013) Mass spectrometry-based detection of protein acetylation. Methods Mol Biol 1077:81–104 42. Zhang K, Tian S, Fan E (2013) Protein lysine acetylation analysis: current MS-based proteomic technologies. Analyst 138:1628 43. Manning AJ, Lee J, Wolfgeher DJ et al (2018) Simple strategies to enhance discovery of acetylation post-translational modifications by quadrupole-orbitrap LC-MS/MS. Biochim Biophys Acta Proteins Proteomics 1866:224–229 44. Smet-Nocca C, Wieruszeski JM, Melnyk O et al (2010) NMR-based detection of acetylation sites in peptides. J Pept Sci 16:414–423 45. de Marco A (2011) Strategies for boosting the accumulation of correctly folded recombinant proteins expressed in Escherichia coli. Methods Mol Biol 752:1–15 46. Wingfield PT (2016) Overview of the purification of recombinant proteins. Curr Protoc Protein Sci 80:6.1.1–6.1.35 47. Sim E, Payton M, Noble M et al (2000) An update on genetic, structural and functional studies of arylamine N-acetyltransferases in eucaryotes and procaryotes. Hum Mol Genet 9:2435–2441

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48. Sandy J, Mushtaq A, Holton SJ et al (2005) Investigation of the catalytic triad of arylamine N-acetyltransferases: essential residues required for acetyl transfer to arylamines. Biochem J 390:115–123 49. Kubiak X, De L, Sierra-Gallay IL, Chaffotte AF et al (2013) Structural and biochemical characterization of an active arylamine N-acetyltransferase possessing a non-canonical cys-his-glu catalytic triad. J Biol Chem 288:22493–22505 50. Mittal R, Peak-Chew SY, Sade RS et al (2010) The acetyltransferase activity of the bacterial toxin YopJ of Yersinia is activated by eukaryotic

host cell inositol hexakisphosphate. J Biol Chem 285:19927–19934 51. Krauser JA (2013) A perspective on tritium versus carbon-14: Ensuring optimal label selection in pharmaceutical research and development. J Label Compd Radiopharm 56:441–446 52. Voytas D, Ke N (1999) Detection and quantitation of radiolabeled proteins and DNA in gels and blots. Curr Protoc Mol Biol 48A.3: A.1–A.10 53. Brunelle JL, Green R (2014) Coomassie blue staining. Methods Enzymol 541:161–167

Chapter 4 A Novel Assay Based on Confocal Microscopy to Test for Pathogen Silencing Suppressor Functions Mustafa Adhab and James E. Schoelz Abstract In plants, RNA silencing is an important mechanism for gene regulation and defense that is targeted by proteins of viral pathogens effecting silencing suppression. In this chapter we describe a new assay to probe silencing suppressor activity using Agrobacterium infiltration of Nicotiana benthamiana and confocal microscopy. The key element in this assay involves the use of a reporter construct that is transiently expressed at a much lower level than free GFP, and this increases the sensitivity of detection of weak silencing suppressors such as the P6 protein of Cauliflower mosaic virus. Although initially developed for virus silencing suppressors, this technique could also prove valuable to characterize the potential for weak silencing suppressors in the effector repertoires of fungi, bacteria, nematodes, and oomycetes. Key words Gene silencing, Virus silencing suppressors, Agroinfiltration, Confocal microscopy, Pathogen effectors, N. benthamiana

1

Introduction Posttranscriptional gene silencing is an evolutionarily conserved process in eukaryotic organisms that targets double-stranded RNAs for degradation [1, 2]. In plants, RNA silencing plays an important role in gene regulation and is also considered to form the initial line of defense against viral pathogens. To counteract this host defense system, most, if not all, plant viruses carry a protein capable of silencing suppression [1, 2]. The first plant virus protein shown to have silencing suppressor activity was HC-Pro of Tobacco etch virus [3, 4]. Since this discovery, several assays have been developed to identify silencing suppressors, and other categories of virus proteins in addition to HC-Pro have been shown to have the capacity for suppression of gene silencing [1, 2]. The most common method for the identification of virus silencing suppressors has involved a transient expression assay developed for Nicotiana benthamiana [1, 5, 6]. In this assay, Agrobacterium tumefaciens is transformed with a binary

Walter Gassmann (ed.), Plant Innate Immunity: Methods and Protocols, Methods in Molecular Biology, vol. 1991, https://doi.org/10.1007/978-1-4939-9458-8_4, © Springer Science+Business Media, LLC, part of Springer Nature 2019

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agrobacterium vector carrying a putative virus silencing suppressor. A second A. tumefaciens isolate carrying a reporter construct such as GFP in a binary vector is also prepared. Typically, both the GFP construct and the virus silencing suppressor are expressed under the control of a strong promoter such as the 35S promoter of Cauliflower mosaic virus (CaMV). When Agrobacterium carrying only the GFP construct is infiltrated into the leaves of N. benthamiana through a technique called agroinfiltration, the GFP gene is expressed at a high level approximately 3–5 days postinfiltration (dpi), and GFP protein expression can be visualized in a darkroom with the use of a handheld UV lamp. Beginning at approximately 5 dpi, the host’s gene silencing machinery starts to target the GFP mRNA for degradation, and GFP expression starts to fade until it is completely extinguished around 8–10 dpi. By contrast, when an Agrobacterium isolate carrying the GFP binary plasmid is mixed with Agrobacterium carrying a virus silencing suppressor in a binary vector and they are co-agroinfiltrated into N. benthamiana leaves, GFP expression remains strong for 17–20 dpi, essentially the life of the leaf. This transient expression assay has worked well in the characterization of a number of silencing suppressors, including the potyvirus HC-Pro protein, the P19 protein of Tomato bushy stunt virus (TBSV), and the coat protein (CP) of Turnip crinkle virus [6]. However, the agroinfiltration assay has not worked for all virus suppressors, including the 2b protein of Cucumber mosaic virus, P25 of Potato virus X, the CP of Citrus tristeza virus [1], and P6 of Cauliflower mosaic virus. Consequently, alternate methods had to be developed to identify these virus silencing suppressors. Figure 1 illustrates a typical agroinfiltration experiment involving agroinfiltration of leaf panels of Agrobacterium carrying a GFP gene alone, co-agroinfiltration of a second leaf panel with an Agrobacterium isolate carrying GFP plus an Agrobacterium isolate carrying TBSV P19, and co-agroinfiltration of a third leaf panel with an Agrobacterium isolate carrying GFP and CaMV P6, either tagged with RFP (P6-RFP) or untagged (P6). The leaves were illuminated with a UV lamp and photographed at 8 dpi. At this timepoint, only the leaf panel co-agroinfiltrated with GFP and P19 expresses GFP. Neither CaMV P6 nor P6-RFP proteins are able to sustain a level of GFP expression that could be detected by the UV lamp. A second approach that has been used to identify silencing suppressors involves a genetic cross between a transgenic Arabidopsis line expressing a putative silencing suppressor and a second transgenic line that carries a silenced reporter gene. In this assay, a virus silencing suppressor is identified through its capacity to restore the expression of the reporter gene by blocking the host’s gene silencing machinery. This approach was first used to demonstrate that the potyvirus HC-Pro protein was a silencing suppressor [3, 4] and has also been used to characterize other virus silencing

Confocal Assay for Silencing Suppressor Function

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Fig. 1 A silencing suppressor assay conducted in whole leaves of N. benthamiana. Agrobacteria carrying GFP, P19 + GFP, P6 + GFP, or P6-RFP + GFP were agroinfiltrated into leaf panels. GFP expression was illuminated with a black-light UV lamp. Photo was taken at 8 dpi

suppressors including the P6 protein of CaMV [7]. However, the drawback of this approach is that it is very time-consuming, as it takes months to first create transgenic plants that express a putative silencing suppressor and then to cross the plants with a silenced reporter gene. In this chapter, we present a method involving transient expression in N. benthamiana together with confocal microscopy. In our system, we utilize a reporter construct that is transiently expressed at a much lower level than free GFP, and this increases the sensitivity of detection of weak silencing suppressors such as P6 of CaMV. The silencing reporter construct we developed for this assay consists of the movement protein (MP) of CaMV tagged at its C-terminus with GFP (MP-GFP) [8]. Figure 2 illustrates the differences in the expression levels of free GFP versus MP-GFP as assessed by confocal microscopy at 3, 5, and 8 dpi. Interestingly, free GFP expression can still be observed with a confocal microscope at 8 dpi (Fig. 2) when it appears to be completely extinguished at the whole-plant level (Fig. 1). By contrast MP-GFP is nearly extinguished at 5 dpi (as assessed by confocal microscopy) and is completely silenced at 8 dpi. The approach we describe here is also appropriate for stronger silencing suppressors such as HC-Pro, P19, and CP of Turnip crinkle virus. Although initially developed for virus silencing suppressors, this technique could also prove valuable for characterizing novel but potentially weak silencing suppressors in the effector repertoires of fungi, bacteria, nematodes, and oomycetes.

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Fig. 2 Confocal images of leaves agroinfiltrated with GFP vs. MP-GFP in the absence of a silencing suppressor. These images illustrate the contrasting levels of expression of GFP vs. MP-GFP at 10 magnification. The constructs GFP and MP-GFP are both expressed from the CaMV 35S promoter, and the images were taken at 3, 5, and 8 dpi

2

Materials Prepare all solutions with ultrapure water and analytical grade reagents.

2.1 Bacteria and Plasmids

1. The A. tumefaciens strain we have used for our experiments is AGL1 [9], although any strain would likely be sufficient. 2. A binary vector in which the target gene is expressed from the 35S promoter. We have used pKYLX7 [10] or the pSITE vector series [11].

2.2

Plants

2.3 Agroinfiltration and Induction

1. Seeds of Nicotiana benthamiana (see Note 1). Seeds can be obtained from the United States Tobacco Germplasm Collection at North Carolina State University. N. benthamiana is selffertile, so seed can be collected from an individual plant to create a seed stock for future experiments. 1. Modified Luria-Bertani (LB) liquid medium: To 1 L of water, add 16 g of tryptone, 10 g of yeast extract, and 5 g NaCl. Autoclave for 25 min at 121  C. Store at room temperature. 2. Modified LB solid medium: To 1 L modified LB liquid medium (Subheading 2.3, step 1), add 15 g agar. Autoclave for 25 min at 121  C. Cool to below 60  C before adding appropriate antibiotics, and then pour the solution into sterile, polystyrene Petri dishes (100 mm  15 mm). Use a Bunsen

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burner to gently flame the surface before the agar solidifies to remove any bubbles and to surface sterilize. Store at 4  C. 3. Induction medium: To 1 L of water, add 3.9 g MES (0.39%), 20 g sucrose (2%), 10 g glucose (1%). Adjust pH to 5.4 with 1 M KOH. Store at 4  C. 4. Acetosyringone (3.5-dimethoxy-4-hydroxyacetophenone): Dissolve in N,N-dimethylformamide to a final concentration of 0.2 M. Store at 20  C (see Note 2). 5. Antibiotic kanamycin (Kan) 100 stock solution: 50 mg/mL kanamycin in ultrapure water (see Note 3). 6. Antibiotic spectinomycin (Spec) 100 stock solution: 100 mg/mL in ultrapure water (see Note 3). 7. Shaker incubator for growth of bacterial cultures. 8. Centrifuge for pelleting Agrobacterium cultures. 2.4 Confocal Imaging

1. The confocal microscope we have used is the Leica model TCP SP8 MP, but any other model should give similar results. The Leica is an inverted spectral confocal microscope with fixed visible laser lines (405–514 nm), tunable white light laser (470–670 nm), three HyD and two PMT detectors, resonant scanner, and Mai Tai DeepSee multiphoton laser tunable to 680–1060 nm for deep tissue imaging.

2.5 Confocal Image Analysis

1. Microscope slides and cover slips for confocal microscopy.

3 3.1

2. We use the following parameters for excitation/emission filter wavelengths: 488 nm/501–530 for GFP, 543/565–615 nm for RFP.

Methods Plant Growth

1. To break dormancy, treat N. benthamiana seeds with commercial bleach at half strength (2.6% NaOCl [v/v]) for 30 min. Distribute seeds on the surface of a commercial potting medium (a peat-based medium is preferred). 2. Thin after germination, and transplant into individual 6-inch pots at approximately 4 weeks. Fertilize plants with a top dressing of Osmocote (14-14-14) at transplanting. 3. Grow N. benthamiana plants in the greenhouse year-round, without the use of supplemental lights or in growth chambers. 4. Agroinfiltrate N. benthamiana plants at an age of 6–8 weeks after seeding. For best results, use the topmost, fully expanded leaves for agroinfiltration (see Note 4).

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Agroinfiltration

1. Beginning with a glycerol stock that has been stored at 80  C, streak out the agrobacterium strain that carries the binary plasmids of interest onto LB agar plates that have the appropriate antibiotics. Allow the agrobacterium isolate to grow at 28  C for 2–3 days (see Note 5). 2. Using a sterile loop, pick a single colony from the plate, and transfer to a 15 mL screw cap tube containing 3 mL of LB broth containing the appropriate antibiotics. Incubate in a shaker incubator at 28  C and 220 rpm until cells reach stationary phase at 1 or 2 days. 3. Transfer 400 μL of the bacterial culture to an Erlenmeyer flask containing 40 mL LB broth containing the appropriate antibiotics. Incubate in a shaker incubator at 28  C and 220 rpm overnight. 4. Transfer the 40 mL culture into a new sterile 50 mL polypropylene conical tube, and close the cap tightly. Pellet the cells by centrifugation for 8 min at 14,000  g. 5. Discard the supernatant, and wash the pellet twice with an equal volume of sterile ultrapure water (see Note 6). Gently resuspend the pellet into 40 mL of agroinfiltration medium, and transfer to an Erlenmeyer flask. Add 40 μL of the 0.2 M acetosyringone. Incubate in a shaker incubator at 28  C and 220 rpm overnight (see Note 7). 6. Adjust the OD600 of the Agrobacterium solution to 1.0. Mix the Agrobacterium carrying the putative silencing suppressor construct in a 1:1 ratio with the reporter construct and the reporter construct at a 1:1 ratio with Agrobacterium carrying the empty binary vector. 7. For agroinfiltration, fill a 3.0 mL or 5.0 mL syringe with the agrobacterium solution. Apply the barrel of the syringe (without the needle) to the abaxial surface of the leaf and gently infiltrate the agrobacterium solution into the plant tissue. 8. Image with confocal microscopy between 2 and 8 dpi.

3.3 Confocal Microscopy Imaging

1. Examine leaves infiltrated with the reporter construct alone daily from 2 to 8 dpi to establish the baseline for gene silencing. Include a second set of leaves infiltrated with a 1:1 mixture of the reporter construct and silencing suppressor to assess the activity of the silencing suppressor. Collect samples for confocal microscopy with a 1.0 cm cork borer. Leaves should be left on the plant to allow for multiple samples to be taken from the same leaf (see Note 8). 2. Initially examine samples with a confocal microscope (see Note 9) at low magnification (10) under bright-field conditions to avoid any potential for bias. Once the epidermal layer is

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brought into focus, then examine the tissue for the presence of the reporter gene fusion construct (see Note 9). Collect at least five images to be analyzed for the spatial distribution of the fluorescent protein. Collect images for leaves agroinfiltrated with the reporter construct by itself as well as leaves agroinfiltrated with the reporter construct plus putative silencing suppressor (see Note 10). 3.4 Confocal Image Analysis

1. Save each image as a TIFF or JPEG file after acquisition from the confocal microscope. 2. Insert each image into a PowerPoint slide, and adjust image size to fill the entire slide. 3. For spatial analysis, on a separate PowerPoint slide, draw a grid of 100 squares. This grid can be copied and overlaid onto each image. Figure 3 illustrates the results at one timepoint for leaves infiltrated with GFP alone, as well as leaves infiltrated with MP-GFP and P6-RFP. 4. Count the number of grids that contain a GFP or RFP signal, and express as a percentage of 100 (see Note 11). Table 1 illustrates typical results that we have obtained in an analysis of the silencing suppressor function P6-RFP coupled with the reporter construct MP-GFP over a period of 8 days.

4

Notes 1. This assay has only been attempted in N. benthamiana, a plant species that is unusually adapted to agroinfiltration and virus silencing suppressor assays. 2. The acetosyringone solution is made up at a 1000 concentration. It is added to the agroinfiltration induction media immediately before the addition of Agrobacterium. It is acceptable to serially transfer the Agrobacterium isolate to a fresh plate rather than repeatedly initiating the culture from a 80 stock. 3. The choice of antibiotics is dependent on the binary vector used in the transient agroinfiltration assay. We have listed the antibiotic concentrations that we frequently use in our agroinfiltration experiments that are appropriate for our binary vector. 4. The age of the plants and the positioning of the leaves are critical for obtaining maximum expression of constructs upon agroinfiltration. We recommend using plants that are between 6 and 8 weeks of age, but plant growth conditions will vary depending on the time of year, so a strict adherence based on the age of the plant from the seeding date may not always be predictive for optimum agroinfiltration conditions. Generally,

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Fig. 3 Expression of free GFP or MP-GFP plus P6-RFP at 5 dpi after agroinfiltration into N. benthamiana leaf tissue, at 10 magnification. An analysis of MP-GFP expressed by itself reveals expression of GFP in only 11 of 100 squares. By contrast, MP-GFP is observed in 87 of 100 squares when co-agroinfiltrated with P6RFP, which itself is observed in 100/100 squares Table 1 Spatial distribution of MP-GFP signal in confocal images in the presence or absence of P6-RFP Treatment

Signal

2 DPI

3 DPI

5 DPI

8 DPI

MP-GFP alone

GFP

36  11

41  22

8  10

45

MP-GFP + P6RFP

GFP

96  5

91  7

82  3

77  18

RFP

96  6

96  4

97  2

91  7

Each of the values presented in the table is the mean of five observations, collected as in Fig. 3

Confocal Assay for Silencing Suppressor Function

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high levels of expression can be obtained in N. benthamiana before the first flowers develop on the plant. Expression levels and ease of infiltration will vary from the top of the plant to the bottom. Smaller, immature leaves may yield high expression levels but can be resistant to the agroinfiltration technique. Conversely, the bottom, fully developed leaves are easy to infiltrate, but may have lower expression levels and should be avoided. Consequently, we choose the top most, fully expanded leaves on the plant. Ease of infiltration may vary depending on the environmental conditions, but under ideal conditions, it should be possible to agroinfiltrate an entire leaf with 5–10 infiltration points on the leaf. In general, wellwatered plants are the easiest to infiltrate. We find it is good to soak the pots in a tray of water for 30–40 min before infiltration. In using multiple plants, it is important to use leaves of comparable age. 5. The agar plate containing the Agrobacterium culture may be stored for up to 1 week. As the culture on the plate ages, it will take longer for the Agrobacterium to become established in the 3 mL culture, so it is important to transfer cultures to fresh plates on a regular basis. 6. It is important to remove traces of LB broth from the Agrobacterium pellet, because in our experience, residual LB broth can induce necrosis in agroinfiltrated leaf tissues. 7. An 8 h incubation is sufficient, but it is also acceptable to initiate this step at the end of the day and to agroinfiltrate the next morning, an incubation of 14–15 h. 8. Maximum expression of the CaMV MP-GFP and MP-RFP constructs is typically observed at 3 dpi, and a significant degree of silencing occurs by 5 dpi. By 8 dpi, the constructs are usually almost completely silenced (Fig. 2). The key to this step is the selection of a reporter construct that is expressed in agroinfiltrated tissue at several orders of magnitude less than the free fluorescent protein. In our experience, it has not been difficult to find these reporter constructs for our silencing assay. In fact, all of the plant and virus proteins that we have tagged with GFP (or RFP) are expressed at a much lower level than the free fluorescent proteins. 9. It is important to first examine agroinfiltrated tissues under bright-field conditions to remove any potential for bias in the analysis of the spatial expression of the reporter construct or in examining the effect of a putative silencing suppressor on the spatial expression of the reporter construct. This step ensures that microscopy fields under examination are truly randomized. In addition, the examination under low magnification allows for a more accurate evaluation of spatial expression patterns over a larger area.

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10. In the development of this silencing assay, we used a reporter construct tagged with GFP (MP-GFP), and the P6 silencing suppressor of CaMV was tagged at its C-terminus with RFP (P6-RFP) [12]. We anticipate that many other reporter constructs in addition to MP-GFP would also be expressed at low levels relative to free GFP and would also be appropriate for this assay. It is not necessary to tag a putative silencing suppressor for this assay, but the use of P6-RFP has been useful for verifying the expression of the silencing suppressor within the tissue expressing the reporter. We have been able to show that an untagged version of P6 also functions as a silencing suppressor in this assay, as well as untagged versions of the P19 protein of TBSV, the coat protein of TCV, and the HC-Pro protein of TEV. 11. Although a visual count of signals within grids is a low-tech approach, it is remarkably reproducible, as images analyzed by three individuals in our lab have yielded similar results. References 1. Qu F, Morris J (2005) Suppressors of RNA silencing encoded by plant viruses and their role in viral infections. FEBS Lett 579:5958–5964. https://doi.org/10.1016/j. febslet.2005.08.041 2. Voinnet O (2005) Induction and suppression of RNA silencing: insights from viral infections. Nat Rev Genet 6:206–220. https://doi.org/ 10.1038/nrg1555 3. Anandalakshmi R, Pruss GJ, Ge X et al (1998) A viral suppressor of gene silencing in plants. Proc Natl Acad Sci U S A 95:13079–13084. https://doi.org/10.1073/pnas.95.22.13079 4. Kasschau KD, Carrington JC (1998) A counter-defensive strategy of plant viruses: suppression of posttranscriptional gene silencing. Cell 95:461–470. https://doi.org/10. 1016/S0092-8674(00)81614-1 5. Johansen LK, Carrington JC (2001) Silencing on the spot. Induction and suppression of RNA silencing in the Agrobacterium-mediated transient expression system. Plant Physiol 126:930–938. https://doi.org/10.1104/pp. 126.3.930 6. Moissiard G, Voinnet O (2004) Viral suppression of RNA silencing in plants. Mol Plant Pathol 5:71–82. https://doi.org/10.1111/j. 1364-3703.2004.00207.x 7. Love AJ, Laird J, Holt J et al (2007) Cauliflower mosaic virus protein P6 is a suppressor

of RNA silencing. J Gen Virol 88:3439–3444. https://doi.org/10.1099/vir.0.83090-0 8. Rodriguez A, Angel CA, Lutz L et al (2014) Association of the P6 protein of Cauliflower mosaic virus with plasmodesmata and plasmodesmal proteins. Plant Physiol 166:1–14. https://doi.org/10.1104/pp.114.249250 9. Lazo GR, Stein PA, Ludwig RA (1991) a DNA transformation-competent Arabidopsis genomic library in Agrobacterium. Biotechnology 9:963–967. https://doi.org/10.1038/ nbt1091-963 10. Schardl CL, Byrd AD, Benzion G et al (1987) Design and construction of a versatile system for the expression of foreign genes in plants. Gene 61:1–11 11. Chakrabarty R, Banerjee R, Chung SM et al (2007) pSITE vectors for stable integration or transient expression of autofluorescent protein fusions in plants: probing Nicotiana benthamiana-virus interactions. Mol Plant-Microbe Interact 20:740–750. https://doi.org/10. 1094/MPMI-20-7-0740 12. Angel CA, Lutz L, Yang X et al (2013) The P6 protein of Cauliflower mosaic virus interacts with CHUP1, a plant protein which moves chloroplasts on actin microfilaments. Virology 443:363–374. https://doi.org/10.1016/j. virol.2013.05.028

Chapter 5 Quantification of Extracellular ATP in Plant Suspension Cell Cultures Sowmya R. Ramachandran, Sonika Kumar, and Kiwamu Tanaka Abstract Extracellular ATP functions as an important signaling molecule in both plants and animals. In plants, ATP is released in the extracellular region of cells in response to environmental perturbations, such as herbivory, cellular damage, or other abiotic and biotic stimuli, which is then perceived by the purinoceptor P2K1 as a damaged-self signal for activation of defense responses. Given its involvement in various physiological processes, quantification of extracellular ATP is important for further understanding of its molecular function. In this chapter, we describe a method for the accurate and reliable determination of extracellular ATP concentrations in plant cell culture media based on the luciferase-luciferin reaction, using either end-point or real-time detection assays. The protocol can be easily performed with any luminometer within 1 h after sample collection. Although we use Arabidopsis suspension cells, the protocol described can be optimized for any cell type. Key words Extracellular ATP, Luciferase-luciferin reaction, Arabidopsis suspension cells, Real-time ATP measurement, End-point ATP measurement

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Introduction Inside the cell, adenosine triphosphate (ATP) is the primary energy currency that fuels cellular processes. In addition, ATP serves as a cofactor and an allosteric regulator of many enzymatic reactions (e.g., for kinases and adenylyl cyclase) and as a component of nucleic acids. In the extracellular region, ATP serves a different function. The first report on the extracellular function of purines comes from studies of heart muscle contractions [1]. Subsequently, extracellular ATP was proposed to function as a purinergic neurotransmitter [2]. Since then, numerous studies have demonstrated that modulation of extracellular ATP is associated with growth regulation, neurotransmission, muscle contraction, and inflammation [3–6]. In plants, the first report of extracellular ATP came from a study on the Venus flytrap in which the rapid movement of the trap

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closure was stimulated upon ATP application [7]. Further studies have demonstrated the involvement of this signal molecule in plant growth, development, and response to stress [8–12]. Very recently, extracellular ATP was reported as an essential signal for plant defense responses [13, 14]. ATP is released into the apoplast via exocytosis and plasma membrane-localized transporters, i.e., an adenine nucleotide transporter and ABC transporters [15–17], upon mechanical stimuli and cellular damage caused by abiotic and biotic stresses [18–20]. In animals, perception of ATP in the extracellular milieu is accomplished by purinergic 2 (P2) receptors that are present on the cell surface [21, 22]. The P2 receptors fall into two classes: the ligand-gated ion channel (P2X) and the G protein-coupled receptor (P2Y) [23, 24]. Plants lack the canonical purinoceptors seen in mammals but perceive extracellular ATP instead through the P2 receptor kinase (P2K), which contains an extracellular legume-type lectin domain [25, 26]. Those receptors are expressed in a spatiotemporal manner and can possess different ligand-binding affinities based on their specific functions. Given the importance of extracellular ATP in cellular signaling, quantification of extracellular ATP is imperative to further our understanding of cellular responses following external stimuli. Traditionally ATP, ADP, and AMP levels have been measured through reverse phase, ion-exchange or ion-pairing HPLC [27]. These techniques are occasionally irreproducible and time-consuming because of the complicated quantitative analysis procedures, and also require large sample sizes [28]. Furthermore, the optimum range of quantification with this method is limited to picomoles [29]. In contrast, a luciferase-luciferin-based enzymatic detection system offers a more sensitive method with the capability to detect concentrations as low as femto- to attomolar levels [30]. The enzymatic reaction catalyzed by luciferase proceeds in the presence of magnesium ions, O2, and ATP. The catalytic reaction of the substrate D-luciferin leads to the excited intermediate D-luciferyl-adenylate that is oxidized into oxyluciferin and emits photons of yellow-green light upon decaying to the stable ground state, as shown in the following equation. ATP þ D‐Luciferin þ O2 ! Oxyluciferin þ AMP þ Pyrophosphate þ CO2 þ Light ð560 nmÞ

The amount of light emitted is proportional to the concentration of ATP in a wide dynamic range. Unlike fluorescence assays that require an external light source, the bioluminescence reaction eliminates nonspecific background signal interference. Thus, detection of ATP using the luciferase-based luminescence provides a reliable, sensitive detection method that can be applied both to in vitro and in vivo cellular systems as well as to different compartments within the cell.

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In this chapter, we describe a method employing two different quantification assays, end-point and real-time detections, for measuring the extracellular ATP concentration in suspension cell cultures. This is a convenient system in terms of simplicity and has a high rate of reproducibility. Cell cultures also provide a clean system for ATP measurement with no interference from other tissue types or epiphytic microorganisms. Although this protocol focuses on Arabidopsis suspension cells, the method is applicable to other tissue types and plant species.

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Materials

2.1 Plant Material and Growth Conditions

1. Arabidopsis suspension cell culture, PSB-D obtained from ABRC https://abrc.osu.edu/ (Stock# CCL84840). 2. Murashige and Skoog minimal organic (MSMO) medium: 4.43 g Murashige and Skoog basal salts, 30 g sucrose, 50 μL of 1 mg/mL zeatin, and 500 μL of 1 mg/mL NAA per 1 L. Adjust the pH to 5.7 with 1 M KOH. Autoclave the medium at 20 psi and 121  C for 20 min, and store the cooled medium at 4  C. 3. Sterile 250 mL conical flasks. 4. Aluminum foil. 5. 6-well cell culture plates. 6. Incubator shaker at 23  C. 7. Laminar flow hood.

2.2 Reagents for Stimulation of ATP Release

1. Salt stress solutions: NaCl 200, 20, and 2 mM prepared in deionized water. 2. Osmotic stress solutions: sorbitol 200, 20, and 2 mM prepared in deionized water. 3. Solvent stimulant solutions: ethanol 0.01, 0.1, and 1% (v/v) and DMSO 0.01, 0.1, and 1% (v/v).

2.3 ATP Measurement

1. Luciferase/luciferin reagent (ENLITEN, Promega). Reconstitute the luciferase/luciferin reagent according to the manufacturer’s instructions. Aliquot the reagent into vials containing 1 mL of reagent, and freeze at 20  C until use (see Notes 1 and 2). One hour before beginning the assay, remove the desired amount of reagent from the 20  C freezer, and keep it at room temperature (see Note 3). 2. ATP stock solution: 100 mM (Sigma). 3. ATP standards dissolved in MSMO medium (see Note 4). 4. Cell strainers with 40 μm pore size.

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5. 96-well cell culture white microplate that is compatible with your luminometer, for example, Greiner Bio-One. 6. Luminometer, for example, multimode plate reader Enspire 2300 (Perkin Elmer). 7. 15 mL sterile conical tubes. 8. Hot water bath or heat block set to 96  C. 9. Aluminum foil. 10. 96-well rubber seals.

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Methods

3.1 End-Point Measurement of Extracellular ATP in Arabidopsis Suspension Cell Culture

In this section, we describe the protocol for the end-point estimation of ATP from suspension cell-conditioned media using the luciferase-luciferin reaction assay. Here, we use salt stress and osmotic stress as representative stimulants to induce release of extracellular ATP into the suspension cell medium. After elicitation, a cell strainer is inserted into the individual wells, and the culture medium is gently collected to minimize ATP release due to mechanical damage of the cells (Fig. 1). Next, the samples are boiled at 96  C for 2 min to inactivate any ATP-hydrolyzing enzymes in the extracellular medium and transferred to a plate reader with a luminometer. The luciferase/luciferin reagent is added to each well using an automatic injector, and the luminescence signal is recorded for a period of 5 s. This method can be used to measure ATP from extracellular fluids and enzymatic reactions.

3.1.1 Plant Material and Growth Conditions

1. Grow Arabidopsis suspension cells in MSMO medium at 23  C with shaking at 150 rpm in the dark. 2. Cell passaging: In the laminar flow hood, transfer 5 mL of 7-day-old suspension cells into a sterile 250 mL conical flask containing 45 mL of MSMO medium. Cover the flask with aluminum foil to keep the cells in the dark. Incubate the cells at 25  C with shaking at 150 rpm. Passage the cells every 7 days to maintain a fresh batch of cells. 3. Transfer 300 μL of 7-day-old cells to each well of a 6-well cell culture plate containing 3 mL of MSMO medium (see Note 5). Seal the plates with parafilm to avoid evaporation of the liquid medium. Cover with aluminum foil and incubate in dark at 23  C, 150 rpm. Cells can be used for ATP assay 4–5 days after subculturing in the cell culture plates. 4. Remove 6-well culture plates from the shaker incubator, and leave them undisturbed on the bench for 2 h before treating cells with a stimulant.

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Fig. 1 Setup for an end-point measurement of extracellular ATP concentrations in sampled cell culture media using a luminometer. (a) Flow diagram showing the different steps involved in ATP measurement. (b) Cell culture plate and a cell strainer. (c) Sampling setup: (left to light) the cell culture plate, samples kept on ice, and a typical 96-well white plate used for ATP measurement 3.1.2 Sample Collection and Preparation

1. Add MSMO medium with 200 mM NaCl or 200 mM sorbitol, and measure the ATP induced over a period of 1 h. The same protocol can be used to measure the ATP response to other stimuli with appropriate controls. 2. To obtain samples, gently place the cell strainers into the wells of the 6-well culture plate without disturbing the cells (Fig. 1) (see Note 6). 3. Pipette 120 μL of supernatant samples from within the cell strainer at different time intervals, and transfer the samples to a 96-well PCR plate placed on ice (see Note 7). 4. Boil the samples in the 96-well plate at 96  C for 5 min to denature ATP-degrading enzymes (see Note 8). Cool to room temperature, and store the samples at 20  C until proceeding to ATP measurements.

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5. Dilute the treated cell suspension 10, by mixing 15 μL of the sample with 135 μL of deionized water. Use 50 μL of this diluted sample for ATP quantification. 3.1.3 ATP Quantification

1. Transfer the reconstituted luciferase/luciferin reagent into a 15 mL sterile tube, and place the injector tubing in the reagent. Fill the injector tubing in the luminometer by following instrument manufacturer’s protocol (see Note 9). 2. Prepare ATP standards from 1 nM to 5 μM in a solution with composition identical to the cell culture media containing the stimulant (see Notes 4 and 10). 3. Transfer 50 μL of the sample and the ATP standard solutions to a 96-well white plate, and insert the plate into a plate reader (see Note 11). 4. Program the instrument to automatically inject the luciferase/ luciferin reagent into each well using the “by well” protocol of the instrument. 5. Immediately after the injection of the reagent, measure relative light units (RLUs) for each well for a period of 5 s. Measure each sample in duplicate, and average the RLUs of the two replicates.

3.1.4 Calculations

1. Plot the values for the ATP standards in a scatter plot graph (Fig. 2). 2. Add a trendline and obtain an equation for the line. 3. Convert the value of the y-value (RLU value) to the corresponding value for the ATP concentration (i.e., the x-value). Repeat these calculations for both the treatment and mock. 4. Calculate standard errors using enough replicates for each treatment (n  6 is recommended). 5. Draw a graph using the calculated average value of ATP concentration  standard errors. Different chemicals used as ATP stimulants were tested and found to have a small effect on the luciferase/luciferin activity (Fig. 3). Upon application of salt stress (200 mM NaCl) and osmotic stress (200 mM sorbitol), there was a significant increase in extracellular ATP concentration (Fig. 4a, b).

3.2 Real-Time Measurement of Extracellular ATP in Arabidopsis Suspension Cell Culture

In this section, we describe a method for real-time detection of extracellular ATP from cell cultures which allows precise detection of dynamic changes in ATP concentrations. Briefly, cells are transferred to individual wells of a 96-well cell culture microplate and acclimatized for 2 h. Next, the luciferase/luciferin reagent is added, and the plate is inserted into the luminometer. The desired

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Fig. 2 Typical standard curve of ATP (100 pM to 100 nM) prepared in MSMO medium and measured using a luminometer

Fig. 3 Direct effect of commonly used chemical stimuli on the luciferase activity. TCA was prepared in water, while all other chemicals were prepared in MSMO medium

stimulant is added to each well, and RLUs are measured at specific intervals. 1. Add 100 μL of cell suspension to individual wells of a 96-well white plate. 2. Keep the plate undisturbed in dark for 2 h to overcome transfer shock. 3. Add 10 μL of luciferase/luciferin reagent to each well without exposing the reagent or wells to light.

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Fig. 4 End-point measurement of extracellular ATP after NaCl addition. Arabidopsis PSB-D cells were treated with or without 200 mM NaCl (a) and 200 mM sorbitol (b), and the ATP concentration in each culture medium was measured at different time intervals as shown

4. Transfer 2.0 M sorbitol solution into a 15 mL falcon tube, and insert injector tubing into the solution. Fill the injector tubing of the luminometer as per instrument recommendations (see Note 9). 5. Inject 10 μL of sorbitol, and immediately measure the signal from each well using the “by plate” protocol option on the instrument. 6. Continue measuring all three wells at 10 s intervals, for a period of 15 min or desired amount of time changing the measurement intervals as needed. 7. For control, inject deionized water (or mock solution), and immediately measure the signal from each well as mentioned in steps 4–6.

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Fig. 5 Real-time measurement of extracellular ATP after addition of sorbitol. Arabidopsis PSB-D cells were treated with or without 200 mM sorbitol, and the ATP response was measured at 10 s intervals for 15 min (N ¼ 3)

8. Obtain a standard curve as described in Subheadings 3.1.3 and 3.1.4. 9. Use the data of the ATP standards for calculations as directed under Subheading 3.1.4. A significant increase in extracellular ATP was detected upon treatment with 200 mM (final concentration) sorbitol in the realtime assay (Fig. 5). A peak in ATP response was detected at about 3 min after application of the stimulant, which gradually declined over 15 min. Overall the ATP released was significantly higher in treatment vs. mock. This protocol provides a simple method to detect ATP release in response to a number of stimuli in a highthroughput manner.

4

Notes 1. Protect the luciferase/luciferin reagent from light and excessive handling/shaking at all times. The reagent loses activity with multiple freeze-thaw cycles and with long periods of storage at 4  C. Keep the reagent at 20  C until use, and only thaw the required amount before the assay. 2. Avoid contaminating the reagent with ATP from external sources. Make sure to wear gloves, and use clean microcentrifuge tubes/plates and a clean surface while handling the reagent.

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3. The luciferase/luciferin reagent performs optimally at room temperature. Check the manufacturer’s recommendations before use. Make sure the reagent is at room temperature before proceeding with the assay. 4. ATP standards should be prepared in the stimuli-containing media in case that the stimulant interferes with the activity of the reagent. Different stimuli such as NaCl can reduce the activity of the luciferase enzyme (Fig. 3). Prepare the ATP standards in a solution with an appropriate concentration of stimuli for accurate extrapolation of results. 5. Other multi-well plates (12- or 24-well plates) may be used if necessary with minor modifications to the protocol. If using a 12-well plate, use a ratio of 1.15 mL:100 μL media/cells. If using a 24-well culture plate, use a ratio of 750 μL:100 μL media:cells. These combinations were found to have similar numbers of cells per mL to 6-well culture plates at 5 days post subculturing. 6. Aspirating cells can lead to high luminescence readings due to a touch response. Avoid disturbing the cells while collecting the supernatant as much as possible. 7. Apyrases or other enzymes may degrade or consume ATP. Store samples on ice until they have been boiled. 8. Boiling denatures extracellular apyrases and other enzymes that degrade ATP. Alternatively, trichloroacetic acid (TCA) can be used to prevent further degradation of the ATP. However, as shown in Fig. 3, TCA drastically reduces the luciferase activity, even at concentrations as low as 0.005% (v/v). 9. Calculate the amount of reagent required along with an extra 0.5 mL for filling the injector tubing. The amount required to fill the tubing may vary from instrument to instrument. If using a different injector instrument, determine the amount of liquid required to fill the tubing before proceeding with the assay. 10. Some components of the media can interfere with the assay by reducing enzyme activity. Check the background RLU of the medium before proceeding. 11. Since the reagent’s capacity to measure ATP changes with time, include ATP standards for each luminescence measurement for the most accurate calculations.

Acknowledgments This project was supported by the National Science Foundation (grant no. IOS-1557813) and USDA NIFA (Hatch project 1015621). PPNS No. 0758, Department of Plant Pathology,

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College of Agriculture, Human and Natural Resource Sciences, Agricultural Research Center, Hatch Project No. WNP00008 and WNP00833, Washington State University, Pullman, WA, 991646430, USA. References 1. Drury AN, Szent-Gyo¨rgyi A (1929) The physiological activity of adenine compounds with special reference to their action upon the mammalian heart. J Physiol 68:213–237 2. Burnstock G (1972) Purinergic nerves. Pharmacol Rev 24:509–581 3. Burnstock G, Williams M (2000) P2 purinergic receptors: modulation of cell function and therapeutic potential. J Pharmacol Exp Ther 295:862–869 4. Cekic C, Linden J (2016) Purinergic regulation of the immune system. Nat Rev Immunol 16:177–192 5. Khakh BS, Burnstock G (2009) The double life of ATP. Sci Am 301:84–92 6. Pavenstadt H, Gloy J, Leipziger J et al (1993) Br J Pharmacol 109:953–959 7. Jaffe MJ (1973) The role of ATP in mechanically stimulated rapid closure of the Venus flytrap. Plant Physiol 51:17–18 8. Lew RR, Dearnaley JDW (2000) Extracellular nucleotide effects on the electrical properties of growing Arabidopsis thaliana root hairs. Plant Sci 153:1–6 9. Reichler SA, Torres J, Rivera AL et al (2009) Intersection of two signalling pathways: extracellular nucleotides regulate pollen germination and pollen tube growth via nitric oxide. J Exp Bot 60:2129–2138 10. Steinebrunner I, Wu J, Sun Y et al (2003) Disruption of apyrases inhibits pollen germination in Arabidopsis. Plant Physiol 131:638–1647 11. Tanaka K, Gilroy S, Jones AM et al (2010) Extracellular nucleotide signaling in plants. Trends Cell Biol 20:601–608 12. Tanaka K, Choi J, Cao Y et al (2014) Extracellular ATP acts as a damage associated molecular pattern (DAMP) signal in plants. Front Plant Sci 5:466 13. Tripathi D, Zhang T, Koo AJ et al (2018) Extracellular ATP acts on jasmonate signaling to reinforce plant defense. Plant Physiol 176:511–523 14. Tripathi D, Tanaka K (2018) A crosstalk between extracellular ATP and JA signaling pathways. Plant Signal Behav 13: e1432229

15. Thomas C, Rajagopal A, Windsor B et al (2000) A role for ectophosphatase in xenobiotic resistance. Plant Cell 12:519–533 16. Kim SY, Sivaguru M, Stacey G (2006) Extracellular ATP in plants. Visualization, localization, and analysis of physiological significance in growth and signaling. Plant Physiol 142:984–992 17. Rieder B, Neuhaus HE (2011) Identification of an Arabidopsis plasma membrane-located ATP transporter important for anther development. Plant Cell 23:1932–1944 18. Dark A, Demidchik V, Richards SL et al (2011) Release of extracellular purines from plant roots and effect on ion fluxes. Plant Signal Behav 6:1855–1857 19. Wu SJ, Liu YS, Wu JY (2008) The signaling role of extracellular ATP and its dependence on Ca2+ flux in elicitation of Salvia miltiorrhiza hairy root cultures. Plant Cell Physiol 49:617–624 20. Weerasinghe RR, Swanson SJ, Okada SF et al (2009) Touch induces ATP release in Arabidopsis roots that is modulated by the heterotrimeric G-protein complex. FEBS Lett 583:2521–2526 21. Webb TE, Simon J, Krishek BJ et al (1993) Cloning and functional expression of a brain G-protein-coupled ATP receptor. FEBS Lett 324:219–225 22. Lustig KD, Shiau AK, Brake AJ et al (1993) Expression cloning of an ATP receptor from mouse neuroblastoma cells. Proc Natl Acad Sci U S A 90:5113–5117 23. Ralevic V, Burnstock G (1998) Receptors for purines and pyrimidines. Pharmacol Rev 50:413–492 24. Abbracchio MP, Burnstock G, Boeynaems JM et al (2006) International Union of Pharmacology LVIII: update on the P2Y G proteincoupled nucleotide receptors: from molecular mechanisms and pathophysiology to therapy. Pharmacol Rev 58:281–341 25. Choi J, Tanaka K, Cao Y et al (2014a) Identification of a plant receptor for extracellular ATP. Science 343:290–294 26. Choi J, Tanaka K, Liang Y et al (2014b) Extracellular ATP, a danger signal, is recognized by

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DORN1 in Arabidopsis. Biochem J 463: 429–437 27. Miyatake K, Sakuraba H, Kitaoka S (1987) Separation and determination of adenine nucleotides by high-performance liquid chromatography. Agric Biol Chem 51:253–255 28. Khlyntseva SV, Bazel’ YR, Vishnikin AB et al (2009) Methods for determination of adenosine triphosphate and other adenine. J Anal Chem 64:657

29. Bhatt DP, Chen X, Geiger JD et al (2012) A sensitive HPLC-based method to quantify adenine nucleotides in primary astrocyte cell cultures. J Chromatogr B Analyt Technol Biomed Life Sci 889-890:110–115 30. Fitzgerald RS, Shriahata M, Chang I et al (2009) The impact of hypoxia and low glucose on the release of acetylcholine and ATP from the incubated cat carotid body. Brain Res 1270:39–44

Chapter 6 Measuring Pectin Properties to Track Cell Wall Alterations During Plant–Pathogen Interactions Gerit Bethke and Jane Glazebrook Abstract Plant cell walls act both as a barrier to pathogen entry and as a source of signaling molecules that can modulate plant immunity. Cell walls consist mainly of three polymeric sugars: cellulose, pectin, and hemicellulose (Mohnen et al., Biomass Recalcitrance: deconstructing the plant cell wall for bioenergy, 2008). In Arabidopsis more than 50% of the primary cell wall is pectin (Zablackis et al., Plant Physiol 107:1129–1138, 1995). There are various types of pectin, but all pectins contain galacturonic acid subunits in their backbone (Harholt et al., Plant Physiol 153:384–395, 2010; Mohnen, Curr Opin Plant Biol 11:266–277, 2008). Many pathogens secrete pectin-degrading enzymes as part of their infection strategy (Espino et al., Proteomics 10:3020–3034, 2010; ten Have et al., Mol Plant-Microbe Interact 11:1009–1016, 1998). Pectin is synthesized in a highly esterified fashion and is de-esterified in the cell wall by pectin methylesterases (Harholt et al., Plant Physiol 153:384–395, 2010; Mohnen, Curr Opin Plant Biol 11:266–277, 2008). During plant–pathogen interactions, both the amount and the patterns of pectin methylesterification in the wall can be altered (Bethke et al., Plant Physiol 164:1093–1107, 2014; Lionetti et al., J Plant Physiol 169:1623–1630, 2012). Pectin methylesterifications influence mechanical properties of pectin, and pectins must be at least partially de-methylesterified to be substrates for pectin-degrading enzymes (Levesque-Tremblay et al., Planta 242:791–811, 2015). Additionally, alterations of pectin methylesterification or pectin content affect pathogen growth (Bethke et al., Plant Physiol 164:1093–1107, 2014; Lionetti et al., J Plant Physiol 169:1623–1630, 2012; Bethke et al., Plant Cell 28:537–556, 2016; Raiola et al., Mol Plant-Microbe Interact 24:432–440, 2011; Vogel et al., Plant Cell 14:2095–2106, 2002; Vogel et al., Plant J 40:968–978, 2004; Wietholter et al., Mol Plant-Microbe Interact 16:945–952, 2003). This chapter explains a simple protocol that can be used in any molecular biology laboratory to estimate total pectin content using a colorimetric assay and pectin composition using antibodies raised against specific pectin components. Key words Plant immunity, Cell wall, Pectin, Methylesterification, Galacturonic acid, Alcohol-insoluble residue

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Introduction Many methods exist to study basic cell wall properties. We and others have previously described methods to measure pectin methylesterase activity [1–4]. Here we describe one possible method to extract crude cell walls (alcohol-insoluble residue)

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(Subheading 3.1) [1, 5], to extract pectin from cell wall extracts (Subheading 3.2) [1, 6, 7], a plate-based assay to measure total uronic acid as a proxy for total pectin content (Subheading 3.3) [8–10] and a dot blot procedure to probe cell wall extracts with antibodies raised against specific pectin components (Subheading 3.4) [1, 8]. Advanced methods that give more in depth structural information of cell wall composition are available, but these require specialized equipment and are not described here [11–14].

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Materials For Subheading 3.1 “Extraction of Crude Cell Walls”: 1. 70% aqueous ethanol: 70:30 v/v ethanol/water. 2. Chloroform/methanol: 1:1 v/v. 3. Acetone. 4. Microcentrifuge. 5. Freeze dryer. 6. Centrifuge tubes. 7. Pipette tips. For Subheading 3.2 “Pectin Extraction 1,2-Diaminocyclohexanetetraacetic Acid (CDTA)”:

Using

1. CDTA extraction buffer: 50 mM Tris-base, 50 mM 1,2-diaminocyclohexanetetraacetic acid (CDTA). Adjust pH to 7.2 using hydrogen chloride. 2. Centrifuge tubes (see Note 2). 3. Pipette tips. 4. Water bath. 5. Optional: freeze dryer. For Subheading 3.3 “Measure Uronic Acid Concentration”: 1. 4 M sulfamic acid in water. 2. 120 mM sodium tetraborate in concentrated sulfuric acid. Add sodium tetraborate to concentrated sulfuric acid by mixing overnight in a glass container using a magnetic stirrer in a fume hood. Use this solution within 1 week. 3. M-hydroxydiphenyl reagent: Dissolve m-hydroxydiphenyl in dimethyl sulfoxide at 100 mg/mL. Add 100 μL of this to 4.9 mL 80% sulfuric acid (80:20 v/v concentrated sulfuric acid/water) just before use. 4. D-(þ)-galacturonic acid. 5. Flat bottom microtiter plate.

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6. Plate seal. 7. Water bath or incubator. 8. Plate reader. For Subheading 3.4 “Probing Dot Blots with Pectin Specific Antibodies”: 1. Nitrocellulose membrane (0.45 μm). 2. Milk powder (e.g., Nestle Carnation™ nonfat). 3. 1 PBS buffer: To 800 mL of distilled water add 8 g sodium chloride, 0.2 g potassium chloride, 1.44 g sodium phosphate dibasic (Na2HPO4), 0.24 g potassium phosphate monobasic (KH2PO4). Adjust the pH to 7.4 using hydrochloric acid and add distilled water to 1 L total volume. 4. Primary antibody raised against cell wall component of interest. Please reference the antibody collection from the Complex Carbohydrate Center at the University of Georgia (https:// www.ccrc.uga.edu/~mao/wallmab/Home/Home.php) or from Paul Knox’s lab at the University of Leeds (http://www. plantprobes.net/index.php). We used LM 8, LM 19, and LM 20 antibodies that preferentially bind xylogalacturonan, unesterified, and esterified homogalacturonan, respectively [15]. 5. Secondary antibody, e.g., goat anti-Rat HRP conjugated antibody (Bethyl A110-105P). 6. Enhanced chemiluminescence kit (e.g., Amersham ECL Prime Western Blotting Detection Reagent from GE). 7. Film, developer, fixer, or charged coupled device (CCD) camera system to detect ECL signal.

3

Methods

3.1 Extraction of Crude Cell Walls (Alcohol-Insoluble Residue (AIR))

1. Optional: Move 3–4 weeks old Arabidopsis plants into the dark for 24–48 h to reduce starch. 2. Harvest rosette leaves and grind tissue (see Note 1). 3. Freeze dry and transfer 60–70 mg of the freeze-dried tissue into a 2 mL tube. 4. Add 1.5 mL of 70% aqueous ethanol and vortex thoroughly. 5. Centrifuge at 10,000  g for 10 min to pellet the alcoholinsoluble residue. Discard the supernatant. 6. Repeat step 5 once more. 7. In the fume hood, add 1.5 mL of chloroform/methanol (1:1 v/v) solution to the residue and shake the tube thoroughly to resuspend the pellet. Centrifuge at 10,000  g for 10 min and discard the supernatant.

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8. Repeat step 7 once or twice more until the supernatant is colorless. 9. Resuspend the pellet in 500 μL of acetone. Centrifuge at 10,000  g for 10 min. Discard the supernatant. 10. Air dry the pellet overnight in the fume hood. This is the AIR. 3.2 Pectin Extraction Using 1,2-Diaminocyclohexanetetraacetic Acid (CDTA)

1. Resuspend crude cell walls (AIR) in CDTA extraction buffer at 100 μL per 1 mg AIR. 2. Put in a 95  C water bath for 15 min; vortex every 5 min (see Note 2). 3. Centrifuge at 10,000  g for 10 min. 4. Remove the supernatant, which contains the pectin. Use this solution directly to measure uronic acid concentration or for dot blot analysis. 5. If the pectin extract needs to be stored, freeze dry the supernatant. It can be stored at 80  C for several months.

3.3 Measuring Uronic Acid Concentration

1. Dilute the pectin extract (Subheading 3.2, step 4) with water (1:5 v/v) (see Note 3). 2. Perform all subsequent steps in the fume hood. Sulfuric acid is very corrosive. Follow proper laboratory safety guidelines and discard waste according local regulations. Glass containers work best; the microtiter plates we used were polypropylene which is appropriate for short term exposure. 3. In a flat bottom microtiter plate, mix 36 μL of diluted pectin extract with 4 μL of 4 M sulfamic acid. 4. Add 200 μL of sulfuric acid containing 120 mM sodium tetraborate; seal with a plate seal and incubate at 80  C for 1 h (see Note 4). 5. Cool samples on ice and measure optical density at 490–525 nm in a plate reader (see Note 5). 6. Add 40 μL of m-hydroxydiphenyl reagent and mix carefully. 7. Measure the optical density at 490–525 nm. 8. Subtract the optical density before m-hydroxydiphenyl reagent addition from the optical density measured after addition of the dye. 9. Calculate the concentration of uronic acid using known amounts of D-(þ)-galacturonic acid as a standard. Use a standard curve with at least five different concentrations ranging from 0.1 to 2 mM galacturonic acid.

3.4 Probing Dot Blots with Pectin-Specific Antibodies

1. Serially dilute pectin solutions (Subheading 3.2, step 4) with CDTA extraction buffer. We usually dilute either 1:3, 1:9, 1:27 or 1:10, 1:20 depending on the antibody. You may need to determine the best dilution for your samples and antibodies.

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2. Spot 1 μL of the diluted pectin solutions on nitrocellulose membranes. 3. Dry the membranes overnight. Membranes can be kept for a few days prior to antibody treatment. 4. Block with 5% milk in 1 PBS for 1–1.5 h. 5. Probe membranes with cell wall component-specific antibody. For LM 8, LM 19, and LM 20 dilute antibodies 1:500 in 5% milk powder in 1 PBS and incubate at room temperature for 1 h (see Note 6). 6. Wash the membrane with 1 PBS three times for 10 min each time. 7. Probe the membrane with a secondary antibody for 1 h. For LM 8, LM 19, and LM 20 use goat anti-Rat HRP conjugated antibody (Bethyl A110-105P) diluted 1:5000 in 5% milk powder in 1 PBS. 8. Wash membranes with 1 PBS three times for 10 min each time. 9. Develop dot blots using an ECL system according to the manufacturer’s instructions. 10. Visualize using film or a CCD camera that can detect the ECL signal.

4

Notes 1. Grinding of tissue can be done using a mortar and pestle or using ball bearings and a paint shaker or similar machine. 2. Use lid locks, safe-lock tubes, or screw cap tubes that can withstand 95  C as some centrifuge tubes might open or deform at high temperatures. Other groups have done the extraction process at room temperature for 4 h [16]. 3. Subsequent dilution may be necessary if extensive browning is observed. Large quantities of sugar will cause browning after addition of sulfuric acid. 4. Do not submerge the plate in water. We wrap the sealed plate in aluminum foil crimping the edges to face upward and place the wrapped microtiter plate in the water bath so the water only reaches about 50% of the plate height. 5. Optimal optical density is at 525 nm. We used a plate reader with a 490 nm filter, which resulted in sufficient signal for the analysis. 6. LM 8, LM 19, and LM 20 were diluted 1:500 in 5% milk powder in 1 PBS. The best dilution will need to be tested for each antibody used. Some antibodies require BSA instead of milk or a different secondary antibody. Check the specifications for the antibody selected.

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Acknowledgments This work was supported by funds from the University of Minnesota and by National Science Foundation award IOS-1353854 to J.G. References 1. Bethke G, Grundman RE, Sreekanta S et al (2014) Arabidopsis PECTIN METHYLESTERASEs contribute to immunity against Pseudomonas syringae. Plant Physiol 164:1093–1107 2. Downie B, Dirk LM, Hadfield KA et al (1998) A gel diffusion assay for quantification of pectin methylesterase activity. Anal Biochem 264:149–157 3. Grsic-Rausch S, Rausch T (2004) A coupled spectrophotometric enzyme assay for the determination of pectin methylesterase activity and its inhibition by proteinaceous inhibitors. Anal Biochem 333:14–18 4. Wolf S, Grsic-Rausch S, Rausch T et al (2003) Identification of pollen-expressed pectin methylesterase inhibitors in Arabidopsis. FEBS Lett 555:551–555 5. Gille S, Hansel U, Ziemann M et al (2009) Identification of plant cell wall mutants by means of a forward chemical genetic approach using hydrolases. Proc Natl Acad Sci U S A 106:14699–14704 6. Zablackis E, Huang J, Muller B et al (1995) Structure of plant cell walls. 34. Characterization of the cell wall polysaccharides of Arabidopsis thaliana leaves. Plant Physiol 107:1129–1138 7. Bethke G, Glazebrook J (2014) Cyclohexane diamine tetraacetic acid (CDTA) extraction of plant cell wall Pectin. Bio-protocol 4:e1357 8. Bethke G, Thao A, Xiong GY et al (2016) Pectin biosynthesis is critical for cell wall integrity and immunity in Arabidopsis thaliana. Plant Cell 28:537–556

9. Filisetti-Cozzi TM, Carpita NC (1991) Measurement of uronic acids without interference from neutral sugars. Anal Biochem 197:157–162 10. van den Hoogen BM, van Weeren PR, LopesCardozo M et al (1998) A microtiter plate assay for the determination of uronic acids. Anal Biochem 257:107–111 11. Mohnen D (2008) Pectin structure and biosynthesis. Curr Opin Plant Biol 11:266–277 12. Anderson CT, Wallace IS, Somerville CR (2012) Metabolic click-labeling with a fucose analog reveals pectin delivery, architecture, and dynamics in Arabidopsis cell walls. Proc Natl Acad Sci U S A 109:1329–1334 13. Mort AJ, Qiu F, Maness NO (1993) Determination of the pattern of methyl esterification in pectin - distribution of contiguous nonesterified residues. Carbohydr Res 247:21–35 14. Naran R, Chen GB, Carpita NC (2008) Novel rhamnogalacturonan I and arabinoxylan polysaccharides of flax seed mucilage. Plant Physiol 148:132–141 15. Verhertbruggen Y, Marcus SE, Haeger A et al (2009) An extended set of monoclonal antibodies to pectic homogalacturonan. Carbohydr Res 344:1858–1862 16. Moller I, Marcus SE, Haeger A et al (2008) High-throughput screening of monoclonal antibodies against plant cell wall glycans by hierarchical clustering of their carbohydrate microarray binding profiles. Glycoconj J 25:37–48

Chapter 7 Method to Study Dynamics of Membrane-Bound Plant Transcription Factors During Biotic Interactions in Tomato Supriyo Chowdhury, Payel Bhattacharjee, Shrabani Basak, Shreya Chowdhury, and Pallob Kundu Abstract Sequestration of a transcription factor in a cellular membrane and releasing it on demand is an additional layer of gene regulation that is considered a rapid mode to reprogram a gene expression cascade when a plasma membrane stress signal is perceived. Better understanding of the dynamic exchange of membranebound transcription factors (MTFs) during biotic stress requires the development of a simple, efficient, and quick assay system. Here we report an Agrobacterium-based transient transformation method to assay the localization of fluorescent protein-tagged MTFs in tomato leaf epidermal peels that are subsequently infected with a pathogenic fungus. Essentially, our method mimics natural infection and facilitates the realistic monitoring of MTF movement during activation of a signaling event. Key words Membrane-bound transcription factor, Sonication, Transient transformation, Signaling, Fungus, Stress, Leaf epidermal peel

1

Introduction Membrane-bound transcription factors (MTFs) in plants are rapidly gaining attention for their involvement in diverse biological pathways and induction of an alternative mode of gene regulation [1]. A conservative estimate suggests that over 10% of all transcription factors, belonging to diverse groups, are membrane-bound in Arabidopsis [2]. This fact further signifies that MTF-based genome-wide gene regulation is essential and underscores the importance of determining its exact biological relevance. The majority of MTFs are anchored to a cellular membrane in an inactive form via a small transmembrane (TM) domain. Protease or proteasome-mediated liberation of a MTF from the membrane and subsequent translocation into the nucleus is the key regulatory process. An important aspect of functional characterization of MTFs is monitoring the dynamics of subcellular localization under different physiological and pathological conditions. Thus,

Walter Gassmann (ed.), Plant Innate Immunity: Methods and Protocols, Methods in Molecular Biology, vol. 1991, https://doi.org/10.1007/978-1-4939-9458-8_7, © Springer Science+Business Media, LLC, part of Springer Nature 2019

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comprehensive knowledge about the molecular interactions during this process requires identification of the precise intracellular position of the transcription factor. However, the locations of most MTFs have not been experimentally determined, and the databases are largely populated with predicted information [3, 4]. A convenient system to characterize the intracellular distribution would confirm the predicted data in many cases, clearly elucidate the functions of TM domains, and greatly facilitate in deciphering the biological relevance of a MTF. Commonly used methods include immunolocalization, subcellular fractionation followed by immunoblot analysis, and use of fluorescent protein-tagged constructs. While immunolocalization is thought to be advantageous because of specificity, sensitivity, and possibility of multiplexing, its high cost, exploitation of animals for raising antibodies, and probable cross-reactivity within a target transcription factor family prevent wide-scale use of this technique for MTF localization in the plant. Subcellular fractionation, although used as a confirmatory test for membrane association of MTF, does not reveal the exact details of membrane release dynamics and requires a specific antibody. In contrast, use of fluorescent protein (FP)-based constructs in conjunction with Agrobacteriumbased transient transformation to monitor protein subcellular localization is a simpler, less expensive, and reproducible methodology which can be adapted for many plants. The present protocol provides a FP-tagging-based method to visualize the localization of MTFs under normal physiological and pathogen stress conditions in tomato. We employ tomato leaf epidermal peels as a convenient tool for transient Agrobacteriummediated delivery of FP-tagged NAC [no apical meristem (NAM), Arabidopsis thaliana transcription activation factor (ATAF1/2), and cup-shaped cotyledon (CUC2)] MTF into the plant cell to study intracellular trafficking dynamics under biotic stress. To enhance Agrobacterium-mediated transformation, we incorporate mild sonication of the epidermal peel while immersed in an Agrobacterium suspension [5], commonly referred to as sonicationassisted Agrobacterium-mediated transformation (SAAT) [6], into our protocol. Enhanced transformation rates using SAAT probably result from micro-wounding of plant tissues caused by ultrasonic cavitations in the liquid medium [7]. Transformed tissues are then treated with pathogenic Alternaria solani, the causal organism of early blight disease of tomato, to simulate a biotic stress condition. Confocal microscopy fluorescent protein imaging analysis is used to visualize the localization of the FP-tagged NAC MTF. This straightforward protocol provides clear images in microscopic analysis and reproducible data.

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Materials Prepare all solutions using ultrapure deionized water obtained by purifying deionized water to attain a resistivity of 18.2 MΩ cm at 25
C. Use analytical or molecular biology grade reagents. Follow safety precautions and waste disposal protocols while preparing reagents or carrying out experiments. Autoclave solutions, if needed, and media at 121
C under 100 kPa pressure for 15–20 min. For filter sterilization, if required, use 0.22 μm syringe filters.

2.1

Plant Material

1. Grow tomato plants (Solanum lycopersicum) from seeds in 9  7.5 cm plastic cups (with drainage hole at bottom) in a mixture of 75% peat moss and 25% horticulture grade perlite having a pH ranging between 5 and 6.5, and maintain them in a glass house at 25  2
C under natural light conditions. 2. Harvest young leaves, the third or fourth fully expanded leaf from the apical bud, from 1.5-month-old tomato plants (see Note 1) (Fig. 1a, b).

2.2 Agrobacterium Strain

1. Obtain Agrobacterium tumefaciens strain LBA4404, containing the disarmed Ti plasmid pAL4404 and harboring the recombinant pK7WGF2 binary vector in which the full-length SlNACMTF3 (Solyc06g073050) of tomato is translationally fused with GFP at the N-terminal (GFP-SlNACMTF3) (see Note 2).

2.3 Alternaria solani Pathogen

1. Obtain the infectious fungus from any reliable national or university collection. 2. Grow it on potato dextrose agar (PDA) medium for 72 h at 25  2
C and 16 h light, 8 h dark cycle.

2.4 Bacterial Culture Medium

1. Add 20 g Luria-Bertani (LB) powder (prepared according to Lennox formulation) in 1 L water; thoroughly dissolve it and autoclave the medium. 2. For LB agar plates, add 1.5% (w/v) agar powder to the above broth prior to autoclaving (see Note 3). 3. Add appropriate antibiotics to the media after cooling it down to ~50
C.

2.5

Antibiotics

1. Prepare 50 mg/mL rifampicin stock solution in ethanol, and use at a final concentration of 25 μg/mL. 2. Prepare 100 mg/mL stock solutions in water for both streptomycin and spectinomycin, and use at a final concentration of 100 μg/mL. 3. Filter sterilize water-soluble antibiotics, and store all antibiotic preparations at 20
C in aliquots.

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Fig. 1 Plant materials used in transient transformation. (a) Ideal plant stage (1.5-month-old) for preparation of leaf epidermal peel (bar ¼ 1 cm), (b) optimal leaf growth (arrow) stage for preparation of leaf epidermal peel (bar ¼ 2 cm), (c) typical leaf epidermal peel (bar ¼ 0.5 cm), (d) leaf epidermal cells viewed under bright field in the microscope (bar ¼ 10 μm)

Acetosyringone

1. Prepare 100 mM stock solution in dimethyl formamide, DMF, and store at 20
C in aliquots.

2.7 Media for Plant Transformation

1. Induction medium: Dissolve 1.05 g K2HPO4, 0.45 g KH2PO4, 0.1 g (NH4)2SO4, 0.05 g sodium citrate, and 500 μL glycerol in water in a beaker; make the volume up to 96.85 mL and autoclave. After autoclaving add the following supplements: 2 mL of autoclaved 0.5 M MES-KOH pH 5.6, 1 mL of autoclaved 20% glucose solution, 100 μL of autoclaved 1 M MgSO4, and 50 μL of 100 mM acetosyringone.

2.6

2. Infiltration medium: Dissolve 440.5 mg Murashige-Skoog (MS) powdered medium, without sucrose, in 100 mL water and autoclave. Cool it to room temperature, and add 400 μL of autoclaved 0.5 M MES-KOH buffer pH 5.6 and 100 μL 100 mM acetosyringone. 3. Co-cultivation medium: Dissolve 220 mg MS medium and 1.5 g sucrose in 50 mL water and autoclave. Add 50 μL of 100 mM acetosyringone after autoclaving. 2.8 DAPI (40 ,6Diamidino-2Phenylindole) Stain

1. Dissolve DAPI hydrochloride in dimethyl formamide at a concentration of 1 mg/mL. Store the stain in dark at 4
C.

2.9 PhosphateBuffered Saline (PBS)

1. Dissolve 800 mg NaCl (136.98 mM), 20 mg KCl (2.68 mM), 144 mg Na2HPO4 (10.14 mM), and 24 mg KH2PO4 (1.76 mM) in 80 mL of water. Adjust pH to 7.6 and final volume to 100 mL. Autoclave if sterilization is required.

2.10

1. 10% glycerol prepared in co-cultivation medium. Any other commercially available mountant can also be used.

Slide Mountant

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Other Supplies

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1. Disposable pre-sterilized plastic Petri plates (90 mm) for bacterial culture and leaf epidermal peel incubation. 2. Autoclaved glassware for bacterial culture. 3. Glass slides, coverslips, clear nail polish, watch glass, and fine brush for microscopic analysis. 4. Fluorescence microscope or confocal microscope with appropriate light source or laser, respectively, for visualization of GFP fluorescence (see Note 4).

3

Methods The methods outlined in this article are as follows: (1) sonicationassisted Agrobacterium-mediated transformation of tomato leaf epidermal peels to study location of GFP-tagged SlNACMTF3 (GFP-SlNACMTF3) under normal condition and (2) localization study of GFP-SlNACMTF3 transformed in leaf epidermal peel upon exposure to the fungal pathogen Alternaria solani.

3.1 SonicationAssisted AgrobacteriumMediated Transformation of Tomato Leaf Epidermal Peels

1. Day 1, streak Agrobacterium harboring the GFP-SlNACMTF3 construct in pK7WGF2 vector on a LBA-rifampicin-streptomycin-spectinomycin plate, and incubate at 28
C (see Note 5). 2. Day 3, inoculate a single Agrobacterium colony carrying GFP-SlNACMTF3 in 10 mL LB with rifampicin, streptomycin, and spectinomycin antibiotics, and allow it to grow with constant shaking at 170 rpm at 28
C, until the OD600 value reaches 0.6–0.8 (usually less than 24 h). 3. Day 4, from the actively growing culture, inoculate 500 μL in 50 mL of induction medium supplemented with rifampicin, streptomycin, and spectinomycin antibiotics and acetosyringone. Allow it to grow under constant shaking (170 rpm) at 28
C and in dark, until OD600 value reaches 0.6–0.8 (14–16 h). 4. Day 5, aliquot the induced bacterial cells in 2 mL centrifuge tubes, and pellet it by centrifugation at 4000  g for 6 min at room temperature. Resuspend pelleted bacterial cells in 1.5 mL infiltration medium. 5. Day 5, extract tomato leaf epidermal peels from young leaves (Fig. 1a, b) (see Note 6) using the following method. Wash leaves with sterile water, remove excess water by blotting with tissue paper, and tweak the edge of the leaf blade using a small forceps to peel off the epidermal layer from the abaxial surface. Carefully remove mesophyll tissues adhering to the epidermal layer using a sharp scalpel blade (see Note 7). Place leaf epidermal peels in water in a watch glass (Fig. 1c, d).

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6. Transfer five epidermal peels of the same set to each tube containing the bacterial suspension. For each set of transformation, six such tubes are used. Place tubes in a floater, and float it at the center of a bath sonicator (we used Takashi Electric Ultrasonic cleaner, model: UD35S-0.7Lit) fitted with an electronic timer. Carry out sonication treatment for about 30 min (see Note 8). 7. Post SAAT treatment, pipette out the infiltration medium, and replace it with 5 mL co-cultivation medium for each tube. Transfer the medium along with the peels to a Petri dish. Incubate at 28
C under diffused light (25 μmol m2 s1) for 48 h (see Note 9) with occasional swirling. 8. Day 7, stain with DAPI if the nucleus needs to be detected. Immerse ~5 peels in 1 mL of co-cultivation medium containing 10 μL of DAPI solution in a 1.5 mL microcentrifuge tube. Incubate for 1 h at 28
C in dark with occasional mixing by inverting the tube. Wash epidermal peels twice with 1 mL of PBS each, and invert the tube several times to remove excess DAPI stain. 9. Mount epidermal peels on glass slides using one drop of mountant and placing a coverslip over the sample. Remove trapped air bubbles by slightly pressing the coverslip, and seal the coverslip on the slide with transparent nail polish. 10. Observe samples and capture images using the following settings: excitation/emission at 488/505–545 nm for GFP fluorescence and 365/420–540 nm for DAPI, 40 objectives, argon laser, and suitable software. Expected results are as follows: GFP fluorescence is detected along the boundary of a cell, and the DAPI staining shows location of the nucleus where GFP fluorescence is not detected. This confirms that full-length SlNACMTF3 is localized in the cell membrane and fails to migrate to the nucleus. 3.2 Studying the Dynamics of MTF Localization by Monitoring GFP-Tagged NACMTF Post Alternaria solani Inoculation

1. After SAAT treatment and 48 h of co-cultivation with Agrobacterium as described in Subheading 3.1, transfer epidermal peels to 5 mL fresh co-cultivation medium in a Petri dish. 2. Gently scrape Alternaria solani mycelia and spores from an agar plate using a sterile blade, and collect in a sterile mortar. Macerate with a pestle, and mix it with water to prepare the mycelial suspension (~500 mg mycelia in 5 mL water) (see Note 10). 3. Add 500 μL of this mycelial suspension to the epidermal peels in the co-cultivation medium. Co-incubate transformed tissues and fungus at 28
C overnight (~16 h) with constant shaking at 120 rpm in an orbital shaker.

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4. The next day, remove epidermal peels from the suspension, stain with DAPI, and mount on glass slides as described in Subheading 3.1. 5. Detect localization of GFP in epidermal peels by confocal microscopy, as described in Subheading 3.1. Expected results are as follows: co-localization of GFP and DAPI fluorescence confirms nuclear entry of the MTF during infection and suggests that fungal inoculation accelerated membrane liberation and nuclear entry of NACMTF3 (see Note 11).

4

Notes 1. Choose any cultivar of indeterminate tomatoes which can be planted in both spring/summer and fall/winter, produces broad leaves, and can be grown in both table and processing trays. We have used Solanum lycopersicum cv. Pusa Ruby which was originally released by IARI, New Delhi. 2. Choose any binary vector with highly expressed GFP protein. However, we have had the best result with pK7WGF2, where we have used an N-terminal enhanced GFP (EGFP) tag with SlNACMTF3. This tag shows better expression than MGFP usually available in the pCAMBIA series of binary vectors. The C-terminal TM domain of SlNACMTF3 is required for sequestration of the TF in the cell membrane [8]. Functionally activated NACMTF (upon stress perception or developmental cue) is proposed to undergo processing in its TM domain and localizes to the nucleus for transcriptional reprogramming. 3. The media can be stored at 4
C for 2 months. 4. We have used a confocal microscope (Leica TCS SP8, Leica Microsystems India Pvt. Ltd). 5. Each experiment should be initiated with freshly streaked Agrobacterium on LB agar plates having appropriate antibiotics. Agrobacterium cultures should be maintained with appropriate antibiotics. However, antibiotics should be left out of the co-cultivation medium. Agrobacterium growth should not exceed an OD600 of 0.8. 6. Plants neither too young nor too mature (ideally plants grown 1.5 months after germination) should be chosen for the epidermal peel preparation (Fig. 1a). For optimal results leaves of similar size and growth stage should be selected. 7. Remove as much mesophyll tissue as possible. This will facilitate avoidance of chlorophyll fluorescence during visualization of GFP.

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8. Sonication time should not exceed 30 min; prolonged sonication may lead to too much tissue damage, rendering samples unusable for biotic stress experiments. 9. It is recommended to have a co-cultivation period of 48 h which would allow expression and enough accumulation of GFP-tagged MTF for easy visualization; however, this duration should be optimized for individual gene constructs. 10. Freshly streaked and rapidly growing fungal cultures should always be used for the preparation of mycelial suspensions for inoculation. While scraping mycelia from PDA to prepare a mycelial suspension, care should be taken to eliminate as much agar as possible for best results. 11. It is not unusual for GFP fluorescence to be also found in the nucleus of some cells in control samples. This is mainly due to the detachment of monomeric GFP, from the fusion protein, which can passively gain entry into the nucleus through nuclear pores. An alternative technique such as subcellular fractionation followed by immunoblot analysis may be performed. References 1. Seo PJ (2014) Recent advances in plant membrane-bound transcription factor research: emphasis on intracellular movement. J Int Plant Biol 56(4):334–342 2. Kim SY, Kim SG, Kim YS et al (2007) Exploring membrane-associated NAC transcription factors in Arabidopsis: implications for membrane biology in genome regulation. Nucleic Acids Res 35 (1):203–213 3. Heazlewood JL, Tonti-Filippini J, Verboom RE et al (2005) Combining experimental and predicted datasets for determination of the subcellular location of proteins in Arabidopsis. Plant Physiol 139(2):598–609 4. Chou KC, Shen HB (2010) Plant-mPLoc: a top-down strategy to augment the power for predicting plant protein subcellular localization. PLoS One 5(6):e11335. https://doi.org/10. 1371/journal.pone.0011335

5. Lee MW, Yang Y (2006) Transient expression assay by agroinfiltration of leaves. In: Julio Salinas J, Sanchez-Serrano JJ (eds) Arabidopsis protocols. Methods in molecular biology, vol 323. Humana Press, Totowa, New Jersey, pp 225–229 6. Trick HN, Finer JJ (1997) SAAT: Sonicationassisted Agrobacterium-mediated transformation. Transgenic Res 6(5):329–336 7. Liu Y, Yang H, Sakanishi A (2006) Ultrasound: mechanical gene transfer into plant cells by sonoporation. Biotechnol Adv 24(1):1–16 8. Bhattacharjee P, Das R, Mandal A et al (2017) Functional characterization of tomato membrane-bound NAC transcription factors. Plant Mol Biol 93:511–532

Chapter 8 Preparation of Plant Material for Analysis of Protein–Nucleic Acid Interactions by FRET-FLIM Maxime Escouboue´, Laurent Camborde, Alain Jauneau, Elodie Gaulin, and Laurent Deslandes Abstract DNA-binding proteins are involved in the dynamic regulation of various cellular processes such as recombination, replication, and transcription. For investigating dynamic assembly and disassembly of molecular complexes in living cells, fluorescence microscopy represents a tremendous tool in biology. A fluorescence resonance energy transfer (FRET) approach coupled to fluorescence lifetime imaging microscopy (FLIM) has been used recently to monitor protein–DNA associations in plant cells. With this approach, the donor fluorophore is a GFP-tagged binding partner expressed in plant cells. A Sytox® Orange treatment converts nuclear nucleic acids to FRET acceptors. A decrease of GFP lifetime is due to FRET between donor and acceptor, indicating close association of the GFP binding partner and Sytox® Orange-stained DNA. In this chapter, we present a step-by-step protocol for the transient expression in N. benthamiana of GFP-tagged proteins and the fixation and permeabilization procedures used for the preparation of plant material aimed at detecting protein–nucleic acid interactions by FRET-FLIM measurements. Key words FRET-FLIM, DNA-binding protein, N. benthamiana cells, Sample fixation, Cell permeabilization, Sytox® Orange

1

Introduction DNA-binding proteins play important roles in various biological processes of prime importance for all living organisms. Numerous fundamental biological processes such as DNA replication, chromosome condensation, recombination, DNA repair, and regulation of gene expression are controlled by protein–DNA interactions. These interactions are far from being completely elucidated despite numerous in vitro and in vivo techniques that have been developed. Some of the well-known in vitro techniques include the footprinting assay, southwestern assay, or electrophoretic mobility shift assay [1]. Although these different techniques are useful for characterizing DNA–protein interactions, they all have limitations since they are executed outside of the natural context in which these

Walter Gassmann (ed.), Plant Innate Immunity: Methods and Protocols, Methods in Molecular Biology, vol. 1991, https://doi.org/10.1007/978-1-4939-9458-8_8, © Springer Science+Business Media, LLC, part of Springer Nature 2019

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interactions normally occur. As alternatives to these in vitro approaches, other techniques exist such as the yeast one-hybrid system or chromatin immunoprecipitation [2, 3]. Nevertheless, despite being useful for the characterization of protein–nucleic acid interactions, these techniques are often time-consuming and can also present experimental limitations (e.g., monitoring of physical associations in heterologous expression systems, false positive interactions or analysis performed on disrupted nuclear material [4]). Other methods such as fluorescence resonance energy transfer (FRET) coupled with fluorescence lifetime imaging (FLIM) represent an extraordinary advance that allows in situ analysis of interactions and microscopic imaging of protein–protein complex formation at the subcellular level [5]. The FRET-FLIM method combines time-resolved fluorescence spectroscopy with imaging microscopy, and is aimed at measuring quantitative parameters of fluorescence within living cells. The fluorescence lifetime (τ) of a fluorescent molecule may be defined as the average time that this molecule remains in its excited state prior to returning to its ground state. For fluorescent molecules commonly used in biology, this phenomenon ranges in the nanosecond timescale. Cremazy et al. [6] developed a FRET-FLIM-based approach dedicated to imaging interactions between DNA and GFP-tagged binding partners in animal cells. With this technique, associations of GFP-tagged proteins with DNA are monitored in cells treated with Sytox® Orange, a fluorescent dye that converts nucleic acids into FRET acceptors, enabling FRET-FLIM measurements. In principle, upon close association (less than 10 nm) of the GFP-tagged binding partner with Sytox® Orange-stained nucleic acids, a significant decrease of the GFP lifetime due to FRET between the donor and the acceptor can be monitored (see Fig. 1). Recently, this FRET-FLIM approach was successfully used to characterize several DNA–protein interactions [7–9] in plant cells. For example, this technique enabled us to demonstrate that the DNA-binding activity of several defensive WRKY transcription factors from Arabidopsis thaliana is inhibited by the acetyltransferase activity of the PopP2 effector from the bacterial pathogen Ralstonia solanacearum [7]. We also showed that plant nucleic acids are targeted by the AeCRN13 effector from the oomycete pathogen Aphanomyces euteiches [6]. A critical parameter with this FRET-FLIM approach performed on plant cells is the permeabilization procedure that is required for the efficient penetration of Sytox® Orange within the cells. We describe here a protocol for the preparation of plant material for characterizing protein–DNA interactions by FRET-FLIM. This method would be followed by a multi-step FRET-FLIM analysis procedure (including the synchronization and calibration of the FLIM system, FLIM measurements and analysis of the streak FLIM data), which is described in detail by Camborde et al. [10].

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Fig. 1 FRET-FLIM measurement principle for monitoring physical associations between DNA and GFP-tagged binding partners. The mean lifetime of the GFP fusion protein (donor) is measured in the absence () and in the presence (þ) of Sytox® Orange, a DNA-binding fluorescent dye serving as acceptor (black stars). In the presence of Sytox®, a decrease of GFP lifetime indicates a Fo¨rster resonance energy transfer (FRET) between donor and acceptor

2

Materials Unless otherwise noted, deionized water and basic molecular biology reagents and equipment are used to prepare the specified solutions.

2.1 Expression and Detection of GFP Fusion Proteins 2.1.1 AgrobacteriumMediated Transient Expression in N. benthamiana Leaves

1. Three- to four-week-old N. benthamiana plants grown in a greenhouse under natural daylight conditions. 2. Agrobacterium tumefaciens GV3101 cells transformed with a binary vector allowing constitutive expression of the protein of interest (POI) fused to GFP (i.e., the binary vectors pB7WGF2 (35S:GFP:POI for N-terminal fusion to the protein) or pB7FWG2 (35S:POI:GFP for C-terminal fusion to the protein)) (see Note 1). 3. Luria-Bertani (LB) medium: 10 g/L Tryptone, 5 g/L NaCl, 5 g/L yeast extract. Combine the reagents and shake until completely dissolved. Adjust the pH to 7.0 with 3 N NaOH (~0.2 mL). Adjust the final volume of the solution to 1 L with H2O. Sterilize by autoclaving. For solid LB medium, prepare LB medium as above, but add 15 g/L agar before autoclaving. 4. Agromix suspension buffer: 10 mM 2-(N-morpholino)ethanesulfonic acid (MES)-NaOH pH 5.6, 10 mM MgCl2, 150 mM acetosyringone (30 ,50 -Dimethoxy-40 -hydroxyacetophenone, a 0.5 M stock solution dissolved in DMSO and stored in aliquots at 20  C).

2.1.2 Protein Extraction from Plant Tissues

1. 1 M Tris–HCl pH 7.5, autoclaved. 2. 3 M NaCl, autoclaved. 3. 0.5 M ethylenediaminetetraacetic acid (EDTA)/NaOH pH 8.0, autoclaved.

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4. 1 M dithiothreitol (DTT), sterilize by filtration, aliquot, and store at 20  C. 5. 10% Triton X-100. 6. Protease inhibitor cocktail for plant cell extracts 100 (SIGMA). 7. Protein extraction buffer: 50 mM Tris–HCl pH 7.5, 150 mM NaCl, 10 mM EDTA pH 8.0, 1 mM DTT, 0.2% Triton™ 100, 1% (v/v) protease inhibitor cocktail. 8. 4 Laemmli denaturation buffer: 0.25 M Tris–HCl pH 7.5, 40% glycerol (v/v), 8% SDS, 0.4 M DTT, 0.04% Bromophenol Blue. Aliquot and store at 20  C. 2.1.3 Immunoblot Detection

1. Standard equipment for SDS-PAGE and semi-dry transfer. 2. Tris-Buffered Saline containing 0.1% Tween® 20 (TBS-T): 10 mM Tris–HCl pH 7.5, 150 mM NaCl, 0.1% (v/v) Tween® 20. 3. Hybond™-ECL™ (GE Healthcare).

nitrocellulose

membrane

(0.45

μm)

4. Skim milk powder. 5. Anti-GFP mouse mAb. 6. Goat anti-mouse IgG-HRP secondary antibody. 7. Chemiluminescent immunoblot blot detection reagents. 8. Chemiluminescence detection system. 2.2 Preparation of Leaf Samples for FRET-FLIM Analysis 2.2.1 Reagents

1. Paraformaldehyde: 16% (v/v) Solution, EM Grade (10 mL prescored ampules, e.g., Electron Microscopy Sciences). 2. 10 Tris-Buffered Saline solution (10 TBS): Tris-base 0.24 M; NaCl 1.37 M; KCl 26.8 mM, pH is adjusted to 7.5 using concentrated HCl (e.g., Euromedex, cat. no. ET220). 3. Sodium cacodylate trihydrate also known as sodium dimethylarsinate trihydrate (C2H6AsNaO2.3H2O) (e.g., Electron Microscopy Sciences). 4. Proteinase K, 20 mg/mL solution (e.g., Ambion or Invitrogen). 5. Sytox® Orange stain: 5 mM solution in DMSO (Invitrogen). 6. Fixation solution: 4% (v/v) paraformaldehyde, 50 mM sodium cacodylate in ultrapure water. Prepare 50 mL and store it at 4  C (see Note 2). This solution is stable for 2 weeks. 7. Washing buffer: 1 TBS. Dilute TBS 10 to a 1 solution with ddH20. Filter the solution through a 0.22 μm filter into a sterile flask.

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8. Proteinase K buffer: 50 mM Tris–HCl pH 7.5, 100 mM NaCl, 1 mM EDTA pH 8.0, 0.5% SDS. Store the solution at room temperature for up to 1 month. 9. Proteinase K solution: 100 μg/mL of proteinase K in proteinase K buffer. This solution should be prepared fresh before each use. Discard after use. 10. Sytox® Orange staining solution: 5 μM Sytox® Orange diluted in 1 TBS solution. Protect the solution from light and use immediately. 2.3 Equipment for Data Acquisition in Microscopy

1. 10 mL syringe. 2. 60-mL polypropylene screw cap container 3. Glass slides, 76  26 mm, and coverslips, 50  22 mm. 4. Immersion oil type F, code 11.513.859 (Leica Microsystems). 5. Inverted microscope (Nikon, model no. TE2000-E). 6. FLIM equipment [10].

3

Methods

3.1 Transient Expression of GFPTagged Protein in N. benthamiana

1. Inoculate one single colony of Agrobacterium tumefaciens containing the 35S:POI-GFP or 35S:GFP-POI constructs in 10 mL LB with appropriate antibiotics. Incubate overnight at 28  C, shaking at 140 rpm. 2. Centrifuge the Agrobacterium cells at 8000  g for 15 min; then gently resuspend the pellet in 2 mL of Agromix resuspension buffer. 3. Measure the optical density (OD) at 600 nm and adjust OD600 to 0.25 with Agromix buffer. 4. Incubate the suspension at room temperature for at least 60 min before leaf infiltration. 5. Infiltrate the suspension into the underside of a N. benthamiana leaf using a 1 mL syringe without needle. For each construct, it is recommended to infiltrate the whole surface of three different leaves. 6. For coexpression of two POI at a time, combine 0.25 OD600 of each Agrobacterium strain (final OD ¼ 0.5). 7. In some cases, use 35S:p19 (0.25 OD600), a suppressor of silencing, to promote the accumulation of the GFP fusion proteins (see Note 3). 8. 24–48 hours after Agro-infiltration, harvest leaf discs using a 7 mm coring tool or equivalent for each protein combination (see Note 4).

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9. For checking the GFP fluorescence intensity of the GFP binding partner (POI), harvest two leaf discs and proceed as described in Subheading 3.2. 10. For checking the accumulation level and the stability of the GFP binding partner (POI) by immunoblotting, harvest four leaf discs and proceed as described in Subheading 3.3. 11. For fixation and permeabilization of the samples for FRETFLIM measurements, harvest eight leaf discs and proceed as described in Subheading 3.4. 3.2 Checking for GFP Fluorescence Intensity of GFP-Tagged Protein

1. Remove the piston of a 10 mL syringe and transfer two leaf discs into the syringe. 2. Reposition the piston and aspirate 3 mL of 1 TBS solution with the 10-mL syringe. 3. Invert the syringe and evacuate the air. Put a finger from one hand over the hole at the top of the syringe. Pull the piston slowly to vacuum-infiltrate 2–3 times. 4. Mount the infiltrated biological samples on a glass slide. 5. Check the fluorescence level and the subcellular localization of the GFP binding partner using an inverted or a confocal microscope. A clearly visible green GFP fluorescence signal inside the nucleus is a prerequisite for the detection of interaction between nucleic acids and GFP binding partner by FRETFLIM.

3.3 Extraction and Immunodetection of GFP-Tagged Protein

1. Grind 4 leaf discs in liquid nitrogen and thaw in 1 mL protein extraction buffer. Incubate on ice for 10 min. 2. Centrifuge the mixture at 20.000  g for 10 min at 4  C. 3. Transfer supernatants carefully to clean chilled microcentrifuge tubes. 4. Transfer 50 μL of the supernatant into a 1.5 mL microcentrifuge tube and add 50 μL of 4 Laemmli denaturation buffer. 5. Denature the samples in 2 Laemmli denaturation buffer at 95  C for 3 min. 6. Separate denatured samples (10 μL) by electrophoresis on a 10% SDS-polyacrylamide gel and transfer to a nitrocellulose membrane. 7. Block the membrane with 2.5% (w/v) skim milk in TBS-T for 1 h at room temperature with gentle shaking. 8. Incubate with anti-GFP mouse antibody diluted 1:3000 in 0.5% (w/v) skim milk in TBS-T overnight at 4  C under gentle agitation. 9. Wash the membrane three times with TBS-T (3  10 min) at room temperature.

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10. Incubate the membrane with HRP-conjugated anti-mouse IgG diluted 1:10000 in 0.5% (w/v) skim milk in TBS-T for 1 h at room temperature. 11. Wash the membrane three times with TBS-T (3  10 min). 12. Detect immunoreactive signals using chemiluminescent immunoblot detection reagents and a chemiluminescence imager. 3.4 Preparation of Leaf Samples for FRET-FLIM Analysis 3.4.1 Fixation of the Samples

For all the subsequent steps, handle the leaf discs as gently as possible with forceps.

1. Harvest eight 7 mm discs with a coring tool from different leaves agro-infiltrated with the same construct. Use at least three different leaves to ensure representative sampling. 2. Cut each leaf disc in half with a razor blade. 3. Remove the piston of a 10 mL syringe and transfer all the halfdiscs into the syringe. Reposition the piston and aspirate 3–5 mL of fixation solution with the 10-mL syringe. 4. Invert the syringe and press on the piston to evacuate almost all the air. Put a finger from one hand over the hole at the top of the syringe. Pull the piston to vacuum-infiltrate 2–3 times. This step should be done as slowly as possible. 5. Re-invert the syringe (with the piston still in position) and put it back into the original packaging. Incubate vertically on ice for 30 min to ensure fixation of plant cells. 6. Transfer the half-disc samples at room temperature to a 60-mL polypropylene screw cap container containing 20 mL of TBS 1 to rinse them and remove the fixation solution.

3.4.2 Permeabilization of the Samples

1. Stack the half-discs and transfer them to a 2-mL microcentrifuge tube with 1 mL of proteinase K solution. Ensure that the samples are fully covered with the proteinase K solution. If necessary, to get full immersion of the discs, increase the volume of proteinase K solution. 2. Invert the tubes several times. Incubate at 37  C for 10 min (see Note 5). 3. Quickly transfer the half-discs to a 60-mL polypropylene screw cap container containing 20 mL of 1 TBS to remove the permeabilization solution. 4. Incubate the samples at room temperature with gentle shaking, about 50 rpm, for 5 min. 5. Repeat this washing step one time. 6. Keep half of the permeabilized half-discs in 1 TBS and store at 4  C overnight in the dark. These unstained samples will be

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used as the control condition for the monitoring of the GFP lifetime of the GFP-tagged protein in the absence of Sytox® (minus Sytox® Orange treatment, see Fig. 1). The second half of the samples will be stained with Sytox® Orange as described in Subheading 3.4.3, step 1 in order to monitor the GFP lifetime of the GFP-tagged POI (donor) in the presence of Sytox® Orange-stained nucleic acids (¼ FRET acceptor). 3.4.3 Sytox® Orange Staining

1. Transfer the remaining half discs to a 10-mL syringe. Vacuuminfiltrate the samples in 3 mL of Sytox® Orange staining solution as described in Subheading 3.4.1, step 4. 2. Carefully transfer the infiltrated discs into a new microcentrifuge tube and incubate the samples in Sytox® Orange staining solution for 30 min at room temperature in the dark. 3. Transfer the half discs to a 60-mL polypropylene screw cap container containing 30 mL of TBS 1 to rinse them. Discard the supernatant. Add 40 mL of TBS 1 and incubate overnight at 4  C in the dark.

3.5 FLIM Measurements

4

Wash fixed leaf discs and mount them on a glass microscopy slide in TBS buffer before observations on an inverted microscope (see Note 6).

Notes 1. The distribution of the pB7WGF2 and pB7FWG2 Gateway vectors is supported by the Functional Genomics unit of the Department of Plant Systems Biology (VIB-Ghent University). 2. This is a highly toxic buffer. It should be handled with protective eyewear and gloves and manipulated in a chemical fume hood. 3. Coexpression with the p19 protein of Tomato bushy stunt virus (a suppressor of RNA silencing) prevents possible RNA silencing and might promote increased transient expression of transgenes. 4. The required expression time depends on (1) the stability of the protein and (2) the binary expression vector used for Agrobacterium-transient expression assays. 24–48 hours after agroinfiltration is generally sufficient to detect GFP-tagged proteins in transformed plant cells. The optimal timing for protein expression should be determined according to the accumulation level of the transiently expressed GFP fusion proteins detected by immunoblotting. 5. Incubation time should not exceed 10 min to avoid protein degradation.

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6. FLIM measurements with the GFP-tagged POI (þ/ Sytox® Orange) are performed in time domain using a streak camera. For each nucleus, average fluorescence decay profiles are plotted and lifetimes are estimated by fitting data with an exponential function using a nonlinear least-squares estimation procedure. For statistical significance of the data, data from at least 30 nuclei should be acquired. All the different steps required for FRET-FLIM analysis performed on fixed and permeabilized samples (synchronization and calibration of the FLIM system, FLIM measurements and analysis of the streak FLIM data) are described in Camborde et al. [10].

Acknowledgments This research was performed in the Laboratoire de Recherche en Sciences Ve´ge´tales and in the Laboratory of Plant-Microbe Interactions, part of the French Laboratory of Excellence ‘TULIP’ (ANR-10-LABX-41; ANR-11-IDEX-0002-02). The described protocol was optimized with the assistance of the core facilities of the TRI-Genotoul Imagery Platform of Toulouse (France). E.G. was supported by an ANR Jeunes Chercheuses/Jeunes Chercheurs grant (APHANO-Effect, ANR-12-JSV6-0004-01). L.D. was supported by a grant from the Agence Nationale de la Recherche (ANR) (RADAR, ANR-15-CE20-0016-01). References 1. Dey B, Thukral S, Krishnan S, Chakrobarty M et al (2012) DNA-protein interactions: methods for detection and analysis. Mol Cell Biochem 365(1–2):279–299 2. Gaudinier A, Zhang L, Reece-Hoyes JS et al (2011) Enhanced Y1H assays for Arabidopsis. Nat Methods 8(12):1053–1055 3. Yamaguchi N, Winter CM, Wu MF et al (2014) PROTOCOLS: chromatin immunoprecipitation from arabidopsis tissues. Arabidopsis Book 12:e0170 4. Reece-Hoyes JS, Marian Walhout AJ (2012) Yeast one-hybrid assays: a historical and technical perspective. Methods 57(4):441–447 5. Fricker M, Runions J, Moore I (2006) Quantitative fluorescence microscopy: from art to science. Annu Rev Plant Biol 57:79–107 6. Cremazy FG, Manders EM, Bastiaens PI et al (2005) Imaging in situ protein-DNA interactions in the cell nucleus using FRET-FLIM. Exp Cell Res 309(2):390–396

7. Le Roux C, Huet G, Jauneau A et al (2015) A receptor pair with an integrated decoy converts pathogen disabling of transcription factors to immunity. Cell 161(5):1074–1088 8. Ramirez-Garce´s D, Camborde L, Pel MJ et al (2016) CRN13 candidate effectors from plant and animal eukaryotic pathogens are DNA-binding proteins which trigger host DNA damage response. New Phytol 210 (2):602–617 9. Testard A, Da Silva D, Ormancey M et al (2016) Calcium- and nitric oxide-dependent nuclear accumulation of cytosolic glyceraldehyde-3-phosphate dehydrogenase in response to long chain bases in tobacco BY-2 cells. Plant Cell Physiol 57(10):2221–2231 10. Camborde L, Jauneau A, Brie`re C et al (2017) Detection of nucleic acid-protein interactions in plant leaves using fluorescence lifetime imaging microscopy. Nat Protoc 12(9):1933–1950

Chapter 9 Analysis of DNA Methylation Profile in Plants by Chop-PCR Pratiti Dasgupta and Shubho Chaudhuri Abstract Plants, when challenged with any unfavorable condition, such as biotic or abiotic stress, adapt to the stress via physiological or structural changes. DNA methylation, an important epigenetic factor, plays an integral role in determining chromatin dynamicity and in turn regulates the process of gene transcription in eukaryotes. DNA methylation resulting in 5-methylcytosine interferes with the transcription process by hindering accessibility of the transcriptional machinery. Transcriptionally active genes are predominantly hypomethylated, whereas repressed genes exhibit hypermethylation. It can thus be interpreted that the presence of methylation in the promoter and upstream regions of loci represses their transcription and vice versa. Chop-PCR is a targeted DNA methylation detection technique that uses partial digestion by methylation-sensitive restriction enzymes (MSREs) followed by PCR amplification. The presence of cytosine methylation at the cleavage sites of the MSREs protects the DNA against digestion and therefore can be amplified using PCR. Enzymatic cleavage occurs unhindered at unmethylated restriction sites and subsequent PCR amplification of the target sequence is not observed. Key words DNA methylation, Chop-PCR, Methylation-sensitive restriction enzymes, Transcription, CTAB method

1

Introduction DNA methylation is an important epigenetic mark that promotes transcriptional silencing along with histone modifications [1–3]. DNA methylation is instrumental in silencing mobile genetic elements or transposons, thereby maintaining genomic stability [4–6]. Evidence has revealed extensive DNA methylation in heterochromatic, centromeric, and pericentromeric regions, indicating the presence of densely methylated transposons and other repetitive sequences. Cytosine DNA methylation in plants is more rich and diverse compared to animals [7]. DNA methylation in plants includes 5-methylcytosine (m5C) and N6-methyladenine (m6A). In animals, methylation is restricted to the symmetrical type (CG), i.e., methylcytosine is predominantly found in CpG islands, although non-CG methylation is prevalent in embryonic stem (ES) cells [1, 8]. Plants exhibit both symmetric and asymmetric

Walter Gassmann (ed.), Plant Innate Immunity: Methods and Protocols, Methods in Molecular Biology, vol. 1991, https://doi.org/10.1007/978-1-4939-9458-8_9, © Springer Science+Business Media, LLC, part of Springer Nature 2019

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cytosine methylation. Apart from the usual methylation of cytosine residues (as in CG methylation carried out by DNA methyltransferases), two unique types of methylation marks are found in plants: symmetric CHG methylation (carried out by Chromomethylase) and asymmetric CHH methylation (carried out by Domains Rearranged methylase) [where H is A, T, or C] [9]. DNA methyltransferases have been reported, for example, MET1, CMT3, and DRMs. MET1, a homolog of mammalian DNMT1, is responsible for the maintenance of symmetric CG methylation. Chromomethylase 3 (CMT3) is a plant-specific DNA methyltransferase, which is responsible for the maintenance of DNA methylation at CHG sites [10]. Domains Rearranged Methylases (DRMs) DRM1 and DRM2 are responsible for de novo DNA methylation at all sequence contexts. The methylation of the asymmetric CHH sites cannot take place in a semi-conservative manner. Therefore, their methylation in dividing cells is maintained by DRMs via RNA-directed de novo DNA methylation [7]. In plants, DNA de-methylation is mediated by 5-methylcytosine glycosylases. DNA glycosylases, such as Repressor of Silencing 1 (ROS1), Demeter (DME), DME-like 2 (DML2), and DML3, remove methylated bases and cleave the DNA backbone at abasic sites. The resulting gap is filled by a DNA polymerase and a DNA ligase. The major DNA methylation occurs at the fifth carbon (C5) of cytosine, resulting in 5-methylcytosine. Methylated cytosine residues serve as recognition sites for various bulky methyl groupbinding proteins. Binding of these proteins blocks the binding of the transcriptional machinery to the methylated sites, thereby repressing transcription of such methylated loci. It can thus be interpreted that the presence of methylation in the promoter and upstream regions of loci represses their transcription and vice versa. Although the inheritance of histone modifications remains unclear, DNA methylation is well established as a heritable epigenetic mark [11]. In some plants, reports suggest DNA methylation is associated with certain levels of stress-memory and, in turn, chromatin reorganization [12]. In plants, the DNA methylation status can be studied at the genome-wide level or for specific candidate genes. It can be determined by various methods, which include sodium bisulfite conversion and sequencing, meDIP (methylated DNA immunoprecipitation), MSAP (Methylation-Sensitive Amplified Polymorphism), and chop-PCR. This chapter deals with the chop-PCR method used for detecting the status of DNA methylation of specific target sequences. Although bisulfite sequencing has been widely used to determine the DNA methylation profile in plants, it involves an expensive and time-consuming workflow. In the case of a targeted DNA methylation profile, chop-PCR proves to be a highly efficient and minimum resource consuming technique [13, 14]. This method essentially compares the level and distribution of DNA methylation

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in different samples. In chop-PCR, genomic DNA isolated from plant tissues is partially digested using various Methylation-Sensitive Restriction Enzymes (MSREs), such as HpaII, MspI, AluI, NlaIII, and DdeI. MSREs are Type II restriction enzymes which are sensitive to DNA methylation, i.e., are unable to digest DNA in the presence of methylated cytosine residues at their cognate sites. Therefore, DNA stretches containing methylated cytosine residues remain intact after MSRE digestion, thereby generating amplicons on PCR amplification. DNA sequences containing unmethylated cytosine residues can be cleaved by the MSREs (at their restriction sites), thereby the PCR amplification yields negative results due to unavailability of template DNA. The presence or absence of bands and their intensities is used to detect the percentage digestion of the sample. No-enzyme reaction digestion control and No-enzyme cut site PCR control are compared against each chop-PCR digestion. The McrBC enzyme may also be used for studying DNA methylation. McrBC recognizes 50 . . .PumC (N40-3000) PumC. . .30 (where Pu is A or G and N is A, T, G, or C) sites and thereby brings about abundant cleavage in case of methylated DNA [15]. After digestion with McrBC, the digested genomic DNA may be used for (a) direct specific gene-based PCR amplification using specific primers; (b) preparing a McrBC-digested DNA library which may be used for methylome profiling using whole genome methylationsequencing. Since McrBC specifically digests methylated DNA, the more highly methylated a sequence is, the less it will be amplified by PCR after digestion [16].

2

Materials

2.1 Plant Growth and Tissue Harvest

1. Seeds of Oryza sativa L. ssp. indica cv IR64. 2. Sterile tap water. 3. Water-soaked sterile gauze in trays (see Note 1). 4. 0.1% (w/v) HgCl2 or alternative seed surface sterilization (see Note 2). 5. 0.25 Murashige and Skoog [8] complete media. 6. Plant growth chamber. 7. Single edge razor blades. 8. Liquid nitrogen. 9. Freezer set at 80  C.

2.2

DNA Isolation

1. Cetyltrimethylammonium bromide (CTAB) extraction buffer (freshly prepared) [8]: 2% hexadecyltrimethylammonium bromide (see Note 3), 1% polyvinyl pyrrolidone (PVP-40), 1.4 M sodium chloride, 100 mM Tris–HCl pH 8.0, 20 mM

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ethylenediaminetetraacetic acid (EDTA) pH 8.0 (see Note 4). Use sterile solutions to prepare the buffer. 2. 7.5 M ammonium acetate: Dissolve 0.5781 g of ammonium acetate per mL in filter-sterilized H2O. Filter sterilize. 3. 3 M sodium acetate: Dissolve 408 g of sodium acetate·3H2O (CH3COONa·3H2O) in 800 mL of H2O. Adjust pH to 5.2 and bring up volume to 1 L. Autoclave. 4. Tris-saturated phenol pH 8.0. 5. Chloroform. 6. Isoamyl alcohol. 7. Absolute ethanol. 8. RNase A: Dissolve 125 mg of RNase A per mL of filter sterile water. Filter the solution using a 0.25 μm filter to make the 125 mg/mL RNase A stock solution. Incubate the RNase A solution at 65  C for 5 min and chill on ice instantly, prior to use. 9. TE buffer: 10 mM Tris–HCl (pH ¼ 8.0), 1 mM EDTA (pH ¼ 8.0). Prepare using sterile solutions. 10. Cold centrifuge. 11. Centrifuge. 12. Vortex. 13. Vacuum desiccator. 14. Microwave oven. 15. Water bath. 16. Gel running apparatus. 17. GelDoc Analyser. 2.3

Chop PCR

1. Restriction Enzymes: HpaII, AluI, MspI, DdeI, NlaIII, McrBC (see Table 1 for restriction cleavage sites.) 2. For PCR reactions: Taq DNA polymerase, 100 mM dNTPs, dimethyl sulfoxide (DMSO). 3. Agarose, filter-sterilized water. 4. TAE buffer: 242 g of Tris base, 100 mL of 0.5 M EDTA (pH ¼ 8.0), 57.1 mL of glacial acetic acid. Make up the volume to 1 L using ddH2O and autoclave to prepare 50 TAE stock buffer. 5. NanoDrop Spectrophotometer. 6. Thermocycler. 7. Heat Block.

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Table 1 Methylation-sensitive restriction enzymes Enzyme HpaII

MspI

AluI

NlaIII

DdeI

McrBC

3

Restriction site 5’…CCGG…3’ 3’…GGCC…5’ 5’…CCGG…3’ 3’…GGCC…5’ 5’…AGCT…3’ 3’…TCGA…5’ 5’…CATG…3’ 3’…GTAC…5’ 5’…CTNAG…3’ 3’…GANTC…5’

5’…PumC (N 40-3000) PumC …3’

Methylation type

Specificity

CG

Unmethylated DNA

CHG

Unmethylated DNA

CHH/CHG

Unmethylated DNA

CHH

Unmethylated DNA

CHH/CHG

Unmethylated DNA

–

Methylated DNA

Methods

3.1 Growth of Rice Plants

1. Surface-sterilize the seeds of Oryza sativa L. ssp. indica cv IR64 with 0.1% (w/v) HgCl2 for 15 min (see Note 2). 2. Wash several times with sterile water. 3. Allow them to germinate on water-soaked sterile gauze in trays at 37  C in the dark for 3 days. 4. Grow the germinated seedlings in water-soaked sterile gauze in trays in the presence of 0.25 Murashige and Skoog [8] complete media at 30  C under 16 h light and 8 h dark photoperiod in a plant growth chamber with 50% relative humidity and 700 μmol photons m2 s1 for the desired period (see Note 5). 5. Harvest 14-day-old rice seedling leaf tissues, removing the roots with a single-edge razor blade. 6. Freeze the tissue samples in liquid nitrogen, and store at 80  C for future use.

3.2 DNA Extraction Using the Cetyltrimethylammonium Bromide (CTAB) Buffer Method

1. Grind 1 g of plant tissue to a fine paste in 3 mL of CTAB buffer using liquid nitrogen. 2. Add ß-mercaptoethanol to the CTAB buffer prior to use. 3. Transfer CTAB/plant extract to 2 mL microcentrifuge tubes and incubate for about 15 min at 55  C in a recirculating water bath. 4. After incubation, centrifuge the CTAB/plant extract at 12,000  g for 5 min. Transfer the supernatant to fresh 2 mL microcentrifuge tubes (~750 μL each).

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5. Add 750 μL of chloroform:isoamyl alcohol (24:1) to each tube and mix gently by inverting. 6. Centrifuge the tubes at 16,200  g for 10 min and transfer the upper layer to fresh microcentrifuge tubes. 7. Digest the extract with RNase A to remove RNA contamination in the genomic DNA isolate. Add RNase A to a final concentration of 100 μg/mL to each tube and incubate the samples at 37  C for an hour. 8. Perform phenol-chloroform extraction [phenol:chloroform mix (1:1)], adding equal volumes. Mix uniformly and centrifuge at 16,200  g for 10 min. Transfer the supernatant to fresh microcentrifuge tubes. 9. Precipitate the DNA using 7.5 M ammonium acetate (to a final concentration of 3 M ammonium acetate) and 3 volumes of ice-cold absolute ethanol. 10. Invert the tubes several times and incubate at 20  C for an hour to precipitate the DNA. 11. After DNA precipitation, centrifuge the tubes at 16,200  g for 15 min at 4  C and discard the supernatant. 12. Wash the DNA pellets using 70% ethanol: Add 1 mL of 70% ethanol to each tube and centrifuge the tubes at 16,200  g for a minute, followed by discarding the supernatant. For greenish/brownish pellets, see Note 6. 13. Dry the DNA pellets in a vacuum desiccator and dissolve each pellet in 50 μL of 1 TE buffer (see Note 7). 14. Use these genomic DNA samples for subsequent chop-PCR reactions. 3.3 DNA Digestion for Chop-PCR

Subject genomic DNA samples to digestion using enzymes AluI (for CHH methylation), HpaII (for CG methylation) and MspI (for CHG methylation). 1. Incubate 200 ng of DNA with 10 U enzyme for 20 min at 37  C. For the McrBC digestion, digest 500 ng of DNA with 10 U enzyme overnight (see Notes 8 and 9). Set up the reactions as follows. For all enzymes except McrBC use 5 μL 10 restriction buffer, 1 μL enzyme (HpaII/MspI/AluI/NlaIII/ DdeI), 200 ng DNA, and bring up the volume to 50 μL using sterile-filtered H2O. For McrBC use 5 μL 10 restriction buffer, 0.5 μL 100 mM GTP, 0.25 μL 20 mg/mL BSA, 1 μL McrBC enzyme, 500 ng DNA, and bring up the volume to 50 μL using sterile-filtered H2O. 2. Set up a no-enzyme control reaction corresponding to each digestion to serve as primary control. For all enzymes except McrBC use 5 μL 10 restriction buffer, 200 ng DNA and bring up the volume to 50 μL using sterile-filtered H2O. For McrBC

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use 5 μL 10 restriction buffer, 0.5 μL 100 mM GTP, 0.25 μL 20 mg/mL BSA, 500 ng DNA and bring up the volume to 50 μL using sterile-filtered H2O. 3. Following digestion, stop the reaction by enzyme inactivation. Inactivate heat-sensitive enzymes (HpaII, AluI, DdeI, NlaIII, McrBC) following digestion by incubating the tubes at 65  C for 30 min. The samples now will serve as the template for the subsequent PCR reactions. Inactivate the non-heat-sensitive enzyme (MspI) by subjecting the samples to immediate phenol-chloroform extraction following digestion. 4. Add sterile water to bring up the reaction volume to 200 μL. 5. Add 100 μL of Tris-saturated phenol and 100 μL of chloroform and mix the contents thoroughly with repeated inverting. 6. Centrifuge the samples in the phenol-chloroform suspension at 16,200  g at room temperature for 15 min. 7. Transfer the supernatant to fresh microcentrifuge tubes. Then precipitate the supernatant with 1/10th volume 3 M Na-acetate (pH ¼ 5.2) and 3 volumes absolute ethanol for an hour at 80  C. 8. Following precipitation of the DNA subject the pellets to a 70% ethanol wash and centrifuge at 16,200  g at room temperature for 1 min and discard the supernatant. 9. Dry the pellets in a vacuum desiccator and dissolve them in 50 μL of 1 TE buffer pH 8.0. This will now serve as the template for subsequent PCR. 3.4 DNA Amplification for Chop-PCR

1. Once the template is obtained, perform the PCR amplification in a thermocycler (initial denaturation at 94  C for 4 min; subsequent denaturation at 94  C for 30 s, annealing at suitable Tm, extension at 72  C for 20 s for 30 cycles; final extension at 72  C for 7 min; hold at 4  C). See Table 2 for PCR reaction mix details. 2. Design primer pairs to include a single cut site for each enzyme (preferred amplicon size ~ 120–150 bp), for better analysis of results. This serves as the test region for each case. Select a no-cut region or control region (DNA sequence stretch with no enzyme cut-site) for each enzyme and amplify the region as secondary control. 3. Following amplification, run the PCR products on a 2.5% agarose gel in 1 TAE buffer for 1 h and perform a densitometric analysis in a gel doc instrument. Repeat the experiment to generate triplicate data sets. 4. Once the quantitation is done, determine the extent of DNA methylation by calculating the percentage of undigested DNA for each amplicon (forward/reverse primer pair) for specific

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Table 2 PCR reaction mix components used for chop-PCR Component

Initial concentration

Final concentration

Taq DNA polymerase buffer

10

1

dNTPs mix

10 mM

0.25 mM

Forward primer

10 μM

0.125 μM

Reverse primer

10 μM

0.125 μM

DMSO

100%

5%

Taq DNA polymerase

2.5 U

0.5 U

DNA template

–

5–7 μL

H2O

–

20 μL

Fig. 1 Principle of chop-PCR. Genomic DNA isolated from plant tissues is partially digested using various Methylation-Sensitive Restriction Enzymes (MSRE), followed by PCR. Presence of methylated cytosine hinders the digestion by MSRE, thereby not generating amplicon bands after PCR. The PCR banding profile generated is used to determine the methylation status of a region

enzyme digestion or cytosine methylation type (CG, CHG, or CHH type). The formula is as follows, with “no enzyme” and “enzyme” signifying the band intensities of the full-length amplicons from the PCR reactions using undigested control and enzyme-digested DNA as templates, respectively: No enzyme  Enzyme  100% No enzyme ½Performed for both control and test region ðUpstream and DownstreamÞ

%Undigested ¼

5. Use the % undigested value to analyze the degree of cytosine methylation according to the principle of chop-PCR (Fig. 1) (see Note 10).

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Fig. 2 HpaII/MspI isoschizomer pair cleavage patterns and digestion profile. HpaII and MspI enzymes are isoschizomers, which recognize the site 50 -CCGG-30 . However, the HpaII enzyme can only digest the DNA if it exists in the unmethylated form (CCGG) and outer cytosine hemi-methylated form mCCGG. MspI, on the other hand is blocked from digestion only if the outer cytosine is methylated (mCCGG). The figure shows four combinations of expected outcomes of HpaII and MspI digestion of plant genomic DNA isolates. The immediate interpretation is shown beside each profile and the final interpretation is highlighted in each case. Thus, by comparing the HpaII and MspI digestion profiles, status of cytosine methylation at CCGG (serves both CG and CHG sites) can be deduced

6. Construct the entire methylation profile by aligning the analyzed data of PCR amplification of each region as shown (Figs. 2 and 3) (see Notes 11 and 12). 7. Keep in mind that digestion with McrBC is expected to produce a profile exactly opposite to the MSRE digestion profile (Fig. 3) (see Note 13).

4

Notes 1. Medium sized trays are used for growing rice seedlings. Preferred dimensions are 30 cm  25 cm. 2. HgCl2 use may be restricted in some institutes. In that case, bleach or chlorine gas generated from bleach and HCl may be used for seed surface sterilization. 3. The CTAB extraction buffer should be freshly prepared during DNA isolation. However, a 10% CTAB stock may be preserved at 37  C for later use.

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Fig. 3 AluI, NlaIII, DdeI, and McrBC cleavage pattern and digestion profile. AluI, NlaIII, and DdeI are class IIB type restriction enzymes with their cognate recognition and digestion sites. Cytosine residues are present at the recognition site of each enzyme, which if methylated forming 5mC, cannot be recognized and cleaved by the enzyme. Thus, these enzymes fail to digest methylated DNA, generating positive amplicons on PCR amplification. McrBC on the other hand is an enzyme specific for methylated DNA. It recognizes two methylated cytosines adjacent to purines in the sequence PumC (N40–3000)mCPu and cleaves at ~30 bp from them at either end. Thus, absence of methylation prevents McrBC digestion, thereby generating an opposite profile in this case

4. Sterile solutions are used to prepare the buffer. The solutions are first added and to it around 5 mL filter sterile H2O is added and pH is adjusted to 5.0 using concentrated HCl. Once the pH stabilizes, CTAB and PVP are added and the contents are mixed thoroughly. Buffer is not autoclaved. Volume is brought up to 50 mL using filter-sterilized water. 5. The MS media is diluted to a final concentration of 0.25 using sterile tap water. 500 mL of this resulting mixture is added to each tray. After seedling emergence, this volume is increased (to 750 mL), since rice plants require water logging for growth. 6. The DNA pellet obtained during DNA isolation by CTAB method at the Na-acetate/absolute ethanol precipitation step may have a greenish/brownish tinge, even after 70% ethanol wash. This may be due to carbohydrate contamination. In this case, the 70% ethanol wash may be repeated. 7. To ascertain the quality of isolated DNA, it is preferable that one determines both the 260 nm/280 nm and 260 nm/ 230 nm absorbance ratio along with the DNA concentration. Moreover, shearing of the genomic DNA during isolation

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should be avoided and a quality check may be performed by agarose gel electrophoresis. 8. Partial digestion is required during chop-PCR. Adding higher amounts of MSREs or increasing the digestion time may lead to over-digestion in chop-PCR, yielding incorrect results. The PCR cycle should be optimized to avoid saturation of amplification. 9. Digestion control and PCR amplification controls must be maintained for each test region digestion in chop-PCR. 10. The absence of PCR amplicon implies no PCR amplification due to absence of template, which in turn indicates the absence of cytosine methylation. Sequence containing methylated cytosine residues, on the other hand, cannot be cleaved by methylation-sensitive restriction enzymes, and therefore bands are obtained on PCR amplification for such regions. 11. Obtaining varying intensity bands for chop-PCR replicates is common. The parameters should be kept constant to avoid such discrepancies. During chop-PCR, low intensity bands in control samples may indicate the following: (a) Incorrect designing of primers (for no enzyme digestion control). (b) Malfunctioning of restriction enzyme or over-digestion. (c) Low level of PCR amplification. In such situations the cycles may be adjusted as required. However, amplification saturation during PCR is undesirable. 12. Absence of bands for test regions in chop-PCR may indicate the absence of DNA methylation at that region. However, in such a case, the presence of primer dimers should be confirmed since its absence will indicate no PCR amplification due to absence of primers. Further, the presence of a band for the no enzyme digestion set at this region should be confirmed. 13. McrBC is a methylation-specific enzyme, it will cleave at its cognate site only when the cytosine residues are methylated. Therefore, absence of PCR amplicons on McrBC-digested chop-PCR indicates presence of DNA methylation and vice versa. References 1. Budhavarapu VN, Chavez M, Tyler JK (2013) How is epigenetic information maintained through DNA replication? Epigenetics Chromatin 6(1):32. https://doi.org/10.1186/ 1756-8935-6-32 2. Jin B, Li Y, Robertson KD (2011) DNA methylation: superior or subordinate in the

epigenetic hierarchy? Genes Cancer 2 (6):607–617. https://doi.org/10.1177/ 1947601910393957 3. Moore LD, Le T, Fan G (2013) DNA methylation and its basic function. Neuropsychopharmacology 38(1):23–38. https://doi.org/10. 1038/npp.2012.112

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4. Feschotte C, Pritham EJ (2007) DNA transposons and the evolution of eukaryotic genomes. Annu Rev Genet 41:331–368. https://doi. org/10.1146/annurev.genet.40.110405. 090448 5. Walter M, Teissandier A, Perez-Palacios R et al (2016) An epigenetic switch ensures transposon repression upon dynamic loss of DNA methylation in embryonic stem cells. elife 5. https://doi.org/10.7554/eLife.11418 6. Ito H (2012) Small RNAs and transposon silencing in plants. Develop Growth Differ 54 (1):100–107. https://doi.org/10.1111/j. 1440-169X.2011.01309.x 7. Vanyushin BF, Ashapkin VV (2011) DNA methylation in higher plants: past, present and future. Biochim Biophys Acta 1809 (8):360–368. https://doi.org/10.1016/j. bbagrm.2011.04.006 8. Lister R, Pelizzola M, Dowen RH et al (2009) Human DNA methylomes at base resolution show widespread epigenomic differences. Nature 462(7271):315–322. https://doi. org/10.1038/nature08514 9. Meyer P (2011) DNA methylation systems and targets in plants. FEBS Lett 585 (13):2008–2015. https://doi.org/10.1016/j. febslet.2010.08.017 10. Lindroth AM, Cao X, Jackson JP et al (2001) Requirement of CHROMOMETHYLASE3 for maintenance of CpXpG methylation.

Science 292(5524):2077–2080. https://doi. org/10.1126/science.1059745 11. Lukens LN, Zhan S (2007) The plant genome’s methylation status and response to stress: implications for plant improvement. Curr Opin Plant Biol 10(3):317–322. https://doi.org/ 10.1016/j.pbi.2007.04.012 12. Kinoshita T, Seki M (2014) Epigenetic memory for stress response and adaptation in plants. Plant Cell Physiol 55(11):1859–1863. https:// doi.org/10.1093/pcp/pcu125 13. Zhang H, Tang K, Wang B et al (2014) Protocol: a beginner’s guide to the analysis of RNA-directed DNA methylation in plants. Plant Methods 10:18. https://doi.org/10. 1186/1746-4811-10-18 14. Bohmdorfer G, Rowley MJ, Kucinski J et al (2014) RNA-directed DNA methylation requires stepwise binding of silencing factors to long non-coding RNA. Plant J 79 (2):181–191. https://doi.org/10.1111/tpj. 12563 15. Sutherland E, Coe L, Raleigh EA (1992) McrBC: a multisubunit GTP-dependent restriction endonuclease. J Mol Biol 225 (2):327–348 16. Vaughn MW, Tanurdzic M, Lippman Z et al (2007) Epigenetic natural variation in Arabidopsis thaliana. PLoS Biol 5(7):e174. https:// doi.org/10.1371/journal.pbio.0050174

Chapter 10 Change in Nucleosome Dynamics During Stress Responses in Plants Amit Paul and Shubho Chaudhuri Abstract The dynamic nature of chromatin is the basis for the regulation of various biological processes in eukaryotic organisms. Nucleosome, the basic unit of chromatin in eukaryotes, undergo various reversible posttranslational modifications (PTM) in response to both external and internal cues. This PTM is recognized by different reader molecules, which facilitates the recruitment of various chromatin remodeling proteins that modulate the chromatin structure. In plants, the promoters of active genes are associated with higher lysine acetylation of histones H3 and H4, and these modifications are recognized by Bromo-domain (BRM) containing chromatin remodelers. This leads to the remodeling of the nucleosome at promoter regions, thereby increasing accessibility of the transcription machinery. It also plays a role in transcriptional repression when enriched in repressed genes. Lysine methylation recruits methyl-binding domain-containing proteins such as LIKE HETEROCHROMATIN PROTEIN1 (LHP1), which facilitates a more condensed chromatin structure that further inhibits access by the transcriptional machinery. In this article, protocols to study the regulation of chromatin conformations and nucleosome dynamics in plants in response to different stress signals are provided. Key words Nucleosome dynamics, Micrococcal endonuclease, Nucleosome scanning PCR

1

Introduction In eukaryotes, the genome is organized into a highly condensed chromatin structure called the chromosome [1]. The basic unit of chromatin is the nucleosome in which DNA is wrapped around a histone octamer. The histone octamer consists of an H3/H4 tetramer to which a dimer of H2A/H2B binds forming a nucleosome. Linker DNA is present between two successive nucleosomes which binds with the histone H1 forming chromatosome. The chromatin undergoes different levels of compaction, which leads to the condensation of eukaryotic DNA into the chromosome. The compaction and de-compaction of chromatin are regulated by various covalent posttranslational modifications of amino acid residues at the N-terminal tail of histones that reversibly regulate various

Walter Gassmann (ed.), Plant Innate Immunity: Methods and Protocols, Methods in Molecular Biology, vol. 1991, https://doi.org/10.1007/978-1-4939-9458-8_10, © Springer Science+Business Media, LLC, part of Springer Nature 2019

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dynamic biological processes such as gene expression, cell cycle progression, and DNA repair [2]. Various posttranslational modifications of histone can modulate gene expression by altering chromatin structure or recruiting a chromatin remodeler [3]. Promoters of the transcriptionally active genes are associated with nucleosome free regions or nucleosome depleted regions. The position of a nucleosome at the transcription start site is very important because it affects RNA polymerase passage through the gene body and therefore regulates transcription [4]. The dynamicity of the nucleosome can be determined by the micrococcal endonuclease (MNase) assay, which selectively cuts at the linker region leaving the core nucleosomal regions intact. This unique property of MNase has enabled us to determine whether a particular region of interest in the chromatin is protected within a nucleosome. Transcriptionally active genes will have more dynamic nucleosomes compared to their inactive counterparts and hence are more accessible to MNase. In order to understand the nucleosome structure of a particular chromatin region, MNase-digested chromatin is analyzed by quantitative PCR using overlapping primers spanning the mononucleosome [5]. Chromatin is broadly categorized as highly condensed heterochromatin and decondensed euchromatin [6]. The degree of chromatin compaction may be altered in response to various environmental as well as endogenous signals. The nuclease (MNase or DNase I) accessibility assay permits evaluation of the in vivo state of chromatin [7]. In this chapter, we provide a method to determine nucleosome dynamics using MNase. We describe plant growth using rice as an example, tissue harvesting and formaldehyde cross-linking, nuclei isolation, MNase digestion, nucleosome scanning, and the nuclease accessibility test.

2

Materials Use sterile distilled water for all solutions and rinses.

2.1

Plant Growth

1. Seeds of Oryza sativa L. ssp. indica cv. 2. Sterile water. 3. Water-soaked sterile gauze in trays. 4. 0.1% (w/v) HgCl2 or alternative seed surface sterilization. 5. 0.25 Murashige and Skoog complete media. 6. Plant growth chamber.

2.2 Tissue Harvesting

1. Single-edge razor blades. 2. 50 mL plastic tubes with perforated caps that attach tightly. 3. Vacuum desiccator. 4. 1% formaldehyde.

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5. 2 M glycine. 6. Liquid nitrogen. 7. Freezer set at 2.3

Nuclei Isolation

80  C.

1. Mortars and pestles. 2. 15 and 50 mL sterile centrifuge tubes. 3. Sterile Miracloth (Millipore Sigma). 4. Cold centrifuge. 5. Microcentrifuge. 6. 2 mL microcentrifuge tubes. 7. A centrifuge with a swinging bucket rotor. 8. Extraction buffer 1 (EB1): 0.4 M sucrose, 10 mM Tris–HCl pH 8.0, 10 mM MgCl2, 5 mM 2-mercaptoethanol, 10 mM spermidine, and 1 mM phenylmethylsulfonyl fluoride (PMSF). 9. Extraction buffer 2 (EB2): 0.25 M sucrose, 10 mM Tris–HCl pH 8.0, 10 mM MgCl2, 5 mM 2-mercaptoetahnol, 1% Triton™ X 100, 10 mM spermidine, 1 mM PMSF. 10. Extraction buffer 3 (EB3): 1.7 M sucrose, 10 mM Tris–HCl pH 8.0, 2 mM MgCl2, 5 mM 2-mercaptoethanol, 0.15% Triton™ X 100, 10 mM spermidine, 1 mM PMSF. 11. Nuclei wash buffer: 50 mM Tris–HCl pH 8.0, 5 mM MgCl2, 10 mM 2-mercaptoetahnol, 20% glycerol, 0.25% Triton X 100. 12. Nuclei storage buffer: 50 mM Tris–HCl pH 8.0, 5 mM MgCl2, 10 mM 2-mercaptoethanol, 25% glycerol. 13. Nanodrop spectrophotometer.

2.4 Micrococcal Digestion and Nuclease Assay

1. Micrococcal digestion buffer: 50 mM Tris–HCl pH 8.0, 5 mM MgCl2, 10 mM 2-mercaptoethanol, 25% glycerol, 1.5 mM CaCl2. 2. Stop buffer: 5% sodium dodecyl sulfate (SDS) and 250 mM ethylenediaminetetraacetic acid (EDTA). 3. Micrococcal endonuclease (MNase). 4. Proteinase K. 5. Vortexer. 6. Incubator at 37  C. 7. Phenol/chloroform 1:1 (v/v). 8. Sodium acetate and absolute ethanol to precipitate DNA. 9. 1.5 mL centrifuge tubes. 10. Tris EDTA (TE) buffer: 10 mM Tris–HCl pH 8.0, 1 mM EDTA. 11. Vacuum dryer.

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12. Tris acetate-EDTA (TAE) buffer: 1 TAE contains 40 mM Tris, 20 mM acetate and 1 mM EDTA. 13. 1.2% agarose gels. 14. Electrophoresis equipment. 15. Gel-extraction kit (see Note 1). 16. Heat block. 17. Variable water bath. 18. 2 Mastermix for real-time PCR.

3 3.1

Methods Plant Growth

1. Surface-sterilize the seeds of Oryza sativa L. ssp. indica cv with 0.1% (w/v) HgCl2 for 15 min. 2. Wash several times with sterile water. 3. Allow them to germinate on water-soaked sterile gauze in trays at 37  C in the dark for 3 days. 4. Grow the germinated seedlings in water-soaked sterile gauze in trays in the presence of 0.25 Murashige and Skoog complete media at 30  C under 16 h light and 8 h dark photoperiod in a plant growth chamber with 50% relative humidity and 700 μmol photons m 2 s 1 for the desired period. 5. Subject a portion of the plants to a stress, such as heat stress (see Note 2).

3.2 Tissue Harvesting and Formaldehyde Cross-Linking

1. Harvest 13-day-old rice seedling aerial tissue, removing the roots with a single-edge razor blade. 2. Wash aerial tissues twice with distilled water and treat them with 40 mL of 1% formaldehyde in 50 mL plastic tubes with firmly attached perforated caps (see Note 3). 3. Vacuum infiltrate the tubes in a vacuum desiccator for 15 min for cross-linking. During this process, shake the tubes to release any trapped air bubbles. 4. Wash the cross-linked tissues with distilled water and treat with 10 mL of 2 M glycine to quench the cross-linking. 5. Finally, wash the tissues with distilled water, freeze them in liquid nitrogen, and keep them at 80  C.

3.3

Nuclei Isolation

1. For each sample, crush 5 g of tissue in liquid nitrogen in a mortar with a pestle. 2. Add 50 mL of EB1 to the crushed tissue and leave it covered until it forms a liquid phase (see Note 4).

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3. Double filter the tissue suspension into a 50 mL sterile plastic centrifuge tube using sterile Mira cloth. 4. Centrifuge the samples at 2600  g for 20 min at 4  C. 5. Discard the supernatant, resuspend the pellet in 1 mL of EB2, and transfer the sample to a 2 mL microcentrifuge tube. 6. Centrifuge the sample at 17,000  g for 10 min at 4  C. Discard the supernatant, and resuspend the pellet in 500 μL of EB2. 7. To a 1.5 mL microcentrifuge tube, add 500 μL of EB3 and over it carefully layer the sample suspension in EB2. 8. Centrifuge the tubes at 3000  g for 30 min at 4  C using a swinging bucket rotor. 9. Discard the supernatant and wash the pellet with 1 mL wash buffer by centrifuging at 17,000  g for 10 min at 4  C. 10. Discard the supernatant and resuspend the pellet in 750 μL of storage buffer. Centrifuge at 17,000  g for 10 min at 4  C. 11. Resuspend the pellet in 1 mL of storage buffer (see Note 5). 12. Take 10 μL of nuclei suspension and dilute it 50-fold with storage buffer. Measure the A260/280OD in a double-beam spectrophotometer to check the quality of nuclei (see Note 6). 3.4 Micrococcal Endonuclease (MNase) Digestion and Nucleosome Scanning Assay to Determine Nucleosome Position

1. Aliquote 250 μL of nuclei suspension in 1.5 mL microcentrifuge tubes and subject them to MNase digestion as follows. Retain a small aliquot of non-digested genomic DNA to compare with digested DNA (see step 9). 2. Add 3 U, 6 U, 10 U, and 15 U of MNase to tubes with 250 μL of nuclei suspension and incubate the set of 4 tubes at 37  C for 10 min (see Note 7). 3. Stop the reaction by adding 50 μL reaction stop buffer and vortex briefly. 4. Add 3 μL of 20 mg/μL Proteinase K to each tube and incubate them overnight at 50  C. 5. The next day, extract the samples with equal volumes phenol: chloroform (v/v) and centrifuge at 17,000  g for 15 min. 6. Take 300 μL of the supernatant and precipitate with 1/10 volume of Na-acetate (30 μL) and 3 volumes (900 μL) of absolute ethanol. 7. Keep at 80  C for 2 h. Spin the sample at 17,000  g to pellet the DNA. 8. Vacuum dry the DNA pellet and dissolve it in 50 μL of TE buffer. 9. Retain a small aliquot of the MNase-digested DNA and electrophorese it with the small amount of genomic DNA retained

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MNase

16

8 4 2 0

Salt-8 hrs

MNase

0 2 4

MNase

8 16 M 16 8 4

Control

2 0

Salt 4 hrs

Fig. 1 Micrococcal endonuclease digestion of chromatin. Nuclei isolated from the control and salinity stressed plants were subjected to MNase digestion (1 U, 3 U, 6 U, 10 U, and 15 U). The reaction was stopped and DNA was extracted once with equal volume of phenol and chloroform. DNA was analyzed on a 1.2% agarose gel

from nuclei suspension not subjected to MNase digestion (see step 1) in a 1.2% agarose gel in 1 TAE buffer (Fig. 1). Assess the quality and repeat order of the nucleosomal DNA ladder generated as multiples of the nucleosome core plus the linker DNA (200 base pairs). 10. Gel-elute the mononucleosomal DNA fraction using a gel extraction kit as described in the manufacturers’ protocol and use the mononucleosomal DNA template for determination of nucleosome positions using primer tiling arrays (Fig. 2). 3.5 Nuclease Accessibility Test to Determine Nucleosome Dynamics

1. Incubate 1 mL nuclei suspension (see Subheading 3.3, step 11) from control and stressed plants at 37  C in a heat block for 5 min. 2. Add 10 U of diluted MNase to each nuclei suspension and incubate them at 37  C in a heat block. 3. After 3 min, remove 200 μL from the nuclei suspension and add it to 50 μL of the reaction stop buffer. Repeat this step at

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Fig. 2 Nucleosome scanning PCR. Designing a tiling array of primers. The sequence of the region of interest is obtained from NCBI to design primers for nucleosome scanning analysis. In this method, overlapping primers (35 bp interval) are designed for the region of interest such that it amplifies 200 bp product. The amplification of 200 bp DNA from the mononucleosomal fraction (M) is compared with that from genomic DNA (G). If the amplification from the mononucleosomal fraction is efficient and comparable to that of genomic DNA, we presume that the region is protected by a well-positioned nucleosome. However, if the primers are outside the nucleosome protected region (linker region), we expect poor amplification from the mononucleosomal fraction since the linker region is sensitive to MNase digestion

subsequent time points (6 min, 10 min and 15 min) of MNase digestion. 4. Add 3 μL of 20 mg/μL Proteinase K to each tube and incubate overnight at 50  C. 5. The next day, extract the samples with 250 μL phenol:chloroform solution and centrifuge at 17,000  g for 15 min. 6. Take 250 μL of the supernatant and precipitate it with 1/10 volume of Na-acetate (25 μL) and 3 volumes (750 μL) of absolute ethanol. Keep it at 80  C for 2 h. 7. Spin the samples at 17,000  g to pellet the DNA (see Note 8). 8. Vacuum dry the DNA pellets, dissolve them in 50 μL of TE buffer, and store them at 20  C. 9. Check 5 μL of the DNA sample by loading it in a 1% agarose gel and running it at 120 V for 30 min to see the digestion. The ideal digestion should form a DNA smear throughout the lane (see Note 9). 10. Analyze the data using the standard curve method as described below: Use the MNase-digested DNA as the template for quantitative PCR. For the MNase accessibility assay, use those primers that were used to determine the position of the

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Fig. 3 Schematic diagram of nuclease accessibility test. The MNase-digested DNA is used as the template for quantitative PCR. For the MNase accessibility assay, we use those primers that were used to determine the position of the nucleosomes. Thus, comparing the accessibility of the MNase enzyme in a given locus in transcriptional “off” and “on” states can indicate the dynamics in nucleosome arrangement. Higher accessibility of MNase results in an increase in the degradation of DNA. This will lead to a decrease of amplicon levels, which can be quantitated using real-time PCR. Tightly arranged nucleosomes inhibit the access of MNase to linker DNA, whereas the MNase accessibility increases in case of dynamic or open chromatin

nucleosomes. It is presumed that the nucleosome arrangement of a given region changes in response to transcription demand, thereby promoting the accessibility of transcription factors. Compare the accessibility of the MNase enzyme in a given locus in transcriptional “off” and “on” states to indicate the dynamics in nucleosome arrangements (Fig. 3). The higher accessibility of MNase results in an increase in the degradation of DNA. This will lead to a decrease of amplicon levels, which can be quantitated using real-time PCR. Analyze the data sets using the standard curve method. Generate a standard curve using DNA from undigested samples (-MNase) diluted 1:5, 1:10, and 1:20 fold (Fig. 3). Plot the Ct values obtained from the digested MNase samples in a graph to calculate the percent of input value. Normalize the percent of input value obtained from the MNase-digested samples using the value obtained from the undigested sample (-MNase) (Fig. 4).

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Fig. 4 Absolute quantification of MNase-digested DNA using standard curve method. The data sets are analyzed using the standard curve method. DNA from undigested samples (-MNase) is diluted 1:2.5, 1:5, and 1:10 and used to generate a standard curve. The Ct values obtained from the digested MNase samples are plotted in a graph as shown, and the percent of input value is calculated. The percent of input value obtained from all the MNase-digested samples is normalized using the value obtained from the undigested sample (-MNase)

4

Notes 1. We use the Qiagen gel extraction kit for DNA extraction from agarose gels. 2. For salinity stress, we exposed 13-day-old rice seedlings to 500 mL of 250 mM NaCl. 3. We perforate tube caps using a heated needle; 10 holes per tube are sufficient. 4. We cover the mortar and pestle with aluminum foil and keep them in a cold room at 4  C until the sample appears to be a liquid suspension. 5. Add 1.5 mM CaCl2 to the storage buffer before suspending the nuclei because MNase needs Ca2+ ions for its optimal activity. 6. During nuclei isolation, if the color of the nuclei pellet has a greenish appearance, this is due to the precipitation of pigments. Wash the nuclei twice with nuclei wash buffer and follow with a single wash with storage buffer. 7. Less smear of MNase-digested DNA in the gel could be due to over-digestion by MNase; try a smaller concentration of MNase. 8. 3 μL glycogen can be added for efficient DNA precipitation. 9. The yield of the DNA could be low due to excessive pipetting of nuclei, thus compromising the integrity of the nuclei.

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References 1. Finch JT, Klug A (1976) Solenoidal model for superstructure in chromatin. Proc Natl Acad Sci 73(6):1897–1901 2. Vermaak D, Ahmad K, Henikoff S (2003) Maintenance of chromatin states: an open-and-shut case. Curr Opin Cell Biol 15(3):266–274 3. Zhang X, Clarenz O, Cokus S, Bernatavichute YV, Pellegrini M, Goodrich J, Jacobsen SE (2007) Whole-genome analysis of histone H3 lysine 27 trimethylation in Arabidopsis. PLoS Biol 5(5):e129 4. Shivaswamy S, Bhinge A, Zhao Y, Jones S, Hirst M, Iyer VR (2008) Dynamic remodeling of

individual nucleosomes across a eukaryotic genome in response to transcriptional perturbation. PLoS Biol 6(3):e65 5. Ozsolak F, Song JS, Liu XS, Fisher DE (2007) High-throughput mapping of the chromatin structure of human promoters. Nat Biotechnol 25(2):244 6. Grewal SI, Moazed D (2003) Heterochromatin and epigenetic control of gene expression. Science 301(5634):798–802 7. Kiefer CM, Hou C, Little JA, Dean A (2008) Epigenetics of β-globin gene regulation. Mutat Res 647(1–2):68–76

Chapter 11 Measuring Cell Ploidy Level in Arabidopsis thaliana by Flow Cytometry Leiyun Yang, Zhixue Wang, and Jian Hua Abstract Cell ploidy levels are regulated by developmental and environmental factors and they also impact the outcome of plant microbe interactions. Here we describe a simple and quick procedure to measure cell ploidy levels in Arabidopsis thaliana leaves by flow cytometry. Cell nuclei are isolated by filtering tissue homogenates from chopped plant tissues. DNA in the nuclei is stained by propidium iodide and the fluorescence emitted from the DNA of each nucleus is read by using a flow cytometer. Distribution of ploidy levels within the plant tissues can be calculated based on the distribution of fluorescence signals. Multiple samples can be prepared and analyzed within the same day. Key words Ploidy, Flow cytometry, Cell cycle, Arabidopsis thaliana, Propidium iodide

1

Introduction Plants undergo a variety of cell cycles during development and in response to environmental signals. The progression of the cell cycle is closely linked to plant–microbe interactions and disease resistance [1]. Cell cycle progression could be altered upon pathogen infection, resulting from a defense response from plants or a manipulation of plants by the pathogen [2–4]. On the other hand, perturbation of cell cycle regulation in plants could activate disease resistance genes and trigger immune responses [5]. Therefore, measuring cell ploidy levels could contribute to the understanding of plant microbe interactions and immune regulation. Flow cytometry is often used for such a purpose as it can measure fluorescence emitted from the stained nuclear DNA when each isolated nucleus passes through, which is proportional to the ploidy level. Many protocols have been developed for staining DNA in the nucleus and measuring the ploidy level by flow cytometry [6–9]. Here we describe a simple and rapid method that is tailored for Arabidopsis leaves and cotyledons. Briefly, plant tissues are chopped with a clean razor blade in buffer to release nuclei, and

Walter Gassmann (ed.), Plant Innate Immunity: Methods and Protocols, Methods in Molecular Biology, vol. 1991, https://doi.org/10.1007/978-1-4939-9458-8_11, © Springer Science+Business Media, LLC, part of Springer Nature 2019

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the homogenates are filtered to remove debris. Propidium iodide is then used to stain DNA while RNA is removed by RNase. Stained nuclei are then run through a flow cytometer and data are collected and analyzed within the same day of sample preparation.

2 2.1

Materials Plant Materials

2.2 Chemicals and Buffers

1. Grow plants at 22
C under long day conditions, either on plates containing ½ MS (pH 5.7) and 1% sucrose for 8 days or on soil for 2–3 weeks (see Note 1). 1. MgSO4 buffer: 1.23 g MgSO4·7H2O, 1.85 g KCl, 0.6 g Hepes. Dissolve in 480 mL ddH2O, adjust pH to 8.0 using KOH and then bring to final volume of 500 mL with ddH2O.Keep at 4
C. 2. Propidium iodide (PI) solution: 10 mg/mL ddH2O. Keep in dark until use. 3. RNase A stock: 20 mg/mL ddH2O. Keep in dark until use. 4. “Aru” buffer (10 mL, see Note 2): 9.65 mL MgSO4 buffer, 100 μL 1 M DTT, 250 μL Triton X-100. Keep on ice after preparation.

2.3 Equipment and Materials (See Note 3)

1. Flow cytometer. 2. 30 μm filters (such as CellTrics® from Sysmex Partec). 3. 5 mL tubes. 4. Single-edge razor blades. 5. 100  15 mm Petri dishes.

3

Methods

3.1 Sample Preparation (Always Keep Samples on Ice)

1. Place a petri dish on ice and add 400 μL precooled “Aru” buffer to the dish (see Note 4). 2. Harvest leaves or cotyledons (see Note 5), and immerse them in the “Aru” buffer. Wash tissues in precooled water to remove soil or other particles when needed before putting them in the buffer. 3. Chop the plant materials in a sample with a new razor blade to release the nuclei, making clean cuts until the plant tissues have no visible chunks (Fig. 1). It usually takes 1–3 min for one sample. Use a new razor blade for each sample. 4. Transfer all the mixture using a 1 mL pipet tip to a 30 μm cell strainer and collect the filtrate with a labeled 5 mL tube. Place the tube on ice (Fig. 2).

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Fig. 1 Leaf tissues of Arabidopsis thaliana after being chopped in “Aru” buffer

Fig. 2 Homogenates after being filtered through a 30 μm filter

5. Wash the chopping plate with 200 μL “Aru” buffer to collect all nuclei, and filter with the same strainer and into the same tube. The final volume should be around 500 μL. 6. Repeat steps 1–5 for each sample until all samples are chopped and filtered into their respective tubes. 7. Add 2.5 μL RNase A and 5 μL PI solutions (see Notes 6 and 7) and tap to mix well. 8. Incubate on ice for around 30 min in the dark before running on the flow cytometer (see Note 8).

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3.2 Running Samples (See Note 9)

1. Turn on the flow cytometer and detect PI emission at the FL2 channel with the 585/40 filter or load a preset program. 2. Check the status of the machine. Empty the waste tank and fill the sheath fluid tank. 3. Mix samples by tapping and immediately load the samples on the machine. 4. We use a BD C6 flow cytometer and run samples at a rate of 11 μL/min (see Note 10). 5. Count 5000–10,000 events for each sample (see Note 11). It may take 2–8 min for one sample. More starting materials will give a shorter run.

3.3

Data Analysis

1. Crop a region containing the intact nuclei signals to remove background signals as much as possible (Fig. 3). 2. A histogram of signals will be generated by the program. Use dashed lines to select the optimal signal regions for each ploidy range. The percentage of each ploidy range will be calculated automatically by the program (Fig. 4). 3. Data can be displayed either as the distribution of cells with different ploidy levels or as ploidy indexes using the formula below: Ploidy index ¼ (%2C nuclei  1) þ (%4C nuclei  2) þ (%8C nuclei  3) þ (%16C nuclei  4) þ (%32C nuclei  5).

4

Notes 1. Cell ploidy levels often increases in leaves as they develop more. For final cell ploidy levels, 8-day-old plants can be used for cotyledons and 20-day-old plants can be used for the first pair of true leaves. For comparison between different genotypes, use comparable tissues at a similar developmental stage. 2. “Aru” buffer should be freshly prepared and used on the same day. 3. The filters can be reused after being thoroughly washed with water. 4. The volume of “Aru buffer” used depends on the tissue weight. Use 600 μL for 25 mg of tissue or less. High background signals may arise from debris when tissues/buffer ratio is too high. 5. Some protocols allow an overnight 4
C and dark incubation of the tissues before chopping. The preincubation is not necessary and it may introduce ploidy level changes.

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Fig. 3 Histogram of FL2-A (relative PI-fluorescent intensity) and FSC-A (size of the particles) using the first pair of true leaves. Four intense regions represent 2C, 4C, 8C, and 16C, respectively, from bottom to top. A gating region (P6) was selected by adjusting the position of the red dashed line to exclude the background signals. Real signals constituted 78.6% of the total captured signals in this case

6. PI has a high affinity to dsRNA in addition to DNA, so it is important that RNAs are completely removed. RNase A and PI can be mixed on the day of sample preparation and 7.5 μL of this mix can be added into the sample. PI and PI/RNase solutions should be kept in the dark. 7. The sample volume is usually around 500 μL after filtering, although it differs from sample to sample as a result of handling variations and/or tissue variations. Transfer the same volume from each sample into a new tube and add a corresponding amount of PI/RNase stock. 8. Some protocols use an animal size standard or plants with known nuclear DNA content as an internal control. This is not necessary for Arabidopsis leaves/cotyledons as the 2C ploidy peak of Arabidopsis is easily identified. 9. Refer to the user’s manual for use of any other flow cytometers. 10. Do not run at a fast rate because it would increase background. 11. Count the same number of events for each sample for best comparison.

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Fig. 4 Histogram of fluorescence signals from a stained Arabidopsis tissue. Four major peaks are from cells with ploidy levels at 2C, 4C, 8C, and 16C, respectively. Dashed lines define each ploidy range for calculating signal percentage in each range. This sample has 17.5% 2C, 22.6% 4C, 37.3% 8C, 19.2% 16C, and 0.6% 32C

Acknowledgments The work in J. Hua’s lab is supported by National Science Foundation IOS-1353738 to J.H. References 1. Bao Z, Hua J (2015) Linking the cell cycle with innate immunity in Arabidopsis. Mol Plant 8 (7):980–982 2. Hamdoun S, Liu Z, Gill M et al (2013) Dynamics of defense responses and cell fate change during Arabidopsis-Pseudomonas syringae interactions. PLoS One 8:e83219 3. Ascencio-Ibanez JT, Sozzani R, Lee TJ et al (2008) Global analysis of Arabidopsis gene expression uncovers a complex array of changes impacting pathogen response and cell cycle during geminivirus infection. Plant Physiol 148:436–454 4. Chandran D, Inada N, Hather G et al (2009) Laser microdissection of Arabidopsis cells at the powdery mildew infection site reveals sitespecific processes and regulators. Proc Natl Acad Sci U S A 107:460–465

5. Bao Z, Yang H, Hua J (2013) Perturbation of cell cycle regulation triggers plant immune response via activation of disease resistance genes. Proc Natl Acad Sci U S A 110 (6):2407–2412 6. Arumuganathan K, Earle ED (1991) Estimation of nuclear DNA content of plants by flow cytometry. Plant Mol Biol Rep 9(3):229–241 7. Dolezˇel J, Greilhuber J, Suda J (2007) Estimation of nuclear DNA content in plants using flow cytometry. Nat Protoc 2(9):2233–2244 8. Galbraith DW (2009) Simultaneous flow cytometric quantification of plant nuclear DNA contents over the full range of described angiosperm 2C values. Cytom Part A 75(8):692–698 9. Dolezˇel J, Bartosˇ JAN (2005) Plant DNA flow cytometry and estimation of nuclear genome size. Ann Bot-London 95(1):99–110

Chapter 12 A Method for Investigating the Pseudomonas syringae-Arabidopsis thaliana Pathosystem Under Various Light Environments Daniel L. Leuchtman, Anthony D. Shumate, Walter Gassmann, and Emmanuel Liscum Abstract Arabidopsis thaliana and Pseudomonas syringae pv. tomato DC3000 (Pst DC3000) comprise an effective model pathosystem for resolving mechanisms behind numerous aspects of plant innate immunity. Following the characterization of key molecular components over the past decades, we may begin investigating defense signaling under various environmental conditions to gain a more holistic understanding of the underlying processes. As a critical regulator of growth and development, exploration into the influence of light on pathogenesis is a logical step toward a systems-level understanding of innate immunity. Based on methods described previously, here we describe a method for investigating plant immune responses under various light environments. Key words Arabidopsis, Pseudomonas syringae, Pathosystem, Infection assay, Pathogen growth assay, Growth curve, Innate immunity, Plant defense, Photobiology, Light treatment

1

Introduction For plants, light is not only a source of energy but also a critical source of information. Key physiological changes are induced by altered patterns in the day length, light quality, or position within the environment [1]. An acutely tuned immune system is also paramount to plant health. Underutilization can result in disease formation, while overutilization comes at a high energetic cost [2]. Several hypotheses offer mechanisms and implications of cross-talk between light and immune signaling components. Among these hypotheses are photosynthetic implications on reactive oxygen species (ROS) accumulation, synthesis of defenserelated compounds, modifications to chloroplast biology, adjustments to the circadian clock, and signaling events mediated by photoreceptors [3–7]. While relationships between proteins

Walter Gassmann (ed.), Plant Innate Immunity: Methods and Protocols, Methods in Molecular Biology, vol. 1991, https://doi.org/10.1007/978-1-4939-9458-8_12, © Springer Science+Business Media, LLC, part of Springer Nature 2019

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traditionally associated with light signaling and those involved in pathogen defense have been demonstrated previously [8, 9], further analysis of these relationships in the context of specific lighting environments could reveal additional signaling overlap and a deeper understanding of these two systems. Several infection methods are used to examine Pst DC3000Arabidopsis interactions including direct leaf injection via syringe pressure infiltration, spraying of inoculum onto soil or plant tissue, and immersion of whole plants within inoculum [10, 11]. As plants used for these methods are typically grown openly in greenhouses or growth chambers, assessing plant-pathogen interactions under various light environments can thus prove problematic due to space and resource limitations. Here, we describe a method for growth and treatment of Arabidopsis plants under various light treatment conditions that incorporates reduced space requirements, shortened growth periods, and high-throughput inoculation. An Arabidopsis seedling flood-inoculation technique developed by Ishiga and colleagues [12] was critical to the development and implementation of our method. Using this approach, Arabidopsis seedlings are grown on MS medium in deep-welled plates [12]. This adaptation allows us to employ simple light-proof containers and transparent colored acrylic sheets to vary light treatment conditions. To this end, we assessed initiation of effector-triggered immunity (ETI) by recognition of effector HopA1 through RPS6 (RESISTANT to PSEUDOMONAS SYRINGAE 6) and EDS1 (ENHANCED DISEASE SUSCEPTIBILITY 1) under normal full-spectrum white (WL), red (RL), blue light (BL), and dark treatment conditions. To illustrate the efficacy of our method, we show results demonstrating that ETI is initiated through RPS6 and EDS1 under BL or RL treatments but fails to initiate in darkness (Fig. 1), confirming that our method may be used to analyze ETI under various light environments.

2

Materials All solutions were prepared using deionized water and molecular biology grade reagents. Solutions may be stored at 4
C unless otherwise specified.

2.1

Plant Growth

1. 50% sucrose: Dissolve 50 g sucrose in water and bring volume to 100 mL. Filter sterilize (0.2 μm pore size). 2. Plant growth media: Half-strength MS (1/2 MS) medium, 0.3% (w/v) Phytagel, Gamborg vitamins, and 1% (w/v) sucrose. Prepare 1/2 MS media and aliquot into autoclave bottles. Swirl bottles to create vortex and add Phytagel. Sterilize via autoclave. Cool 1/2 MS to approximately 55
C in hot water bath. Working in a laminar flow hood, add Gamborg

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Fig. 1 Col, rps6-3, and eds1-2 that were left untreated in WL (a) or incubated in darkness (b), blue light (c), or red light (d) 3 days prior and up to 4 days following inoculation. Bacterial growth was measured on day 0 and day 4 after inoculation with DC3000(hopA1). Plants were inoculated with a bacterial suspension at a density of 1  105 CFU/mL. Values represent averages of CFU/mg leaf tissue from triplicate samples with error bars denoting standard deviation. Statistical significance was determined using a Student’s t-test comparing Col to either rps6-3 or eds1-2 on D4 ( p, *0.05, **0.01, ***0.001)

vitamins to 1 and sucrose to 1% (v/v). Pour 1/2 MS into deep-well plates (100 mm  25 mm). Dry overnight in sterile hood with lids on. 3. Seed Sterilization I: 50% (v/v) ethanol, 0.5% Triton X-100. Make fresh. 4. Seed Sterilization II: 95% (v/v) ethanol. 5. 100% ethanol. 6. Sterilized filter paper. 7. Sterilized forceps. 8. Benchtop microcentrifuge. 9. 3M Micropore 2.5 cm surgical tape.

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10. Plastic container and lids coated with matte black paint. Must be completely opaque. Approximately 300 mm  175 mm  100 mm. 11. Transparent acrylic sheets for desired light treatment. 2.2 Bacterial Isolation and Quantification

1. 100 mg/mL cycloheximide dissolved in 50% (v/v) ethanol. 2. Pseudomonas agar (PA) plates: Pseudomonas agar F. Prepare Pseudomonas agar according to package directions. Cool to approximately 55
C in hot water bath. Add cycloheximide to 100 μM and antibiotics to select for strain of interest. Allow PA plates to solidify. 3. Sterile 10 mM MgCl2. 4. 3% H2O2. 5. Sterile water for rinsing plant material following surface sterilization with H2O2. 6. Sterilized mortar and pestles: Covered in foil and autoclaved or incubated at 250
C overnight. Cool to room temperature. 7. Absorbance spectrophotometer for measuring optical density of bacterial suspension. 8. Sterilized 1.5 mL microfuge tube pestles. 9. Labeled weigh boats for weighing plant material.

3 3.1

Methods Seed Sterilization

1. Aliquot seed into sterile 1.5 mL microfuge tube and add 1 mL of Seed Sterilization I. Shake for 3 min, and centrifuge for 30 s at max speed in benchtop centrifuge (~16,000  g). 2. Decant. Add 1 mL of Seed Sterilization II. Mix by hand, taking care to dislodge all pelleted seed, and centrifuge for 30 s at maximum speed in the benchtop centrifuge (~16,000  g). Repeat two additional times. Suspend in Seed Sterilization II. 3. Label a sterile Petri dish on top and bottom. Insert sterile filter paper and aliquot sterilized seed. Allow to dry for approximately 0.5 h (see Note 1).

3.2 Planting and Light Treatment

1. Dip the forceps in ethanol, and use each prong to individually select a seed and transfer onto the plant growth plate. Leave optimal growth space for each seed (5–6 seeds per plate). Seal the plate using Micropore tape. 2. Place plates in dark boxes with lids and incubate 2–5 days at 4
C. 3. At room temperature, replace lids with red acrylic glass, and incubate for 1 h.

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Fig. 2 Col plants grown for approximately 2 weeks and then left untreated (WL) or incubated in darkness (NL), blue light (BL), or red light (RL) for 3 days

4. Replace red acrylic glass with dark box lid and incubate at room temperature for 2 days in darkness (see Note 2). 5. Remove lid, and place in walk-in growth chamber under 12 h light/12 h dark cycle at 22
C and light intensity of 50–70 μmol m2 s1. 6. Plants are ready for infection within 2–3 weeks or when 6–8 true leaves are present (Fig. 2). 7. Begin light treatment 3 days prior to infection by covering the container with an acrylic glass sheet. 3.3

Infection

1. Streak a glycerol stock of bacteria onto PA plate 2 days prior to infection. Spread the bacterial stock to create a lawn, but without covering the entire plate. Incubate overnight at 28
C. 2. Using a pipette tip, swipe 5–6 cm along the growing edge of the bacterial lawn. 3. Transfer bacteria obtained into 200 μL of 10 mM MgCl2 and vortex. 4. Dilute 10 μL bacterial suspension into 990 μL of 10 mM MgCl2, and measure the optical density (OD600) by absorbance at 600 nm. Multiply the value by a dilution factor of 100 to obtain the stock absorbance. 5. In a sterile 50 mL conical tube, prepare 40 mL intermediate stock at 1  106 colony-forming units (CFU)/mL by diluting the primary stock using the following formula: 0.001/(primary stock OD600)  40 mL  1000 ¼ μL of primary stock needed in 40 mL of 10 mM MgCl2. 6. Aliquot 45 mL of 10 mM MgCl2 into two separate 50 mL conical tubes. Add 5 mL of intermediate stock and 12.5 μL of Silwet L-77. This will yield a bacterial suspension at a final concentration of 1  105 CFU/mL, 0.025% Silwet L-77. Mix by inversion. 7. Decant the suspension onto plant growth plates, and gently shake for approximately 15 s. Decant the suspension onto the next plate. This can be done in tandem with the two separate suspensions from step 6 to increase time efficiency. Return plant growth plates to light treatment boxes, and allow them to dry for 1 h with lids on (see Note 3).

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8. Reseal plant growth plates designated for bacterial quantification 2, 3, or 4 days post-infection with Micropore tape, and return to light treatment boxes. 9. A minimum of 12 healthy plants per genotype per bacterial strain (4 plants from at least 3 separate plates) is needed for quantification. 3.4 Bacterial Isolation from Plant Tissue Following Infection (Day 0)

1. Cut four plants below the rosette to remove them from the growth plate, and record their collective weight. 2. Add the plant material to the mortar and pestle with 1 mL of 10 mM MgCl2. Take care to transfer all weighed plant material. Grind well to form a slurry. 3. Transfer 20 μL of parent slurry into 80 μL of 10 mM MgCl2, and vortex, resulting in an intermediate slurry. Continue the dilution series by transferring 20 μL of intermediate slurry to 180 μL of 10 mM MgCl2. Plate 50 μL of the parent slurry and each dilution. Results will be 5  102-, 1  102-, and 1  103-fold dilutions relative to the original parent slurry. Repeat for the other two replicates within each genotype and bacterial strain combination. 4. Incubate plates at 28
C for 2 days. Choose a dilution for quantification of colonies. Bacterial colony counts between 20 and 80 are ideal. 5. Multiply bacterial counts by the dilution factor, and divide by mg fresh weight. Results are expressed in CFU/mg fresh weight.

3.5 Bacterial Isolation from Plant Tissue Post-infection (Beyond Day 0)

1. Remove and weigh four plants from each plate in a single weigh boat as directed in step 1 of the previous section. 2. Add 15–20 mL of H2O2 solution into the weigh boat and agitate for 3 min. Decant the H2O2 solution taking care not to lose any plant material. 3. Rinse surface-sterilized plants with sterile water three times, decanting water between each rinse. 4. Add the plant material to the mortar and pestle with 1 mL of 10 mM MgCl2. Grind well and add 4 mL of 10 mM MgCl2 to form a slurry. 5. Aliquot and spread 50 μL of the slurry to plate a 1  102-fold dilution. Perform additional serial dilutions by transferring 50 μL of the parent slurry into 450 μL of 10 mM MgCl2, mixing, and plating 50 μL to obtain dilutions at 1  103fold and beyond. Typically, dilutions ranging from 1  104 to 1  109 are assessed.

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6. Incubate plates at 28
C for 2 days. Choose a dilution for quantification of colonies. Bacterial colony counts between 20 and 80 are ideal. 7. Multiply bacterial counts by the dilution factor, and divide by mg fresh weight. Results are expressed in CFU/mg fresh weight.

4

Notes 1. Rubbing the Petri plate with a dryer sheet will reduce static cling of seeds to surfaces once dry. 2. Red-light treatment followed by darkness stimulates uniform germination and extends hypocotyls beyond agar surface. 3. Minimize the time plant growth plates are out of light treatment systems.

References 1. Fiorucci AS, Fankhauser C (2017) Plant strategies for enhancing access to sunlight. Curr Biol 27(17):R931–R940. https://doi.org/10. 1016/j.cub.2017.05.085 2. Huot B, Yao J, Montgomery BL et al (2014) Growth-defense tradeoffs in plants: a balancing act to optimize fitness. Mol Plant 7 (8):1267–1287. https://doi.org/10.1093/ mp/ssu049 3. Hua J (2013) Modulation of plant immunity by light, circadian rhythm, and temperature. Curr Opin Plant Biol 16(4):406–413. https://doi.org/10.1016/j.pbi.2013.06.017 4. Karpinski S, Szechynska-Hebda M, Wituszynska W et al (2013) Light acclimation, retrograde signalling, cell death and immune defences in plants. Plant Cell Environ 36 (4):736–744. https://doi.org/10.1111/pce. 12018 5. Ballare CL (2014) Light regulation of plant defense. Annu Rev Plant Biol 65(1):335–363. https://doi.org/10.1146/annurev-arplant050213-040145 6. Kangasjarvi S, Tikkanen M, Durian G et al (2014) Photosynthetic light reactions—an adjustable hub in basic production and plant immunity signaling. Plant Physiol Biochem 81:128–134. https://doi.org/10.1016/j. plaphy.2013.12.004 7. Trotta A, Rahikainen M, Konert G et al (2014) Signalling crosstalk in light stress and immune

reactions in plants. Philos Trans R Soc Lond Ser B Biol Sci 369(1640):20130235. https:// doi.org/10.1098/rstb.2013.0235 8. Jeong R-D, Kachroo A, Kachroo P (2010) Blue light photoreceptors are required for the stability and function of a resistance protein mediating viral defense in Arabidopsis. Plant Signal Behav 5(11):15041509. https://doi. org/10.4161/psb.5.11.13705 9. Jeong RD, Chandra-Shekara AC, Barman SR et al (2010) Cryptochrome 2 and phototropin 2 regulate resistance protein-mediated viral defense by negatively regulating an E3 ubiquitin ligase. Proc Natl Acad Sci U S A 107 (30):13538–13543. https://doi.org/10. 1073/pnas.1004529107 10. Yao J, Withers J, He SY (2013) Pseudomonas syringae infection assays in Arabidopsis. Methods Mol Biol 1011:63–81. https://doi.org/ 10.1007/978-1-62703-414-2_6 11. Tornero P, Dangl JL (2001) A highthroughput method for quantifying growth of phytopathogenic bacteria in Arabidopsis thaliana. Plant J 28(4):475–481 12. Ishiga Y, Ishiga T, Uppalapati SR et al (2011) Arabidopsis seedling flood-inoculation technique: a rapid and reliable assay for studying plant-bacterial interactions. Plant Methods 7:32. https://doi.org/10.1186/1746-48117-32

Chapter 13 Isolation and Characterization of Plant Metabolite Signals that Induce Type III Secretion by the Plant Pathogen Pseudomonas syringae Conner J. Rogan and Jeffrey C. Anderson Abstract Pseudomonas syringae is a bacterium that can cause disease on a wide range of plant species including important agricultural crops. A primary virulence mechanism used by P. syringae to infect host plants is the type III secretion system (T3SS), a syringe-like structure that delivers defense-suppressing proteins directly into plant cells. Genes encoding the T3SS are not transcribed in P. syringae prior to contact with a potential host plant and must be expressed during initial stages of infection. Specific organic and amino acids exuded by plants were recently identified as signals that can induce expression of T3SS-associated genes. Here we describe a technique to produce exudates from intact Arabidopsis seedlings and evaluate the exudates for the presence of these bioactive metabolites. We provide procedures for exudate production as well as downstream assays to assess T3SS gene expression using a GFP transcriptional reporter. We also describe methods for preparing high-quality protein and RNA from exudate-treated bacteria to directly assess changes in mRNA and protein abundance. These methods could be used to investigate mechanisms regulating P. syringae perception of plant metabolites as well as the release of these substances by the plant, and more generally to investigate host signals perceived by other phytopathogens. Key words Pseudomonas syringae, Arabidopsis, Extracellular metabolites, Plant exudate, Type III secretion

1

Introduction Pseudomonas syringae is a gram-negative bacterium that causes disease on a diversity of plants including important crops such as tomato and serves as an important model for molecular studies of plant-pathogen interactions [1]. P. syringae infects its host by entering into leaf tissue through natural openings such as stomata and colonizing the apoplastic space between plant cells, growing to high levels using nutrients released from host plant cells as energy sources [1]. During tissue invasion and apoplast colonization, P. syringae must evade host immune responses [2]. It does this in part by

Walter Gassmann (ed.), Plant Innate Immunity: Methods and Protocols, Methods in Molecular Biology, vol. 1991, https://doi.org/10.1007/978-1-4939-9458-8_13, © Springer Science+Business Media, LLC, part of Springer Nature 2019

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deploying a type III secretion system (T3SS), a syringe-like structure that delivers proteins termed effectors into the cytoplasm of the plant cell [3]. Many effectors are known to suppress host immune signaling pathways, creating a more favorable environment for bacterial growth [2]. Although the T3SS is required for P. syringae virulence, genes encoding the T3SS are not expressed prior to infection and must be upregulated when the bacterium comes into contact with the host [4]. Early studies found that genes encoding the T3SS can be induced by culturing P. syringae in several different minimal growth media, each containing a simple sugar and lacking complex mixtures of N- and C-containing compounds typically found in rich media [4–7]. The nature of these minimal media conditions suggested that expression of T3SSassociated genes is regulated by general environmental conditions encountered by P. syringae during infection rather than specific plant-derived signals. However, in some cases the level of T3SS gene expression was found to be higher in planta relative to levels observed in minimal media, indicating that additional T3SSinducing signals may be present in plant tissue [4, 6]. Recent studies of Arabidopsis mkp1 (mapk phosphatase 1), a loss-of-function mutant that is more resistant to P. syringae infection, provided new insight into the nature of host signals recognized by P. syringae and their importance in influencing infection outcomes [8, 9]. This work revealed that exudates prepared by soaking Arabidopsis seedlings in water strongly induce the expression of T3SS genes in DC3000. Metabolomics analysis of these seedling exudates identified several organic and amino acids including citric acid, aspartic acid, and 4-hydroxybenzoic acid as the bioactive metabolites present in these samples [9]. Importantly, exudates prepared from mkp1 seedlings had decreased levels of these bioactive compounds, and exogenous application of these compounds to mkp1 plants completely suppressed the enhanced resistance phenotype of the mkp1 mutant [9]. These results provided evidence that release of virulence-inducing metabolites is genetically regulated and opened the door for additional studies of the underlying mechanisms that control both extracellular release of metabolites from plant tissue and metabolite perception by P. syringae. Here we present our detailed protocol for preparing exudates from Arabidopsis seedlings and testing for the presence of bioactive compounds that induce the expression of virulence genes in P. syringae. Included are methods and environmental conditions required to grow seedlings suitable for exudate production (Fig. 1). We outline procedures for characterizing the basic chemical characteristics of unknown compounds present in plant exudates and provide a road map for selecting possible downstream analytical methods to identify bioactive compounds that may be present (Fig. 1). We provide a protocol for treating P. syringae with

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3.1 Arabidopsis seedlings

3.2 Seedling exudate production

3.3 seedling exudate

organic extraction

Test metabolite solubility

3.4 Incubate P. syringae with exudates aqueous

3.4.1

organic

3.4.2

GFP reporter assay for T3SS-inducing activity

Bacterial protein and RNA isolation Immunoblot/ qRT-PCR (see Notes 29, 30)

GC-MS/LC-MS analysis to identify metabolites (see Note 24)

Fig. 1 Workflow for preparing and testing exudates from Arabidopsis seedlings for presence of metabolites that induce type III secretion (T3SS)-associated genes in Pseudomonas syringae. Numbers correspond to specific subsections of Methods. Inset is photograph of 2-week-old seedlings. White arrows point to seedlings ideal for exudate production. Asterisk highlights abnormally large seedling with water-soaked leaves that should be avoided

exudates and monitoring for effects on T3SS gene expression using a GFP transcriptional reporter assay (Fig. 2). Lastly, we describe our methods for extracting high-quality bacterial RNA and protein to investigate exudate-induced effects on mRNA transcript and protein abundance (Fig. 2). The overall workflow described in this chapter could be used to screen for Arabidopsis mutants with altered metabolite release phenotypes (similar to mkp1 studies) or to identify P. syringae mutants with altered responses to host signals. Furthermore, the analytical method described could be applied to studies of exudates from other plant materials

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Fig. 2 Assays for detecting type III secretion-inducing bioactive metabolites in seedling exudates. Exudates prepared by soaking Arabidopsis seedlings in water were tested for the presence of bioactive compounds that induce T3SSassociated genes in P. syringae DC3000. (a) Time course of GFP fluorescence from a DC3000 avrRpm1promoter:GFP transcriptional reporter strain. (b) Detection of AvrPto levels in DC3000 treated with seedling exudate. Upper panel is immunoblot developed using an anti-AvrPto polyclonal antibody. Lower panel is total protein stained by Coomassie Brilliant Blue (CBB) to show equal loading

(e.g., root exudate, leaf washes, leaf apoplastic fluids) and other plant species, as well as to investigate possible host signals perceived by other phytopathogenic bacteria.

2

Materials Prepare all solutions using ultrapure water with a resistivity of
18 MΩ cm at 25  C.

2.1 Seedling Propagation

1. Arabidopsis Seedling Propagation Media: Dissolve 4.3 g of Murashige and Skoog basal salt mixture and 10 g of sucrose in 800 mL of water. Adjust the pH to 5.8 using a 5 M solution of NaOH. Add water to a final volume of 1000 mL, and divide the solution equally into two 1 L autoclavable media bottles. Add 2.9 g of agar (see Notes 1 and 2) and a magnetic stir bar to each bottle. Autoclave the prepared medium, and then cool to ~60  C. 2. Arabidopsis Seedling Propagation Vitamins: Prepare a 1000 vitamin solution by dissolving 100 mg of Murashige and Skoog vitamin powder in 2 mL of ultrapure water. Use a 1 mL syringe fitted with a 0.2 μm sterile filter to directly add 0.5 mL of the vitamin solution to each bottle of medium (see Note 3). Mix the vitamins into the medium using a magnetic stirrer (see Note 4). 3. Arabidopsis Seedling Propagation Plates: Pour the prepared agar medium into 100-mm disposable polystyrene Petri dishes. Use an autoclaved graduated cylinder or 50 mL conical

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centrifuge tube to dispense exactly 25 mL of medium to each plate (see Note 5). Replace the lids on the dishes, and allow the agar to solidify (see Note 6). Prepared plates can be stored in plastic sleeves at 4  C for up to 2 months. 4. 0.5% sodium hypochlorite (1:10 dilution of household bleach). 5. 1 mL plastic Luer-lock syringe. 6. 0.2 μM cellulose acetate membrane sterile syringe filters. 7. Triton™ X-100. 8. 95% ethanol. 9. Micropore (3M) surgical tape. 2.2 Preparation of Seedling Exudates

1. Fine-tip forceps.

2.3 Biochemical Fractionation of Exudates Based on Solubility Characteristics

1. Chloroform.

2.4 Bioassay of Exudates for Presence of T3SSInducing Metabolites

1. King’s B agar medium: 1% (w/v) peptone, 5.7 mM K2HPO4, 8.3 mM anhydrous MgSO4, 1% (v/v) glycerol, 15 g bacteriological agar [10].

2.5 Isolation of Protein and RNA from P. syringae

1. Tri Reagent® (MilliporeSigma).

2. 24-well polystyrene multi-well plates or 50 mL conical polypropylene tubes (see Note 7).

2. Methanol. 3. Freeze-dry system for lyophilization of samples. 4. SpeedVac centrifugal evaporator.

2. Modified hrp-inducing minimal medium: 10 mM K2HPO4/ KH2PO4 pH 6.0, 7.5 mM (NH4)2SO4, 3.3 mM MgCl2, 1.7 mM NaCl, 100 mM fructose [7].

2. Chloroform. 3. 70% ethanol. 4. Isopropanol.

3

Methods Perform method Subheadings 3.1, 3.2, and 3.3 in a sterile laminar flow hood or biosafety cabinet to prevent microbial contamination of agar plates and seedling exudates.

3.1 Arabidopsis Seed Sowing and Seedling Growth Conditions

1. Dispense seeds into a 1.5 mL microcentrifuge tube (see Note 8). Add 1 mL of 0.5% (v/v) sodium hypochlorite containing 0.02% Triton™ X-100 to the seeds, and gently mix for 20 min. Allow the seeds to settle, and remove the bleach solution (see Note 9).

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2. In a sterile hood, wash the seeds at least five times with 1 mL of sterile water. After the final wash, leave the seeds in 1 mL of water, and place at 4  C for a minimum of 2 days (see Notes 10 and 11). 3. Use a 1 mL pipette to sow the cold-treated seeds onto the surface of agar plates at a density of 40–50 seeds per plate (see Note 12). Allow ~1 cm distance between each seed to allow for proper seedling growth (see Note 13). Wrap the sides of the plate tightly with Micropore tape to minimize water loss from plates during seedling growth. 4. Place the seedling growth plates in a controlled environment chamber set to 21–22  C and 10 h day length (see Note 14). 3.2 Preparation of Seedling Exudates

1. Carefully remove four 2-week old seedlings from an agar plate using fine-tip forceps (see Note 15). Wash the seedlings by floating in 100 mL of sterile water for approximately 15 s to remove residual agar and media from the roots. Transfer the washed seedlings to 1 mL of water in a single well of a sterile 24-well polystyrene plate (see Notes 16–19). Replace the lid, and incubate the plate for 4–16 h under the same conditions used for seedling growth. 2. Transfer the exudate from each seedling well to a 2.0 mL microcentrifuge tube. Use exudates immediately or place at 80  C for longer-term storage (see Note 7). Measure the fresh weight of remaining seedlings to confirm that they are equal in size and number between different treatment conditions.

3.3 Determine the Solubility Properties of Unknown Bioactive Compounds in Plant Exudates

1. Thaw exudates completely, and incubate on ice. Remove an aliquot of the exudate to a clean tube and freeze; this will serve as an unfractionated input control for subsequent assays of samples post-fractionation. Add an equal volume of chloroform to the remainder of the exudate sample and vortex to mix completely. 2. Centrifuge the chloroform: exudate mixtures at 16,000  g in a microcentrifuge at room temperature for 5 min to facilitate a clear separation of the resulting organic and aqueous phases. Carefully remove the upper aqueous phase, and transfer to a clean 1.5 mL centrifuge tube. Add 500 μL of water to the remaining chloroform phase, vortex the sample, and centrifuge again using the same conditions as above (see Note 20). Remove the aqueous phase, and combine with the previous collected aqueous phase. 3. Freeze the collected aqueous phase samples in liquid nitrogen, and lyophilize to dryness using a freeze-dry system (see Note 21). Evaporate the chloroform from the remaining organic

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phase to dryness using a SpeedVac system (see Note 22). Dried samples can be stored at 80  C until use (see Notes 23 and 24). For immediate testing, re-solubilize all dried samples to original volume of the unfractionated exudate sample, and test for presence of bioactive compounds using methods described in Subheading 3.4. 3.4 Assay Plant Exudates for Bioactive Compounds that Induce Virulence Gene Expression in Pseudomonas syringae

1. Streak out appropriate P. syringae strains (see step 1 in Subheading 3.4.1) stored as 80  C glycerol stocks onto King’s B (KB) agar plates supplemented with appropriate antibiotics. Grow the bacteria for 2–3 days at room temperature until a confluent lawn is visible. 2. In a single well of a 24-well polystyrene plate, add 450 μL seedling exudate and 450 μL of hrp-inducing minimal media. Include a control treatment that substitutes water for seedling exudate. Replace the lid, and place the plate in an incubating shaker for 30 min at 22  C to equilibrate treatment solutions to the correct temperature. 3. Scrape a pea-sized amount of bacteria from a 2- to 3-day-old culture growing on KB agar with a pipet tip, and resuspend in 1 mL of water in a 1.5 mL centrifuge tube. Vortex until the bacteria are thoroughly mixed in solution, and then pellet the bacteria by centrifugation at 16,000  g for 30 s. Remove the water, and then repeat the wash step two more times (see Note 25). 4. After the final wash, prepare a 1:20 dilution of the bacteria, and measure the optical density at 600 nm (OD600) of the diluted solution using a spectrophotometer. Based on the OD600 reading, adjust the density of washed bacteria in solution to an OD600 ¼ 1.0 (see Note 26). Remove the 24-well plate from the shaker, and add 100 μL to each treatment well for a final OD600 of 0.1. Return the assay plate to the 22  C incubator, and incubate with shaking at 130 rpm.

3.4.1 Measure T3SSInducing Activity of Seedling Exudates Using an avrRpm1promoter:GFP Transcriptional Reporter

1. Incubate DC3000 carrying an avrRpm1promoter:GFP:: pBBR1MCS-2 transcriptional reporter plasmid [11, 12] with exudates as outlined in Subheading 3.4. Include treatments of DC3000 carrying an empty pBBR1MCS-2 plasmid as a negative control, and prepare at least three wells per treatment condition. 2. At the desired timepoint, remove 100 μL from each treatment well, and measure GFP fluorescence of each sample at an excitation wavelength of 485 nm and emission wavelength of 535 nm in 96-well assay plates using a fluorescence plate reader. Measure the OD600 of samples using a spectrophotometer, and use these values to normalize fluorescence readings to culture density.

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3. To account for possible non-specific fluorescence, subtract the normalized fluorescence values obtained for empty vector controls from the normalized avrRpm1promoter:GFP measurements to determine GFP fluorescence that is specific to activity of the avrRpm1 promoter. 3.4.2 Isolation of RNA and Protein from Pseudomonas syringae for Downstream qRT-PCR and/or Immunoblotting Analyses

1. Incubate wild-type DC3000 with exudates as outlined in Subheading 3.4. At the desired timepoint, transfer the treated bacteria in each well into a 1.5 mL centrifuge tube, and centrifuge at 16,000  g for 5 min. Remove the resulting supernatant and discard (see Note 27). Freeze the remaining pelleted bacteria in liquid nitrogen, and store the samples at 80  C until use. 2. Place tubes containing bacterial pellets in a Dewar flask containing liquid nitrogen. Using long-handle forceps, remove one tube at a time from the nitrogen, and quickly open the lid to alleviate any pressure buildup as the tube warms (see Note 28). Immediately add 500 μL of TRIzol® reagent to the bacterial pellet, replace the cap, and sonicate the tube in a sonicating water bath at room temperature for 30 s or until the pellet has thawed and dissolved completely. Incubate the tubes on ice after sonication until all samples are prepared. 3. Add 100 μL of chloroform to each sample, and mix well by vortexing. Incubate the samples at room temperature for 10 min, and then centrifuge the samples at 16,000  g for 10 min at room temperature. 4. Remove the upper aqueous phase, and transfer to a clean 1.5 mL tube. Add 250 μL of isopropanol, mix well by vortexing, and place at 20  C for at least 2 h to precipitate RNA. 5. To the remaining organic phase, add 150 μL of 100% ethanol and 750 μL of isopropanol. Vortex well to mix, and place at 20  C overnight to precipitate proteins. 6. For protein isolation, centrifuge tubes containing the precipitated proteins at 16,000  g for 5 min. Remove the supernatant, and wash the pelleted proteins twice with 200 μL of 80% acetone. Use a sonicating water bath to break up the pellets into a fine suspension during each wash step. Protein suspensions can be stored in 80% acetone at 20  C until use (see Note 29). 7. For RNA isolation, centrifuge the samples at 16,000 x g for 10 min to pellet the precipitated RNA. Remove the supernatant, and wash the RNA pellet twice with 500 μL of 70% ethanol. RNA pellets can be stored in 70% ethanol at 20  C or used immediately by air-drying the pellets for 10 min until dry and then solubilizing the RNA into an appropriate buffer for downstream analyses (see Note 30).

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Notes 1. Agar remains insoluble prior to autoclaving and is difficult to keep evenly distributed in solution. Adding the agar after aliquoting the media into individual bottles ensures that equal amounts are added. 2. Agar from different vendors can have different gelling properties. If a different source is used, it may be necessary to empirically determine the correct amount of agar for optimal seedling growth. For instance, agar that is less solid tends to cause seedling leaves to embed in the agar. This can cause unwanted water-soaking of the leaf tissue. 3. Syringe-end filters contain a certain amount of air space that will retain a portion of the filtered solution. Therefore, in this case it is necessary to prepare a slightly larger volume of vitamin mixture to account for this loss. 4. Stir the media slowly to avoid introducing bubbles that could adversely affect the surface quality of the solidified agar. 5. Differences in seedling growth can occur on plates that have different amounts of media, most likely due to differences in amount of available nutrients and water. Measuring out an exact amount of media for each plate minimizes this variability. 6. Plates can be air-dried with the lids removed for several minutes in order to remove any condensation that might form. However, do not air-dry the plates for extended periods of time in order to maximize the amount of water retained in the agar. Also, all plates should be air-dried for the same length of time to ensure equivalent growth environments between plates. 7. Larger amounts of exudate for biochemical fractionation or metabolite identification studies can be prepared by transferring seedlings to water in a 50 mL conical tube at a ratio of 3–5 seedlings per mL. 8. Limit the maximum number of seeds in a tube to 100 μL. The seeds will expand at least twofold once imbibed with water, and this increased volume will limit the efficacy of subsequent washes to remove the bleach solution. Use multiple tubes to sterilize larger quantities of seeds. 9. Centrifuging the seeds at 1000  g for 10 s between washes helps to make sure that all seeds have settled to the bottom of the tube. 10. Cold treatment helps to break seed dormancy resulting in more rapid and uniform germination. 11. Seeds can also be cold-treated after sowing onto the agar plates.

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12. Aspirate 400 μL of water containing 50–100 seeds into the pipet tip. Do not dispense the seeds using the plunger mechanism. Rather, hold the pipette slightly sideways, and gently tap the end of the tip onto the surface of the agar plate to dispense the seeds. Avoid sowing the seeds below the agar surface as this may alter seedling growth. If many seeds are dispensed at once, try to secure the disposable tip more tightly onto the pipette, or decrease the number of seeds within the tip while sowing. 13. Plates that are sown too densely may result in seedlings that are stressed due to nutrient or water limitation particularly after several weeks of growth. 14. We grow our seedlings on shelving that is lit by one broad spectrum T12 bulb and two 4000K T8 bulbs with an average light intensity of 1600 lux at seedling level. 15. Select seedlings that are uniform in size, color, and leaf tissue quality. This is particularly important when preparing exudates for comparative studies of different plant genotypes or treatment conditions. Avoid seedlings that have any visible tissue water-soaking caused by leaves that are in direct contact with the agar surface. In our experience water-soaked tissue releases increased amounts of metabolites, and this can cause unwanted variability between exudate samples. 16. Do not pinch the seedlings with forceps as this can cause unwanted metabolite release from damaged tissue. Instead, place the forceps tips under opposing leaves, and gently lift the seedling up. 17. Discard any seedlings with roots that break when lifting from the agar. If seedlings are difficult to remove, use a micro-spatula to remove the entire agar gel from the Petri dish, and place the agar in a dish containing a thin layer of water. Seedlings can then be easily removed without damaging root tissue. 18. Use a pipet tip to gently push down any leaves that may not be in contact with the water or roots that may be stuck to the side of the well above the water line. 19. If preparing exudates from different plant genotypes or from plants grown under different environmental conditions, prepare at least four replicate wells per genotype or treatment condition. 20. A small bubble of water will likely remain on top of the chloroform after removing the aqueous phase. The purpose of adding water and repeating the phase separation is to remove metabolites that are present in this residual aqueous solution so as to limit their contamination of the organic phase.

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21. Samples can be lyophilized directly in 1.5 mL centrifuge tubes by using an 18 gauge hypodermic needle to create a small hole in the lid of each tube. 22. A standard 50  C freeze-dry system is not sufficient to freeze chloroform, and damage could occur to vacuum pumps if used for removing chloroform. A SpeedVac centrifugal evaporator is a better choice for rapid evaporation of chloroform from samples. 23. Dried exudate samples can be resuspended in additional solvents such as methanol or acetonitrile to test for the ability of these solvents to solubilize compounds of interest. After resuspending samples in test solvents, centrifuge for 10 min at
16,000 x g to pellet insoluble material. Remove the solvent containing soluble compounds to a clean tube. Wash the remaining pellet 2–3 times to remove residual material. Evaporate the remaining solvent from both soluble and insoluble fractions using a SpeedVac evaporator. 24. Mass spectrometry-based approaches to identifying both hydrophilic and lipophilic metabolites are discussed in [13, 14]. 25. After pelleting the bacteria, a thin layer of loose material is often visible on the top of the pellet. We attribute this layer to exopolysaccharides secreted by the bacteria, and always remove this layer during the first wash step by gentle pipetting. 26. OD600 ¼ 1.0 is approximately equal to 1  109 colonyforming units per mL of DC3000. 27. Pelleted bacteria can be difficult to see in the bottom of the tube. Remove the supernatant slowly, and leave ~20–30 μL in the bottom of the tube to avoid accidently disrupting the pellet. 28. To avoid breaking hinges of the microcentrifuge tubes due to low temperatures, remove tubes from liquid nitrogen, and open the lids just enough to alleviate pressure buildup. Place the tubes in a centrifuge tube rack, and touch the hinges of each tube with a finger for a few seconds to warm. The caps can then be opened completely without breaking the hinges. 29. For SDS-PAGE analysis, remove the acetone solution and air-dry the protein pellet. Add 8 μL of 5x SDS-PAGE sample buffer to each pellet, and incubate for 20 min at 65  C to solubilize proteins. Add 12 μL of water to each tube to reach a final 2 sample buffer concentration. Vortex the samples, and incubate for an additional 20 min at 65  C. Briefly centrifuge the samples to collect any condensation that forms, and vortex to mix. Protein samples are now ready for loading onto SDS-PAGE gels.

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30. A protocol for measuring abundance of transcripts from T3SSassociated genes using quantitative RT-PCR is provided in reference [9].

Acknowledgments Research in the Anderson laboratory is funded by NSF grant 1557694 and Oregon Agricultural Research Foundation grant 8518A. References 1. Xin XF, He SY (2013) Pseudomonas syringae pv. tomato DC3000: a model pathogen for probing disease susceptibility and hormone signaling in plants. Annu Rev Phytopathol 51:473–498 2. Abramovitch RB, Anderson JC, Martin GB (2006) Bacterial elicitation and evasion of plant innate immunity. Nature Rev Mol Cell Bio 7:601–611 3. Bu¨ttner D, He SY (2009) Type III protein secretion in plant pathogenic bacteria. Plant Physiol 150:1656–1664 4. Tang X, Xiao Y, Zhou JM (2006) Regulation of the type III secretion system in phytopathogenic bacteria. Mol Plant Microbe Interact 19:1159–1166 5. Salmeron JM, Staskawicz BJ (1993) Molecular characterization and hrp dependence of the avirulence gene avrPto from Pseudomonas syringae pv. tomato. Mol Gen Genet 239:6–16 6. Rahme L, Mindrinos M, Panopoulos N (1991) Genetic and transcriptional organization of the hrp cluster of Pseudomonas syringae pv. phaseolicola. J Bacteriol 173:575–586 7. Huynh TV, Dahlbeck D, Staskawicz BJ (1989) Bacterial blight of soybean: regulation of a pathogen gene determining host cultivar specificity. Science 245:1374–1377 8. Anderson JC, Bartels S, Gonzalez Besteiro MA et al (2011) Arabidopsis MAP Kinase

Phosphatase 1 (AtMKP1) negatively regulates MPK6-mediated PAMP responses and resistance against bacteria. Plant J 67:258–268 9. Anderson JC, Wan Y, Kim Y-M et al (2014) Decreased abundance of type III secretion system-inducing signals in Arabidopsis mkp1 enhances resistance against Pseudomonas syringae. Proc Natl Acad Sci 111:6846–6851 10. King EO, Ward MK, Raney DE (1954) Two simple media for the demonstration of pyocyanin and fluorescin. J Lab Clin Med 44:301–307 11. Kovach ME, Elzer PH, Hill DS et al (1995) Four new derivatives of the broad-host-range cloning vector pBBR1MCS, carrying different antibiotic-resistance cassettes. Gene 166:175–176 12. Chang JH, Urbach JM, Law TF et al (2005) A high-throughput, near-saturating screen for type III effector genes from Pseudomonas syringae. Proc Natl Acad Sci 102:2549–2554 13. Dettmer K, Aronov PA, Hammock BD (2007) Mass spectrometry-based metabolomics. Mass Spectrom Rev 26:51–78 14. Kueger S, Steinhauser D, Willmitzer L et al (2012) High-resolution plant metabolomics: from mass spectral features to metabolites and from whole-cell analysis to subcellular metabolite distributions. Plant J 70:39–50

Chapter 14 Quantification of Cauline Leaf Abscission in Response to Plant Pathogens O. Rahul Patharkar Abstract Abscission is a process that allows plants to shed unwanted organs. Plants can use abscission as a defense mechanism to shed leaves that are heavily infected with pathogenic bacteria. By shedding infected leaves, plants completely eliminate the bacteria from the plant body, thus preventing further spreading of the disease. A lot is known about how plants limit the growth of pathogenic bacteria in vegetative leaf tissues. Much less is known about how plants defend themselves in non-vegetative developmental stages and how they use organ level responses such as leaf abscission for defense. Organ level defense responses can be effectively studied in the Pseudomonas syringae-triggered leaf abscission system in Arabidopsis. This method article describes detailed procedures for quantitative analysis of cauline leaf abscission including dissecting abscission zones for extraction of RNA and proteins for quantitative gene or protein expression analysis. The method described for molecular analysis of abscission zones could also be used in other cases where tissue is extremely limiting. Key words Leaf abscission, Defense response, Flowering plants, Pseudomonas syringae, DC3000, Bacterial infection, Limiting tissue, RNA extraction, Protein extraction, Quantitative abscission

1

Introduction As a general rule, multicellular organisms have sophisticated immune systems. Organisms with weakened immune systems often perish as they fall victim to opportunistic diseases. Plants, like other living organisms, have numerous ways of protecting themselves from disease. Plants do not have adaptive immune systems like we humans do. However, plants are capable of defense strategies that humans are not. For example, Arabidopsis and other plants can shed (abscise) leaves that are heavily infected with pathogenic bacteria [1]. Shedding an infected leaf eliminates all of the disease in the leaf from the plant body preventing the spread of the disease to healthy parts of the plant. Losing a leaf or two does not necessarily prohibit a plant’s survival since plants can simply grow more leaves when conditions are favorable.

Walter Gassmann (ed.), Plant Innate Immunity: Methods and Protocols, Methods in Molecular Biology, vol. 1991, https://doi.org/10.1007/978-1-4939-9458-8_14, © Springer Science+Business Media, LLC, part of Springer Nature 2019

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Arabidopsis has long been a favorite model organism for plant research. Many tools, resources, and a vast body of literature make molecular genetic research in Arabidopsis relatively easy compared to other plant species. A particularly well-studied research area is that of the Arabidopsis-Pseudomonas syringae interaction in vegetative stage rosette leaves [2–4]. Much less is known about the Arabidopsis-Pseudomonas syringae interaction in other parts of the plant at different developmental stages. Recently, we showed that Arabidopsis can shed its aerial stem leaves (cauline leaves) to protect itself against infection by Pseudomonas syringae pv. tomato strain DC3000 [1]. Cauline leaf abscission in Arabidopsis is regulated by a core abscission module of genes that regulates both leaf abscission and floral organ abscission and also by genes necessary for pathogen defense in leaves [1, 5, 6]. Since Pseudomonas syringae is spread by raindrop momentum, insects, and wind, shedding infected leaves reduces the sources for future infection [7–10]. This method article provides a detailed protocol to study Pseudomonas syringae-triggered cauline leaf abscission from statistical experimental design to growing plants and pathogens, treating plants with pathogen, analyzing abscission, performing molecular analysis with limited tissue, and processing data. Abscission happens at a specialized region of cells called the abscission zone (AZ) [11]. AZs typically reside at the base of the organ that will be shed [11]. AZs in Arabidopsis leaves are quite small (roughly 0.25 mm
0.3 mm
1.0 mm). At first glance this may appear to be a major limitation for doing things like gene or protein expression analysis. However, with the right modifications to existing techniques, these methods are quite straightforward. On the plus side, working with isolated AZs makes data interpretation much simpler since they are a simpler tissue than an entire leaf which is composed of several independent tissues. The methods described here for handling small pieces of tissue could also be used for other non-AZ tissues such as the shoot apex, root apex, or floral receptacle.

2

Materials Prepare solutions with ultrapure (Milli-Q or equivalent). It is not necessary to prepare soil with ultrapure water.

2.1 Quantification of Pseudomonas syringae-Triggered Cauline Leaf Abscission

1. Growth chamber set on 16 h light/8 h dark, 23  C, 50–70% relative humidity, cool white (4100 K) or daylight (5000 K) bulbs that deliver 100–150 μE m2 s1 light at plant height. 2. Standard flats for growing plants with inserts that have 36 pots per flat (Hummert Standard Inserts—Series 3600) (see Note 1). 3. Pro-Mix BX (Premier Tech Horticulture), a general-purpose plant-growing media containing perlite, vermiculite, and peat

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moss. Autoclave an appropriate amount of soil for 30 min., 121  C, on a gravity cycle to ensure pests like fungus gnat eggs are destroyed. 4. Fertilizer solution: dissolve 7.0 g Peters Peat Lite Special 20-10-20 fertilizer (Scotts, Marysville, OH) in 4.0 L water. 5. Arabidopsis seeds: accession Columbia (Col-0) seeds for plants that abscise normally when infected with Pseudomonas syringae pv. tomato strain DC3000, NahG seeds for transgenic plants that do not abscise normally when infected as compared to Col-0. 6. King’s B agar plates: combine 10 g proteose peptone #2 (DIFCO), 1.5 g anhydrous K2HPO4, 15 g glycerol, and 10 g agar, and adjust volume to 1.0 L with water. Autoclave 15 min. Add 5 mL 1M MgSO4, and mix by swirling gently (see Note 2). Once media is about 55–60  C, add desired antibiotic, and mix by swirling gently. Pour approximately 20 mL agar media into standard 15 cm plates. Store solidified plates at 4  C. 7. 80  C glycerol stock of Pseudomonas syringae pv. tomato strain DC3000 (DC3000) made by suspending freshly grown bacteria in 15% glycerol, flash freezing with liquid nitrogen, and then storing in a 80  C freezer. 8. 10 mM MgCl2 in water. 9. 1 mL syringe without a needle. 2.2 Harvesting Cauline Leaf Abscission Zones

1. Thin double edge razor blades (standard single edge razor blades can also work). 2. Fine forceps (Jeweler’s forceps #5). 3. Glass microscope slide for a cutting surface. 4. Liquid nitrogen. 5. Foam tube rack to support 0.5 mL microcentrifuge tubes upright in liquid nitrogen. 6. Shallow Styrofoam box to hold liquid nitrogen that will cool tubes. 7. 0.5 mL microcentrifuge grinding tubes with matching pestles (Research Products International, SKU: 199225).

2.3 Leaf Abscission Zone RNA Extraction

1. 40 μg/μL linear polyacrylamide (LPA): Dissolve 80 mg acrylamide (not bis-acrylamide) in 1.0 mL water in a 2.0 mL tube. Add 10 μL 10% ammonium persulfate and 2 μL TEMED. Mix and incubate at 37  C for 1 h until the solution becomes viscous. Add 0.8 mL isopropanol and mix. Spin at 10,000
g for 5 min. Discard the supernatant. Wash the pellet with 70% ethanol, and pour away the ethanol. Briefly spin and pipet away the residual ethanol. Air dry 5 min. Resuspend the pellet in 2.0 mL water. Store at 4  C (see Note 3).

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2. Trizol þ LPA: 0.5 μL LPA per 100 μL Trizol® (Invitrogen). 3. Chloroform. 4. Standard 0.5 mL microcentrifuge tubes. 5. High-salt solution: 0.8 M sodium citrate, 1.2 M sodium chloride. Add diethyl pyrocarbonate to 0.01% (w/vol). Shake 4 h. Autoclave. 6. Isopropanol. 7. 75% ethanol made with RNase-free water. 8. RNase-free water. 2.4 Leaf Abscission Zone Protein Extraction

1. 1
SDS sample buffer: 2% sodium dodecyl sulfate, 40 mM Tris–HCl pH 8.5, 10% glycerol. Add 2% 2-mercaptoethanol immediately before use (see Note 4). 2. Acrylamide mini-gels (Bio-Rad Mini-Protean or equivalent).

3

Methods Follow standard safety procedures for all hazardous reagents. All steps are carried out at room temperature unless otherwise noted.

3.1 Quantification of Pseudomonas syringae-Triggered Cauline Leaf Abscission

1. Prepare a randomized complete block experimental design (each block consists of all subjects in the experiment) for planting various Arabidopsis genotypes or treatments. Determine how many plants you will plant based on desired number of genotypes or treatments and replicates and the maximum that can be handled. Approximately 24 total plants would be a good maximum number to handle at a time to ensure mistakes do not happen frequently (see Note 5). Randomization of subjects can be accomplished on http://emerald.tufts.edu/~gdallal/ random_block_size.htm. The randomized plan will be used to plant seeds in a randomized order to minimize the effect of nuisance factors like watering level or distance from chamber blowers. See Fig. 1 for a four replicate example. Typically 7–15 replicates will be necessary to find statistical differences. 2. Prepare a flat of soil by measuring the necessary dry soil with pots or a pre-calibrated container and wetting it with 1.0 L fertilizer solution per 36 pot flat. Mix soil and fertilizer solution well (see Note 6). Distribute the soil as evenly as possible into the pots. Add water directly to the flat (below pots) so that the soil will become saturated with water after standing but not overly flooded. 3. Sow seeds that were imbibed in water and cold-treated overnight at 4  C according to the random block design from step 1. Put flats in the growth chamber with a plastic cover for 3–4 days until seeds germinate. Bottom water as needed when pots are getting dry.

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Fig. 1 Schematic diagram for using a random block design to plant an experiment with three genotypes and four replicates. First, desired genotypes or treatments are entered as shown in the top left panel along with subjects per block (in this case 3) and number of blocks/replicates (in this case 4). After pressing the “Generate Plan” button, the subjects will be randomized as shown in the top right panel. Separations between blocks were manually added to the randomized output. Finally, seeds of the three genotypes can be sown into a 36 pot flat as shown in the bottom panel. Notice that each replicate has all genotypes in the experiment very close to one another, thus minimizing the overall growth effect that position in the flat causes

4. When plants are flowering and have an inflorescence 10 cm tall, inoculate a King’s B plate with cells from the 80  C DC3000 glycerol stock. Grow the plate at room temperature or 28  C. It will take 2–3 days to see bacteria growing. Make sure plants are well-watered at this point because you will not want stomata to be closed due to drought stress when leaves are to be inoculated. 5. Harvest the cells on the plate 3 days after inoculating by scraping them off with a pipet tip, and resuspend them in 10 mM MgCl2 (see Note 7). Determine the concentration of cells by measuring the absorbance at 600 nm with a spectrophotometer or a NanoDrop. Dilute the cell suspension to 3.3
107 colony-forming units per milliliter with 10 mM MgCl2 (A600 ¼ 0.0100 on a NanoDrop or A600 ¼ 0.0790 on a GeneQuant Pro; see Note 8). 6. Several hours before the dark cycle begins in the growth chamber, infiltrate cauline leaves 1 and 2 (the first two cauline leaves to develop, Fig. 2) by gently squeezing the bacterial inoculum into the bottom of the cauline leaf with a needleless 1 mL syringe (Fig. 3) (see Note 9). This procedure should be performed at least a few hours before the dark cycle begins in the

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Fig. 2 Cauline leaves are numbered in the order they develop. While it should theoretically be possible to assay any cauline leaf, leaf 1 and 2 are ideal for studying Pseudomonas syringae-triggered leaf abscission because they are big enough to infiltrate easily

Fig. 3 Syringe infiltrating cauline leaves. (a) Bacterial suspension is squeezed into the leaf, through the stomata, by the pressure of a needless syringe placed against a leaf that is supported by your finger. (b) The bottom half of the cauline leaf appears water-soaked after it has been infiltrated

growth chamber since Arabidopsis closes its stomata shortly before the dark cycle begins. When done correctly, you will see the leaf become waterlogged (Fig. 3). Typically it is necessary to infiltrate both halves of the leaf separately. It is necessary

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Fig. 4 Gently pulling a cauline leaf on a free standing plant can reproducibly apply a small force to cauline leaves to assess whether abscission has occurred. (a) The free standing main stem is approximately at the 8 cm mark on the ruler. (b) The cauline leaf is gently pulled so that the main stem moves 1 cm to the 9 cm mark. In this example, abscission has not occurred since the cauline leaf remains attached. The yellow bracket indicates starting and ending positions of the main stem

for the part of the leaf that touches the AZ to be infiltrated for full abscission to occur. It is desirable to practice infiltration without damaging leaves prior to performing a large-scale experiment since this technique requires a certain touch. Consider practicing infiltration on rosette leaves since young flowering plants have more rosette leaves than cauline leaves. Gently blot off any residual inoculum on the outside of the leaf with Kimwipes (see Note 10). 7. Three days later, score abscission. Leaves that have abscised fall off of the plant when they are gently touched (see Note 11). Apply a reproducible “touching” force to all leaves by grasping the leaf at the distal end and gently pulling the leaf horizontally away from the stem until the stem moves 1 cm (Fig. 4). If the leaf falls off with this extremely gentle pull, abscission has occurred. In many cases when abscission has occurred, the leaf will fall off even before it is pulled. 8. Record abscission for each leaf (yes or no) organized by genotype and block (replicate). Calculate how frequently a genotype abscises with a given treatment (Fig. 5). If both leaf 1 and 2 abscise on a plant, 100% of the leaves abscised. If one of two leaves abscised, then 50% abscised, and if none abscised, then 0% abscised. Thus these data are discrete. Continuous data can be generated with a breakstrength meter (see Note 12). Figure 5 illustrates how the data can be analyzed.

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Fig. 5 Example of data analysis of an experiment with three genotypes and four replicates (blocks). Typically it will take 7–15 replicates to find statistical differences. However, all replicates do not have to be performed at the same time when using the random block experimental design 3.2 Harvesting Cauline Leaf Abscission Zones

1. Label 0.5 μL grinding tubes. 2. Cut samples alternately (e.g., cut one wild-type AZ, freeze it, and then cut the AZ of one mutant AZ). 3. Cut AZs with a thin double-edged razor blade by making four cuts (Fig. 6). Drop the AZ into the grinding tube using forceps as needed, and immediately place in liquid nitrogen so that the tube remains upright (Fig. 7). 4. After AZs have been collected for the entire experiment (WT and mutant), pulverize the AZs with a pestle cooled in liquid nitrogen. Work quickly, and do not allow the AZ to thaw prior to the addition of Trizol or SDS sample buffer. Note the grinding tubes will have collected a small amount of frozen gas that will need to be boiled off slowly and carefully first (see Note 13).

3.3 Leaf Abscission Zone RNA Extraction

1. Once tissue is ground to powder, remove the pestle with one hand (do not set pestle down), and immediately add 100 μL of TrizolþLPA directly to the bottom of the grinding tube (where it will freeze) with the other hand. Then quickly put the pestle back into the grinding tube. Once the Trizol begins to thaw, grind the mixture of Trizol and AZs until the Trizol has thawed completely. 2. After all samples are ground, mix with the pestle again, and remove the pestle keeping as much Trizol in the tube as possible. Collect pestles in a small beaker with water for washing. 3. Spin at 10,000
g for 5 min. Pipet the supernatant (about 95 μL) to a new 0.5 mL tube leaving cell debris behind.

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Fig. 6 Dissection of the cauline leaf abscission zone can be accomplished with four cuts of a new razor blade against a glass microscope slide. (a) Make the first and second cuts along the indicated numbered lines. (b) Make the third and fourth cuts along the indicated lines. (c) The released AZ is indicated and is approximately 1.0 mm
0.7 mm
0.4 mm. Optionally, the AZ can be trimmed further if an extremely high ratio of AZ to non-AZ tissue is desired; however, the indicated AZ dissection should be sufficient for most purposes

Fig. 7 Collection, freezing, and pulverizing AZs are performed in a single 0.5 mL tube per sample to prevent any sample loss. (a) A foam sponge rack can support 0.5 mL tubes in a shallow pool of liquid nitrogen. Additionally, the foam absorbs liquid nitrogen to further freeze the sides of the tubes without having the tubes floating in liquid nitrogen. (b) The complete foam rack in a Styrofoam box setup with tubes and pestles cooling in liquid nitrogen is shown

4. Incubate the sample for at least 5 min at room temperature, or freeze the samples at 20  C for processing at a later date. 5. Add 20 μL chloroform. Shake hard by hand to make a milky homogenate. Wait 2–3 min. Spin at 10,000
g for 5 min. 6. Transfer the upper aqueous phase (about 50 μL) to a new 0.5 mL tube. Add 30 μL isopropanol þ 20 μL high-salt solution (see Note 14). Vortex briefly. Wait 10 min. Spin at >10,000
g for 5–10 min. 7. Carefully remove the supernatant without disturbing the almost invisible pellet (see Note 15). Spin briefly (5 sec pulse), and remove residual supernatant.

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8. Add 200 μL 75% ethanol. Mix gently by inversion. Spin down briefly (5 s pulse). 9. Carefully remove the supernatant without disturbing the white pellet. 10. Spin down briefly (5 s pulse). Pipet out the residual ethanol. 11. Allow the pellet to dry for 1 min on its side. 12. Add 5μL RNase-free water. Vortex on speed 4. Then vortex at maximum speed several times. Store at 80  C if not using immediately. The RNA is now ready for Illumina library preparation or cDNA synthesis followed by qPCR (see Notes 16–18). 3.4 Leaf Abscission Zone Protein Extraction

1. Once tissue is ground to powder, remove the pestle with one hand (do not set the pestle down), and immediately add 20 μL of 1
SDS sample buffer directly to the bottom of the grinding tube (where it will freeze) with the other hand, and then quickly put the pestle back into the grinding tube. Grind the mixture of SDS sample buffer and AZs until the SDS sample buffer has thawed completely. 2. After all samples are ground, mix with the pestle again, and remove the pestle keeping as much SDS sample buffer in the tube as possible. Collect pestles in a small beaker with water to wash later. 3. Heat protein samples at 65  C for 5–10 min. Spin for 30 s at 10,000
g. 4. Load 5 μL per lane onto an acrylamide mini-gel with 15 lanes and a maximum gel thickness of 0.75 mm (see Notes 19 and 20). Transfer proteins to PVDF membranes, and perform Western blots according to standard procedures.

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Notes 1. Other size pots can also be used; however, 36 pot inserts provide a good balance of spacing and adequate soil per plant. 2. Adding MgSO4 before autoclaving will result in cloudy plates with precipitated salts; however, these cloudy plates do not perform noticeably worse than properly prepared plates. 3. LPA acts as a carrier for nucleic acid precipitation and will produce a visible pellet to prevent sample loss. 4. Bromophenol blue is omitted so that it is easier to watch the grinding process. The extract will be a shade of green and will be possible to load on a gel without adding any dye. If desired, a small amount of bromophenol blue can be added to the sample buffer (added until light blue by eye), but making the

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sample buffer very dark blue is not recommended since it will make the grinding process difficult to observe. 5. It is much better to perform a small to medium number of samples to ensure consistency and precision. Because we use the randomized complete block experimental design, replicates performed at different times can be combined so that the final replicate number can be enormous if necessary, which allows for great statistical power. Also, inflorescences can become tangled and hard to access when too many plants are grown in a flat. 6. Pro-Mix BX already has fertilizer in it; however, adding a small quantity of fertilizer grows noticeably bigger, healthier plants. Also, note that there can be variation in batches of soil where the batches perform differently. If plants appear slightly purple after they begin flowering, they may require a little more nitrogen. 7. Bacteria can also be streaked from an existing plate rather than a glycerol stock. Grow bacteria for 2 days if streaked from an existing plate. Generally speaking, it is not desirable to serially grow bacteria from plates for repeated rounds since mutations and contamination are possible. 8. DC3000 cell suspensions at A600 ¼ 0.001 (NanoDrop) will also reliably trigger abscission at a slightly lower frequency than A600 ¼ 0.01. Cell suspensions at A600 ¼ 0.001 will not trigger a hypersensitive response, rather leaves will gradually yellow and then abscise. A600 ¼ 0.01 DC3000 cell suspensions will trigger a nonspecific hypersensitive response after 24 h. DC3000 cell suspensions at A600 ¼ 0.0001 do not reliably trigger abscission or cause obvious disease symptoms in Col-0 cauline leaves. 9. Infecting cauline leaves by spraying them with high concentrations of DC3000 with Silwet L-77 did not result in noticeable disease symptoms or abscission. 10. Failure to blot away excess inoculum on the outside of the leaf can result in reduced abscission, possibly due to the infection progressing too quickly for the abscission program to be executed properly. In these cases, AZ cells will enlarge, but full abscission may not occur. 11. In most growth chambers, touching leaves is necessary to make abscised leaves fall off because there is insufficient wind to blow the leaves off. 12. Continuous data can be generated with a breakstrength meter which measures the force required to pull leaves off of the plant. However, to do so requires the construction of a breakstrength meter [12]. Additionally, data acquired with a

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breakstrength meter is not completely normally distributed. A lot of the measurements either indicate very tight attachment or extremely loose attachment (leaf has abscised) with less measurements in between. 13. The tubes in the liquid nitrogen are so cold that gradually air will liquefy and collect in the bottom of the tube. When tubes are temporarily removed from the liquid nitrogen, for handling purposes, the liquefied air will boil rapidly and could result in your AZs flying out of the tube. To prevent AZ loss, temporarily cap the tube loosely with an extra cap (cut off from a spare tube) that has holes poked in it with a 26 gauge needle, and allow the liquefied air to boil away prior to handling open tubes outside of liquid nitrogen. This spare cap with holes acts as a screen that keeps the AZs in while allowing gas to escape. 14. High-salt solution along with isopropanol keeps some polysaccharides in solution so that the final RNA is purer. However, adding 25 μL isopropanol þ 25 μL high-salt solution as suggested in the Trizol® protocol will not precipitate LPA; thus the RNA pellet will be impossible to track. 15. In isopropanol the pellet is almost invisible. Holding the tube at just the right angle to the light may allow you to visualize the pellet. Once ethanol is added, the pellet will be white and easy to see. 16. The integrity of the RNA can be visualized by mixing 0.5 μL RNA with an 0.5 2
RNA denaturing buffer (7 M Urea, 2
TAE buffer, 10% glycerol, 0.01% bromophenol blue) and incubating at 65  C for 5 min. Then run the RNA on a standard 1
TAE gel with 1.2% agarose. The key to clearly visualizing very small amount of RNA is to use wells that are very small so that your sample is as concentrated as possible on the gel. Wells that are 2 mm
0.75 mm are ideal. 17. HAESA (At4g28490) can serve as a good positive control for a gene that increases as abscission progresses, and At5g46630 can serve as a reference gene [5]. 18. The expected yield is approximately 400–500 ng total RNA per AZ. While a single AZ provides sufficient RNA for qPCR, up to 10 AZs can be extracted in a single 100 μL Trizol extraction. 19. Small wells and thin gels are key for having concentrated protein bands. It is possible to visualize low abundance proteins such as transcription factors on Western blots with protein extract from a quarter of an AZ with this protocol [13, 14]. 20. The remainder of the sample can be loaded onto three other gel lanes or saved at 20  C.

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Acknowledgments I would like to thank Catherine Espinoza Patharkar for reading and editing the manuscript and John Walker for supporting this work. References 1. Patharkar OR, Gassmann W, Walker JC (2017) Leaf shedding as an anti-bacterial defense in Arabidopsis cauline leaves. PLoS Genet 13: e1007132 2. Jones JDG, Dangl JL (2006) The plant immune system. NATNEWS 444:323–329 3. Abramovitch RB, Anderson JC, Martin GB (2006) Bacterial elicitation and evasion of plant innate immunity. Nat Rev Mol Cell Biol 7:601–611 4. Zipfel C (2008) Pattern-recognition receptors in plant innate immunity. Curr Opin Immunol 20:10–16 5. Patharkar OR, Walker JC (2016) Core mechanisms regulating developmentally timed and environmentally triggered abscission. Plant Physiol 172:510–520 6. Patharkar OR, Walker JC (2017) Advances in abscission signaling. J Exp Bot. https://doi. org/10.1093/jxb/erx256 7. Hirano SS, Upper CD (2000) Bacteria in the leaf ecosystem with emphasis on Pseudomonas syringae—a pathogen, ice nucleus, and epiphyte. Microbiol Mol Biol Rev 64:624–653 8. Upper CD, Hirano SS, Dodd KK et al (2003) Factors that affect spread of Pseudomonas

syringae in the phyllosphere. Phytopathology 93:1082–1092 9. Bashan Y (1986) Field dispersal of Pseudomonas syringae pv. tomato, Xanthomonas campestris pv. vesicatoria, and Alternaria macrospora by animals, people, birds, insects, mites, agricultural tools, aircraft, soil particles, and water sources. Can J Bot 64:276–281 10. Lilley AK, Hails RS, Cory JS et al (1997) The dispersal and establishment of pseudomonad populations in the phyllosphere of sugar beet by phytophagous caterpillars. FEMS Microbiol Ecol 24:151–157 11. Addicott FT (1982) Abscission. University of California Press, Oakland, CA 12. Lease KA, Cho SK, Walker JC (2006) A petal breakstrength meter for Arabidopsis abscission studies. Plant Methods 2:2 13. Patharkar OR, Walker JC (2015) Floral organ abscission is regulated by a positive feedback loop. Proc Natl Acad Sci 112:2906–2911 14. Patharkar OR, Macken TA, Walker JC (2016) Serine 231 and 257 of Agamous-like 15 are phosphorylated in floral receptacles. Plant Signal Behav 11:e1199314

Chapter 15 Methods for Replicating Leaf Vibrations Induced by Insect Herbivores Sabrina C. J. Michael, Heidi A. Appel, and Reginald B. Cocroft Abstract Testing plant responses to natural sources of mechanical vibration requires methods that can precisely reproduce complex vibrational stimuli. Here we describe a method for conducting high-fidelity vibrational playbacks using consumer audio equipment and custom-written signal processing software. Key words Plant mechanosensing, Plant–insect interactions, Vibrational communication, Herbivory

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Introduction There is great current interest in the role of acoustic information in plant–environment interactions. Recent trends in the field include a focus on sources of acoustic energy relevant to plants in natural environments [1–8], study of the sensory receptors involved in responses to acoustic stimuli and of downstream responses to activation of these receptors [9–18], the agricultural use of sound to influence plant growth and development [16–19], and adoption of experimental methods from the study of plant-borne vibrational communication in animals [5, 20, 21]. The terminology in this literature is in flux. The term “plant acoustics” has often been used, though “acoustic” typically refers to airborne sound and its detection by sensory structures that have evolved to detect pressure waves. Plants lack specialized sound-detecting structures, and although airborne sound does induce mechanical vibrations in plant tissues [22], we are unaware of any published work on whether natural sources of airborne sound are relevant to plants. Some authors have used the phrase “sound vibration” to draw attention to the function of plant mechanoreceptors in sensing vibrations in plant tissue [13, 14, 18], and the term “biotremology” has been proposed to encompass study of the biological role of any form of mechanical wave [23]. The study of plant responses to acoustic or

Walter Gassmann (ed.), Plant Innate Immunity: Methods and Protocols, Methods in Molecular Biology, vol. 1991, https://doi.org/10.1007/978-1-4939-9458-8_15, © Springer Science+Business Media, LLC, part of Springer Nature 2019

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vibrational stimuli is also closely related to the study of plant responses to wind or touch, or “thigmomorphogenesis” [11]. In this chapter we follow recent literature, using “acoustic” as a general reference to information carried by some form of mechanical wave, and “vibration” to refer to the stimuli perceived by plants. The activity of herbivores, whether feeding, moving, or signaling, is one of the most prevalent biotic sources of mechanical vibrations in plants in nature [20]. The importance of herbivoreinduced vibrations in insect–plant interactions is highlighted by the finding that the feeding vibrations of herbivores can cause priming of plant defenses [5]. Other types of ecological interactions may also involve plant responses to mechanical vibrations, and a host of questions remain to be addressed about how plants perceive herbivore vibrations, distinguish them from nonrelevant vibrations, and respond both locally and systemically. To address questions about the influence of vibrations produced by herbivores or other natural sources, it is necessary to be able to measure and experimentally reproduce the mechanical vibrations of plant tissues. This vibrationmeasurement-and-reproduction step has been rate-limiting for the study of vibration-mediated interactions between plants and their environment, given the need for specialized and expensive instruments. Airborne sound has often been used as a stimulus, but although it does induce vibrations in plants, plant structure greatly influences the properties of those vibrations, yielding an uncertain relationship between the stimulus that is produced by the experimenter and the one that is experienced by the plant (see below). Our laboratories are developing vibrational playback methods based on relatively inexpensive, off-the-shelf consumer equipment. Vibration recording and measurement is a broad topic that is outside the scope of this chapter, and here we only discuss those sensors and procedures that are necessary for properly calibrating vibrational playbacks. Our focus is on experimentally testing plant responses to mechanical vibrations, while ensuring that the stimuli experienced by the test plants have the desired properties. Although many of the issues that arise when conducting vibrational playbacks on plants have been discussed elsewhere [20], the high throughput required for testing plant responses requires carefully calibrated vibrational stimuli on multiple plants at the same time. Here we describe a method that achieves calibrated, multichannel vibrational playbacks using a combination of signal processing software and consumer audio equipment.

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Materials Overview: The basic setup described here (Fig. 1a) for playing back herbivore vibrations to plants consists of a computer with signal processing software; an audio interface device; an amplifier; a set of

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Fig. 1 The experimental setup: (a) layout; (b) diagram of method of attachment of a plant leaf to the vibration playback device

vibration transducers; a vibration sensor; an oscilloscope; cables for connecting the equipment; vibration isolation; and a playback device and a method for connecting it to the plant (Fig. 1b). 1. Computer with appropriate hardware drivers installed (see Note 1). 2. Software: (a) For signal processing: Matlab (Mathworks, Inc.) including the Signal Processing toolbox (see Note 2). A customwritten script for prefiltering the playback stimuli and adjusting their amplitude is the centerpiece of highfidelity vibrational playbacks. We guide the user through a custom-written Matlab script with a graphical user interface to be used for vibrational playbacks. The script can be downloaded at the following web address: https://gre envibes.missouri.edu/vibe-ware/.

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(b) For recording and editing signals: Audacity (https:// www.audacityteam.org/), a user-friendly free shareware program. (c) For playing back stimuli: a digital audio workstation for playing back stimuli on multiple channels simultaneously (see Note 3). 3. Multichannel audio interface (see Note 4). 4. Vibration transducers for playback: a small 4- or 8-ohm speaker (see Note 5) with suitable modification as described in Note 6. 5. Amplifier: a multichannel amplifier for driving the vibration playback devices to deliver vibrational stimuli to several plants at a time (see Note 7). 6. Small, calibrated vibration sensor such as a miniature accelerometer (see Note 8). 7. Vibration isolation (see Note 9). 8. Oscilloscope: a digital or analog oscilloscope (see Note 10). 9. Cables: for each channel in the setup below, use one ¼00 (6.35 mm) male to RCA male mono cable for connecting the interface outputs to the amplifier input; a length of speaker wire, 5–100 (1.5–3 m) depending on the configuration of the lab space, for connecting each amplifier output to the corresponding speaker; one BNC female—BNC female cable for connecting the accelerometer power supply output to the oscilloscope input via one BNC T-connector; and one BNC female to ¼00 (6.35 mm) male mono cable for connecting the oscilloscope via the T-connector to the input channel of the interface. 10. Adhesive: accelerometer mounting wax, soft dental wax, or other nonpermanent adhesive to couple the playback device and the vibration sensor to the playback device and/or plant (Fig. 2; see Note 11). 11. A clamp for holding and positioning the plant pot (see Note 12).

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Methods The two basic challenges in conducting vibrational playbacks are (1) to compensate for the filtering effect of the playback device and substrate and (2) to play back the stimulus at an appropriate amplitude [20]. When these issues are addressed using appropriate hardware and signal processing software, any of a wide variety of vibration playback devices can produce high-fidelity results like those shown in Fig. 3a, b. If these issues are not addressed, then regardless of the vibration playback device used, the stimulus delivered to the plant will depart in unknown ways from the stimulus the

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Fig. 2 Modifying a small audio speaker to make it a suitable vibrational playback device. (a) The speaker before modification; (b) the speaker with the paper membrane removed; (c) the speaker with a graphite rod added and an accelerometer attached

experimenter intended to deliver. Likewise, filtering by plant structures will markedly change the spectrum of stimuli played back as airborne sound (Fig. 3c). Here we assume that the investigator plans to use natural stimuli with a broad frequency range, such as excerpts from vibration recordings of herbivore activity on plants. However, the methods described here are equally suitable for other stimuli including those generated using software. 3.1

Input Calibration

1. Before calibrating the input gain, ensure that the computer is communicating properly with the interface. Check all connections and switches; make sure the computer is set to record and play back through the desired device; and check the sound card settings on your computer (see Note 13). 2. Calibrate the input channel gain on the interface to a level of 1.0 to adjust the amplitude of a playback. Perform this procedure initially; periodically check; and re-do when using a different sensor or if the gain control setting is incidentally changed. (a) For the speaker–accelerometer combination, attach the accelerometer to the vertical rod on the modified speaker using wax (Fig. 2c) when calibrating the playback device (see Note 14).

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Fig. 3 Comparison of a recording of vibrations produced by a feeding caterpillar, and a recording of the played-back vibrations using the modified speaker shown in Fig. 2b, c and the procedure shown in Fig. 4. (a) Waveforms of the original recording (after conversion to acceleration; see Note 17) and the recorded playback (duration of waveforms ¼ 10 s). (b) Frequency-vs.-amplitude spectra

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Fig. 4 The graphical user interface (GUI) created in Matlab for prefiltering vibrational playback stimuli and adjusting their amplitude. The initial settings can be adjusted in panel a. The system filter is calculated in panel b. The calculated system filter is applied to the experimental playback stimuli and the amplitude is also adjusted and applied in panel c. See Subheading 3.3 for detailed instructions

To calibrate the audio interface input gain for channel 1, generate a pure-tone stimulus of several hundred Hz (this can be done in Audacity), and loop it to play continuously. To play the tone while recording it, in Audacity select Preferences ! Recording and check “Overdub: play other tracks while recording new one.” Then adjust the gain for input channel 1 until the amplitude of the tone in Audacity matches that on the oscilloscope. Note that the scale in Audacity is þ/ 1 V, so that if the oscilloscope shows a peak amplitude of 300 mV, adjust the input gain until the peak amplitude in Audacity is 0.3.

ä Fig. 3 (continued) of the same two signals, showing the close match of the recorded playback and the original recording. (c) Amplitude spectra of airborne sound (a noise burst) played from a speaker 20 cm from an Arabidopsis thaliana plant, and of the induced vibrations measured on one of the leaves using a laser vibrometer, showing the marked differences between the airborne sound and the leaf vibrations

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3.2 Output Calibration

1. Set the output gain on the amplifier and interface. Start with a low gain setting on the amplifier, about ¼ of the maximum on each amplifier channel that will be used (see Note 15). 2. Once the input gain is calibrated, use the provided Matlab script (see Subheading 2, step a) to prefilter the playback stimuli to compensate for the frequency response of the playback device and substrate, and to adjust the amplitude of the playbacks (Fig. 4). Run the procedure for each channel used in the experiment (see Note 16).

3.3 Playback Stimulus Prefiltering and Amplitude Adjustment

1. Use the custom software (see Subheading 2, step a) to measure and compensate for the frequency response of the playback device and match the amplitude of the playback stimuli to that of the original stimuli. It is necessary to match the average peak or RMS amplitude of those stimuli, or to match other amplitude levels called for by the experimental design (see Note 17). In this case, this step was done before the leaf is attached (see Note 18). 2. Ensure that the following files are in the same folder: For example, VIBE_PB_GUI_PC.m, VIBE_PB_GUI_PC.fig, and all stimulus files in .wav format. 3. Choose the channel corresponding to the playback device for which stimuli are currently being prepared, and to which the accelerometer is attached (Fig. 4a(i)). 4. Enter a frequency range that slightly exceeds the relevant frequency range of the stimuli (Fig. 4a(ii)) (see Note 19). 5. Enter the desired amplitude level for the playbacks (Fig. 4a(iii)) (see Note 17). 6. Enter the appropriate amplitude units. For an accelerometer, set to m/s2 (Fig. 4a(iv)). 7. Select the type of amplitude measurement, peak (used for most stimuli) or RMS (Fig. 4a(v)) (see Note 17). 8. Select the type of sensor used for calibrating the playbacks and the gain used (Fig. 4a(vi)). 9. If desired, save the above settings for future use using the File menu (top left, just below the title). 10. Click on “Play and record original signal”(Fig. 4(i)). The program will generate a short burst of random noise and play it through the playback device, while recording from the accelerometer. After a few seconds, the waveform of the recorded signal should appear, along with the amplitude spectra of the random noise and the recorded signal. If there is clipping of the recorded signal, the program will lower the amplitude and play the signal again until the signal is not clipping. After the waveform and spectra appear, and assuming the recorded signal

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looks like the one shown in Fig. 4, proceed to the next step. If the signal does not appear to be correct refer to Note 20. 11. Click on “Play and record 1 filtered signal” (Fig. 4b(ii)). The program will play back the original noise signal modified by the calculated filter. The waveform and spectra will appear, and the spectrum of the recorded signal should be closer to that of the original playback. It may not match the original very closely, however; especially for frequencies that are strongly attenuated by the playback system, the calculated system filter will be imprecise. 12. Click on “Play and record 2 filtered signal” (Fig. 4b(iii)). The spectra of the recording and the original playback should now be very similar, because the original noise has been modified to compensate for the system filter (e.g., the amplitude of frequencies attenuated by the playback system will have been correspondingly increased). If the played-back signal does not appear to be correctly filtered, refer to Note 20. 13. Click on “select file(s)” (Fig. 4c(i)); a browser window will open and the stimuli to be played from the current channel can be selected. Once a stimulus has been filtered and its amplitude adjusted (the default is to within þ/ 1 dB across the desired frequency range), it will be saved and the waveform and spectra displayed; if there are additional stimuli to be filtered, it will move onto the next after a few seconds. If some of the playback stimuli adjusted for the current channel will be played back at different amplitudes, click on “select file(s)” again after changing the amplitude value (see Fig. 4a(iii)). 14. Once all of the stimuli for the current channel have been saved, move the accelerometer to the next playback device or plant. Plug the Channel 1 output (from interface device) to the next channel on the amplifier, and select the corresponding channel (see Fig. 4a(i)), which will cause a channel identifier to be added to each filtered file (see Note 21). 3.4 Attaching Playback Device to Plant

1. Once all of the playback stimuli have been filtered and amplitude-adjusted for each channel, attach plants to the playback devices using wax, either accelerometer mounting wax or dental wax, the best method tested thus far for coupling the playback device to the plant. Take care when attaching the leaf to the playback device and (especially) when detaching it at the end of the playback, to avoid injury to the leaf (see Note 22). Because this method involves physical contact with the plant, include a control treatment in which leaves are likewise attached to silent playback devices or a sham. 2. To position the plant to allow contact with the playback device, use a clamp (Fig. 1b) to hold an empty plant pot and then insert the pot containing the playback plant into the empty pot.

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3. Use the clamp to adjust the height and orientation of the plant and to position the target leaf. Position the end of the dowel so that it is lightly touching the target leaf (see Note 23). If the leaf needs to be bent toward the dowel, the force exerted by the leaf petiole will tend to detach the leaf from the wax. 4. Once the leaf is in position, apply gentle pressure for 2–3 s from the opposite side of the leaf using a cotton swab, so that the leaf is lightly but firmly adhered to the wax to achieve a nondestructive attachment. Test the connection by gently lifting the edge of the leaf 1–2 mm from below. 3.5 Creating Playback Files

1. To deliver the same stimulus at regular intervals: (a) Open wav file containing the filtered and adjusted playback stimulus. (b) Select “Generate” and then “Silence.” (c) Choose the desired time of silence needed before and after the playback stimulus. (d) Use loop play in Reaper to generate a continuous playback track (see below). 2. To create a playback file in Audacity with multiple filtered and adjusted playback stimuli and various silent gaps: (a) Follow the steps above in Subheading 3.5, step 1 to add silence before and after stimulus. Use “Loop play” in Reaper to generate a continuous playback track (see below). (b) If using multiple stimuli in one playback channel, simply copy one stimulus exemplar from the filtered/adjusted file. To paste, first have the Audacity file to be used for the playback open, and zoom in on the waveform until the individual samples are visible. Paste the stimulus where the individual sample is at zero. 3. Load the adjusted playback stimuli for playing back to the plants into Reaper: (a) Insert a separate track (Select “Track” and then “Insert new track”) for each playback channel. (b) Load the playback files one at a time (and only one per track) by selecting “Insert” and then “Insert Media File.” (c) Select the “Route” icon on each track and deselect the “Master Send” option, then select the output channel for each track. For example, the first track should be sent to “Output Channel 1”; the second to “Output Channel 2”; etc.

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Notes 1. Apple computers usually allow trouble-free connection to commercial audio devices. ASUS and other PC computers are also suitable with the appropriate drivers. 2. We use Matlab for methods described in this chapter, but there is a free shareware program, GNU Octave (https://www.gnu. org/software/octave/), which mimics Matlab, including the signal processing toolbox. However, we have not tested this Matlab script in Octave. 3. For playing back stimuli: a digital audio workstation for playing back stimuli on multiple channels simultaneously, such as Reaper (which offers an inexpensive education license and was used in this chapter), ProTools, or Cubase. 4. An audio interface functions as an external sound card for the computer and provides multichannel input and output. In this chapter we used the Tascam US 2020. 5. Virtually any small 4- or 8-ohm speaker can be used to produce high-fidelity vibrational playbacks, with the proper modifications and signal processing. The procedures described in the chapter are based on an inexpensive 8-ohm speaker. 6. We used small audio speakers, modified to reduce the emission of airborne sound and to allow coupling of the playback signal to the plant (Figs. 1b and 2). When using a speaker as a playback device, first remove the paper membrane to reduce the production of airborne sound. Then attach a graphite rod (or a wooden dowel) to the speaker to allow coupling between the moving speaker coil and the plant (Fig. 2b). Some airborne sound will still be produced by the speaker, but the vibrations transmitted to the plant via the rod will be substantially higher in amplitude than any vibrations induced in the plant by the airborne sound. 7. An ever-changing array of makes and models of amplifiers is available commercially, typically with a flat frequency range of 20–20,000 Hz (and usually the ability to amplify signals with frequencies outside this range). The procedures described in the chapter use a Behringer HA8000 amplifier, which is used primarily for headphones but is also capable of driving 8 ohm speakers. 8. The basic requirements for a vibration sensor to be used in the filtering and amplitude adjustment steps are that the vibration sensor is calibrated (i.e., the relationship between vibration amplitude and sensor output has been measured); and that it is small, to minimize mass loading of the structure it is attached to, because mass loading changes a structure’s vibration-

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transmitting properties. We suggest using a miniature accelerometer with an output of 100 mV/G and a frequency range from 50 to 70% necrosis/ooze

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>70 to 90% necrosis/ooze

8

>90% necrosis/ooze

3.7 Disease Severity Rating

1. Rate the percentage of fruitlet area displaying water-soaking, necrosis and/or ooze to assign each fruitlet to a severity category according to a disease rating scale (Fig. 1b, c and Table 1; see Note 11).

3.8 Bacterial Growth Measurement

1. Prepare microcentrifuge tubes for serial dilutions by adding 900 μL of 10 mM MgCl2 per tube (see Notes 12 and 13). 2. Label an Agdia bag with strain and replicate number for each apple half to be processed. Add 1 mL of 10 mM MgCl2 to each bag. 3. Use a bag filled with 1 mL 10 mM MgCl2 to zero a balance. 4. Using forceps cleaned with 70% ethanol, transfer each apple half into the lower corner of the Agdia bag, farthest from the meshless channel. 5. Weigh and record the weights of the bags with apples in them. 6. Smash the apples in the bags with a plastic soft face hammer until no large pieces remain. Avoid forcing pulp toward the top of the bag or into the meshless pipetting channel, as it will make pipetting the juice difficult. 7. Thoroughly macerate the apple tissue with a drill press equipped with the Agdia homogenizer attachment. Repeatedly apply pressure to the tissue in the bag, carefully crushing the tissue from the top of the bag toward the bottom and away from the channel (see Note 14). Homogenize each sample for about 30 sec, and be consistent about the amount of time each sample is macerated.

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8. Mix the liquid around in the bag to ensure a homogeneous sample. Starting at the side of the bag furthest from the channel, roll the bag toward the meshless channel to fill the channel with the liquid homogenate. The apple pulp should remain within the mesh. 9. Using a wide-bore 1000 μL pipet tip, remove 100 μL of homogenate from the channel, taking care to avoid the pulp (see Note 15). If the tip gets clogged by pulp when filling with the liquid from the channel, dispose of it and start with a fresh tip. 10. Dispense the homogenate with a fresh tip into a prepared microcentrifuge tube of 900 μL of 10 mM MgCl2 for the first dilution of the series. Vortex well, and transfer 100 μL into the next tube of the series. 11. Repeat the dilution steps as many times as needed to achieve the required cell dilutions, using a fresh tip each time. Vortex each tube well after the addition of the 100 μL aliquot before removing the next aliquot. 12. Plate the resulting dilutions on LB plates supplemented with any necessary antibiotics, and incubate plates at 28  C (to reduce the number of plates required, see Note 16). 13. After 2 days, count the colonies at an appropriate dilution, and use the weight of the apple and the dilution factor to calculate CFU/g of apple tissue (see Notes 17 and 18).

4

Notes 1. The mesh sample bags produced by Agdia (Product # ACC 00930) are the only ones we have used with this protocol to date. 2. It is best if trees have not been treated with a chemical thinning agent, although suitable fruits may still be found on such trees. 3. We routinely use fruit of the highly susceptible cultivar “Gala,” but we have also used other cultivars for successful fire blight infections, including “Golden Delicious,” “Rome,” and “Fuji.” The timing of symptom development can vary between cultivars and should be determined prior to setting up largerscale experiments. Of the cultivars we have tested to date, we have found that “Gala” has the best combination of fire blight susceptibility and survival during cold room storage. 4. Apple fruitlets remain viable and susceptible longest when stored at temperatures very near 0  C. Fruitlets kept in a standard 2  C cold room remain suitable for experiments for at least 4 months after harvest. Store fruitlets in an onion

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produce mesh bag, which is then placed into a loosely wrapped plastic bag as well to help the fruitlets retain moisture. Once fruitlets begin to lose moisture, showing wrinkles at the peduncle end, disease responses may be attenuated. 5. Only wash enough for use within 3 weeks. Apple fruitlets will spoil more quickly if they are not thoroughly dried after bleaching. 6. Alternatively, bacteria may be grown in liquid medium until late log phase (typically 12–16 h in LB), and the cells pelleted and washed twice with 10 mM MgCl2, as described. It is not necessary to prepare and apply E. amylovora inoculum in a biosafety cabinet. Solutions, tubes, and tips should be autoclaved before use, but may thereafter be used on the bench. 7. There should not be any brown tissue remaining at the stylar end after trimming, as this can be a source of fungal contamination. 8. The hole should be large enough to hold 20 μL of inoculum. The technique should be practiced ahead of time to ensure the correct sized holes will be created. 9. Within about 30 min, the inoculum solution will be absorbed into the fruit tissues and will no longer be visible. 10. Light may be provided during incubation, but does not appear to be necessary for normal bacterial multiplication in immature apples. 11. Fire blight disease symptoms develop on the apples over the course of a week or more (Fig. 1b). The exact timing of disease development can vary substantially, depending on bacterial strains used, apple fruitlet age, and ambient humidity conditions. 12. The goal is to dilute the homogenate sufficiently to allow counting of between 20 and 200 colonies when plating. This range of colony numbers allows for accurate bacterial population estimation. 13. Optimal sampling time points can vary based upon the bacterial strains and apple cultivars of interest and should be decided based on rate of visible disease progression (Fig. 1b). A zero time point (immediately after inoculation) is advised, to confirm accurate starting populations and ensure consistency between experiments. For initial (day 0) bacterial populations, a 1000-fold dilution should suffice for dilution plating. On subsequent days, populations may reach as high as 5  109 CFU/g, requiring up to eight 10-fold dilutions to produce countable numbers of colonies (see Note 12). Preliminary tests can be used to clarify the number of dilutions required at various dpi.

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14. To prevent pulp or juice from squirting out of the bag while homogenizing, the opening of the bag can be folded over to make a loose seal. Use gentle pressure on the pulp at first when larger apple pieces are present, and stronger pressure on the pulp as the tissue becomes liquefied. 15. Insert the wide-bore tip into the homogenate in the bag as little as possible to minimize carry-over on the exterior of the tip. When dispensing the homogenate into the first tube of MgCl2 in the dilution series, again insert tip as little as possible to avoid dispensing any liquid clinging to the exterior of the tip, for best accuracy. 16. One can greatly reduce the numbers of plates and amount of media used by employing the “drip plate” method. Quickly arrange four to five 10–20 μL droplets of your dilutions in a row on a typical 100 mm  15 mm LB petri plate and then tip the plate so that the suspensions move down the plate to spread out the droplets, being very careful not to let the droplets run into each other or run into the plate edge. The arrangement of the droplets and tilting of the plate should be done as quickly as practical, since the droplets will begin to soak into the agar and will become less spreadable within a short period of time. It is possible to test 4–5 dilutions on a single petri dish in this manner. 17. Symptoms typically appear at 4–5 dpi, and the changes in bacterial population occur mainly in the first 5 dpi. Therefore, to quantify changes to early and mid-log phase growth, it is advisable to use more frequent and earlier time points. In our experience, populations at 3 dpi are approximately 2.2  108 cfu/g; at 5 dpi, they are about 1.2  109 cfu/g; and by 7 dpi, they reach approximately 5  109 cfu/g. 18. Background (contaminating) bacterial growth is generally minimal on dilution series plates, but can be essentially eliminated by using spontaneous antibiotic-resistant derivatives of wild-type E. amylovora strains.

Acknowledgments We thank Dr. Megan Dewdney for originally recommending the apple fruit inoculation approach. We also thank Drs. Steven A. Lee and Henry K. Ngugi, and Mr. Brian L. Lehman, for their contributions in developing the inoculation and incubation procedure. We thank Dr. Kari Peter and Brian Lehman for the information about apple fruitlet fungicide treatments. This work was supported in part by a United States Department of Agriculture National Institute of Food and Agriculture (USDA NIFA) Predoctoral Fellowship

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(Grant # 2017-67011-26030) to S.M.K, a grant from the State Horticultural Association of Pennsylvania to T.W.M and H.K.N., the Penn State Department of Plant Pathology and Environmental Microbiology, and USDA NIFA Hatch Appropriations under Project #PEN04649 and Accession #1016093. References 1. van der Zwet T, Orolaza-Halbrendt N, Zeller W (2012) Fire blight: history, biology, and management. Am Phytopathol Soc Press, St. Paul 2. Denning W (1794) On the decay of apple trees. Trans Soc Promotion Agric Arts Manufact Inst State NY 1:219–222 3. Arthur JC (1885) Proof that bacteria are the direct cause of the disease in trees known as pear blight. Am Assoc Adv Sci Proc 34:294–298 4. Coxe W (1817) Pears. In: A view of the cultivation of fruit trees, and the management of orchards and cider, M. Carey and Son, Philadelphia 5. Oh C-S, Beer SV (2005) Molecular genetics of Erwinia amylovora involved in the development of fire blight. FEMS Microbiol Lett 253:185–192 6. Vrancken K, Holtappels M, Schoofs H et al (2013) Pathogenicity and infection strategies of the fire blight pathogen Erwinia amylovora in Rosaceae: state of the art. Microbiology 159:823–832 7. Kim JF (2001) Revisiting the chlamydial type III protein secretion system: clues to the origin of type III protein secretion. Trends Genet 17:65–69 8. Smits TH, Rezzonico F, Kamber T et al (2010) Complete genome sequence of the fire blight pathogen Erwinia amylovora CFBP 1430 and comparison to other Erwinia spp. Mol PlantMicrobe Interact 23:384–393 9. Sebaihia M, Bocsanczy AM, Biehl BS et al (2010) Complete genome sequence of the plant pathogen Erwinia amylovora strain ATCC 49946. J Bacteriol 192:2020–2021 10. Moreau M, Degrave A, Vedel R et al (2012) EDS1 contributes to nonhost resistance of Arabidopsis thaliana against Erwinia amylovora. Mol Plant-Microbe Interact 25:421–430 11. Degrave A, Moreau M, Launay A et al (2013) The bacterial effector DspA/E is toxic in Arabidopsis thaliana and is required for multiplication and survival of the fire blight pathogen. Mol Plant Pathol 14:506–517 12. Dellagi A, Brisset M-N, Paulin J-P et al (1998) Dual role of desferrioxamine in Erwinia

amylovora pathogenicity. Mol Plant-Microbe Interact 11:734–742 13. Parker KG, Leupschen NS, Jones AL (1974) Inoculation trials with Erwinia amylovora to apple rootstocks. Plant Dis Rep 58:243–247 14. Momol MT, Norelli JL, Piccioni DE et al (1998) Internal movement of Erwinia amylovora through symptomless apple scion tissues into the rootstock. Plant Dis 82:646–650 15. Jensen PJ, Rytter J, Detwiler EA et al (2003) Rootstock effects on gene expression patterns in apple tree scions. Plant Mol Biol 53:493–511 16. Sedlak J, Paprstein F, Korba J et al (2013) Development of a system for testing apple resistance to Erwinia amylovora using in vitro culture techniques. Plant Prot Sci 51:1–5 17. Lee SA, Ngugi HK, Halbrendt NO et al (2010) Virulence characteristics accounting for fire blight disease severity in apple trees and seedlings. Phytopathology 100:539–550 18. Zhao Y, Blumer SE, Sundin GW (2005) Identification of Erwinia amylovora genes induced during infection of immature pear tissue. J Bacteriol 187:8088–8103 19. Bellemann P, Geider K (1992) Localization of transposon insertions in pathogenicity mutants of Erwinia amylovora and their biochemical characterization. J Gen Microbiol 138:931–940 20. Bogdanove AJ, Kim JF, Wei Z et al (1998) Homology and functional similarity of an hrplinked pathogenicity locus, dspEF, of Erwinia amylovora and the avirulence locus aveE of Pseudomonas syringae pathovar tomato. Proc Natl Acad Sci U S A 95:1325–1330 21. Esau K (1977) Anatomy of seed plants. Wiley, Santa Barbara 22. Dennis FG Jr (2000) The history of fruit thinning. Plant Growth Regul 31:1–16 23. MacDaniels LH (1940) Memoir Number 230—the morphology of the apple and other pome fruits. Cornell University, Ithaca 24. Klee SM, Mostafa I, Chen S et al (2018) An Erwinia amylovora yjeK mutant exhibits reduced virulence, increased chemical sensitivity and numerous environmentally dependent proteomic alterations. Mol Plant Pathol 19:1667–1678

Chapter 18 Generating Transgenic Arabidopsis Plants for Functional Analysis of Pathogen Effectors and Corresponding R Proteins Sharon Pike, Walter Gassmann, and Jianbin Su Abstract Inducible expression of a pathogen effector has been proven to be a powerful strategy for dissecting its virulence and avirulence functions. However, leaky expression of some effector proteins can cause drastic physiological changes, such as growth retardation, accelerated senescence, and sterility. Unfortunately, leaky expression from current inducible vectors is unavoidable. To overcome these problems, a highly efficient Arabidopsis transformation protocol is described here, which allows the generation of hundreds to over a thousand T1 plants for selecting appropriate lines. In addition, since transgenic silencing is frequently observed, a principle for screening stable transgenic plants is also introduced. Key words Arabidopsis transformation, Pathogen effectors, AvrRps4, Inducible expression, Leaky expression, RPS4, Gene silencing

1

Introduction The development of dexamethasone (DEX) and estradiol (Est) inducible gene expression systems [1, 2] greatly facilitates functional characterization of pathogen effectors at the whole plant level. For example, the requirement of the Arabidopsis resistance (R) protein RPS2 for the Pseudomonas syringae AvrRpt2-mediated hypersensitive response was confirmed by DEX inducible expression of avrRpt2 in the Arabidopsis Columbia (Col-0) background which has RPS2 [3]. Recently, DEX-inducible avrRpt2 lines in the rps2 background demonstrated a virulence function of AvrRpt2 in which AvrRpt2 cleaves Aux/IAA, the key negative regulators of auxin signaling, to promote pathogenicity [4]. Similarly, suppression of the apoplastic reactive oxygen species (ROS) burst and callose deposition by AvrRps4 was demonstrated by using DEX inducible avrRps4 lines in the rps4-2 background [5]. AvrRps4 is

Walter Gassmann (ed.), Plant Innate Immunity: Methods and Protocols, Methods in Molecular Biology, vol. 1991, https://doi.org/10.1007/978-1-4939-9458-8_18, © Springer Science+Business Media, LLC, part of Springer Nature 2019

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cleaved into a 133 amino acid N-terminus (AvrRps4N) and an 88 amino acid C-terminus (AvrRps4C) in planta [5]. In order to study the avirulence function of AvrRps4, we attempted to generate DEX-inducible myc-AvrRps4FL (full length AvrRps4), myc-AvrRps4N and myc-AvrRps4C lines in the Col-0 background. To study the dynamic interaction between AvrRps4 and one of its corresponding R proteins, RPS4 [6], we also generated an HA-tagged genomic clone for RPS4, designated as genomic RPS4-HA. We successfully obtained many T1 RPS4-HA lines with varying levels of protein expression. Unfortunately, all the T1 lines (about 200 each) for AvrRps4FL and AvrRps4C showed growth inhibition, early senescence or a sterile phenotype caused by leaky expression (data not shown). As leaky expression is unavoidable for current inducible vectors, the only way to generate normal plants with inducible expression of AvrRps4 is to increase the total number of T1 lines. Recently, AGROBacterium-mediated Enhanced Seedling Transformation (AGROBEST) was developed for Agrobacterium-mediated transient gene expression in young Arabidopsis seedlings [7]. Two factors considered to be key for the success of AGROBEST were buffering with 2-(N-morpholino)ethanesulfonic acid (MES) pH 5.5 and adding Agrobacterium (AB) vir gene induction medium salts to Murashige and Skoog (MS) medium [7], possibly resulting in optimized vir gene induction by MES-AB salt [8]. Here, we describe a detailed protocol for high-efficiency Arabidopsis transformation, by which hundreds to a thousand independent T1 lines can be easily obtained by initially dipping only 8–12 plants. We optimized the widely used sucrose/Silwet L-77 protocol [9, 10], by incorporating ¼ strength MS salts, 2 mM MES (pH 5.7) and 0.5 μg/mL 6-Benzylaminopurine (BAP) into the dipping buffer. Moreover, we use young plants with 2–3 short branches before formation of visible siliques to reach high transformation efficiency. This is a method that can be applied to other genes, especially to genes that are frequently silenced.

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Materials

2.1 Plant Growth Materials

1. Seeds of Arabidopsis: Columbia accession (Col-0) obtained from ABRC. 2. Seed sterilization solutions: 70% Ethanol (v/v); 20% sodium hypochlorite (bleach), 0.05% Tween® 20 (v/v). 3. Autoclaved ddH2O rinse. 4. 50 mL test tubes. 5. Refrigerator at 4 oC.

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Fig. 1 Schematic of dipping procedures and representative images of DEX-inducible AvrRps4C T1 lines. (a) Schematic of dipping procedures for agrobacterium-mediated Arabidopsis transformation. The height of plants is around 10–15 cm, with two to three short branches and several flowers at the top. After dipping, the pot is laid on its side with the lid tightly closed overnight. Subsequently, the lid is kept loosely closed for at least 12 h before it is removed and the pots are turned upright. (b) Representative images for DEX-inducible AvrRps4C T1 lines; most of them are stunted due to leaky AvrRps4C expression

6. ½ Murashige and Skoog (MS) Petri plates for initial plant growth: 2.165 g/L commercially available MS basal salt mixture, 0.4–0.5% agar (v/v) (see Note 1). 7. Plant growth shelves: constant light at 22 oC. 8. Potting soil in pots set in flats. 9. Flats with transparent domes. 10. Fine forceps. 11. Greenhouse or growth chamber: 16 h/8 h light/dark cycles. 12. High nitrogen/phosphate (3:1 N/P) ratio fertilizer. 13. Container for transformation solution with sufficient width and depth to permit submerging of the stems during dipping (Subheading 3.3, step 1, Fig. 1a). 14. Fine mesh strainer for harvesting seed. 2.2 Agrobacteria Strains and Vectors

1. Glycerol stocks of agrobacterium GV3101 strain harboring a construct of interest at 80  C. We are presenting results using pTA7002-AvrRps4FL, pTA7002-AvrRps4N, pTA7002C AvrRps4 and pBIB-RPS4-HA as examples. 2. Luria-Bertani (LB) (Miller) 1% agar Petri plates with 50 μg/ mL gentamicin and 50 μg/mL kanamycin. Before use, recover these strains on LB agar plates (see Note 2).

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2.3 Stock Solutions and Buffers

1. Commercially available MS salt mixture. 2. 1000 stock solutions: 50 mg/mL gentamicin in ddH2O, 100 mg/mL timentin in ddH2O, and 50 mg/mL kanamycin in ddH2O, all kept at 20  C. 20 mg/mL hygromycin in ddH2O, kept at 4  C in dark. Sterilize all stock solutions by 0.2 μm syringe filtration. 3. 200,000 BAP stock solution: 1 mg/mL stock in 1 N HCl, stored at 4  C (see Note 3). 4. 100 MES stock: dissolve 42.6 g MES in 80 mL ddH2O and adjust the pH to 5.7 with KOH. Bring up to 100 mL final volume, filter sterilize, and keep at room temperature. 5. Transformation solution: 10 mL of 100 MES stock, 50 g of sucrose, 1.1 g of MS salt mixture, 5 μL BAP, 200 μL Silwet L-77, freshly prepared (see Note 4).

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Methods

3.1 Preparation of HEALTHY ARABIDOPSIS PLANTS

1. Sterilize seeds by soaking them in 70% ethanol in a 15 mL tube for 3 min. Carefully pipette off the ethanol solution. Add the bleach/Tween® 20 solution and soak for 10 min. Rinse 5 with sterile deionized water. After the last rinse, add about 2 mL water and keep seeds in the dark at 4  C for 3–5 days (see Note 5). 2. Evenly distribute the seeds on the ½ MS plates and keep them under constant light for 6–8 days (see Note 6). 3. Transfer the seedlings carefully with a fine forceps to a pot, burying the root in well-watered soil (see Note 7). 4. After transferring seedlings from the ½ MS plate to soil, cover the flat with a transparent lid for at least 3 days, then loosen the lid a small amount and allow the plants to adapt to the lower humidity for another 2 days. Then remove the lid and water plants regularly from the bottom. 5. Fertilize plants with the high nitrogen/phosphate (N/P) ratio fertilizer every two weeks (see Note 8). 6. When plants have grown to the optimal stage for transformation (see Note 9) (Fig. 1a), on the day before dipping, water plants well to compact the soil.

3.2 Preparation of Agrobacterium Cultures

1. Streak the agrobacterium strains on LB agar plates with appropriate antibiotics four days before the planned dipping. Grow them for 2 days at 28  C. Pick 20–40 single colonies with a sterile toothpick or loop and transfer to 5 mL of LB liquid medium with appropriate antibiotics. Mix well and shake at 200 rpm, 28  C overnight.

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2. Transfer 2 mL of culture to 200 mL of LB liquid medium with appropriate antibiotics, shaking at 200 rpm, 28  C overnight. If C58C1 strains are used, transfer 4 mL of culture to 400 mL of LB medium. 3. Harvest agrobacterium cells by centrifugation at 3000  g for 15 min at room temperature. 4. Discard the supernatant into a container with bleach to kill the residual cells in the supernatant. 5. Rinse the pellet with 20 mL of transformation solution to remove residual LB medium; then discard the supernatant into a container with bleach. 6. Resuspend the pellet in 20 mL of transformation solution either by vigorously shaking by hand or by vortexing. Then add about 300 mL of transformation solution to adjust the OD600 to 0.8. OD600. Values between 0.8 and 1.2 were found to work well experimentally. 3.3 Dipping Transformation

1. Before dipping, set out a clean tray and a transparent lid. Pour the transformation solution into a container with sufficient depth and width to submerge plant stems when the pot is inverted (Fig. 1a). 2. Invert plants and submerge in the transformation solution for 5 min (Fig. 1a), then quickly transfer plants to the tray, laying the pot on its side (Fig. 1a), and tightly cover with the transparent lid. 3. The day after dipping, loosen the lid for at least 12 h (Fig. 1a) and then remove the lid and turn the pot upright. 4. Continue to water the plants for another 3 weeks; then remove all the plant stems with siliques and transfer them to an envelope. Dry for 2 days at 37 C. 5. Collect seeds with a fine mesh strainer. Usually, 6–8 mL seeds can be harvested from 8–12 plants.

3.4 Selection of T1 Transgenic Plants

1. Transfer 3 mL of seed to a 50 mL tube, surface-sterilize and cold-treat (see Subheading 3.1, step 1). After washing, the volume of the swollen seeds should have increased to 15 mL. 2. Transfer 1 mL of swollen seeds to a ½ MS plate (9 cm in diameter) with 100 μg/mL timentin and 20 μg/mL hygromycin (or the appropriate concentration of antibiotic compatible with the selectable marker of your construct) and spread the seeds evenly on the plate (see Note 10). 3. Transfer approximately 72 hygromycin resistant seedlings to soil for each construct and evaluate the phenotype (Fig. 1b) (see Note 11).

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Fig. 2 Western-blot screening and selection of T1 and T3 genomic RPS4-HA lines by Western blot. (a) Screening of T1 genomic RPS4-HA lines. 10 μg total protein was loaded for each well. Anti-HA antibody was used for the detection of RPS4-HA fusion protein. Red triangles indicate T2 plants of lines showing 3:1 segregation on hygromycin plates that were selected for further analysis. Protein expression of RPS4-HA was hardly detected in some lines and they were removed. (b) Confirmation of T3 genomic RPS4-HA lines by Western blot. 10 μg total protein was loaded for each well. 10-day-old homozygous lines were harvested for confirmation

4. Confirm protein expression by Western blot if an antibody is available or the protein is tagged. Remove any lines lacking protein expression (Fig. 2a). 5. Harvest seeds for each independent line. 3.5 Characterization of T2 and T3 Plants

1. Sterilize seeds harvested from T1 plants and sow them on ½ MS plates with 20 μg/mL hygromycin. 2. Count the segregation ratio for each line. If the ratio of resistant to sensitive lines is around 3:1, the line is considered to be a single T-DNA locus line. Randomly transfer 12 seedlings for each line to soil for these putative single T-DNA insertion lines (see Note 12). 3. Harvest T2 seeds from each individual plant. 4. Sterilize seeds harvested from T2 plants and sow them on ½ MS plates with 20 μg/mL hygromycin. 5. Count the segregation ratio for each line. If all the seedlings show resistance to hygromycin, the line is considered to be a homozygous line. 6. Confirm protein expression again in these homozygous lines. If the protein expression is very low compared to its corresponding T1 line, the line is likely silenced and is not suitable for subsequent analysis. For example, RPS4-HA in line #23 was very high in T1, but is hardly detected in T3; however, #8-7, #18-10, and #19-7 are good lines for further experiments (Fig. 2a, b). Even with seed from the same T2 plant, some T3 lines also show gene silencing, such as # 8-9 and #19-9 (Fig. 2b). Thus, one should be very careful when

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working with transgenic plants. We highly recommend checking protein expression at every generation and harvesting seeds separately from each plant.

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Notes 1. For growing Arabidopsis on ½ MS agar plates, the agar concentration used here is only half of that normally used, allowing easier transfer of seedlings and avoiding severe damage to the roots. 2. Agrobacterium strain C58C1 was also tested in this protocol. It works as well as GV3101, but grows more slowly. When C58C1 is used, we usually inoculate a two-fold higher volume compared to GV3101. 3. BAP stock solution (1 mg/mL) is stable in HCl for several years at 4  C. BAP is one of the first generation synthetic cytokinins, and promotes blossoming. The total number of flowers is greatly increased when plants are treated with BAP. 4. Because BAP is dissolved in 1 N HCl, the pH of a solution is only slightly reduced after its addition. For example, adding 5 μL BAP to 1 L transformation solution will reduce the pH by about 0.005–0.01. 5. Seeds are left in water at 4  C more than 5 days but less than 3 weeks to stratify them for more even germination. 6. 1000–2000 seeds can be plated on a 150  15 mm Petri plate. 7. To prevent soil loss when the pot is inverted during dipping, there must be sufficient root growth and rosette cover. We transfer 4 seedlings to each 9  9 cm pot that is 6 or 9 cm deep. 8. A high N/P ratio fertilizer such as Miracle-Gro All Purpose Plant Food makes plants grow stronger and delays flowering. 9. Plants used for dipping in this protocol are younger than those that are widely used. We tested transformation efficiency with plants at different ages, and found that younger plants (Fig. 1a) work much better than older ones. Using younger plants also avoids the need for removing siliques before dipping. We don’t yet understand the molecular basis behind this observation. Because agrobacterium is alive in the plant, we suspect that all the newly emerged buds or branches can be easily infected, which would increase the total number of transformation events. 10. In most cases, 0.5 mL dry seeds were sterilized. We usually sow 2–3 plates for each construct. Normally, there are 30–100 resistant seedlings in one plate.

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11. Usually, 72 seedlings are transferred for each construct. For DEX-AvrRps4C, more than 600 lines were transferred to soil, only 2% of them looked normal (Fig. 1b). The ratio of normallooking plants for DEX-AvrRps4FL was relatively higher, about 8%. 12. If the resistant vs sensitive ratio is around 3:1, it does not always mean the line contains only one T-DNA insertion. If two or more T-DNAs inserted themselves into the same chromosome in close proximity (single locus), the resistant vs sensitive ratio is also around 3:1.

Acknowledgments Work in the Gassmann lab is supported by NSF grants IOS-112114 and IOS-1456181. References 1. Aoyama T, Chua NH (1997) A glucocorticoidmediated transcriptional induction system in transgenic plants. Plant J 11:605–612 2. Zuo J, Niu QW, Chua NH (2000) Technical advance: an estrogen receptor-based transactivator XVE mediates highly inducible gene expression in transgenic plants. Plant J 24:265–273 3. McNellis TW, Mudgett MB, Li K, Aoyama T, Horvath D, Chua NH et al (1998) Glucocorticoid-inducible expression of a bacterial avirulence gene in transgenic Arabidopsis induces hypersensitive cell death. Plant J 14:247–257 4. Cui F, Wu S, Sun W, Coaker G, Kunkel B, He P et al (2013) The Pseudomonas syringae type III effector AvrRpt2 promotes pathogen virulence via stimulating Arabidopsis auxin/indole acetic acid protein turnover. Plant Physiol 162:1018–1029 5. Sohn KH, Zhang Y, Jones JD (2009) The Pseudomonas syringae effector protein, AvrRPS4, requires in planta processing and

the KRVY domain to function. Plant J 57:1079–1091 6. Gassmann W, Hinsch ME, Staskawicz BJ (1999) The Arabidopsis RPS4 bacterialresistance gene is a member of the TIR-NBSLRR family of disease-resistance genes. Plant J 20:265–277 7. Wu HY, Liu KH, Wang YC, Wu JF, Chiu WL, Chen CY et al (2014) AGROBEST: an efficient Agrobacterium-mediated transient expression method for versatile gene function analyses in Arabidopsis seedlings. Plant Methods 10:19 8. Gelvin SB (2006) Agrobacterium virulence gene induction. Methods Mol Biol 343:77–84 9. Zhang X, Henriques R, Lin SS, Niu QW, Chua NH (2006) Agrobacterium-mediated transformation of Arabidopsis thaliana using the floral dip method. Nat Protoc 1:641–646 10. Clough SJ, Bent AF (1998) Floral dip: a simplified method for Agrobacterium-mediated transformation of Arabidopsis thaliana. Plant J 16:735–743

Chapter 19 Identification of Novel Pararetroviral Promoters for Designing Efficient Plant Gene Expression Systems Ankita Shrestha, Ahamed Khan, and Nrisingha Dey Abstract Plant-infecting viruses, particularly the Pararetroviruses, have been used for many years as versatile genetic resources to design efficient plant expression vectors. The Pararetroviruses (members of the Caulimoviridae) typically contain two transcriptional promoters (the sub-genomic transcript promoter and the fulllength transcript promoter) and 6–7 overlapping open reading frames (ORFs) with a genome size of 7–9 kB. Their promoter elements have been extensively exploited during the last two decades to construct effective gene expression systems. At the same time, the caulimoviral promoters have also been genetically manipulated with different molecular approaches to develop synthetic “chimeras” exhibiting precise functionality. Native and “tailor-made” synthetic promoters of Pararetroviruses are particularly attractive for formulating unique gene expression cassettes that perform extremely well in gene-stacking and genepyramiding in plant cells. In this chapter, we will mainly discuss important protocols associated with identifying novel/unique pararetroviral promoters that have optimal lengths with appropriate activities for developing efficient plant gene expression systems. Key words Caulimoviral promoter, TSS, Electroporation, Infiltration, Transient expression

1

Introduction The Caulimoviridae family of viruses typically consists of members of plant pararetroviruses which replicate via an RNA intermediate evolved from retroelements; however, unlike animal retroviruses, they usually do not integrate into the host genome [1, 2]. They are classified primarily on the characteristics of the viral genome (RNA/DNA) and have been extensively studied for their genomic expression strategies which include the transcription of 35S and 19S RNA [3]. The leader sequence of 35S RNA contains several open reading frames (ORFs) and each genus is distinguished from others by the arrangement of ORFs that synthesize functional viral proteins. One of the most widely studied viral promoters, the

Ankita Shrestha and Ahamed Khan have contributed equally to this work. Walter Gassmann (ed.), Plant Innate Immunity: Methods and Protocols, Methods in Molecular Biology, vol. 1991, https://doi.org/10.1007/978-1-4939-9458-8_19, © Springer Science+Business Media, LLC, part of Springer Nature 2019

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CaMV35S promoter from Cauliflower mosaic virus possesses excellent transcriptional activity [4]. Likewise, several other members of Caulimoviridae, for example, Figwort mosaic virus (FMV), Mirabilis mosaic virus (MMV), Dahlia mosaic virus (DaMV), Peanut chlorotic streak virus (PClSV), Cassava vein mosaic virus (CsVMV), and Horseradish latent virus (HRLV) have also been exploited to study in planta gene expression patterns using transgenic platforms [5–12]. Due to the constitutive nature of the caulimoviral promoters, they have been employed for strong gene expression in plant biotechnology and to study gene regulatory mechanisms [13–18]. Usually, these plant-infecting viruses are potent infectious agents that cause disease symptoms in plants. However, they do not primarily cause huge economic crop losses. On the contrary, we can explore their infection mechanisms in a beneficial way to develop technologies for controlled gene expression. The mechanisms implemented by these viruses to successfully infect and remain within specific host cells across a diverse range of environmental conditions are quite remarkable. A unique feature of these Caulimoviruses is that they are capable of efficiently utilizing the host machinery for their survival, even in extremely competitive cellular environments [6]. To accomplish these tasks, viruses contain unique arrangements of cis-elements in their regulatory systems which allow the binding of multiple stress-inducible transcription factors (TFs) that favor quick transcription of viral genes (Fig. 1). This is attributed to cis-elements in the promoter that recruit specific TFs. The pattern of cis-elements is unique to each caulimoviral promoter and is defined by its immediate surroundings [6]. These distinctive properties of viral promoters can be utilized as sources of efficient gene expression in planta (Fig. 1). The idea of using caulimoviral promoters in gene expression platforms has been extended to design chimeric promoters containing consensus DNA sequences from two or more native promoters [19–21]. Leader-deletion analysis is a classical approach used to identify and evaluate the strength of any native promoter by deleting specific DNA sequences in a sequential manner. Introducing these promoters into plants to evaluate their transcriptional activities has been achieved by developing new platforms to transform plant cells. Procedures such as electroporation and agro-infiltration aid in introducing foreign DNA (promoter sequences linked to reporter genes) into plant cells and permit rapid investigations into the regulation of gene expression in plants. Here, we describe methods that employ plant-infecting caulimoviral promoters as tools to develop new gene expression systems in a plant-based platform. The aim of this chapter is to describe the basic protocols that are employed for rapid identification and evaluation of a useful promoter fragment having an optimized length particularly in transient approaches. An accompanying chapter describes approaches to characterize regulatory elements in these

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Fig. 1 A proposed model for plant–virus interactions depicting the use of viral regulatory systems for gene expression in plants. (a) Plants perceive external stress stimuli (both biotic and abiotic) via embedded signal sensors on their cellular surfaces and trigger defense responses by upregulating stress-inducible TFs, which in turn increases the abundance of these TFs. (b) The viral regulatory system consists of cognate cis-elements that scavenge the endogenous plant TFs from the cellular pool to enhance the production of necessary proteins. (c) Foreign genes can be expressed under the control of such viral promoters and mimic this viral survival strategy in the host cell

promoters more fully (see the next Chapter). We intend that these chapters will be user-friendly to researchers working in the field of plant molecular biology, particularly in developing new gene expression systems.

2

Materials (Use ultrapure water for all solutions).

2.1 Identification and Isolation of Viral DNA

1. Plant tissue sample infected with the respective virus. 2. Homogenizer, cheese cloth, beakers, and funnel. 3. Grinding buffer (100 mL): 0.2 M Tris pH 7.0, 0.02 M ethylenediaminetetraacetic acid (EDTA), 1.5 M urea. 4. Triton X-100: to be added to 10 mL grinding buffer at a 2% v/v final concentration. 5. Centrifuge tubes (30 mL Oak Ridge High-Speed PPCO Centrifuge Tubes) 6. Cooling ultracentrifuge.

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7. Sucrose (15% w/v): Add 15 g sucrose to 100 mL of sterile water. 8. Chloroform. 9. Resuspension buffer: 0.1 M Tris pH 7.4, 2.5 mM MgCl2. 10. EDTA solution: 0.5 M. 11. DNaseI, Restriction endonucleases, cloning vector (any pUC derivative). 12. Low melting agarose 13. Agarose gel electrophoresis apparatus. 14. Protease K. 15. Sodium dodecyl sulfate (SDS) 1% w/v: Add 1 g of SDS to 100 mL of sterile water. 16. Phenol. 17. Ethanol. 2.2 Cloning of Promoter Sequences

1. Promoter constructs cloned into the vector pUC119, the protoplast expressing vector pUCPMAGUS [5], and the plant expression vector pKYLX71GUS [21]. 2. The reference control pUCPMAGUS-CaMV35S. 3. The CaMV35S-GFP reporter gene to check the efficiency of protoplast transformation and agro-infiltration assays. 4. Deletion constructs for each promoter using specific primers and PCR amplification to tailor EcoRI and HindIII restriction sites at the 50 - and 30 -ends, respectively.

2.3 PCR Amplification, Cloning, and Transformation

1. Taq mix (50 μL volume per reaction): 10 mM Taq DNA polymerase, 10 mM Taq Polymerase buffer (10), 10 mM MgCl2, and 10 mM dNTP mix stored at 20  C. 2. Restriction enzymes: EcoRI, HindIII, XhoI, SacI stored at 20  C. 3. Ligase: T4 DNA ligase and ligase buffer stored at 20  C. 4. Incubator set at 37  C. 5. Plasmid isolation Mini-prep kit. 6. Gel Extraction kit. 7. Thermal cycler. 8. Ultrapure low melting agarose. 9. Petri dishes (90  15 mm) 10. DNA sequencer: Beckman Coulter CEQ-8000 sequencer or equivalent. 11. DNA separation gel and separation buffer stored at 4  C. 12. Tris-acetate-EDTA (TAE) buffer (50X): 242 g/L Tris base, 100 mL/L of 0.5 M EDTA pH 8.0, 57.1 mL/L glacial acetic

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acid, autoclaved and stored at room temperature (RT). Dilute to 1 using milli-Q water for use in horizontal electrophoresis. 13. Luria-Bertani (LB Miller) broth medium: Add 20 g/L LB broth to milli-Q water, dissolve and autoclave. 14. LB agar petri plates: Add 30 g/L agar to LB broth prior to autoclaving. Add 100 μg/mL ampicillin when partially cooled, before pouring the plates. 2.4 Transient Expression in Protoplasts

1. Tobacco cell suspension cultures (Nicotiana tabacum L. cv. Xanthi). 2. Tobacco cell suspension culture medium: 4.33 g/L Murashige and Skoog basal salts, 204 mg/L of potassium phosphate (KH2PO4), 0.5 mg/L pyridoxine-HCl, 0.5 mg/L nicotinic acid, 0.5 mg/L thiamine-HCl, 0.2 mg/L 2,4-dichlorophenoxyacetic acid (2,4-D), 0.1 mg/L kinetin, pH 5.7 with 1 M KOH. Autoclave and store at RT. 3. Protoplast culture medium (AOKI): 91.1 g/L mannitol, 1.95 g/L 2-morpholinoethane sulfonic acid (MES), 1.47 g/ L CaCl2·2H2O, 27 mg/L K2HPO4, 101 mg/L KNO3, 120 mg/L MgSO4, 2 mg/L potassium iodide (KI), 30 g/L sucrose, pH 5.6 with 1 M KOH. Autoclave and store at RT. 4. MMC solution: 91.1 g/L mannitol, 1.95 g/L MES, 1.47 g/L calcium chloride hydrate (CaCl2·2H2O), pH 5.6 with 1 M KOH. Autoclave and store at 4  C. 5. Macerating enzyme solution: 0.75% Cellulase Onozuka RS (Yakult Honsa, Japan) and 0.075% pectinase (Sigma). Dissolve both enzymes in autoclaved MMC solution and filter-sterilize using a 0.22 μm pore size. 6. Sucrose solution (25%): Sucrose 250 g/L, MES 1.95 g/L, and CaCl2·H2O 1.47 g/L. Autoclave and store at 4  C. 7. Sterile Pasteur pipettes. 8. 250 mL conical (Erlenmeyer) flasks: autoclave with stoppers, foil, or lids. 9. 1.5, 15, and 50 mL sterile centrifuge tubes. 10. Incubator set at 26  C. 11. Electroporation buffer: 91.1 g/L mannitol, 5.21 g/L potassium chloride (KCl), 975 mg/L of MES in 100 mL ultrapure H2O, pH 5.6 with 1 M KOH. Autoclave and store at 4  C. 12. Mannitol agarose solution coated Petri dishes: Dissolve 90 g/L of mannitol and 10 g/L of low melting agarose in ultrapure water and autoclave. Coat sterile 25 mm Petri dishes with 1 mL of this solution. 13. Electroporator: Gene Pulser II apparatus (model 165-2017; Bio-Rad, USA) or equivalent.

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2.5 Transient Expression in Tobacco

1. Plant materials: Wild-type tobacco Nicotiana tabacum cv. Samsun NN 2. Greenhouse conditions: Photoperiod 16/8 h (light/dark), light intensity 220 μmol/m2/s, temperature 28  3  C, and humidity 70–75%. 3. Agrobacterium strain: Agrobacterium tumefaciens strain LBA4404. 4. Antibiotic rifampicin stock: Dissolve 25 mg of rifampicin powder in 1 mL of absolute methanol. Filter-sterilize using a 0.22 μm filter and store at 20  C. Use at a final concentration of 100 μg/mL. 5. Infiltration medium: Add 500 mM MES, 20 mM Na3PO4 and 5 mg/mL D-glucose and 150 μM acetosyringone (3,5-dimethoxy-40 -hydroxy-acetophenone). Bring up the volume to 50 mL with ultrapure water.

2.6 Transcription Start Site (TSS) Determination by Primer Extension Assay

1. Plant RNA extraction kit, for example, Spectrum™ Plant Total RNA Kit (Sigma). 2. cDNA synthesis kit, for example, First Strand cDNA Synthesis Kit (Thermo Fischer Scientific). 3. γ-32P ATP, high-specific-activity. 4. T4 polynucleotide kinase: 10 Units/μL. 5. Absolute ethanol: 99% v/v. 6. Formamide loading dye: 95% deionized formamide, 0.025% (w/v) bromophenol blue, 0.025% (w/v) xylene cyanol FF, 5 mM EDTA 0.5 M, pH 8.0. 7. 1 M Tris–HCl pH 7.5: Dissolve 157.60 g Tris–HCl in 800 mL dH2O. Adjust pH to 7.5 with concentrated HCl. Bring the final volume to 1 L with dH2O. 8. Primer annealing buffer: 50 mM Tris–HCl pH 7.5, 1.25 M KCl, 5 mM EDTA. 9. 3 M sodium acetate pH 5.2: 100 mL. 10. T4 polynucleotide kinase labeling buffer (10): 0.5 M Tris–HCl pH 7.6, 1 mM EDTA pH 8.0, 0.1 M MgCl2, 50 mM dithiothreitol, 1 mM spermidine HCl. 11. Tris EDTA buffer: 10 mM Tris–HCl pH 7.5, 1 mM EDTA 0.5 M, pH 8.0. 12. 40% (w/v) acrylamide and bisacrylamide solution (19:1): 20 g/L N, N0 methyl-bisacrylamide, 380 g/L acrylamide. Filter sterilize with a 0.45 μm filter and store in dark bottle at 4  C (see Note 1). 13. Mini Quick Spin Oligo Columns (Roche).

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14. Sanger Sequencing Kit, for example, Thermo Sequenase Dye Primer Manual Cycle Sequencing Kit (USB, Inc). 15. Phosphor-imager. 16. Speed vacuum evaporator 17. Scintillation counter. 18. Gel Dryer. 19. X-ray film. 20. Autoradiography cassette (Hypercassette™, Amersham Bioscience, or equivalent).

3

Methods

3.1 Identification and Isolation of Viral DNA

The isolation of Caulimoviral genome (dsDNA) from infected plant material is generally performed using the standard “gradient fractionation” technique [22–24]. However, slight modifications in this technique are required depending on the virus isolate, particularly due to differences in the culture titer volume of the virus found in the infected plant material. Here, we describe the identification and isolation of viral DNA from the CaMV, the most common representative of the Pararetrovirus group of viruses which typically consists of circular double-stranded DNA of ~8000 base pairs.

3.1.1 Isolation of Viral DNA

1. Harvest systemically infected leaf tissue (l-5 g) from the plant. 2. Add 10 mL of prechilled grinding buffer and Triton X-100 (final concentration of 2% v/v) to the tissue (see Note 2). 3. Homogenize the tissue for 30–45 s at maximum speed using a suitable homogenizer. 4. Filter the crude homogenate through cheesecloth and treat with chloroform (10% v/v) for 15 min at room temperature. 5. Centrifuge the extract at 16,000 x g for 10 min and collect the supernatant. 6. Layer the supernatant onto a sucrose gradient prepared by adding 5 mL of 15% sucrose to a centrifuge tube. Centrifuge the tube at high-speed ~60,000 x g for 2 h and discard the supernatant. 7. Resuspend the pellet containing the virus in 150 μL resuspension buffer per gram of plant tissue. Add DNaseI to a final concentration of 10 μg/mL and incubate at 37  C for 10 min (see Note 3). 8. To stop the DNaseI reaction add EDTA to a final concentration of 10 mM.

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9. To this crude virus preparation add Protease K to a final concentration of 0.5 mg/mL and 1% SDS. Incubate at 65  C for 10 min (see Note 4). 10. Extract the viral DNA using equal volume of phenol followed by ethanol precipitation. Dissolve the precipitated DNA in 100 μL of 10 mM Tris pH 8.0 and 1 mM EDTA. 3.1.2 Viral DNA Characterization

1. For characterizing the isolated viral DNA, treat with any one of the restriction endonucleases such as EcoRI, BamHI, or XbaI and run the digested DNA on an agarose gel. 2. Clone the obtained fragments into a suitable sequencing vector. Analyze the DNA fragments and identify the endonuclease sites present in the viral genome by sequencing the clones. 3. To obtain the complete sequence of the viral genome, follow the standard protocol of restriction endonuclease mapping [23].

3.2 PCR, Cloning and Transformation

Promoter deletion analysis is a useful tool to gain insight into the mechanisms of gene regulation and identify important regulatory regions involved in transcription [25, 26]. This technique involves sequential deletion of the native promoter to generate a series of 50 and 30 -end deleted fragments. These promoter fragments are cloned upstream of a reporter gene {β-Glucuronidase (GUS), Green Fluorescent Protein (GFP), or Luciferase (LUC)}; the expression of the reporter gene is directly correlated with the promoter activity and is used to assess the strength of each fragment in transient or transgenic systems. 1. Promoter leader-deletion analysis: For generation of deletion constructs, generate a series of fragments by uniformly deleting a few nucleotides (preferably 50 bp) from the 50 - and 30 -end of the native promoter. 2. Design primer sets for each of the fragments that contain EcoRI and HindIII sites in the forward and reverse primers, respectively. 3. Amplify each promoter fragment using appropriate primers to introduce the EcoRI site at the 50 - end and HindIII site at the 30 - end of the amplified products. 4. Carry out the PCR amplification in a total reaction volume of 50 μL using 50–100 ng of template plasmid DNA (native promoter clone), 5 pM of each primer, 200 μM dNTPs, and 2 U of a high-fidelity DNA polymerase. The PCR conditions are as follows: initial denaturation (94  C for 3 min) followed by 33 cycles of: denaturation (94  C for 30 s), annealing (55–58  C depending on the Tm of primer sets for 30 s),

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extension (72  C for 1 min) and a final extension of 72  C for 10 min. 5. Electrophorese the PCR amplicons on a 1% agarose gel and elute using a gel extraction protocol (for example, Qiagen). Digest the eluted product with EcoRI and HindIII restriction enzymes for 1–1.5 h and ligate in the corresponding sites of pUC119 vector (see Note 5). Perform all ligation reactions using the T4 DNA Ligase as per manufacturer’s protocol. 6. Transform the ligated product into E. coli TB1 chemically competent cells as follows: Add 10–12 μL of the ligation product to 100 μL of competent cells, incubate on ice for 30 min, heat-shock for 90 s at 42  C, and incubate again on ice for another 3–4 min. Add 1 mL of LB broth medium and incubate at 37  C with constant shaking at 200 rpm. Centrifuge the cells at 4000  g for 5 min. Plate the cells on LB-agar medium containing 100 μg/mL ampicillin and incubate overnight at 37  C. 7. Subject the clones to DNA sequencing and confirm the sequence integrity of each 50 - and 30 -end-deleted promoter fragment. 8. Subsequently, clone the EcoRI and HindIII digested PCR products in the corresponding sites of the protoplast expression vector pUCPMAGUS. 9. Transform the ligated product in E. coli TB1 chemically competent cells as described in Subheading 3.2, step 6 (see Note 6). 10. Sub-clone all promoter fragments into the plant expression vector pKYLX71GUS as EcoRI and HindIII fragments. Ligate using the T4 DNA Ligase as per manufacturer’s protocol (see Note 7). 11. Transform the ligated product into E. coli TB1 chemically competent cells as described in Subheading 3.2, step 6 (see Note 6). 3.3 Isolation of Plasmid DNA Containing 50 - and 30 End-Deleted Promoter Fragments

1. Pick the transformed cells on the LB-agar selection media and inoculate them in 5 mL of LB-broth medium supplemented with appropriate antibiotic. Incubate the LB-broth containing tubes overnight at 37  C with constant shaking (200 rpm).

3.4 Transient Expression in Protoplasts

1. Maintain tobacco cell suspension cultures (Xanthi cv. Brad) by subculturing to fresh medium every 4 days. Harvest the cells for protoplasts from 3-day-old cultures by centrifugation at 110  g for 5 min.

2. Isolate the plasmids from the overnight cultures and digest using EcoRI and HindIII restriction enzymes to check the size of inserts.

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2. Decant the culture medium and add 30 mL enzyme solution. Place the mixture in a sterile 250 mL conical flask, cover, and incubate at 26  C in dark with constant slow shaking at 80 rpm. 3. Transfer the suspension to a 50 mL sterile centrifuge tube and centrifuge at 100  g for 5 min. Carefully recover the protoplasts by removing the enzyme solution. 4. To wash the protoplasts once, add 30 mL of MMC solution to the protoplasts and centrifuge at 100  g for 3 min. Decant the MMC solution and add 10 mL of fresh MMC solution to the centrifuge tube. 5. Layer the protoplasts carefully on a 25% sucrose cushion in a 15 mL sterile centrifuge tube using a Pasteur pipette (see Note 8). Centrifuge at 100  g for 5 min. Allow the protoplasts to settle down till a visible ring is observed at the interface of sucrose and MMC solution. Carefully recover the top fraction which contains the protoplasts and transfer to a fresh tube. To the above fraction, add MMC solution and bring the volume to 10 mL. Resuspend the pellet slowly. Repeat twice. 6. Resuspend the protoplasts in electroporation buffer. Make sure the suspended solution is neither too thick nor too thin (see Note 9). Place the suspension on ice. 7. Aliquot 750 μL of the above protoplast solution (containing ~2  105 protoplasts) in an electroporation cuvette (0.4 cm gap; 200 V and 950 μF). Add 5–10 μg of plasmid DNA (pUCPMA) that contains GUS reporter and mix with a pipette. 8. Electroporate and place the sample on ice. Take pUCPMAGUS-CaMV35S as the reference control for this assay. Transfer the protoplasts to a new microcentrifuge tube and centrifuge at 200  g for 3 min. Carefully remove the electroporation buffer and wash once with 600 μL of MMC solution. 9. Remove the MMC solution and add 1 mL of AOKI medium. Resuspend by mixing gently with a pipette and pour on to a 25 mm plate coated with agarose mannitol agar. Seal each plate with Parafilm and cover with aluminum foil. Incubate the plates at 28  C in dark. Harvest the protoplasts after 36 h to measure GUS activity. 3.5 Transient Expression in Tobacco

The plant expression vector (pKYLX71GUS) containing the respective promoters cloned in the EcoRI and HindIII sites and having the GUS gene cloned downstream of each of these promoter fragments is used to study the relative strength of each promoter deletion construct. The fragments are transformed by the freezethaw method into the Agrobacterium tumefaciens strain LBA4404 and suspensions of Agrobacterium harboring the plant expression cassette are prepared and infiltrated into young tobacco (Nicotiana tabacum) leaves (see Note 10). The details are as follows.

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1. From a frozen glycerol stock of Agrobacterium, grow a 10 mL culture (LB broth supplemented with 25 μg/mL rifampicin) overnight at 28  C with constant shaking at 180 rpm. The following morning, dilute the culture in 100 mL LB broth (supplemented with 25 μg/mL rifampicin) and grow until the OD600 reaches 0.6–0.8. 2. Precool the centrifuge to 4  C. When cells are ready to harvest, chill flasks on ice for 15–30 min. Centrifuge at 4500  g for 15 min at 4  C. Remove the supernatant and resuspend the pellet in each bottle in 20 mL of prechilled 20 mM CaCl2. Keep on ice for 20–30 min. Centrifuge at 4500  g for 10 min. Remove supernatant and gently resuspend the pellet in 2 mL of prechilled 20 mM CaCl2 containing 15% glycerol (v/v). Pipette 100 μL aliquots into microcentrifuge tubes, freeze in liquid nitrogen, and store at 80  C.

3.5.2 Freeze-Thaw Method for Transformation

1. Thaw the stored Agrobacterium competent cells on ice prior to transformation. Mix 200 ng of plasmid DNA with 100 μL competent cells (see Note 11). 2. Incubate the cells on ice for 10 min. Add liquid nitrogen and keep for 30 s. Thaw the frozen cells at 37  C for 5 min. 3. Add 1 mL of LB-broth and shake constantly at 180 rpm at 28  C for 4–5 h. 4. Centrifuge at 4500  g for 5 min and discard the supernatant. Resuspend the pellet and plate 100 μL on LB agar-plates containing rifampicin along with appropriate selection antibiotics. 5. Incubate the plates for 2–3 days at 28  C. Pick single colonies for plasmid analysis. Prepare glycerol stocks of each confirmed positive transformant and store at 80  C.

3.5.3 Agro-Infiltration in Tobacco Leaves

1. Grow the transformed Agrobacterium cultures (from glycerol stocks) overnight in 10 mL LB-broth medium supplemented with appropriate antibiotics (kanamycin at 50 μg/mL). Subculture in fresh medium the following morning. Grow until the OD600 reaches 0.6–0.8. 2. Harvest the cells by centrifugation at 4500  g for 5 min and resuspend in fresh LB-broth. Centrifuge again at 4500  g for 5 min and finally resuspend in infiltration medium (see Subheading 2.5, item 5) corresponding to an OD600 of 0.1. The cultures are now ready for infiltration. 3. Pick 2–3 young leaves (third leaf from the apex) of mature greenhouse grown tobacco plants for infiltration (see Note 12). Carefully infiltrate the cultures into the abaxial surface of the leaf using a 2 mL syringe without needle. Infiltrate at least 3 leaves for each construct. 4. Harvest the leaves for GUS assay/histochemical staining after 48–72 h of agro-infiltration.

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Fig. 2 Schematic diagram of the primer extension assay used for determination of Transcription Start Site (TSS) of any promoter. The lanes A, T, G, and C represent Sanger sequencing reactions electrophoresed on a 6% denaturing polyacrylamide gel next to the reaction product. The TSS is mapped as the residue (A/T/G/C) corresponding to the reaction product and indicated as +1 3.6 Transcription Start Site (TSS) Determination by Primer Extension Assay

Primer extension is a classical method used to map the 50 end (s) of RNA, thus determining the TSS of the promoter (see Note 13). The method typically uses a single-stranded DNA primer that is allowed to hybridize to the 50 end of the mRNA of the target gene at a position downstream of the TSS (defined as +1) [27] (Fig. 2).

3.6.1 Primer Tagging

Add 1 μL of 5 pM primer (oligonucleotide) to 2 μL of 10 T4 PNK buffer and 3 μL of γ-32P ATP along with 1 μL of T4

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polynucleotide kinase (10 U) (see Note 14). Bring the final volume up to 20 μL. Keep the reaction mix for 1 h at 37  C. Heat to inactivate the enzyme at 70  C for 10 min. 3.6.2 Probe Purification

Load 20 μL of tagged Probe on Microspin™ G-25 Columns. Centrifuge for 5 min, 1500  g at room temperature. Collect the flow-through in 1.5 mL microcentrifuge tubes containing radiolabeled primer (unincorporated oligonucleotide will remain in the resin). Tagged primer can be stored at 20  C.

3.6.3 Probe and RNA Precipitation

Precipitate the RNA and primer by adding 0.3 M sodium acetate and 2.5 volumes of ethanol. Incubate overnight at 80  C to precipitate RNA and probe. Centrifuge at 14,000  g for 45 min to pellet RNA and primer. Discard the supernatant and allow the pellet to dry. Dissolve the pellet by adding 8 μL of nuclease free water.

3.6.4 Primer Annealing

Add 2 μL of primer annealing buffer (PAB) to a total of 8 μL reaction mixture. Mix and spin briefly (see Note 15). Anneal the primers to their target RNA templates by increasing temperature to 70  C; hold for 10 min and then slowly cool back to room temperature in the metallic box on a heat dryer (see Note 16).

3.6.5 Reverse Transcriptase Reaction

Perform reverse transcriptase reaction following the manufacturers’ protocol. Incubate the reaction for 5 min at 25  C and gradually increase to 37  C for a total incubation time of 1 h. Denature the reaction at 70  C for 5 min and store at 4  C.

3.6.6 Ethanol Precipitation

Add 6 μL of 3 M sodium acetate and 150 μL of chilled ethanol to precipitate the product. Keep the reaction mixture at 20  C for 30 min. Centrifuge for 30 min at 4  C to pellet the product. Discard the supernatant (ethanol) and dry the pellet with a speed vacuum evaporator. Check the presence of tagged-precipitated product using a Geiger counter. Add 4 μL of formamide loading dye to the pellet and resuspend it.

3.6.7 Preparation of 8% Urea Polyacrylamide Gel

1. Set up the gel plate according to the manufacturers’ instructions. 2. Prepare the solution by adding 40% of (w/v) acrylamide and bis-acrylamide solution, 8% urea and 5 TBE in 80 mL of water. To dissolve the urea, heat in a microwave oven for 20 s (see Note 17). 3. Filter the solution through 3 mm Whatman™ paper. 4. Adjust the volume to 100 mL and add 200 μL of 25% APS and 80 μL of TEMED. Immediately pour the solution to cast the gel and allow it to solidify for 4 h. Set up the running apparatus and perform a free run for 2 h at 1700 V in 0.5 TBE.

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3.6.8 Sample Preparation and PAGE Separation

1. Boil the sample in boiling water for 3 min and immediately put it on ice. 2. Rinse the well using a 10 mL syringe and load 5 μL of each sample along with the sequencing products (Sanger sequencing reaction products). 3. Run the gel for 2–3 h till the bromophenol blue dye migrates to the bottom. Take out the gel by placing it against a 4 mm Whatman™ paper. Wrap the gel with plastic wrap to avoid the contamination of the cassette with isotope. Dry the gel and expose it to X-ray film with intensifying screen overnight at 80  C.

3.7

4

Conclusion

This chapter deals with the isolation, cloning, and initial characterization (TSS) of a pararetroviral promoter that can be exploited in the development of efficient plant gene expression systems; the next chapter will discuss the details of determining gene expression achieved by promoters and their transcriptional regulation.

Notes 1. Use acrylamide and bis-acrylamide with proper precautions since they can seriously damage health. Solutions of acrylamide and bis-acrylamide in ultrapure water are also available commercially for use in protein and nucleic acid electrophoresis, for example, Acrylamide/Bis-acrylamide, 30% solution (Sigma Aldrich). 2. Add Triton X-100 to a final concentration of 2% to lyse the chloroplasts and prevent virus particle aggregation. 3. DNaseI prevents the viral DNA from getting contaminated by any nuclear DNA. 4. Addition of SDS during digestion with Protease K significantly decreases the proportion of linear molecules in the virus preparation. 5. Do not exceed the incubation time while performing restriction digestion since most restriction enzymes possess star activities which might cut the DNA at nonspecific sites. 6. Prepare/procure competent cells that have good transformation efficiencies. This will increase the chances of getting positive transformants. 7. Take appropriate concentrations (and not volumes) of vector and insert during ligation. Preferably, ligate using a 1:3 (vector: insert) ratio overnight at 16  C.

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8. Add the protoplasts very gently on top of the sucrose cushion aligning the Pasteur pipette by the wall of the centrifuge tube. Avoid applying any pressure on the cushion. 9. Prior to electroporation, check the consistency of the suspended protoplasts by applying a pulse without any DNA. The time lapse for a single pulse should not be too long nor too short. Usually a pulse of 40–45 ms is recommended for an appropriate protoplast thickness. 10. Methods described under Subheadings 3.4 and 3.5 are rapid ways to assay promoter constructs, an alternative is to generate stable transgenic plants [28, 29]. 11. Do not over-thaw the competent cells. Always keep a control plate (without any DNA) to check for the presence of any contaminants in Agrobacterium cultures. 12. For infiltration, do not use leaves that are too old/matured. Preferably, use leaves with fewer venations and larger surface areas to facilitate the rapid infiltration of buffer. 13. Primer extension is a classic technique, alternatively use 50 -Rapid Amplification of cDNA Ends (RACE) PCR. 14. Use γ-32P ATP (radioactive) with utmost precaution since direct exposure poses severe hazards. Make sure to use radioactive materials only when guarded with a radiation shield and to follow the guidelines of your research institution. 15. PAB should not be added into the reaction mixture until the RNA is dissolved since the high salts of PAB will hinder in the process. 16. Annealing temperature can vary from 45 to 60  C with different primers. 17. Prolonged exposure to acrylamide and bis-acrylamide through direct inhalation can cause serious damage to health. References 1. Bousalem M, Douzery EJ, Seal S (2008) Taxonomy, molecular phylogeny and evolution of plant reverse transcribing viruses (family Caulimoviridae) inferred from full-length genome and reverse transcriptase sequences. Arch Virol 153(6):1085 2. Pfeiffer P, Hohn T (1983) Involvement of reverse transcription in the replication of cauliflower mosaic virus: a detailed model and test of some aspects. Cell 33(3):781–789 3. Ow DW, Jacobs JD, Howell SH (1987) Functional regions of the cauliflower mosaic virus 35S RNA promoter determined by use of the firefly luciferase gene as a reporter of promoter

activity. Proc Natl Acad Sci U S A 84 (14):4870–4874 4. Stavolone L, Kononova M, Pauli S et al (2003) Cestrum yellow leaf curling virus (CmYLCV) promoter: a new strong constitutive promoter for heterologous gene expression in a wide variety of crops. Plant Mol Biol 53(5):703–713 5. Dey N, Maiti IB (1999) Structure and promoter/leader deletion analysis of mirabilis mosaic virus (MMV) full-length transcript promoter in transgenic plants. Plant Mol Biol 40 (5):771–782 6. Khan A, Shrestha A, Bhuyan K et al (2018) Structural characterization of a novel full-

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length transcript promoter from Horseradish Latent Virus (HRLV) and its transcriptional regulation by multiple stress responsive transcription factors. Plant Mol Biol 96:1–18 7. Banerjee J, Sahoo DK, Raha S et al (2015) A region containing an as-1 element of Dahlia Mosaic Virus (DaMV) subgenomic transcript promoter plays a key role in green tissue-and root-specific expression in plants. Plant Mol Biol Report 33(3):532–556 8. Maiti IB, Shepperd RJ (1998) Promoter (FLt) for the full-length transcript of peanut chlorotic streak caulimovirus (PCLSV) and expression of chimeric genes in plants. Google Patents 9. Ranjan R, Dey N (2012) Development of vascular tissue and stress inducible hybrid–synthetic promoters through DOF-1 motifs rearrangement. Cell Biochem Biophys 63 (3):235–245 10. Bhattacharyya S, Dey N, Maiti IB (2002) Analysis of cis-sequence of subgenomic transcript promoter from the Figwort mosaic virus and comparison of promoter activity with the cauliflower mosaic virus promoters in monocot and dicot cells. Virus Res 90(1):47–62 11. Pattanaik S, Dey N, Bhattacharyya S et al (2004) Isolation of full-length transcript promoter from the Strawberry vein banding virus (SVBV) and expression analysis by protoplasts transient assays and in transgenic plants. Plant Sci 167(3):427–438 12. Deb D, Shrestha A, Maiti IB et al (2018) Recombinant promoter (MUASCsV8CP) driven Totiviral Killer Protein 4 (KP4) imparts resistance against fungal pathogens in transgenic tobacco. Front Plant Sci 9:278 13. Castillo E, Martinelli F, Zakharov-Negre F et al (2017) Effects of transgenic expression of Brevibacterium linens methionine gamma lyase (MGL) on accumulation of Tylenchulus semipenetrans and key aminoacid contents in Carrizo citrange. Plant Mol Biol 95(4-5):497–505 14. Chatterjee A, Das NC, Raha S et al (2017) Enrichment of apoplastic fluid with therapeutic recombinant protein for efficient biofarming. Biotechnol Prog 33(3):726–736 15. Koziel MG, Beland GL, Bowman C et al (1993) Field performance of elite transgenic maize plants expressing an insecticidal protein derived from Bacillus thuringiensis. Nat Biotechnol 11(2):194 16. Neuhaus J-M, Ahl-Goy P, Hinz U et al (1991) High-level expression of a tobacco chitinase gene in Nicotiana sylvestris. Susceptibility of transgenic plants to Cercospora nicotianae infection. Plant Mol Biol 16(1):141–151

17. Patro S, Maiti S, Panda SK et al (2015) Utilization of plant-derived recombinant human β-defensins (hBD-1 and hBD-2) for averting salmonellosis. Transgenic Res 24(2):353–364 18. Shrestha A, Khan A, Dey N (2018) Cis-trans engineering: advances and perspectives on customized transcriptional regulation in plants. Mol Plant 11:886–898. https://doi.org/10. 1016/j.molp.2018.05.008 19. Acharya S, Ranjan R, Pattanaik S et al (2014) Efficient chimeric plant promoters derived from plant infecting viral promoter sequences. Planta 239(2):381–396 20. Patro S, Kumar D, Ranjan R et al (2012) The development of efficient plant promoters for transgene expression employing plant virus promoters. Mol Plant 5(4):941–944 21. Ranjan R, Patro S, Kumari S et al (2011) Efficient chimeric promoters derived from fulllength and sub-genomic transcript promoters of Figwort mosaic virus (FMV). J Biotechnol 152(1):58–62 22. Gardner R, Shepherd R (1980) A procedure for rapid isolation and analysis of cauliflower mosaic virus DNA. Virology 106(1):159–161 23. Richins RD, Scholthof HB, Shepherd RJ (1987) Sequence of figwort mosaic virus DNA (caulimovirus group). Nucleic Acids Res 15(20):8451–8466 24. Stenger D, Mullin R, Morris T (1988) Isolation, molecular cloning, and detection of strawberry vein banding virus DNA. Phytopathology 78(2):154–159 25. Cominelli E, Galbiati M, Albertini A et al (2011) DOF-binding sites additively contribute to guard cell-specificity of AtMYB60 promoter. BMC Plant Biol 11(1):162 26. Karthikeyan AS, Ballachanda DN, Raghothama KG (2009) Promoter deletion analysis elucidates the role of cis elements and 5’UTR intron in spatiotemporal regulation of AtPht1; 4 expression in Arabidopsis. Physiol Plant 136 (1):10–18 27. Carey MF, Peterson CL, Smale ST (2013) The primer extension assay. Cold Spring Harb Protoc 2013(2):164–l73 28. Fisher DK, Guiltinan MJ (1995) Rapid, efficient production of homozygous transgenic tobacco plants with Agrobacterium tumefaciens: a seed-to-seed protocol. Plant Mol Biol Report 13(3):278–289 29. Zhang X, Henriques R, Lin S-S et al (2006) Agrobacterium-mediated transformation of Arabidopsis thaliana using the floral dip method. Nat Protoc 1:641. https://doi.org/ 10.1038/nprot.2006.97

Chapter 20 Biochemical and Molecular Characterization of Novel Pararetroviral Promoters in Plants Ahamed Khan, Ankita Shrestha, and Nrisingha Dey Abstract Special attention needs to be given to defining and studying the regulatory apparatus of different pararetroviral promoters under various physiological conditions because they have significant sequence heterogeneity and unique distributions of stress-responsive cis-elements. Transcriptional regulation studies of a pararetroviral promoter involve both gene expression analyses and investigation of its structural/regulatory framework. The expression of reporter genes such as β-Glucuronidase (GUS) or Luciferase (LUC) transcriptionally fused to a promoter usually determines the strength or function of a target promoter. In parallel, DNA–protein interaction studies are employed to assess the functional relevance of predicted transcription factor binding sites in target pararetroviral promoter sequences. In this chapter, we will describe protocols used to determine the transgene integration and expression in transgenic plant systems. Alongside, we will also discuss the fusion reporter assays that can determine the promoter activity and DNA–protein interaction studies that aid in the evaluation of its transcriptional regulation. Key words GUS, Luciferase, EMSA, Real-time PCR, Histochemical

1

Introduction Pararetroviral promoters typically contain multiple cis-acting elements that have specific binding sites for cognate plant transcription factors (TFs) that usually regulate gene transcription. The apparent similarity in the genomic organization and replication strategy of the Caulimoviruses make them excellent models to study the transcriptional framework associated with host-specific adaptations. Several studies have revealed the mechanisms that regulate the transcription and unique expression patterns of caulimoviral promoters [1, 2]. Such studies are crucial for improving our basic understanding of gene regulation [3] and expand the toolbox of available promoters for use in plant biotechnology.

Ahamed Khan and Ankita Shrestha contributed equally to this work. Walter Gassmann (ed.), Plant Innate Immunity: Methods and Protocols, Methods in Molecular Biology, vol. 1991, https://doi.org/10.1007/978-1-4939-9458-8_20, © Springer Science+Business Media, LLC, part of Springer Nature 2019

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The reporter genes such as β-Glucuronidase (GUS), Green Fluorescent Protein (GFP), or Luciferase (LUC) placed downstream of the promoters are used to define the transcriptional activity of a promoter [4, 5] in both transient and transgenic platforms. In addition, DNA–protein interactions play an important role in controlling the transcriptional regulation of any gene in a plant cell. TFs are specialized trans-regulatory proteins that recognize and bind to specific cis-elements (enhancers and silencers) which in turn activate or suppress gene transcription. Each TF has two functional domains, a protein–protein interaction domain and a DNA-binding domain (helix-turn-helix, leucine zipper, zinc finger, or helix-loop-helix) which aid in their interaction with the promoter or other TFs [6]. In this chapter, we describe the determination of transgene copy number and gene expression analysis in a transgenic platform. We also describe fusion reporter gene assays employed to define Caulimoviral promoter activity using biochemical and fluorometric approaches. Subsequently, we discuss a protocol for electrophoretic mobility shift assays (EMSA), a rapid way of studying DNA–protein interactions, as a first step toward determining the selective binding of TFs to cis-elements which in turn regulate the transcriptional activity of a pararetroviral promoter.

2

Materials

2.1 Genomic DNA Isolation and Quantitative Real-Time PCR (qPCR) to Determine Transgene Copy Number

1. Plant genomic DNA extraction kit, for example, DNeasy Plant Mini Kit (Qiagen). 2. DNA quantification kit, for example, Qubit Fluorometric Quantitation (Thermo Fischer Scientific). 3. Spectrophotometer. 4. Oligonucleotides (both forward and reverse) specific to GUS and a SCR (single copy reference) gene (see Note 1). 5. SYBR Green Master Mix, for example, SYBR™ Green PCR Master Mix (Thermo Fischer Scientific). 6. Real-Time PCR system (7500 Fast Real-Time System, Applied Biosystems) or equivalent.

2.2 RNA Isolation and qPCR to Determine Promoter Strength

1. Plant total RNA extraction kit, for example, Plant RNeasy extraction kit (Qiagen, CA, USA). 2. Diethyl pyrocarbonate (DEPC) (see Note 2). 3. TRIZOL® reagent. 4. cDNA synthesis kit, for example, First Strand cDNA Synthesis Kit (Thermo Fischer Scientific). 5. SYBR Green Master Mix, for example, SYBR™ Green PCR Master Mix (Thermo Fischer Scientific).

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6. Real-Time PCR, for example, 7500 Fast Real-Time System (Applied Biosystems). 7. Oligonucleotides (both forward and reverse) specific to GUS. 2.3 β-Glucuronidase (GUS) Assay

1. 0.5 M EDTA, pH 8.0: Add 46.52 g of EDTA in sterile milli-Q water. Adjust pH to 8.0 using 1 M NaOH and bring the volume to 250 mL. 2. 2 M KPO4: Dissolve 63.2 g of K2HPO4 and 5 g of KH2PO4 in sterile milli-Q water. Adjust pH to 7.8 with 1 M KOH and increase the volume to 200 mL. 3. 10 N NaOH: Dissolve 100 g of NaOH in sterile milli-Q water and bring the volume to 200 mL. Store at room temperature (RT) in a plastic bottle. 4. 1 M dithiothreitol (DTT): Dissolve 1.545 g of DTT in 0.01 M sodium acetate (NaOAc). Adjust the pH to 5.2 and increase the volume to 10 mL. Filter sterilize and store at
80  C (see Note 3). 5. Assay buffer (100 mL): Add 5 mL of 2 M KPO4 (pH 7.8), 200 μL of 0.5 M EDTA (pH 8.0), 1 mL of Triton X-100, 12.5 mL of 80% glycerol, and 15.4 mg of DTT in 81.1 mL autoclaved milli-Q water. Store at RT. 6. Methylumbelliferyl β-D-glucuronide (MUG) solution (1mM): Add 3.52 mg of MUG and 100 μL DMSO in 9.9 mL of assay buffer. Store at
20  C. Prepare fresh MUG solution before each use. 7. Stop solution, 0.2 M Na2CO3: Dissolve 21.2 g of Na2CO3 in autoclaved water. Adjust the volume to 1000 mL. 8. Bradford reagent. 9. Spectrophotometer (Unico UV-2000 Spectrophotometer, SpectraLab Scientific Incorporation, USA) or equivalent. 10. Fluorometer (TKO 100, Hoefer Scientific Instruments) or equivalent.

2.4 Histochemical GUS Staining

1. 2 0.1 M Phosphate buffer, pH 7.0: Dissolve 12.0 g Na2PO4 and 14.19 g Na2HPO4 in 800 mL distilled H2O. Adjust pH to 7.0 (using 1 M NaOH or 1 M HCl) and bring the volume to 1000 mL. 2. X-Gluc solution: Dissolve 1 mg 5-bromo-4-chloro-3-indolyl β-D-glucuronide in 0.1 mL methanol. Add 1 mL 2 phosphate buffer, 20 μL of 0.1 M potassium ferrocyanide, 20 μL of 0.1 M potassium ferricyanide, 10 μL of 10% (w/v) Triton X-100 solution, and 0.85 mL water. 3. Fixative solution: 4% formaldehyde in 1 phosphate buffer. Prepare fresh before each use.

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4. Washing solution: 70% (v/v) ethanol. 5. 50% and 100% glycerol. 6. Bright-field microscope (Olympus SZX12) or equivalent. 2.5 Luciferase Reporter Gene Assay

1. Murashige and Skoog (MS) medium: 4.33 g/L commercially available basal salt mixture, 3% sucrose. Autoclave and store at RT. 2. Luciferase assay kit, for example, Luciferase Assay System (Promega Corporation) containing Luciferase Assay Reagent (LAR) and Cell Lysis Reagent. 3. D-Luciferin stock solution: 150 mM D-luciferin in milli-Q water. Store as 1-mL aliquots in the dark at
80  C. 4. D-Luciferin working solution: 5 mM D-luciferin in 0.01% Triton X100, freshly prepared before each assay. 5. Colorimetric assay for protein concentration following detergent solubilization, for example, Bio-Rad DC™ protein assay kit. 6. Purified firefly luciferase standard: Prepare serial dilutions (1:108, 1:109, 1:1010, 1:1011) in Cell Lysis Reagent supplemented with l mg/mL BSA. 7. Luminometer (TD-20/20 Luminometer, Turner Design) or equivalent.

2.6 Recombinant Protein Purification

1. Protein expression vectors: His Tag6 pET series (pET28, pET29, pET30, pET32). 2. Antibiotic stocks: Prepare ampicillin (100 mg/mL) and kanamycin (50 mg/mL) stocks in sterile water and filter sterilize with a 0.22 μm filter. Store at
20  C. 3. Luria-Bertani (LB) Miller Broth bacterial growth medium: 25 g/L commercially available mixture. Autoclave and store at RT. 4. Bacterial expression system: Escherichia coli strains competent cells. 5. Ni-NTA Resin. 6. 1 M isopropyl β-D-1-thiogalactopyranoside (IPTG) stock: Dissolve 2.38 g of IPTG in 8 mL of autoclave distilled water, bring to a final volume of 10 mL. Filter sterilize with a 0.22 μm syringe filter and store in 1 mL aliquots at
20  C. 7. 4 M NaCl: Dissolve 234 g of NaCl in 1 L of water. Autoclave and store at RT. 8. 1 M imidazole solution: Dissolve 68.08 g of imidazole in 1000 mL of sterile dH2O.

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9. Plastic 5 mL columns for gravity purification, for example, UBPBio. 10. Bacterial Cell Lysis buffer: 50 mM Tris–HCl pH 8.0, 300 mM NaCl, 0.5% Triton X-100, 0.05% β-mercaptoethanol (β-ME), 1 mM PMSF, 1% Protease Inhibitor Cocktail powder (Sigma) (see Note 3). 11. Equilibration buffer: 50 mM Tris–HCl (pH 8.0), 300 mM NaCl, 0.05% β-ME, 1 mM PMSF, 1% Protease Inhibitor Cocktail, 10 mM Imidazole. 12. Wash buffer: Equilibration buffer containing 20 mM imidazole. 13. Elution buffer: Equilibration buffer containing 250 mM imidazole. 14. Dialysis buffer: 20 mM Tris–HCl (pH 7.5), 150 mM NaCl, 2.5 mM MgCl2, 1 mM β-ME, 10% v/v glycerol. 15. 30% Acrylamide-Bis Solution (29:1): Dissolve 29 g of acrylamide and 1 g N,N0 -methylenebisacrylamide powder in 100 mL of autoclaved water. Filter stock through a filter paper (use Whatman® qualitative filter paper, Grade 1) and store at 4  C or order commercially available solution (see Note 4). 16. 1.5 M Tris, pH 8.8: Dissolve 36.3 g of Tris in 100 mL water, adjust the pH to 8.8, and increase the volume to 200 mL. 17. 0.5 M Tris, pH 6.8: Dissolve 6 g Tris in 40 mL water, adjust the pH to 6.8, and bring the volume to 100 mL. 18. Ammonium persulfate (APS) 10%: Dissolve 100 mg of APS in 1 mL of autoclaved water. 19. Sodium dodecyl sulfate (SDS) 10%: Dissolve 10 g of SDS in 100 mL of deionized water and mix it until dissolved. Do not autoclave. 2.7 EMSA (Electrophoretic Mobility Shift Assay)

1. γ-32P ATP, high-specific-activity. 2. T4 Polynucleotide Kinase (10 U/μL) Thermo Scientific™ Cat#: EK0031. 3. T4 Polynucleotide Kinase Forward Reaction Buffer (5): 500 mM Tris–HCl pH 7.6, 100 mM MgCl2, 5 mM DTT, 1 mM spermidine, and 1 mM EDTA. 4. 10 Oligo annealing buffer: 10 mM Tris pH 8.0, 100 mM NaCl, 1 mM EDTA. 5. 10 TBE Tris/Borate/EDTA (TBE): Dissolve 121.1 g of Tris base, 61.8 g of boric acid, and 7.4 g of EDTA disodium salt. Bring the final volume to 1 L. 6. Poly deoxyinosinic-deoxycytidylic (dI-dC) acid sodium salt.

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7. Bovine serum albumin (BSA) 10 mg/mL: Dissolve 10 mg of BSA in 1 mL of autoclaved water. 8. 40% Acrylamide:Bis Solution: Dissolve 380 g of acrylamide and 20 g of N,N0 -methylbisacrylamide in 500 mL of water. Bring the final volume to 1 L. Filter sterilize the stock through 0.45 μm pore filter. Store in dark bottles at 4  C (see Note 4). 9. 10 EMSA binding buffer: 100 mM Tris (pH 7.5), 10 mM MgCl2, 1 mM EDTA, 10% v/v glycerol, 1 mg/mL BSA, and 250 ng/μL of poly dI-dC. 10. Gel dryer. 11. 3 mm Whatman paper. 12. X-ray film.

3

Methods The following protocols can be employed with a variety of plant materials expressing the reporter gene construct generated by protoplast transfection, transient expression in tobacco, or stable transformation by biolistic or Agrobacterium [3, 7–9], for further characterization of any pararetroviral promoter.

3.1 Genomic DNA Isolation and Quantitative Real-Time PCR (qPCR) to Determine Transgene Copy Number

When using stable transgenic plants transformed with new viral promoter constructs, the transgene copy number must first be determined. This can be done by a SYBR green-based qPCR using a DNA template which reveals its copy number [10, 11]. It is a quick alternative to the conventional Southern Hybridization technique that involves radioactive probes [12] and is relatively time consuming. The details of this technique are described below: 1. Extract the total genomic DNA from approximately 100 mg of leaf tissue using a plant genomic DNA extraction kit. 2. Quantify the DNA and prepare appropriate serial dilutions following the manufacturer’s instructions. Prepare dilutions of primers in sterile water at a final concentration of 10 μM (see Note 5). 3. Set up the qPCR reaction using the SYBR Green Master Mix. Perform the reactions using standard PCR conditions: 10 min at 95  C; 40 cycles of 10 s at 95  C; 1 min at 60  C; 5 s at 72  C. Analyze the data obtained as per manufacturer’s instructions and determine the Ct values for both target and reference genes (see Note 6). 4. Calculate the transgene copy number following the standard 2ΔΔCt method [13].

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3.2 RNA Isolation and Quantitative RealTime PCR

3.2.1 RNA Extraction Using TRIZOL Reagent

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An important means to assess gene expression is through the determination of the messenger ribonucleic acid (mRNA) transcript levels corresponding to a particular gene. It involves the isolation of total cellular RNA and subsequent analysis of the targeted mRNA in real-time quantitative reverse transcription polymerase chain reaction [14, 15]. This process is highly sensitive and reliable which allows rapid detection of transcript levels in plant samples. 1. Use only DEPC (see Note 2)-treated supplies and utensils (microfuge tubes, pipette tips and tip boxes, mortar and pestles, spatulas and water) for RNA isolation. 2. Harvest 1 g young leaf tissues and freeze them immediately in liquid nitrogen. Homogenize the tissue in prechilled mortars with a pestle until finely powdered. Transfer the ground tissue to 1.5 mL microfuge tubes and add 1 mL of Trizol reagent to the homogenized tissue. 3. Vortex and incubate at room temperature for 10 min. Centrifuge the samples at 13000  g for 10 min at 4  C to remove extracellular materials. 4. Transfer the supernatant to new tubes and add 200 μL of chloroform per 1 mL of TRIZOL. Shake vigorously for 15 s and incubate at room temperature for 3 min. 5. Centrifuge the samples at 13000  g for 10 min at 4  C for phase separation. Carefully transfer the aqueous phase (ca 50–60% of TRIZOL volume) to new tubes and add 500 μL of isopropanol per 1 mL of TRIZOL. Incubate at room temperature for 10 min. 6. Centrifuge the samples at 13000  g for 10 min at 4  C to pellet. Remove the supernatant and wash the pellets with 70% (v/v) ethanol diluted with DEPC-treated water. 7. Vortex and centrifuge at 7500  g for 5 min at 4  C. Discard the supernatant and air dry the pellets for 5–10 min at room temperature. Do not over-dry the pellets. 8. Dissolve the dried pellets in DEPC-treated water by incubating at 37  C for 15 min (see Note 7).

3.2.2 Quantitative RealTime PCR (qRT-PCR)

The qRT-PCR is essentially performed to check the relative quantification of GUS transcript in the total RNA sample isolated from tobacco leaves, the steps of which are described below: 1. For cDNA synthesis, perform reverse transcription using a cDNA synthesis kit as described in the manufacturer’s protocol. 2. Perform the qRT-PCR following the manufacturers’ instructions (for example, of the iTaq universal SYBR Green one-step

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kit, Bio-Rad, USA) in a total reaction volume of 20 μL. Use the tobacco α-tubulin as the loading control for all independent samples. 3. Calculate the relative abundance of GUS transcript using the comparative threshold cycle (Ct) method [16]. 3.3 β-Glucuronidase (GUS) Assay

1. Put protoplasts into 1.5 mL tubes and spin at 100 x g for 4 min (refer to the previous Chapter). In the case of transgenic leaf samples, crush using sterile sand and then place in 1.5 mL tubes followed by a spin at 10,000  g for 4 min. Remove the supernatant and add 110 μL of GUS assay buffer to the pellet and sonicate to shear the cells (pulse at 25% amplitude for 2 s ON and 3 s OFF) while keeping on ice. Centrifuge at 12,000  g for 10 min at 4  C. Retain the supernatant that contains the protein. 2. Aliquot 75 μL of MUG solution in a microfuge tube and keep at 37  C for 10 min in a dry bath. Add 25 μL of protein sample (from step 1) to the pre-warmed tube and note the time (see Note 8). This is the test sample. Simultaneously, prepare the sample for the reference control (CaMV35S). 3. For the 0 min reading, immediately pipette out 25 μL solution (from step 2) and add to a new tube containing 200 μL of Stop solution. For the 10 min reading, add a 25 μL aliquot (from step 2) to another tube (at the appropriate time) containing 200 μL Stop solution. Repeat the same for the 20 min reading. Measure the fluorescence at 0, 10, and 20 min using a fluorimeter. 4. Measure the protein concentration of each protein sample (from step 1) using Bradford Reagent [17]. Calculate the GUS activity of the test sample and reference control and express as nmol MUG/min/mg protein. Test all constructs in at least three independent experiments.

3.4 Histochemical GUS Staining

β-Glucuronidase (GUS) is a reporter gene that is frequently used in plant molecular biology to study gene expression in plant systems. The GUS gene fusion system is based on the detection of the enzymatic activity of GUS in protein extracts using fluorimetric and histochemical assays, respectively. The commonly used substrate for histochemical GUS staining in tissues is 5-bromo-4chloro-3-indolyl glucuronide (X-Gluc) which yields a blue precipitate at the site of enzyme activity and reveals the pattern of gene expression in plant tissues. 1. For staining, harvest fresh (infiltrated or transgenic) plant tissue samples. Vacuum infiltrate the tissues by dipping in X-Gluc solution in dark at 37  C for 3–4 h or overnight or until a blue

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distinct color appears. The staining time should not exceed more than 24 h. 2. Remove the tissues from the staining solution and rinse once with sterile water. 3. In case of leaf samples, incubate in 70% ethanol (v/v) overnight or until the chlorophyll is completely removed (bleached) (see Note 9). Rinse the samples with sterile water. 4. Mount the tissue samples in 100% glycerol on microscope slides and examine using a light microscope. 3.5 Luciferase Reporter Gene Assay

The luciferase reporter assay is commonly used to study gene expression, particularly to evaluate the transcriptional activity of a promoter. It is a rapid and relatively economical “quantitative measurement” of any promoter activity [4]. The firefly luciferase (LUC) gene acts as a real-time reporter to instantaneously examine the regulation of gene expression in plants [18]. The routine luciferase assay can be performed as follows: 1. Harvest the plant tissue and immediately freeze in liquid nitrogen. Grind the frozen plant tissue to a fine powder with a prechilled mortar and pestle. 2. Transfer approximately 100 mg powder into a microcentrifuge tube and place on ice. Next, add two volumes of ice-cold lx cell lysis reagent to the powder. 3. Vortex the suspension and allow the tube to thaw at RT. 4. Centrifuge the tube at 13,000  g at 4  C for 10 min. Transfer the supernatant to a fresh microcentrifuge tube placed on ice. 5. Thaw appropriate volume of LAR for assaying all samples. Add 100 μL LAR to 30 μL of plant extract and incubate for 3 s in a luminometer cuvette. 6. Measure the emitted light over a period of 10 s (starting at 3 s gradually up to 10 s with a 2 s interval, i.e., at 3, 5, 7, and 9 s) in the luminometer. 7. Perform a Bradford assay to determine the protein concentration in each of the plant extracts. Prepare the standard using 100 pg of purified luciferase (Sigma) dissolved in 30 μL of LAR. 8. Convert the luminescence intensity to “specific luciferase activity” and express as “pg luciferase per μg protein.”

3.6 DNA–Protein Interaction Studies

DNA–protein interactions are critical, particularly in regulating transcription and determining the levels of gene expression. Such interactions can be studied by identifying and studying the protein component complexed with the regulatory region of the promoter. For analysis of any DNA–protein complex, the gel shift or

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electrophoretic mobility shift assay (EMSA) and DNaseI footprinting assay are two widely used methods that can reveal sequence-specific binding of proteins, particularly TFs, to a promoter region [19]. EMSA is one of the most rapid and reliable techniques which determines the formation of DNA–protein complexes. Also, the use of mutational and competition EMSA can reveal important information about the binding specificities of cognate TFs to the cis-regulatory sequences present on the promoter. 3.6.1 Expression and Purification of His-Tagged Recombinant Proteins

1. Transform the plasmid (pET series vector containing the gene of interest) in BL21 competent cells and plate on LB agar containing appropriate antibiotics. 2. Pick up a single colony and inoculate in 10 mL of LB broth; grow overnight at 37  C with constant shaking at 200 rpm. 3. Inoculate the overnight culture (1/100 volume) in appropriate LB broth (100 mL or 500 mL) and allow to grow till the OD600 reaches 0.5–0.6. 4. Immediately induce the bacterial culture by adding 0.5 mM IPTG (see Note 10). Grow the induced culture at 28  C for 8 h (see Note 11). Harvest the culture by centrifugation at 10,000  g for 5 min. Store the pellet at
80  C. 5. Thaw the obtained pellet on ice for 5–10 min. Resuspend the pellet in 10 mL of lysis buffer supplemented with 100 μL of lysozyme (20 mg/mL) and 10 μL of benzonase nuclease. Incubate on ice for 30 min. Vortex the culture at 10 min intervals. 6. Sonicate on ice at 30–35 A˚ (pulse 5 s ON/6 s OFF) for 3 min until the cell suspension becomes translucent (see Note 12). 7. Centrifuge at 13,000  g for 10 min at 4  C. Transfer the supernatant which contains the soluble protein into a new centrifuge tube. Set aside 100 μL of the supernatant for SDS-PAGE analysis. 8. Equilibrate the resin (Ni-NTA, Cobalt) in equilibration buffer. Add the equilibrated resin to the supernatant and keep it on a rocker at 20 rpm for 2 h at 4  C. Recombinant His-tagged proteins will selectively bind to the resin. 9. Pass the supernatant along with resin through Gravity Flow Columns until the resin settles at the bottom. Allow the buffer to completely drain out. Keep 50 μL of flow-through for SDS-PAGE analysis. 10. Wash the resin thrice with two bed volumes of wash buffer. Keep 50 μL of sample after each wash for SDS-PAGE analysis. Elute the recombinant protein in three volumes of resin (2–3 times) with elution buffer.

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11. Dialyze the eluted protein fraction against dialysis buffer overnight at 4  C (see Note 13). Concentrate the dialyzed recombinant protein samples using Amicon Ultra centrifugal filter units. 12. Check the quality of protein by SDS-PAGE and perform a Bradford assay to quantify the protein. 3.6.2 EMSA (Electrophoretic Mobility Shift Assay)

The EMSA is performed to determine the formation of DNA–protein complexes. The recombinant protein which was purified in the above step is used for this assay, the details of which are described below: 1. Primer tagging: Add 1 μL of 5 pmol oligonucleotide primer to 2 μL of 10 T4 PNK buffer and 3 μL of γ-32P ATP along with 1 μL of T4 polynucleotide kinase (10 U) (see Note 14). Bring the final volume up to 20 μL. Keep the reaction mixture for 1 h at 37  C. Heat inactivate for 10 min at 70  C (see Note 15). 2. Probe purification: Load 20 μL of tagged probe on Microspin™ G-25 columns and centrifuge for 5 min, 1500  g at room temperature. Collect the flow-through in 1.5 mL microfuge tubes containing radiolabeled primer (unincorporated oligonucleotide will remain in the resin). The tagged primer can be stored at
20  C. 3. Primer annealing: Mix equal volumes of tagged forward primer and untagged reverse primer (5 pmol), 4 μL of 10 annealing buffer and bring the volume up to 40 μL. Heat the sample at 95  C for 15 min and allow it to cool down slowly at room temperature to 4  C. Store at
20  C until use. 4. 5% polyacrylamide gel preparation: Clean the glass plate with ethanol and assemble the casting plates with 1.5 mm spacer and comb. Cast the gel by mixing 3.12 mL of 40% acrylamide, 20.05 mL of water, 1.25 mL 10 TBE, 500 μL of 10% APS, and 80 μL TEMED. Add TEMED immediately before casting the gel (see Note 4). Mix by gently swirling and pour the solution immediately between the plates. Insert a comb (1.5 mm) with wide teeth and allow the gel to polymerize; then remove the comb. Mount the gel in an electrophoresis tank and add 0.5 TBE to the tank (see Note 16). 5. Sample preparation: Place the recombinant protein with 1 binding buffer on ice. Add radiolabeled probe (20,000 cpm) in a total reaction volume of 30 μL. Incubate for 30 m at RT. This allows the binding between DNA and protein to occur. Pre-run the gel for 30 m at 180 V. Electrophorese the samples in the 5% gel with 0.5 TBE at 180 V for 2 h. Remove the gel from the tank and place on 4 mm Whatman filter paper. Wrap the gel with plastic wrap and dry using a vacuum dryer at

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75  C for 1 h. Place the dried gel in a cassette with an intensifying screen and expose to an X-ray film overnight at
80  C. 6. Look for a band shift on the film which indicates if the protein binds the DNA probe

4

Notes 1. The SCR should be an endogenous single-copy gene naturally present in the same genomic DNA sample that is to be assayed. With SYBR Green qPCR it is fairly easy to compare amplification in reactions using different primer pairs, as long as the template DNA is the same and reactions have similar amplification efficiency. 2. DEPC is highly toxic till autoclaved and hence should be handled carefully. 3. PMSF and DTT should be freshly prepared before each use because PMSF is highly unstable in aqueous solutions (110 min at pH 7.0, 55 min at pH 7.5, and 35 min at pH 8.0, all at 25  C) while DTT rapidly degrades in air as a result of oxidation. PMSF has a tendency to crystallize when stored in isopropanol at
20  C. It is highly recommended to dissolve PMSF immediately before use. Dissolve gently by tapping and avoid using a vortex. 4. Use acrylamide, bis-acrylamide, and TEMED with proper precautions since they can seriously damage health. 5. Take appropriate controls such as a “no-template control” for PCR reactions. 6. PCR conditions may vary depending on the annealing temperature of the primers. 7. It is advisable to check the integrity of the isolated RNA on an 1% agarose gel followed by quantification in a spectrophotometer. 8. Perform all steps including measurement of fluorescence in the dark. 9. If the chlorophyll bleaching is not complete, use equal volumes of absolute ethanol and acetone for a maximum of 2 h. 10. IPTG concentration may vary depending on the nature of the protein. A concentration that works well for one protein might be too high or low for another protein. Using higher concentrations of IPTG might result in protein accumulating in inclusion bodies due to excessive protein expression. Try to carry out protein induction at very low IPTG concentrations and then gradually increase concentrations.

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11. Protein induction time and cell growth temperature may vary depending on the nature of the protein. If the protein is not getting induced at one temperature, try checking for optimal induction using a series of temperatures such as 18  C for 16 h, 25  C for 8–12 h, 30  C for 6–12 h, 37  C for 3–4 h. 12. Avoid extensive sonication and heating of the sample. 13. Prechill the dialysis buffer at 4  C for at least 45 min before use. 14. Use γ-32P ATP (radioactive) only when guarded with a radiation shield. Direct exposure may cause serious health damage. 15. During primer tagging, short spins can help prevent sample evaporation when heated at 95  C for 15 min. 16. The percentage of gel may vary depending on the size of the probe and protein. References 1. Khan A, Shrestha A, Bhuyan K, Maiti IB, Dey N (2018) Structural characterization of a novel full-length transcript promoter from Horseradish Latent Virus (HRLV) and its transcriptional regulation by multiple stress responsive transcription factors. Plant Mol Biol 96:1–18 2. Sarkar S, Das A, Khandagale P, Maiti IB, Chattopadhyay S, Dey N (2017) Interaction of Arabidopsis TGA3 and WRKY53 transcription factors on Cestrum yellow leaf curling virus (CmYLCV) promoter mediates salicylic acid-dependent gene expression in planta. Planta 247:1–19 3. Davey MR, Anthony P (2010) Genetic transformation—biolistics. In: Altpeter F, Sandhu S (eds) Plant cell culture. https://doi.org/10. 1002/9780470686522.ch12 4. Millar AJ, Short SR, Hiratsuka K, Chua N-H, Kay SA (1992) Firefly luciferase as a reporter of regulated gene expression in higher plants. Plant Mol Biol Report 10(4):324–337 5. Patro S, Kumar D, Ranjan R, Maiti IB, Dey N (2012) The development of efficient plant promoters for transgene expression employing plant virus promoters. Mol Plant 5 (4):941–944 6. Harrison SC (1991) A structural taxonomy of DNA-binding domains. Nature 353 (6346):715–719 7. Fisher DK, Guiltinan MJ (1995) Rapid, efficient production of homozygous transgenic tobacco plants withagrobacterium tumefaciens: a seed-to-seed protocol. Plant Mol Biol Report 13(3):278–289 8. Acharya S, Ranjan R, Pattanaik S, Maiti IB, Dey N (2014) Efficient chimeric plant

promoters derived from plant infecting viral promoter sequences. Planta 239(2):381–396 9. Ranjan R, Dey N (2012) Development of vascular tissue and stress inducible hybrid–synthetic promoters through DOF-1 motifs rearrangement. Cell Biochem Biophys 63 (3):235–245 10. Bubner B, Baldwin IT (2004) Use of real-time PCR for determining copy number and zygosity in transgenic plants. Plant Cell Rep 23 (5):263–271 11. Truman W, Sreekanta S, Lu Y, Bethke G, Tsuda K, Katagiri F, Glazebrook J (2013) the calmodulin binding protein 60 family includes both negative and positive regulators of plant immunity. Plant Physiol 113:1741–1751 12. Southern EM (1975) Detection of specific sequences among DNA fragments separated by gel electrophoresis. J Mol Biol 98 (3):503–517 13. Li Z, Hansen JL, Liu Y, Zemetra RS, Berger PH (2004) Using real-time PCR to determine transgene copy number in wheat. Plant Mol Biol Report 22(2):179–188 14. Jain M, Nijhawan A, Tyagi AK, Khurana JP (2006) Validation of housekeeping genes as internal control for studying gene expression in rice by quantitative real-time PCR. Biochem Biophys Res Commun 345(2):646–651 15. Nicot N, Hausman J-F, Hoffmann L, Evers D (2005) Housekeeping gene selection for realtime RT-PCR normalization in potato during biotic and abiotic stress. J Exp Bot 56 (421):2907–2914 16. Livak KJ, Schmittgen TD (2001) Analysis of relative gene expression data using real-time

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quantitative PCR and the 2
ΔΔCT method. Methods 25(4):402–408 17. Bradford MM (1976) A rapid and sensitive method for the quantitation of microgram quantities of protein utilizing the principle of protein-dye binding. Anal Biochem 72 (1-2):248–254 18. Spolaore S, Trainotti L, Casadoro G (2001) A simple protocol for transient gene expression in

ripe fleshy fruit mediated by Agrobacterium. J Exp Bot 52(357):845–850. https://doi.org/ 10.1093/jexbot/52.357.845 19. Grierson C, Du JS, Zabala M, Beggs K, Smith C, Holdsworth M, Bevan M (1994) Separate cis sequences and trans factors direct metabolic and developmental regulation of a potato tuber storage protein gene. Plant J 5 (6):815–826

Chapter 21 Sampling and Handling of Soil to Identify Microorganisms with Impacts on Plant Growth Robert J. Kremer Abstract Sampling and handling of soils and rhizosphere soil are very critical steps for obtaining representative microbial cultures and genomic material from these environments. Attention to position in the landscape of a sampling site, previous management of the site, time of year, and depth of sampling is important to assure representative samples. Detailed protocols are provided to assist environmental microbiologists and molecular biologists in proper sampling and handling of soils that serve as the source of cultures and DNA for subsequent use in important experiments carried out in the laboratory. Key words Soil profile, Landscape position, Composite soil sample, Soil microbial community, Soil survey, Rhizosphere

1

Introduction Field sampling of soils for characterization of microorganisms associated with plant growth impacts may be the step most vulnerable to errors affecting the accuracy in estimating the actual size, composition, and functions of the soil microbial community. Soil microbial characterization has advanced tremendously during the past two decades as molecular and genomics methods were adapted to obtain vast taxonomic categorizations of microorganisms in various soil ecosystems [1] that previously was not possible using traditional culture-based techniques. Thus, properly sampled soils based on defined protocols that account for the complex properties of the soil environment has become a very critical research methods component to assure the best representation of the soil microbial community when using molecular techniques [1]. Timing of collection of soils is important to assure a diverse microbial community for sampling. Seasonal variability may be low in undisturbed soils of forests and grasslands where most soil properties such as carbon and nutrient contents remain relatively constant [2]. However soils in agricultural fields subjected to

Walter Gassmann (ed.), Plant Innate Immunity: Methods and Protocols, Methods in Molecular Biology, vol. 1991, https://doi.org/10.1007/978-1-4939-9458-8_21, © Springer Science+Business Media, LLC, part of Springer Nature 2019

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tillage, fertilizer amendments, and manure additions through application or livestock grazing will vary widely in diversity after these disturbances. A way to overcome temporal variation in soil microbial structure in temperate regions is to sample in early to mid-spring or late fall to avoid recent tillage or additions of organic matter or fertilizers [2]. Sampling at these times may also avoid moisture and temperature extremes that often occur during the summer growing season. A review of research objectives will narrow the number of sampling options at the collection site. For example, a survey for sites to assemble a collection of bacteria with impacts on growth of a specific crop plant necessitates the identification of production fields in which the crop is currently or was previously grown. If the objective is to assemble a set of microbial accessions from several ecosystems to screen growth effects on one or more plants, a survey is still recommended to relate soil properties to differences in effects of similar microorganisms obtained from different sites. The number of collection sites can be estimated if the type of crop and soil management in agricultural fields or if vegetation differences in natural ecosystems are of concern. If effects of management such as tillage or no-tillage, use of certain crop varieties (i.e., transgenic or non-transgenic), and conventional or organic production practices are presumed to affect activity of growth-promoting (or -suppressive) bacteria, a search of the crops produced under these conditions within a designated area will be required before setting out on a collection trip. It is also important to decide whether “bulk” (root-free) or rhizosphere (including segments of plant roots) or both types of samples will be collected. Landscape or topographic variability at the field sites also need to be considered because specific soil types routinely occupy different landscape positions (Fig. 1). It is recommended to consult a soil survey for the site(s) selected for sampling. Soil surveys contain soils data for individual counties, parishes, or other land resource areas of the United States and territories and most are available online from U.S.D.A., Natural Resources Conservation Service (https://www. nrcs.usda.gov/wps/portal/nrcs/site/soils/home/). Unless sampling at state or federal experiment field sites, agronomic production fields are private property and it is always recommended to gain permission before entering these sites to collect soils. Similar approaches should be followed if samples are collected in other ecosystems including grasslands, forests, and wetlands. Once on-site it is important to conduct a visual survey of the area that may involve a brief walk-through within and around the field or site. The extent of the survey may depend on the topography of the site. If the field is relatively level, or “flat,” then sampling may be relatively easy assuming the soils are generally uniform within the landscape. However, if the landscape consists of a number of sloping areas, a decision must be made which sites best

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Fig. 1 The major landscape positions used to describe zones within the topography [11]. From Brady, Nyle C., Weil, Raymond R., The Nature and Property of Soils, 14th Ed., ©2008. Reprinted by permission of Pearson Education, Inc., New York, New York

represent the dominant soils for sampling or if a “zone sampling” approach should be followed where landscape positions are defined as sampling zones. Soil properties will differ within the landscape position and, along with soil management and season, influence microbial community structure [3]. For example, the bacterial community diversity in maize (Zea mays) differs significantly in soils of contrasting textures even though the soils are within the same field [4]. A quick field inspection involving removal of soil cores with a soil probe to a depth of 5–10 cm and checking for any obvious changes in color or texture should reveal any sites that differ in soil type, from which separate samples should be collected. Also notes should be taken to describe the site and general soil properties as these will allow informative interpretation of contrasting plant-associated microbial data collected from various field sites. Gathering overall field and basic soils information will then allow development of a site-specific sampling plan. For initial collection of soil microorganisms with plant growth effects, a modest sample plan can be developed that will yield a representative soil sample for each soil type and management system present at the field site. Assuming two or more soil types are present at the site, zone sampling along landscape positions can be followed. Within each soil type-landscape position, a minimum of three soil cores should be collected at each sampling point, which are combined to provide a composite sample for that sampling site. Such composite sampling provides a sample with the least variability in soil properties including microbial characteristics [2]. This

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Fig. 2 Hypothetical soil profile illustrating the major horizons that may be present in a typical soil in temperatehumid regions [11]. From Brady, Nyle C., Weil, Raymond R., The Nature and Property of Soils, 14th Ed., ©2008. Reprinted by permission of Pearson Education, Inc., New York, New York

sampling approach can be repeated within the sampling zone to collect as many “replicate” microbial cultures as the project deems necessary. Samples of the surface soil, that is, the upper 5–10 cm of the soil profile, will yield the most abundant and diverse microbial communities. The exact sampling depth can vary at each site within each soil type due to topsoil loss from cultivation and can be detected using the soil probe and examining the soil cores as indicated above. Soils from undisturbed sites may have a distinct organic or “O” horizon of accumulated organic matter over the surface or “A” horizon that should be sampled separately. The differences in horizonation or soil layers of a hypothetical soil profile are detailed in Fig. 2. Generally accessions of plantassociated microorganisms are isolated from the surface A horizon; however, isolates from the organic layers such as litter on the forest floor or the thatch of turf sites and from subsoil depths (20–60 cm) may also be obtained depending on the objectives of the particular

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project. At greater depths nutrients and carbon substrates are lower and suitable for more oligotrophic microorganisms that consistently function in these environments. Roots of intact plants are sampled to obtain rhizosphere soil if isolation of rhizobacteria is desired. With these general guidelines in mind, a step-by-step method is presented to obtain a collection of soil samples representative of a chosen site for soil microbiome analyses. For more detail on landscape positions and landforms, characterization of horizons within a soil profile, and descriptions for soil sample handling and sampling techniques, consult Field Book for Describing and Sampling Soils [5]. The field book is available from the USDA-NRCS website.

2

Materials 1. County soil survey maps (showing soil series and landscape features at field site). 2. Clinometer or Abney level for measuring landscape slope. 3. Tape measure or measuring wheel. 4. Field notebook. 5. Digital camera. 6. Wire stake flags. 7. GPS device or good GPS application on mobile phone. 8. Soil probe (or tube sampler), stainless steel with depth marked on sample collection end (see Note 1). 9. Ethanol, 70%, in spray bottle. 10. Paper towels. 11. Pail, plastic, no galvanized metal. 12. Sieve, 4–6 mm mesh (see Note 2). 13. Sample bags, plastic or wax-lined paper, quart-size. 14. Permanent-ink marking pen. 15. Cooler or ice chest. 16. Bagged ice or ice packs. 17. Shovel. 18. Clippers for removing roots or root parts. 19. Camel hair or artist’s brush.

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Methods Whether the isolation of microorganisms with plant growth effects will be from bulk (“root-free”) soil, rhizosphere soil, or plant roots, the tools and soil survey information are similar and will be required regardless of the intended source of isolation. Often microbial isolation from one or more sources may be planned in advance or may be decided after surveying the field site, thus assembling materials necessary for all possible sample types will aid in an efficient collection trip.

3.1

Soil Sampling

1. After arrival at the site, make a visual survey for possible landscape differences that may indicate different soil types. Consult soil survey maps from NRCS or access them via internet in the field to help in delineating differences in topography and where soil series are mapped. 2. Estimate slope roughly with a visual assessment or use a clinometer, which reads slopes 90 (Fig. 3). Record observations in the field notebook including general landscape features and slope measurements, vegetation, type of management (especially important if sampling in a crop production field), topsoil depth, and texture. 3. Photograph the sampling site with a good digital camera to provide a good supplemental record of the source of the microbial accessions. 4. For each landscape position (Fig. 1), lay out a transect across the slope using a tape measure or measuring wheel, the length of which will vary depending on number of soil samples to be collected for microbial isolation (Fig. 4). For two or more samples, select intervals of 10–20 m between sampling points to provide representative soils for isolation (see Note 3). 5. Mark the length of the transect at both ends using steel wire stake flags. Flag intervals of 10–20 m to determine the location for each replicate soil sample to be collected and take GPS coordinates for each designated sampling point with an appropriate device or cell phone application and record them. 6. At each sample point remove any accumulated litter or the “O” horizon (Fig. 2) and either set aside to replace after extracting the soil core or retain as a separate sample for microbial isolation (see Note 4) 7. At each flagged sample point establish three subsample points in a triangular arrangement within a 3-m radius of the point [6]. 8. Before sampling and between each sampling point, surface sterilize the soil probe sampling tube by spraying with 70% ethanol and allow to evaporate or wipe dry with a paper towel.

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Fig. 3 A clinometer is an instrument used for rapid measurement of slopes across the landscape positions. The instrument shown is a standard compass with a built-in clinometer available at most outdoor equipment suppliers. On the instrument, turn the compass dial to show the cardinal points as shown. Hold the compass at eye level with arms stretched out in front so that the clinometer needle, (“C”), is pointing vertically at 0 in the scale in the bottom of the compass housing. While standing in the field parallel with the landscape slope, tilt the bottom of the compass to align with the slope of the terrain. Read the angle of slope at the point of the clinometer needle. The scale in this example shows 8% slope, the degree of deviation of the landscape surface from horizontal. This is considered a “gentle slope” and may be associated with shoulder or backslope landscape features [5]. Photo by R.J. Kremer

Fig. 4 A hypothetical sampling area with landscape positions delineated based on slope angle or degree. Transects (“A” and “B”) are depicted as straight lines perpendicular to the slope across landscape positions. Sample points are shown at regular intervals along each transect. (15-day experiments use 1 plant per pot. Suggested size of pots: 8.3  8.3  7 cm square-shaped pots. 4. Inoculate control plants with boiled cultures (uninoculated), containing the same inoculum boiled for 15 min at 100  C. 5. Place the pots with inoculated seedling in a covered tray in the greenhouse at 30  C with a 16-h light/8-h dark cycle. 6. Water the pots twice a week with 50 mL of one-tenth strength Hoagland’s solution and with water as needed. Supplement with potassium nitrate (0.5–5 mM KNO3) depending upon the treatment.

3.3.6 Plant Growth Promotion Assay

1. Five to 30 days after inoculation, or when growth promotion is observed, determine the plants-growth parameters. 2. Briefly wash the roots in buffered saline and measure the plant parameters using the WinRHIZO pro software (see Note 4). Measure at least 10 plants per experiment. 3. Assess the phytostimulatory effect of inoculation compared to inoculation with bacterial suspension boiled for 15 min. For plants cultivated in pots several growth parameters can be measured: plant height, inflorescence length, number of seeds, shoot and root fresh and dry weight (see Note 5) (dry for 72 h at 60  C). 4. For statistical analyses use the Wilcoxon test. 5. Record images of the plants.

4

Notes 1. Turface is important to prevent vermiculite adhering to the roots, which facilitates sampling and measurements. Turface also maintains better moisture conditions. Quartz sand can be used alternatively. 2. Soil samples must be freshly collected and well homogenized before a subsample is used for bacterial isolation. If necessary, use

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refrigerated (4  C) transport between the collection point and the laboratory. Avoid using broken or damaged roots for the procedure; the surface disinfection could jeopardize recovery of the endophytic population. The diversity of bacteria isolated is proportional to the diversity of media used, including selection of spore-forming bacteria as described by Beneduzi et al. [6] . We found it useful to transfer fast-growing colonies from isolation plates onto 150  15 mm plates in a 96-well plate grid format. 3. Some bacteria do not survive centrifugation, so do not over centrifuge. 4. If WinRhizo equipment is not available you can take images of the plants and measure parameters using free software such as EZ-Rhizo (http://www.psrg.org.uk/plant-biometrics.html). 5. Root and shoot dry weight are also very useful parameters to assess plant growth promotion. References 1. Pankievicz VCS, do Amaral FP et al (2015) Robust biological nitrogen fixation in a model grass-bacterial association. Plant J 81:907–919. https://doi.org/10.1111/tpj. 12777 2. Baldani I, Reis VM et al (2014) The art of isolating nitrogen-fixing bacteria from non-leguminous plants using N-free semisolid media: a practical guide for microbiologists. Plant and Soil 384:413–431. https://doi. org/10.1007/s11104-014-2186-6 3. Souza EM, Pedrosa FO (2015) Inorganic Nitrogen Metabolism in Azospirillum spp. In: Cassa´n FD, Okon Y, Creus CM (eds) Handbook for Azospirillum, Technical Issues and Protocols. Springer International Publishing Switzerland, Springer 4. Machado HB, Funayama S et al (1991) Excretion of ammonium by Azospirillum brasilense mutants resistant to ethylenediamine. Can J Microbiol 37:549–555 5. Hoagland DR, Arnon DI (1950) The waterculture method for growing plants without soil. Berkeley, California 6. Beneduzi A, Peres D, da Costa PB (2008) Genetic and phenotypic diversity of plant-

growth-promoting bacilli isolated from wheat fields in southern Brazil. Res Microbiol 159:244–250. https://doi.org/10.1016/j. resmic.2008.03.003 7. Magalha˜es Cruz L, Souza EM et al (2001) 16S ribosomal DNA characterization of nitrogenfixing bacteria isolated from banana (Musa spp.) and pineapple (Ananas comosus (L.) Merril). Appl Environ Microbiol 67:2375–2379. https://doi.org/10.1128/AEM.67.5.23752379.2001 8. Altschul SF, Gish W et al (1990) Basic local alignment search tool. J Mol Biol 215:403–410. https://doi.org/10.1016/ S0022-2836(05)80360-2 9. Balsanelli E, Tuleski TR, Baura VA et al (2013) Maize root lectins mediate the interaction with Herbaspirillum seropedicae via N-acetyl glucosamine residues of lipopolysaccharides. PLoS One 8:e77001. https://doi.org/10.1371/ journal.pone.0077001 10. Martins PK, Nakayama TJ, Ribeiro AP et al (2015) Floral-dip: a simple and rapid. Biotechnol Rep (Amst) 6:61–63. https://doi.org/10. 1016/j.btre.2015.02.006

INDEX A

C

Abscission ........................................................ vi, 127–138 Abscission zone (AZ)................. 128–131, 133, 135, 138 Accelerometer...................................................... 144, 145, 148, 149, 152–155 Acetosyringone.................................................. 15, 37–39, 64, 65, 71, 212 Acetylation......................................... v, 13, 14, 20, 23–29 Agrobacterium tumefaciens................................. 9, 11, 15, 17, 20, 33, 36, 63, 71, 73, 212 Alternaria solani ......................................... 62, 63, 65, 66 Amplification, DNA ........................................................ 85 Amplifier .............................................................. 142, 144, 148, 149, 151, 155 Amplitude ...........................................143, 145, 147–149, 151, 153–155, 230 Antibody ..................................................... 14, 16–20, 25, 56–59, 62, 72, 74, 118, 204 Aphanomyces euteiches ..................................................... 70 Apoplast ..........................................................44, 115, 160 Apple fruitlet ........................................................ 187–197 Arabidopsis thaliana .........................................11, 62, 70, 101–105, 147, 159–183, 188 Aspartic acid .................................................................. 116 Assay, infection ..................................................... 159–183 Avirulence ...................................................................... 200 Azospirillum spp................................................... 248, 254

Calibration.....................70, 77, 145, 148, 152, 154, 155 Callose ........................................................................... 199 Carbohydrate.............................................................57, 88 Cauliflower mosaic virus (CaMV).......................... 34, 35, 42, 208, 213 Caulimoviridae ..................................................... 207, 208 Cell cycle................................................................. 92, 101 Cell wall ..................................................v, 24, 55–59, 160 Chromatin compaction ..........................................................91, 92 euchromatin .............................................................. 92 heterochromatin........................................................ 92 Citric acid ...................................................................... 116 Clinometer............................................................ 241–243 Coenzyme A (CoA) .....................................13, 23, 25–29 Communication, vibrational ........................................ 141 Community, microbial ........................................ 237, 239, 240, 244, 247 Compound, bioactive .........................116, 118, 120–122 Coomassie..................................................................26, 27 Cotyledon .............................................................. 62, 101, 102, 104, 105

B Bacterium diazotroph ...................................................... 247, 248 endophytic ...................................................... 247–256 epiphytic ......................................................... 247–256 growth-promoting .................................... vi, 238, 247 growth-suppressive ................................................. 238 β-Glucuronidase (GUS) ...................................... 214, 216, 217, 224–226, 230, 231 Bioinformatics ......................................................... v, 1–11 Biological nitrogen fixation (BNF) ..................... 247, 248 Bioluminescence.................................................... 44, 161, 172, 173, 178, 182 Biotremology................................................................. 141 BLAST searches................................................................. 3 Brassicaceae........................................................... 159, 160

D Darkness ...................................................... 108, 111, 113 Dexamethasone (DEX)........................................ 199, 201 40 ,6-Diamidino-2-phenylindole (DAPI) ....................... 64 Digestion ............................................................. 9, 81, 84, 86–89, 92, 93, 95–97, 220 Disease resistance .............................................1, 101, 189 Dot blot .....................................................................56–59

E Ecosystem ............................................................. 237, 238 Effectors.......................................................v, vi, 1–11, 14, 24, 25, 27, 28, 35, 70, 108, 116, 160 Electrophoresis ...................................................... 4, 6, 20, 26, 74, 89, 94, 210, 211, 220, 233 Electrophoretic mobility shift assay (EMSA) ............... 69, 224, 227, 228, 231, 233 Endonuclease, micrococcal......................... 92, 93, 95, 96 Enhanced disease susceptibility 1 (EDS1)................... 108 Epidermal peel.................................................... 62, 64–67

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AND

PROTOCOLS

Epigenetics ................................................................79, 80 Epithem cell.......................................................... 160, 170 Erwinia amylovora ......................................... vi, 187–189, 192, 193, 196, 197 Estradiol (Est) ............................................................... 199 Exopolysaccharides ....................................................... 187 Expression, inducible ........................................... 199, 200 Extracellular ATP (adenosine triphosphate)............43–52 Exudates ..............................................116, 117, 119–124

F Field, agricultural ................................................. 237, 238 Filter, digital .................................................................. 153 Filtering ......................................3, 9, 105, 144, 151, 154 Fire blight ............................................................. 187–197 Fixation ...................................................... 72, 74, 75, 248 Flow cytometry .................................................... 101–105 Fluorescence lifetime imaging microscopy (FLIM)...........................................................69–77 Fluorescence resonance energy transfer (FRET)...........................................................69–77 Formaldehyde cross-linking .....................................92, 94 Fungicides.................................................... 189, 191, 192 Fungus ..................................................... 63, 66, 129, 180

G Galacturonic acid.......................................................56, 58 Gene expression ................................ 14, 24, 69, 92, 116, 117, 121, 122, 199, 200, 207–221, 224, 229–231 Gene silencing ........................................... 33, 34, 38, 204 Genome sequencing ......................................................... 2 Golden Gate ....................................................... 2–6, 8, 11 G protein-coupled receptor............................................ 44 Gram-negative ............................................. 115, 159, 178 Green fluorescent protein (GFP) ..................... 17, 34–37, 39–42, 63, 65–68, 70, 71, 73, 74, 76, 117, 118, 121, 214, 224 Growth, bacterial ....................................... 109, 116, 187, 189, 192, 194, 197, 226 Growth chamber ...............................................37, 81, 83, 92, 94, 108, 111, 128, 130, 131, 137, 163, 168, 169, 191, 193, 201, 250, 252, 254 Guttation ............................................................. 159, 161, 164, 169, 170, 181, 182

H HAESA (HAE).............................................................. 138 Herbaspirillum spp. ...................................................... 254 Herbivores ............................................................ 141–156 Histone .............................. 14, 20, 23, 24, 79, 80, 91, 92 Homogenates ...................................................... 102, 103, 135, 195–197, 199–206, 213, 251 Hydathodes .......................................................... 159–183

Hydroponic ................................................................... 255 4-Hydroxybenzoic acid................................................. 116

I Immunity, effector-triggered (ETI) ..............24, 108, 160 Immunity, PAMP-triggered (PTI)............................... 160 Immunoprecipitation......................................v, 13–20, 80 Infiltration ......................................................... 17–19, 41, 64, 66, 73, 108, 133, 212, 217, 221 Insect .......................................................vi, 128, 141–156 Isoschizomer ................................................................... 87

K Kinase........................................14, 25, 43, 212, 219, 233

L Landscape ............................................238, 239, 241–244 Leaf, cauline.......................................................... 127–138 Leaf discs .................................... 18, 19, 73–76, 175–177 Library ......................................................... 2, 4, 6, 11, 81 Ligand-gated ion channel............................................... 44 Light ............................................................... vi, 5, 44, 49, 63, 65, 73, 83, 94, 107–113, 124, 135, 138, 160, 163, 170, 171, 173, 180, 196, 201, 212, 231, 250, 252 blue ........................................................ 108, 109, 111 red .......................................................... 109, 111, 113 white .......................................................................... 37 Liquid nitrogen .................................................18, 19, 74, 81, 83, 93, 94, 120, 122, 129, 131, 135, 138, 163, 167, 179, 217, 229, 231 Localization, subcellular ......................................... 11, 23, 61, 62, 74 Luciferase (LUC) ..............................................44, 46, 48, 49, 52, 178, 214, 224, 226, 231 Luciferin .................................................................. 45, 46, 48, 49, 52

M mapk phosphatase 1 (mkp1)........................................... 116 Mating, triparental ....................... 11, 162, 165, 177, 178 Mechanosensing ............................................................ 141 Media, minimal ........................................... 116, 119, 121 Membrane cellular........................................................................ 61 nitrocellulose ............................ 16, 20, 57, 59, 72, 74 protein blot.............................................................. 204 Metabolites ...................................................... vi, 115–126 Methylation, DNA ................................................ v, 79–89 Methylcytosine ..........................................................79, 80 Methylesterase ................................................................. 55 Microbiome .............................................................. vi, 241 Micronutrient ....................................................... 249, 250

PLANT INNATE IMMUNITY: METHODS Microorganism ...............................................45, 237–245 Microscopy confocal...................................................33–42, 67, 74 fluorescence ............................................................... 65 Mycelia.......................................................................66, 68

N Nicotiana benthamiana .......................................... 14, 15, 17, 33–37, 39, 40, 71, 73 Nicotiana tabacum ..................................... 211, 212, 216 Nucleosome...............................................................91–99 Nucleus .............................................................. 61, 66–68, 74, 77, 101

O Open reading frame (ORF).......................................... 207 Optical density (OD) .............................................. 17, 58, 59, 73, 109, 111, 121, 169, 172 Organ, floral .................................................................. 128 Oryza sativa.........................................81, 83, 92, 94, 249 Oscilloscope.........................................143, 144, 147, 152

P Pararetroviruses .................................................... 207, 213 Pathogen-associated molecular pattern (PAMP) .............................................................. 160 Pectin .........................................................................55–59 Peduncles ..................................................... 191, 192, 196 Pellet, DNA ................................................. 84, 88, 95, 97 Permeabilization................................................. 70, 74, 75 Photoreceptor ............................................................... 107 Photosynthesis............................................................... 107 Plant, graminaceous ...................................................... 248 Plant immunity.............................................................. v, vi Playback ............................................... 142–150, 152–156 Ploidy ............................................................... vi, 101–105 Polymutant .................................................................... 178 Posttranslational modifications (PTMs) ........... 13, 14, 91 Precipitation ............................................................ 16, 84, 85, 88, 99, 135, 214, 219 Promoter ............................................. vi, 6, 8, 34, 36, 80, 92, 122, 207–221, 223–235 Propidium iodide (PI) .................................................. 102 Protein DNA-binding ............................................................ 69 recombinant ....................................28, 226, 232, 233 Protoplasts .................................................. 210, 211, 215, 216, 221, 228, 230 Pseudomonas syringae ............................................. v, 1–11, 24, 27, 107–113, 115–126, 128, 130, 132, 161, 178, 199 Pulp .............................................................. 194, 195, 197

AND

PROTOCOLS Index 259

R Radiolabeling..................................................25, 219, 233 Ralstonia solanacearum ..................................... 14, 25, 70 Reactive oxygen species (ROS) ........................... 107, 199 Receptor ...........................................................14, 44, 141 Receptor-like kinase ........................................................ 44 Reporter gene..................................................34, 39, 178, 208, 210, 214, 224, 226, 228, 230, 231 RESISTANT to PSEUDOMONAS SYRINGAE 6 (RPS6) ............................................................ 108 Restriction enzyme, methylaton-sensitive .................... 81, 83, 86, 89 Rhizosphere......................................................... 238, 241, 242, 244, 245, 248, 251 RNA, double-stranded (dsRNA) ..................33, 105, 213 Root pressure ................................................................ 159 Rootstock ...................................................................... 187

S Sampling, soil ...................................................... 239, 241, 242, 244, 245, 251, 255 Screen, high-throughput .............................................. 192 Seedling .............................................................83, 86, 94, 99, 108, 116–121, 123, 124, 168, 188, 200, 202–205, 252, 254, 255 Sequencing, bisulfite ....................................................... 80 Setaria viridis .............................................. 248, 250, 254 Signaling ...............................................................v, 14, 16, 20, 24, 25, 44, 107, 116 Silencing suppressor ........................................................ v, 33–42 transcriptional............................................................ 79 Soil .................................................vi, 102, 108, 128–130, 135, 137, 163, 166, 180–182, 201–206, 237–245 Soil profile ............................................................ 240, 241 Solanum lycopersicum ................................................63, 67 Sonication ...................................62, 65, 66, 68, 122, 235 Sorghum bicolor.............................................................. 249 Spores..............................................................66, 251, 256 Staining, histochemical ................................................. 217 Stimulus, acoustic ......................................................... 141 Stomata .......................................115, 131, 132, 159, 160 Survey .................................... 26, 29, 238, 241, 242, 244 Suspension cell ..........................................................43–52 Syringe ...............................................................17, 38, 63, 73–75, 108, 116, 118, 119, 123, 129, 131, 132, 160, 202, 217, 220, 226 Sytox® Orange .............................................70–73, 76, 77

T Thigmomorphogenesis ................................................. 142 Ti plasmid ........................................................................ 63

PLANT INNATE IMMUNITY: METHODS

260 Index

AND

PROTOCOLS

Touch.............................................................v, vi, 52, 133, 137, 142, 150, 168, 176 Transcription factors (TFs)..................................v, 14, 24, 25, 61–68, 70, 98, 138, 208, 223 Transcription start site (TSS) ............................... 92, 212, 213, 218–220 Transformation high-efficiency ......................................................... 200 stable ........................................................................ 228 transient ......................................................62, 64, 228 Transpiration ................................................................. 159 Transposons....................................................79, 178, 179 Triticum aestivum ......................................................... 249 Type III secretion .......................... 6, 115–126, 160, 180

Viral protein P6 ......................................................... 34, 35, 42, 105 P19............................................ 20, 34, 35, 42, 73, 76 Virulence........................................................2, 14, 23–29, 116, 121, 122, 188, 189, 199, 200 Virus................................................................... 20, 33–35, 39, 41, 76, 209, 213, 220

W Wave, mechanical .......................................................... 141

X

Uronic acid ................................................................56, 58

Xanthomonas axonopodis pv. manihotis (Xam)................................................................. 161 Xanthomonas campestris pv. campestris .......... vi, 159–183 Xanthomonas euvesicatoria ............................................. 24 Xylem ...................................................159, 161, 170, 172

V

Z

U

Vasculature............................................................ 159–161 Vibration feeding ..................................................................... 142 leaf .............................................................. vi, 141–156

Zea mays................................................................ 239, 249