Plant Signal Transduction. Methods and Protocols

Plant Signal Transduction

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

For other titles published in this series, go to www.springer.com/series/7651

METHODS

IN

MOLECULAR BIOLOGY™

Plant Signal Transduction Methods and Protocols

Edited by

Thomas Pfannschmidt Institut für Allgemeine Botanik und Pflanzenphysiologie, Lehrstuhl Pflanzenphysiologie, Friedrich-Schiller-Universitaet Jena, Jena, Germany

Editor Dr. Thomas Pfannschmidt Institut für Allgemeine Botanik und Pflanzenphysiologie Lehrstuhl Pflanzenphysiologie Friedrich-Schiller-Universität Jena Jena, Germany

ISBN: 978-1-58829-943-7 e-ISBN: 978-1-59745-289-2 ISSN: 1064-3745 e-ISSN: 1940-6029 DOI: 10.1007/978-1-59745-289-2 Library of Congress Control Number: 2008938154 © Humana Press, a part of Springer Science+Business Media, LLC 2009 All rights reserved. This work may not be translated or copied in whole or in part without the written permission of the publisher (233 Spring Street, New York, NY 10013, USA), except for brief excerpts in connection with reviews or scholarly analysis. Use in connection with any form of information storage and retrieval, electronic adaptation, computer software, or by similar or dissimilar methodology now known or hereafter developed is forbidden. The use in this publication of trade names, trademarks, service marks, and similar terms, even if they are not identified as such, is not to be taken as an expression of opinion as to whether or not they are subject to proprietary rights. While the advice and information in this book are believed to be true and accurate at the date of going to press, neither the authors nor the editors nor the publisher can accept any legal responsibility for any errors or omissions that may be made. The publisher makes no warranty, express or implied, with respect to the material contained herein. Cover illustration: Derived from Figure 2 in Chapter 7 Printed on acid-free paper 9 8 7 6 5 4 3 2 1 springer.com

Preface Plants are sessile organisms that cannot escape from the conditions in their direct environment and thus have to deal with all biotic and abiotic factors in it. During evolution, plants developed a sophisticated network of responses which are controlled at the molecular level and which enable plants to develop, survive and propagate under a wide range of conditions. A number of receptors or receptor systems were identified in recent years which detect e.g. light, hormones or pathogens and which convert the respective stimulus into an appropriate signal. This, in turn, is transduced via a signaling cascade or network which specifically activates or represses a respective response. Plant physiologists, biochemists, geneticists and molecular biologists from academic institutions as well as industry work worldwide on this emerging field of research to analyze and understand how plants sense the environment and how they manage it to respond to it properly. Such knowledge is of immense interest for basic research in academia, agriculture or ecosystem research and will contribute to the solution of many problems of a growing mankind. This volume is aimed to give plant scientists of all major directions well-established methods on hand which they can easily use to analyze questions or problems in plant signal transduction on the molecular level. Special emphasis is given to specific problems occurring when working with plants. The experimental procedures are proven protocols known to work on well-characterized model organisms. They will be a help either to establish a particular technique in the lab or to develop respective modified protocols for other plants. It is impossible to cover all technical aspects in current plant signal transduction research; therefore, the volume focuses on two major aspects of wide importance: in planta analyses and proteins involved in signal transduction. The volume starts with an overview chapter by Jorge Casal who summarizes important developments and describes the multiplicity of players and interactions in plant signaling networks. This includes also topics which could not be covered methodologically and thus helps the interested reader to find out more about specific questions for instance on RNA biology. The next two chapters were contributed by Cordelia Bolle who describes a great number of protocols for phenotyping of mutants to understand possible influences on light and hormone signaling as well as on developmental responses. These are followed by a chapter on high-through put RT-PCR written by Mario Klatte and Petra Bauer useful for a number of questions including genotyping of mutants. Axel Mithöfer and colleagues then describe a method how to measure cellular changes in the important second messenger Ca2+. Detection of changes in the cellular redox state using a redox-sensitive green fluorescent protein is then described by Andreas Meyer and Thorsten Brach. Related to this Neil Baker and colleagues provide a protocol how to measure the accumulation of various reactive oxygen species within plant tissues. The chapter written by Toru Hisabori and colleagues introduces the part of protein analyses and describes the detection and purification of thioredoxin target proteins. It is followed by a chapter of Julia Vainonen and colleagues who contribute a protocol on mass spectrometry and analysis of an important post-translational modification protein phosphorylation. Claus Schwechheimer and colleagues then give a comprehensive overview on the field of protein degradation providing an extensive literature account on this fast developing topic. Volker Wagner and Maria Mittag give a protocol how to observe and quantify circadian rhythmicity of proteins. This is followed by three chapters on protein interactions. Christina Chaban and colleagues give a detailed protocol for protein v

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interaction via molecular analysis by means of bimolecular fluorescence complementation. This is complemented by a chapter by Kotaro Yamamoto and colleagues describing fluorescence cross correlation spectroscopy of plant proteins. Finally, Klaus Harter and colleagues provide a protocol on protein interaction detection via the mating based splitubiquitin-system. Analysis of plant kinases via yeast complementation assays is described by Markus Teige and colleagues. This chapter is complemented by a description of in vivo characterization of plant phosphatases by Irute Meskiene and colleagues. The last five chapters are focused on protein-nucleotide interactions. Benjamin Fode and Christiane Gatz start with a protocol on chromatin immunoprecipitation to study transcription factors. This is complemented by a chapter on purification of DNA-binding proteins and characterization of their properties by fluorescence based electrophoretic mobility assays written by Sebastian Steiner and Thomas Pfannschmidt. Dierk Wanke and Klaus Harter then provide a protocol on the analysis of plant promoter sequences via the yeast one-hybrid system. In a second chapter Wanke and colleagues also provide an extensive overview of how to analyse promoters by bioinformatics means. In the final chapter then Jan Schöning and Dorothee Staiger describe methods to analyse RNA-protein interactions. These chapters cover a great extent of methods useful to test many basic questions in plant signaling. I hope that many researchers will find them helpful for their own specific questions and for improvement of their experimental approaches. Finally, I would like to thank all authors for their willingness to share their protocols and their time to summarize all the little details necessary to perform the described experiments in a proper and successful way. Without their commitment, the volume would never have been made possible. I am also very grateful to John Walker, the series editor, for all his useful hints and his help and the editorial office of Methods in Molecular Biology of Humana Press for the help in editing and formatting this book. Thomas Pfannschmidt

Contents Preface. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Contributors. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1 Multiple Dimensions in Plant Signal Transduction: An Overview . . . . . . . . . . Jorge J. Casal 2 Phenotyping of Arabidopsis Mutants for Developmental Effects of Gene Deletions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Cordelia Bolle 3 Phenotyping of Abiotic Responses and Hormone Treatments in Arabidopsis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Cordelia Bolle 4 Accurate Real-time Reverse Transcription Quantitative PCR . . . . . . . . . . . . . . Marco Klatte and Petra Bauer 5 Probing Spatio-temporal Intracellular Calcium Variations in Plants . . . . . . . . . Axel Mithöfer, Christian Mazars, and Massimo E. Maffei 6 Dynamic Redox Measurements with Redox-Sensitive GFP in Plants by Confocal Laser Scanning Microscopy . . . . . . . . . . . . . . . . . Andreas J. Meyer and Thorsten Brach 7 Imaging of Reactive Oxygen Species in vivo . . . . . . . . . . . . . . . . . . . . . . . . . . . . Steven M. Driever, Michael J. Fryer, Philip M. Mullineaux, and Neil R. Baker 8 Identification of Thioredoxin Targeted Proteins Using Thioredoxin Single Cysteine Mutant-immobilized Resin . . . . . . . . . . . Ken Motohashi, Patrick G. N. Romano, and Toru Hisabori 9 Determination of in vivo Protein Phosphorylation in Photosynthetic Membranes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Julia P. Vainonen, Alexander V. Vener, and Eva-Mari Aro 10 Examining Protein Stability and Its Relevance for Plant Growth and Development . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Claus Schwechheimer, Björn C. Willige, Melina Zourelidou, and Esther M. N. Dohmann 11 Probing Circadian Rhythms in Chlamydomonas rheinhardtii by Functional Proteomics. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Volker Wagner and Maria Mittag 12 Bimolecular Fluorescence Complementation (BiFC) to Study Protein-protein Interactions in Living Plant Cells . . . . . . . . . Katia Schütze, Klaus Harter, and Christina Chaban 13 Fluorescence Cross-Correlation Spectroscopy of Plant Proteins . . . . . . . . . . . Hideki Muto, Masataka Kinjo, and Kotaro T. Yamamoto

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The Determination of Protein-protein Interactions by the Mating-based Split-ubiquitin-system (mbSUS) . . . . . . . . . . . . . . . . . . Christopher Grefen, Petr Obrdlik, and Klaus Harter 15 Functional Complementation of Yeast Mutants to Study Plant Signalling Pathways. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Norbert Mehlmer, Elisabeth Scheikl-Pourkhalil, and Markus Teige 16 Phosphatase Activities Analyzed by in vivo Expressions. . . . . . . . . . . . . . . . . . Alois Schweighofer, Zahra Ayatollahi, and Irute Meskiene 17. Chromatin Immunoprecipitation Experiments to Investigate in vivo Binding of Arabidopsis Transcription Factors to Target Sequences. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Benjamin Fode and Christiane Gatz 18 Fluorescence-based Electrophoretic Mobility Shift Assay in the Analysis of DNA-binding Proteins . . . . . . . . . . . . . . . . . . . . . . . . . . . . Sebastian Steiner and Thomas Pfannschmidt 19 Analysis of Plant Regulatory DNA sequences by the Yeast-One-Hybrid Assay . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Dierk Wanke and Klaus Harter 20 Analysis of Plant Regulatory DNA Sequences by Transient Protoplast Assays and Computer Aided Sequence Evaluation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Kenneth W. Berendzen, Klaus Harter, and Dierk Wanke 21 RNA-protein Interaction Mediating Post-transcriptional Regulation in the Circadian System . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Jan C. Schöning and Dorothee Staiger Index . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Contributors EVA-MARI ARO • Plant Physiology and Molecular Biology, Department of Biology, University of Turku, Turku, Finland ZAHRA AYATOLLAHI • Max F. Perutz Laboratories, University of Vienna, Vienna, Austria NEIL R. BAKER • Department of Biological Sciences, University of Essex, Colchester, Essex, UK PETRA BAUER • Department of Biosciences - Botany, Saarland University, Saarbrücken, Germany KENNETH W. BERENDZEN • ZMBP Pflanzenphysiologie, Universität Tübingen, Tübingen, Germany CORDELIA BOLLE • Ludwig-Maximilians-Universität, Department für Biologie I, Lehrstuhl für Botanik, München, Germany THORSTEN BRACH • Heidelberg Institute of Plant Sciences, University of Heidelberg, Heidelberg, Germany JORGE J. CASAL • IFEVA, Facultad de Agronomía, Universidad de Buenos Aires y CONICET, Buenos Aires, Argentina CHRISTINA CHABAN • ZMBP/Pflanzenphysiologie, Universität Tübingen, Tübingen, Germany ESTHER M. N. DOHMANN • Department of Developmental Genetics, Center for Plant Molecular Biology, Tübingen University, Tübingen, Germany. STEVEN M. DRIEVER • Department of Biological Sciences, University of Essex, Colchester, Essex, UK BENJAMIN FODE • Albrecht-von-Haller-Institut für Pflanzenwissenschaften, Allgemeine und Entwicklungsphysiologie, Göttingen, Germany MICHAEL J. FRYER • Department of Biological Sciences, University of Essex, Colchester, Essex, UK CHRISTIANE GATZ • Albrecht-von-Haller-Institut für Pflanzenwissenschaften, Allgemeine und Entwicklungsphysiologie, Göttingen, Germany CHRISTOPHER GREFEN • ZMBP/Pflanzenphysiologie, Universität Tübingen, Tübingen, Germany KLAUS HARTER • ZMBP/Pflanzenphysiologie, Universität Tübingen, Tübingen, Germany TORU HISABORI • Chemical Resources Laboratory, Tokyo Institute of Technology, Yokohama, Japan MASATAKA KINJO • Laboratory of Supramolecular Biophysics, Research Institute for Electronic Science, Hokkaido University, Sapporo, Japan MARCO KLATTE • Department of Biosciences - Botany, Saarland University, Saarbrücken, Germany MASSIMO E. MAFFEI • Department of Plant Biology and Centre of Excellence CEBIOVEM, University of Turin, Turin, Italy CHRISTIAN MAZARS • UMR CNRS-UPS 5546, Pôle de Biotechnologie Végétale, Castanet-Tolosan, France NORBERT MEHLMER • Department of Biochemistry, Max F. Perutz Laboratories, University of Vienna, Vienna, Austria IRUTE MESKIENE • Max F. Perutz Laboratories, University of Vienna, Vienna, Austria

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ANDREAS J. MEYER • Heidelberg Institute of Plant Sciences, University of Heidelberg, Heidelberg, Germany AXEL MITHÖFER • Max-Planck Institute for Chemical Ecology, Department Bioorganic Chemistry, Jena, Germany MARIA MITTAG • Institut für Allgemeine Botanik, Friedrich-Schiller-Universität Jena, Jena, Germany KEN MOTOHASHI • Chemical Resources Laboratory, Tokyo Institute of Technology, Yokohama, Japan PHILIP M. MULLINEAUX • Department of Biological Sciences, University of Essex, Colchester, Essex, UK HIDEKI MUTO • Laboratory of Supramolecular Biophysics, Research Institute for Electronic Science, Hokkaido University, Sapporo, Japan PETR OBRDLIK • IonGate Biosciences GmbH, Frankfurt, Germany THOMAS PFANNSCHMIDT • Junior Research Group: “Plant acclimation to environmental changes – Protein analysis by MS”, Institut für Allgemeine Botanik und Pflanzenphysiologie, Lehrstuhl Pflanzenphysiologie, Friedrich-Schiller-Universität Jena, Jena, Germany PATRICK G. N. ROMANO • Chemical Resources Laboratory, Tokyo Institute of Technology, Yokohama, Japan ELISABETH SCHEIKL-POURKHALIL • Department of Biochemistry, Max F. Perutz Laboratories, University of Vienna, Vienna, Austria JAN C. SCHÖNING • Department of Molecular Cell Physiology, University of Bielefeld, Bielefeld, Germany KATIA SCHÜTZE • ZMBP/Pflanzenphysiologie, Universität Tübingen, Tübingen, Germany DOROTHEE STAIGER • Department of Molecular Cell Physiology, University of Bielefeld, Bielefeld, Germany CLAUS SCHWECHHEIMER • Department of Developmental Genetics, Center for Plant Molecular Biology, Tübingen University, Tübingen, Germany; Center of Life and Food Sciences Weihenstephan, Technische Universität München, Freising, Germany ALOIS SCHWEIGHOFER • Max F. Perutz Laboratories, University of Vienna, Vienna, Austria SEBASTIAN STEINER • Junior Research Group: “Plant acclimation to environmental changes – Protein analysis by MS”, Institut für Allgemeine Botanik und Pflanzenphysiologie, Lehrstuhl Pflanzenphysiologie, Friedrich-Schiller-Universität Jena, Jena, Germany MARKUS TEIGE • Department of Biochemistry, Max F. Perutz Laboratories, University of Vienna, Vienna, Austria JULIA P. VAINONEN • Plant Physiology and Molecular Biology, Department of Biology, University of Turku, Turku, Finland ALEXANDER V. VENER • Department of Cell Biology, Linköping University, Linköping, Sweden VOLKER WAGNER • Institut für Allgemeine Botanik, Friedrich-Schiller-Universität Jena, Jena, Germany DIERK WANKE • ZMBP Pflanzenphysiologie, Universität Tübingen, Tübingen, Germany BJÖRN C. WILLIGE • Department of Developmental Genetics, Center for Plant Molecular Biology, Tübingen University, Tübingen, Germany; Center of Life and Food Sciences Weihenstephan, Technische Universität München, Freising, Germany KOTARO T. YAMAMOTO • Department of Biological Sciences, Faculty of Science, Hokkaido University, Sapporo, Japan

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MELINA ZOURELIDOU • Department of Developmental Genetics, Center for Plant Molecular Biology, Tübingen University, Tübingen, Germany; Center of Life and Food Sciences Weihenstephan, Technische Universität München, Freising, Germany

Chapter 1 Multiple Dimensions in Plant Signal Transduction: An Overview Jorge J. Casal Abstract Each process involved in the generation of plant body form and function is under the control of signals from the exogenous and/or endogenous plant environment. These controls are necessary for adequate plant adjustment to the prevailing conditions, but at the same time they impose the need for sophisticated mechanisms to achieve adequate sensitivity towards signals and stability against noise. To cope with this challenge plants use multiple signals, multiple receptors even for the same signal and interactive signal transducers with multiple targets. Here we provide an overview of this multiplicity and its functional significance. Key words: Signals, receptors, signalling networks, transcription factors, hormones, regulatory RNA, second messengers, chromatin remodelling.

1. Introduction The study of signal transduction may involve the definition of the signal, the identification of the receptor(s), the downstream signalling transducers (if any) and the end-point targets and the analysis of the molecular interactions among these components. Specific lines of investigation can focus on selective aspects of the aforementioned list (e.g., the identification of the receptor partner(s)). The vision of the apparently simple line defined by the sequence involving the signal, its receptor, the signal transduction elements and the targets has changed significantly in recent years. Multiple dimensions can be found easily at each one of the steps of the aforementioned line. The focus of this

T. Pfannschmidt (ed.), Plant Signal Transduction, vol. 479 Humana Press, a part of Springer Science + Business Media, LLC 2009 DOI: 10.1007/978-1-59745-289-2_1

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introductory chapter is on the description of this multiplicity and its functional significance. Clearly, it is not possible to deal with all the areas of plant signal transduction in this chapter. Numerous recent reviews cover specific issues. We give priority to the areas that have irrupted with overwhelming strength in the recent years; provide examples of the complexity that is emerging and of how this complexity helps in coping with the challenge imposed by the requirements of plant life.

2. A World of Signals 2.1. Definitions

The signal is a feature of the physical and/or chemical environment that (1) carries information about the environment, (2) can be perceived by the organism (or by part of the organism) and (3) can modify aspects of its physiology and/or development. A given feature of the environment is not a signal if the plant is not able to perceive that condition and respond to it. The signal can originate outside or inside the organism that is able to perceive it. Certain aspects of light and temperature environments experienced by the plant provide examples of such exogenous signals. Hormones provide examples of endogenous signals. Signals from the external environment often do not act directly on their targets but are transduced by changes in the status of endogenous signals. Implicit in the concept of signal is the ability of that feature to carry information about the outside or endogenous environment. In this regard, light used by the photochemical reactions of photosynthesis is the source of energy and is not considered a signal for these reactions. When the shoot of a seedling emerges from the soil, it becomes exposed to light that is perceived by specific receptors such as phytochromes and cryptochromes. The latter change in the light environment is a signal because it provides indication that the shoot is no longer fully covered by the soil, and this information triggers profound modifications of plant body shape and function, which favour light capture and photosynthesis (1). This example is a case of a light signal providing information about the light environment but there are cases where the signal involves changes in a given feature of the environment and information about another feature. The seeds of many weeds require alternating temperatures to germinate. The differences between daytime and night-time temperatures decrease with the depth within the soil and with soil cover by a foliage canopy. The seeds that require alternating temperatures can use this requirement to avoid germination if placed at a distance to the soil surface that the growth of its axis is unlikely to cover and/or to avoid germination under the

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shade from neighbours that would reduce the light available for photosynthesis (2). The temperature regime can therefore be a signal that provides information about seed position within the soil or about the presence of competitors. The daily duration of the light period (photoperiod) controls the time of flowering or vegetative storage organ formation. The photoperiod depends on the season and therefore, under specific conditions the photoperiodic requirement can help the plant to avoid flowering under extreme temperatures, low water availability or any other stressful conditions whose occurrence correlates in time with the photoperiod. 2.2. Signals that Target the Same Cells where they Originate

If the signal is endogenous to the plant it can in turn have its origin in the same cell, tissue or organ where the subsequent response takes place or in different and eventually distant cells, tissues or organs. When the photosynthetic tissues are exposed to excess light, several reactive oxygen species can originate in the chloroplast. These species cause damage to membranes and proteins but they can also act as signals that induce the expression of selected components of the defence system against oxidative injury. These signals can act in the regulation of nuclear gene expression in the same cell where they originate (3).

2.3. Signals that Target Nearby Cells

Many signals target cells located close to the cell of their origin. For instance, in the shoot apical meristem of Arabidopsis, the CLAVATA 3 mRNA is expressed in stem cells and the CLAVATA 3 protein is transported through the secretory pathway and localized to the apoplast, where it signals the organizing centre in the underlying cells (4) . Another example is obser ved during Arabidopsis embryogenesis, when post-transcriptional RNA silencing signals move from cell to cell by diffusion through the plasmodesmata and this movement correlates with the plasmodesmata aperture (5). A third example is provided by the photoreceptor phytochrome B expressed in mesophyll cells, which is able to down-regulate the expression of the flowering promoter FLOWERING LOCUS T (FT) in the vascular bundles, while phytochrome B expressed in the vascular bundles is not effective. This indicates that an unknown phytochrome-derived signal moves from the mesophyll to the vascular bundles (6).

2.4. Long-Distance Signals

Classic examples of long-distance signals are provided by hormones. It has been known for many decades that auxin produced in the shoot apex and travelling down the stem can inhibit the outgrowth of axillary shoot meristems (7). In plants grown in soil with low water content, abscisic acid synthesized in the root can travel with the transpiration stream to the leaves and cause the closure of stomata and a reduction in the rate of transpiration (8). Classical physiological studies involving grafting and localized irradiations have demonstrated that the photoperiodic induction

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of flowering requires a signal produced in the leaves under inductive conditions and translocated to the apex, where it initiates the transition to flowering. Recent studies have provided strong support for the role of the FT protein as the signal that moves from the leaves to the apex under inductive conditions in Arabidopsis. We describe the evidence in favour of this contention because of its value as an example of the molecular analysis of a long-distance signal that had remained elusive for several decades. The FT gene is expressed in the vascular tissues of the leaves (9) but the FT protein interacts with the transcription factor FLOWERING LOCUS D (FD) in the shoot apex (10, 11). By using a miRNA construct to silence FT expression, Mathieu et al. (12) have shown that in order to induce flowering, FT mRNA is necessary in vascular tissues but not in the apex. The expression of Myc FT under the control of a vasculature-specific promoter rescues the late-flowering phenotype of ft mutants (13). This construct yields Myc FT mRNA expression restricted (within levels of detection) to vascular tissues, but the Myc-tagged protein shows a gradient out of the vascular tissues (13). A Myctagged FT protein fusion containing a nuclear localization signal is biologically active when expressed ectopically but it fails to induce early flowering if the gene is placed under the control of a vasculature-specific promoter, indicating that a protein unable to leave the vascular tissue is not effective unless expressed in target tissues (13). FT fusion proteins expressed specifically in phloem cells move to the apex and move long distances between grafted plants (14). Similarly, the expression of a fusion containing FT, the tobacco Etch virus (TEV) protein, three copies of the yellow-fluorescent protein (YFP) and a nuclear localization signal, in vascular tissues, does not yield a YFP signal in the apex and fails to induce flowering, despite the effectiveness of this construct when expressed ectopically (12). However, early flowering occurs when this construct is co-expressed with a TEV protease gene in vascular tissues to cleave the fusion protein and release FT (12). Finally, expression of FT fused to GFP in the companion cells or FT in the minor veins of the phloem is able to induce flowering but the expression of FT fused to GFP in the minor veins is not effective, suggesting that the fusion protein is too large to migrate from the minor veins to the meristem (14). These experiments uncouple FT protein movement from its biological function, supporting the notion that under inductive conditions the FT protein is produced in the vascular tissue of the leaf and migrates to the apex via the phloem. The FT protein and neither FT mRNA nor an FT downstream player is the signal carrying the information from the leaf to the apex. Further support to this conclusion comes from experiments with short-day plants. In rice, the protein encoded by Hd3a (an orthologue of FT) moves from the leaf to the shoot apical

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meristem and induces flowering (15). In Cucurbita moschata, expression of FT in a domain very distant from the apex induces flowering under long days (16). In grafting experiments, stocks of the long-day plant Cucurbita maxima induce flowering of Cucurbita moschata scions under long days and the phloem exudates of the stocks contain detectable FT protein but no detectable FT mRNA (16).

3. The Receptors and their Downstream Action 3.1. Two NegativeRegulation Steps in Series

Perhaps intuitively, one could imagine that the most common sequence in signal transduction involves activation of the receptor by the relevant signal followed by activation of downstream players and finally activation of the physiological or developmental process. More often than not, however, signalling involves two steps where a player negatively regulates a downstream partner. Protein destruction in the 26S proteasome plays a key role in negative regulation in diverse signalling pathways. Targeting for degradation in the 26S proteasome involves binding of ubiquitin by steps catalyzed by the enzymes E1, E2 and E3, where the latter provides protein selectivity to the system (17). TRANSPORT INHIBITOR RESPONSE 1 (TIR1) was first considered to be involved in the control of auxin transport but later identified as an auxin receptor (18, 19). TIR1 is an F-box protein that integrates a type of E3 complex. Auxin binds to the base of the TIR1 pocket and the rest of the pocket is occupied by an AUX/ IAA protein. Auxin acts as a “molecular glue” by occupying the hydrophobic cavity at the interface between TIR1 and AUX1 and facilitating their interaction (20). When the levels of auxin are low, the transcription of the genes stimulated by this hormone via the action of ARF transcription factors remains blocked by the action of AUX/IAA transcriptional repressors. Increased levels of auxin facilitate the binding of AUX/IAA targets by TIR1, and these targets become ubiquitinated and subsequently destroyed in the 26S proteasome. A comparable picture is observed for gibberellin signalling, where GIBBERELLIN INSENSITIVE DWARF1 (GID1) binds DELLA proteins in a gibberellin-dependent manner, causing the degradation of DELLA in the proteasome and the release of the gibberellin-response phenotype that was repressed by the DELLA proteins (21). Light signalling provides another example. In the nucleus, several transcription factors required for photomorphogenesis are ubiquitinated and targeted to degradation in the 26S proteasome by the action of the E3 ligase CONSTITUTIVE PHOTOMORPHOGENESIS

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1 (COP1) (22, 23, 24). The phytochrome and cryptochrome photoreceptors cause COP1 migration to the cytoplasm (25). In response to the reduced levels of nuclear COP1, the pools of transcription factor build up and bind their target sequences in genes that respond to light, favouring the transition between the pattern typical of seedlings grown in darkness (skotomorphogenesis) and the pattern typical of light-exposed seedlings (photomorphogenesis). In the case of TIR1, GID1 and the photoreceptors, binding of the hormone or exposure to light activates the receptor. However, binding of ethylene to its receptors related to bacterial two-component regulators results in their inactivation. Ethylene binding involves the N-terminal domain of the receptor (26). Single loss-of-function mutants of ethylene receptors show at most a weak phenotype but double, triple and quadruple receptor mutants show ethylene-response phenotypes in the absence of ethylene, indicating that ethylene responses are negatively regulated by the receptors and binding of the ligand releases the expression of the ethylene-induced pattern of cell growth (27). In an evolutionary sense, the ability to regulate basic processes such as cell growth in response to endogenous or exogenous stimuli could originate if one step involved in the process becomes dependent on the signal for its own activation. Alternatively, a similar regulation could be achieved if one step required by the process becomes repressed by a factor that in turn is repressed by the signal. A recent analysis of the acquisition of the gibberellin-DELLA mechanism of growth regulation suggests that this mechanism evolved through independent recruitment of the following: (1) the ability of gibberellins to enhance the interaction between the receptor (GID1) and the DELLA family of candidate transcription factors, which results in the destruction of DELLA in the 26S proteasome; and (2) the ability of DELLA to inhibit growth (28). The acquisition of negative regulation could have advantages because the basic process may already be working while the module involving negative regulation by the signal of the process negative regulator is evolving. 3.2. A Metabolic Enzyme Working as a Receptor

The search for receptors requires an open mind. Some receptor functions can remain hidden by apparently unrelated biochemical functions. An interesting case is that of glucose sensing by Hexokinase 1 (29). In addition to its metabolic functions outside the nucleus, this enzyme has a separable regulatory function, which involves the formation of a nuclear complex containing two proteins previously known for their unrelated metabolic activities: the vacuolar H+-ATPase B1 and the 19S regulatory particle of proteasome subunit. This complex would directly modulate the expression of targets of glucose signalling. Mutants at each one of the latter two loci share phenotypic consequences with hexokinase

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1 mutants (29). As noted by Cho et al. (29), metabolic enzyme genes are normally neglected in signal-transduction studies and complex components with a known function are not analysed for unrelated functions, but they could be part of other complexes with a completely different role. 3.3. Multiple Receptors may Perceive a Signal

In many cases a signal can be perceived by multiple receptors. These receptors can belong to the same family and there is often strong redundancy. This redundancy is revealed by the need to combine mutations in multiple members of the family to observe a clear phenotype, as is the case for ethylene receptors (27). The analysis of binding affinities by ethylene receptors argues against a role in the perception of different ranges of ethylene (26). Phytochromes show conditional redundancy, i.e., only in certain contexts the impact of loss-of-function mutation of one member of the family is stronger in the loss-of function background of another member (30). However, there is also obvious labour division because phytochrome A is functionally more important in environments rich in far-red light (places under the shade of green leaves) than in environments rich in red light, whereas phytochrome B has the opposite functional pattern (31). Phytochrome A is therefore able to perceive the difference between darkness and light of dense canopies (low proportion of red compared with far-red light) whereas phytochrome B is able to perceive the difference between dense canopies and open places (high proportion of red light) (32). In addition to perception by the products of multiple members of a gene family, a signal can be perceived by non-homologous genes. Light is perceived by multiple photoreceptors and cryptochromes play a key role with phytochromes in the control of plant body shape and function. Non-homologous receptors may also show redundancy (30). Blue light is perceived by cryptochromes and phytochrome A and in Arabidopsis plants grown under this light condition, only the triple phyA cry1 cry2 mutant (lacking phytochrome A and cryptochromes 1 and 2) shows the hypocotyl length of a dark-grown seedling. Three ABA receptors of widely divergent nature have recently been reported: one is FCA, which is an RNA-binding protein involved in the control of flowering; the second is the Mg-chelatase H subunit, also involved in the synthesis of chlorophyll and in retrograde signalling from the plastid to the nucleus; and the third is the G protein-coupled receptor GCR2 (33). These receptors appear to be involved in different physiological processes controlled by abscisic acid.

3.4. A Receptor can be Insufficient to Perceive a Signal

In the case of the photoperiodic signal that controls flowering, the simultaneous measure of light and time is necessary. In Arabidopsis thaliana, flowering is accelerated by long days. The expression of the CONSTANS gene is under the control of a

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circadian rhythm (34). When the days are short, the phase of high CO expression occurs in darkness and the CO protein is degraded (35). When the days are longer, the presence of light partially overlaps with the phase of the rhythm when CO mRNA is high (36) and the activities of photoreceptors like cryptochrome 2 and phytochrome A stabilize the CO protein. CO activates the expression of FT, which is a strong promoter of flowering. The “coincidence” between light and an endogenous rhythm was proposed several decades ago as a mechanism for the measurement of day length. With different actors, the same principle operates in the control of daily fluctuations of hypocotyl growth in young seedlings to achieve maximum growth rates during the last hours of the night (37). Control by light alone would lead to rapid rates during the whole night and a mechanism simply counting the number of hours in darkness before growth is released from inhibition would not allow plants to adjust to different durations of the night throughout the year. The expression of the bHLH transcription factor genes PIF4 and PIF5, which positively regulate extension growth, is controlled by a circadian rhythm. The abundance of the PIF4 and PIF5 proteins is negatively regulated by light perceived by phytochromes. Therefore, during the day, the levels of PIF4 and PIF5 remain low and growth is partially arrested; during the first portion of the night PIF4 and PIF5 mRNA abundance is low. Only during the last part of the night there is a coincidence between high PIF4 and PIF5 expression and darkness to allow PIF4 and PIF5 protein stability and the promotion of hypocotyl growth. 3.5. Multiple Events Triggered by a Receptor

A given receptor can perform multiple initial events defined by the biochemical nature and/or the sub-cellular localization of those events. Phytochrome A, for instance, is present in the cytoplasm in dark-grown seedlings and exposure to light triggers the migration to the nucleus but a very significant pool remains in the cytoplasm (38). The inhibition of hypocotyl growth by phytochrome A requires nuclear localization and the double fhy1 fhl mutant, where phytochrome A does not accumulate in the nucleus (39, 40), fails to show this response. However, fhy1 fhl retains some responses apparently mediated by phytochrome A suggesting that the cytoplasmic pool is responsible for these effects (40). Phytochrome-mediated induction of transcription factor migration from the cytoplasm to the nucleus has also been reported and this implies a cytoplasmic function of the receptor (38). In the nucleus, both phytochromes A and B trigger the phosphorylation of the bHLH transcription factor PIF3, which becomes tagged for degradation in the proteasome (41). In the nucleus, phytochromes A and B also induce the migration of COP1 to the cytosol (25). Signal-output analyses have revealed that phytochrome A can

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initiate two at least partially divergent pathways, one that requires sustained excitation with far-red light and another that saturates with infrequent excitation and lower fluence rates. The former requires a region of the promoter of target genes that is fully dispensable for the latter, providing evidence that different environmental signals can propagate by pathways with discrete end-points even if perceived by the same receptor (42). Even if we narrow down the analysis to a single target, this can be affected at different points, which include the regulation of the rate of transcription of its gene, the stability of the transcripts, the rate of protein synthesis, the turnover of the protein, the regulation of its sub-cellular localization and the regulation of its activity. Very often the propagation of a given signal can intersect and modify the line that connects DNA to its active protein at more than one point. The transition from full darkness to light, for instance, enhances the expression of the bZIP transcription factor gene HY5, which plays a key role in photomorphogenesis (43). Light also increases the stability of the HY5 protein by reducing COP1 nuclear activity (22, 23) and modifies HY5 phosphorylation status (44). An interesting case is that of AUX/ IAA because auxin increases their mRNA levels but at the same time it causes the degradation of the protein (45). 3.6. Signalling by Non-Protein Molecules 3.6.1. Second Messengers

In addition to proteins, signal transduction can incorporate non-protein molecules that participate in the propagation of the signal downstream the receptor, which are called second messengers, because the first messenger is the ligand that binds the receptor. The role of second messengers is an issue of frequent discussion in plant physiology because it is not easy to move from pharmacological approaches to genetic evidence. Calcium is a typical second messenger and it has been proposed to mediate responses to abscisic acid. Seedlings of Arabidopsis mutated in a class of Ca2+dependent Ca2+-release channel located at the vacuole membrane have altered electrophysiology and germination responses to abscisic acid but normal stomatal responses to abscisic acid (46). Recent reports showing the opposite relationship between Ca2+ transients and physiological responses illustrate the complexity of Ca2+-mediated signalling. In guard cells of Arabidopsis, more cytosolic Ca2+ transients are observed under low than under high concentrations of carbon dioxide, without dramatic effects on transient amplitude or duration (47). Stomata aperture responses to carbon dioxide concentrations are reduced in the cells in which Ca2+ transients are abolished by a Ca2+ chelator. Ca2+ transients appear to be required not for the early stomatal movement but for maintaining the aperture according to carbon dioxide levels (47). Since, experimentally induced Ca2+ transients trigger rapid stomatal closure, Young et al. (47) conclude that the carbon dioxide signal must be involved in the modulation of Ca2+ sensors.

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3.6.2. Regulatory RNA

In addition to proteins and second messengers, regulatory RNA may also have a role in signal transduction. Small regulatory RNA can down-regulate the expression of target genes at the transcriptional and post-transcriptional levels. The microRNA (miRNA) and the small interfering RNA (siRNA) are two types of small regulatory RNA with different origins (48). miRNA derives from a DNA sequence different from that of the target regulated locus. RNA polymerase II generates a primary transcript with a stem-loop RNA secondary structure. A dicer endonuclease excises a miRNA duplex of approximately 21 nucleotides from this structure, and one strand of the miRNA duplex binds the ARGONOUTE protein and guides the complex to the partially complementary target mRNA sequence, where ARGONOUTE catalyses the cleavage of the target mRNA. The expression of the loci that give origin to miRNA is regulated by endogenous and exogenous signals. The miRNA produced by the JAW locus triggers cleavage of several TCP mRNAs, and the restriction of TCP gene expression to the correct domains is necessary for normal leaf development (49). miR172 increases its expression with the progression of development and regulates flowering time and floral organ identity by down-regulating AP2-like target genes at the translational, rather than RNA cleavage levels (50). Not all the targets of miRNA are transcription factors. Arabidopsis infection by Pseudomonas syringae induces the expression of a miRNA that targets the mRNA of auxin receptors and the reduced auxin signalling restricts pathogen growth, leading to increased plant resistance (51). The homeostasis of phosphate is controlled by the miRNA miR399, which is normally not expressed but is induced in the roots by phosphate starvation (52). miR399 represses the expression of a ubiquitin-conjugating E2 enzyme that directly or indirectly down-regulates the phosphate uptake and transport systems (52). These are only some examples that support the emergent idea of a significant function of regulatory RNA of different origin in signal transduction.

3.7. Multiple Targets

Many aspects of the regulation of gene expression (defined as the processes that yield the active gene product in its place of action) can be targets of specific signalling cascades. Transcription factors (or transcription-factor combinations) provide specificity to the control of transcription in the context of a given physiological or developmental response. It is therefore not surprising to find modifications in transcription factor abundance, localization or activity as intermediates between signalling cascades and the modification of expression of structural genes involved in the diverse physiological processes. The aforementioned cases of HY5 and AUX/IAA transcription factors provide examples for light and auxin signalling. However, many of the transcription factors that change their own expression in response to a signal often have weak effects on the physiological output of that signal (53).

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Signalling can also affect chromatin structure. A typical example is provided by the control of flowering by the epigenetic memory of winter (54). In Arabidopsis, flowering is repressed by the MADS-box transcription factor FLOWERING LOCUS C (FLC) (55). The expression of FLC is promoted by FRIGIDA, among other genes, which may maintain chromatin in a state that allows transcription. In turn, the genes of the so-called autonomous pathway, which include RNA-binding proteins, repress FLC likely acting at the post-transcriptional level. Further repression of FLC is achieved by the action of FVE, which also affects chromatin status. The balance between these opposing controls yields a level of FLC expression that shows strong natural variation. Prolonged exposure to cold temperatures accelerates flowering in the Arabidopsis accessions with strong FLC expression through the vernalization process. Vernalization reduces the expression of FLC and this reduction persists during mitotic divisions until flowering. Prolonged cold increases the expression of VERNALIZATION INSENSITIVE 3 (VIN3), which is involved in the acetylation of histone and the methylation in histone H3 Lys9 and Lys27 that participate in the early steps of FLC repression (56). The VERNALIZATION 1 (VRN1) and VERNALIZATION 2 (VRN2) genes are required to maintain the inhibitory status of chromatin on FLC once the plants return from cold to non-vernalizing temperatures (54). Therefore, endogenous and vernalization signals modify the “histone-code” that activates or represses FLC gene transcription. Even complexes of general function offer room for regulatory specificity. The nuclear protein PHYTOCHROME AND FLOWERING TIME 1 (PFT1) is involved in the induction of flowering by low-red to far-red ratios (which reduce the proportion of active phytochrome B) and has a negligible role in the photoperiodic induction of flowering (57). In an independent study, PFT1 has recently been identified as one of the subunits of the MEDIATOR complex of Arabidopsis thaliana (58). Mediator is involved in the exchange of information between the transcription factors that bind DNA and the core promoters of transcription. Therefore, PFT1 could provide selective regulation of Mediator function in specific developmental contexts.

4. Signalling Networks 4.1. Topology of Biological Networks

Signalling depends on numerous interactions: proteins that interact with other proteins, proteins that interact with DNA, etc. Cellular processes can therefore be represented by networks of interactive players or “nodes” and their interactions or “links” that connect the nodes. The “degree” is the number of connections of a node. The

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nature of the components under consideration defines the different types of biological networks and previous studies in different biological systems include protein–protein interactions, protein binding to DNA sequences, protein phosphorylation, metabolic interactions, genetic interactions, co-expression networks, etc. (59). Networks of very different nature (including those describing social relationships, the Internet or intracellular interactions) share common architectural features (60). In the so-called “scalefree” networks the distribution of links follows the power law, where most nodes have few links and a few nodes, called “hubs”, have a larger number of links. These generalizations are helping to uncover the functional significance of the complexity often found in the control of biological processes. 4.2. Functional Consequences of Network Topology

Important functional features of the cell’s circuits, such as robustness and vulnerability, can be accounted for by the topology of biological networks (61). Robustness is the property that allows a system to maintain its functions despite perturbations caused by fluctuations in the internal and/or external environments (62). Vulnerability is the fragility shown by the system, i.e., the probability of collapse. Extreme vulnerability against rare perturbations is considered a trade-off of robustness (62). Networks are robust because they contain many nodes with few links and random perturbations are likely to affect these nodes, precluding failure propagation. Networks can collapse in response to rare events because hubs represent highly vulnerable spots. Transcriptional networks are important in the control of plant development. For instance, coordinate regulation through positive regulatory loops among homeotic transcription factors has also been described during early floral organogenesis in Arabidopsis thaliana (63). However, the functional dynamics of transcriptional networks needs to be further investigated in multi-cellular organisms in general and plants in particular.

5. Conclusions Figure 1.1 provides a model of the players involved in plant signal transduction. As a model, this figure represents a simplification of the reality. Proteins shape plant body form and function directly through their action as enzymes and structural proteins (arrow 1). In addition, proteins also have a wide array of regulatory functions and can modify other proteins in different ways. These modifications can occur through direct protein–protein interactions, which result in altered protein activity (arrow 3). Proteins can also modify the kinetics of second messenger abundance and

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Fig. 1.1. Model demonstrating the interactions among players involved in signal transduction in plants. 1, Proteins shape plant body form and function directly as enzymes and structural proteins. 2, Signals from the endogenous and external environment affect the activity of selected proteins (receptors). 3, Proteins can directly affect the activity of other proteins. 4, Proteins can modify the kinetics of second messenger abundance and localization, which in turn also alter protein activity. 5, Proteins regulate the rate of transcription by acting as transcription factors. 6, Proteins regulate the rate of transcription by modifying the status and accessibility to DNA. 7, Proteins regulate mRNA stability and the rate of translation. 8, Regulatory RNAs modify transcription. 9, Regulatory RNAs modify mRNA stability.

localization, which in turn can also alter protein activity (arrow 4). Signals from the endogenous and external environment affect the activity of selected proteins that act as receptors of these signals (arrow 2). Proteins may also control protein abundance by regulating the rate of transcription acting as transcription factors (arrow 5), or by modifying the status and accessibility to DNA (arrow 6), the processing of the transcripts, their stability and the rate of translation (arrow 7). Proteins can also regulate the production of regulatory RNA, which in turn can modify transcription (arrow 8) or transcript stability (arrow 9). This model shows that signals can modulate pant body shape and function through multiple loops that are not mutually exclusive. We can unfold a given loop into a linear pathway but signalling as a whole deviates from linearity. In principle, each loop can involve a network of interactive components (protein–protein, protein–DNA, etc.). Multiple signals, receptors, transducing elements and targets shape plant physiology and development, adding another dimension to the model in Fig. 1.1. The challenge ahead has two faces. On the one hand, a reductionist approach will be necessary to gain insight into the modes in which the signalling current moves via specific cascades. In this sense, the complexity of the networks is a burden that must be handled with care. On the other hand, understanding plant signalling ultimately means understanding the system as a whole and in this context, the complexity of the network is of central significance and richness and cannot be neglected. The chapters that follow offer a careful description of laboratory tools available to face this challenge.

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Acknowledgements I thank University of Buenos Aires, Consejo Nacional de investigaciones Científicas y Técnicas (CONICET), Agencia Nacional de Promoción Científica y Tecnológica (ANPCYT), Guggenheim Foundation and FIRCA for financial support of the experiments conducted in my laboratory. References 1. Casal, J.J., Luccioni, L., Oliverio, K.A., and Boccalandro, H.E. (2003) Light, phytochrome signalling and photomorphogenesis in Arabidopsis. Photochem. Photobiol. Sci. 2, 625–636. 2. Benech Arnold, R.L., Ghersa, C.M., Sanchez, R.A., and Garcia Fernandez, A.E. (1988) The role of fluctuating temperatures in the germination and establishment of Sorghum halepense (L.) Pers. Regulation of germination under leaf canopies. Funct. Ecol. 2, 311–318. 3. Fey, V., Wagner, R., Bräutigam, K., and Pfannschmidt, T. (2005) Photosynthetic redox control of nuclear gene expression. J. Exp. Bot. 56, 1491–1498 4. Rojo, E., Sharma, V.K., Kovaleva, V., Raikhel, N.V., and Fletcher, J.C. (2002) CLV3 is localized to the extracellular space, where it activates the Arabidopsis CLAVATA stem cell signaling pathway. Plant Cell 14, 969–977. 5. Kobayashi, K. and Zambryski, P. (2007) RNA silencing and its cell-to-cell spread during Arabidopsis embryogenesis. Plant J. 50, 597–604. 6. Endo, M., Nakamura, S., Araki, T., Mochizuki, N., and Nagatani, A. (2005) Phytochrome B in the mesophyll delays flowering by suppressing FLOWERING LOCUS T expression in Arabidopsis vascular bundles. Plant Cell 17, 1941–1952. 7. Leyser, O. (2005) The fall and rise of apical dominance. Curr. Opin. Genet. Dev. 15, 468–471. 8. Jiang, F. and Hartung, W. (2007) Long-distance signalling of abscisic acid (ABA): The factors regulating the intensity of the ABA signal. J. Exp. Bot. doi:10.1093/jxb/erm127. 9. Takada, S. and Goto, K. (2003) TERMINAL FLOWER2, an Arabidopsis homolog of HETEROCHROMATIN PROTEIN1, counteracts the activation of FLOWERING LOCUS T by CONSTANS in the vascular

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30. Casal, J.J. (2000) Phytochromes, cryptochromes, phototropin: Photoreceptor interactions in plants. Photochem. Photobiol. 71, 1–11. 31. Quail, P.H., Boylan, M.T., Parks, B.M., Short, T.W., Xu, Y., and Wagner, D. (1995) Phytochromes: Photosensory perception and signal transduction. Science 268, 675–680. 32. Yanovsky, M.J., Casal, J.J., and Whitelam, G.C. (1995) Phytochrome A, phytochrome B and HY4 are involved in hypocotyl growth responses to natural radiation in Arabidopsis: Weak de-etiolation of the phyA mutant under dense canopies. Plant Cell Environ. 18, 788–794. 33. Hirayama, T. and Shinozaki, K. (2007) Perception and transduction of abscisic acid signals: keys to the function of the versatile plant hormone ABA. Trends Plant Sci. 12, 343–351. 34. Suarez-Lopez, P., Wheatley, K., Robson, F., Onouchi, H., Valverde, F., and Coupland, G. (2001) CONSTANS mediates between the circadian clock and the control of flowering in Arabidopsis. Nature 410, 1116–1120. 35. Valverde, F., Mouradov, A., Soppe, W., Ravenscroft, D., Samach, A., and Coupland, G. (2004) Photoreceptor regulation of CONSTANS protein in photoperiodic flowering. Science 303, 1003–1006. 36. Yanovsky, M.J. and Kay, S.A. (2002) Molecular basis of seasonal time measurement in Arabidopsis. Nature 419, 308–312. 37. Nozue, K., Covington, M.F., Duek, P.D., Lorrain, S., Fankhauser, C., Harmer, S.L., and Maloof, J.N. (2007) Rhythmic growth explained by coincidence between internal and external cues. Nature 448, 358–61 38. Nagy, F. and Schäfer, E. (2000) Nuclear and cytosolic events of light-induced, phytochrome-regulated signaling in higher plants. EMBO J. 19, 157–163. 39. Hiltbrunner, A., Viczian, A., Bury, E., Tscheuschler, A., Kircher, S., Toth, R., Honsberger, A., Nagy, F., Fankhauser, C., and Schafer, E. (2005) Nuclear accumulation of the phytochrome A photoreceptor requires FHY1. Curr. Biol. 15, 2125–2130. 40. Röster, J., Klein, I., and Zeidler, M. (2007) Arabidopsis fhl/fhy1 double mutant reveals a distinct cytoplasmic action of phytochrome A. Proc. Nat. Acad. Sci. 104, 10737–10742. 41. Al-Sady, B., Ni, W., Kircher, S., Schäfer, E., and Quail, P.H. (2006) Photoactivated phytochrome induces rapid PIF3 phosphorylation prior to proteasome-mediated degradation. Mol. Cell 23, 439–446.

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42. Cerdán, P.D., Staneloni, R.J., Ortega, J., Bunge, M.M., Rodríguez-Batiller, M.J., Sánchez, R.A., and Casal, J.J. (2000) Sustained but not transient phytochrome A signaling targets a region of an Lhcb1*2 promoter not necessary for phytochrome B action. Plant Cell 12, 1203–1211. 43. Oyama, T., Shimura, Y., and Okada, K. (1997) The Arabidopsis HY5 gene encodes a bZIP protein that regulates stimulusinduced development of root and hypocotyl. Genes Dev. 11, 2983–2995. 44 . Hardtke , C.S . and Deng , X.-W. (2000) The cell biology of the COP/DET/FUS proteins. Regulating proteolysis in photomorphogenesis and Beyond? Plant Physiol. 124, 1548–1557. 45. Kepinski, S. and Leyser, O. (2002) Ubiquitination and auxin signaling: A degrading story. Plant Cell 14, S81–S95. 46. Peiter, E., Maathuis, F.J.M., Mills, L.N., Knight, H., Pelloux, J., Hetherington, A.M., and Sanders, D. (2005) The vacuolar Ca2+-activated channel TPC1 regulates germination and stomatal movement. Nature 434, 404–408. 47. Young, J.J., Mehta, S., Israelsson, M., Godoski, J., Grill, E., and Schroeder, J.I. (2006) CO2 signaling in guard cells: Calcium sensitivity response modulation, a Ca2+-independent phase, and CO2 insensitivity of the gca2 mutant. Proc. Nat. Acad. Sci. 103, 7506–7511. 48. Rajagopalan, R., Vaucheret, H., Trejo, J., and Bartel, D.P. (2006) A diverse and evolutionarily fluid set of microRNAs in Arabidopsis thaliana. Genes Dev. 20, 3407–3425. 49. Palatnik, J.F., Allen, E., Wu, X., Schommer, C., Schwab, R., Carrington, J.C., and Weigel, D. (2003) Control of leaf morphogenesis by microRNAs. Nature 425, 257–263. 50. Aukerman, M.J. and Sakai, H. (2003) Regulation of flowering time and floral organ identity by a microRNA and its APETALA2-like target genes. Plant Cell 15, 2730–2741. 51. Navarro, L., Dunoyer, P., Jay, F., Arnold, B., Dharmasiri, N., Estelle, M., Voinnet, O., and Jones, J.D.G. (2006) A plant miRNA contributes to antibacterial resistance by repressing auxin signaling. Science 312, 436–439.

52. Chiou, T.-J., Aung, K., Lin, S.-I., Wu, Ch.Ch., Chiang, S.-F., and Su, Ch.-L. (2006) Regulation of phosphate homeostasis by microRNA in Arabidopsis. Plant Cell 18, 412–421. 53. Khanna, R., Shen, Y., Toledo-Ortiz, G., Kikis, E.A., Johannesson, H., Hwang, Y.-S., and Quail, P.H. (2006) Functional profiling reveals that only a small number of phytochrome-regulated early-response genes in Arabidopsis are necessary for optimal deetiolation. Plant Cell 18, 2157–2171. 54. Henderson, I.R. and Dean, C. (2004) Control of Arabidopsis flowering: The chill before the bloom. Development 131, 3829–3838. 55. Michaels, S.D. and Amasino, R.M. (1999) FLOWERING LOCUS C encodes a novel MADS domain protein that acts as a repressor of flowering. Plant Cell 11, 949–956. 56. Sung, S. and Amasino, R.M. (2004) Vernalization in Arabidopsis thaliana is mediated by the PHD finger protein VIN3. Nature 427, 159–164. 57. Cerdán, P.D. and Chory, J. (2003) Regulation of flowering time by light quality. Nature 423, 881–885. 58. Bäckström, S., Elfving, N., Nilsson, R., Wingsle, G., and Björklund, S. (2007) Purification of a plant mediator from Arabidopsis thaliana Identifies PFT1 as the Med25 subunit. Mol. Cell 26, 717–729. 59. Zhu, X., Gerstein, M., and Snyder, M. (2007) Getting connected: analysis and principles of biological networks. Genes Dev. 21, 1010–1024. 60. Barabási, A.L. and Albert, R. (1999) Emergence of scaling in random networks. Science 286, 509–512. 61. Barabási, A.L. and Oltavi, Z.N. (2004) Network biology: Understanding the cell’s functional organization. Nature Rev. Genet. 5, 101–113. 62. Kitano, H. (2004) Biological robustness. Nature Rev. Genet. 5, 828–837. 63. Gómez-Mena, C., de Folter, S., Costa, M.M.R., Angenent, G.C., and Sablowski, R. (2005) Transcriptional program controlled by the floral homeotic gene Agamous during early organogenesis. Development 132, 429–438.

Chapter 2 Phenotyping of Arabidopsis Mutants for Developmental Effects of Gene Deletions Cordelia Bolle Abstract With the completion of the Arabidopsis thaliana genome sequencing project the next major challenge is the assignment of biological functions to the more than 25,000 genes. Reverse genetics is a powerful tool to elucidate gene function in Arabidopsis. Increasingly sophisticated genetic approaches are being developed for reverse genetics with the long-term goal of understanding how the coordinated activity of all proteins rises to a complex organism. Identification of a biological function for each gene is often doomed to fail as many loss- or gain-of-function lines exhibit no obvious phenotypes under normal propagation conditions. Here we provide an overview on how to phenotype plants during their development. This phenotypic streamlined analysis is based on a series of defined growth stages (germination, seedling, vegetative, and reproductive stages), which can be used for the profiling of mutants. Key words: Phenotype, reverse, genetics, signaling, development, mutant analysis.

1. Introduction The present-day plant biologist faces the daunting challenge of assigning functions to the boundless “function-not-known” genes identified by recent plant genome sequencing projects (1). Many proteins have also been vaguely implied to be involved in signal transduction because of their homologies to other proteins or protein families, although it is not clear from the sequence, in which signaling pathway or process they might be involved. To ascertain the function of a particular gene a reverse genetics approach is commonly undertaken (2, 3). The first step in such an undertaking is to procure mutations in the gene of interest from

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the different collections available (e.g., http://www.arabidopsis. org/servlets/Search?action=new_search&type=germplasm). At present several different strategies are employed to generate genespecific knock-out mutations. These include gene silencing with techniques such as RNAi or antisense, insertional mutagenesis, and chemical-induced point mutations identified by TILLING (4). Furthermore, lines overexpressing specific genes can provide valuable information. A common problem faced by the scientist after generating plant lines homozygous for the mutations is that no obvious mutant phenotype can be detected. Under these circumstances it may be necessary to assay the mutant plants in many different developmental processes to uncover phenotypes. Developmental phenotypes are often overlooked due to the inexperience with the different developmental stages and due to uncontrolled growth conditions that increase the natural variation within plant growth. Here we present a streamlined protocol of how to use a broad spectrum of developmental assays to assess alterations from wild-type development, from germination to the adult plant and the dried seeds.

2. Materials 1. Plates for tissue culture: Round, sterile plastic Petri dishes with a diameter of 145 mm are best for most experiments, otherwise small Petri dishes can also be used. Some producers add a grid on the bottom lid, which helps with orientation and alignment. 2. Media plates: Half-strength (0.5X) Murashige and Skoog (MS) medium (5) is autoclaved together with 0.8% (w/v) agar at 121°C for 15 min. This medium contains usually no sucrose as sucrose interferes with many signal transduction pathways. Once the medium has been cooled down to 60°C it can be poured into the Petri dishes. After the media are solidified and have been dried of excessive condensated water they can be stored for up to 2–3 months at 4–10°C. Before using plates from the cold, they should be dried again in a sterile hood to remove excess water. For vertical plates increasing the concentration of agarose in the media to 1% (w/v) is advisable. 3. Sealing the plates: Bandage tape such as Leukopor or Micropore should be used to close the lids of the Petri dishes because in contrast to parafilm it allows gas exchange. 4. Sterilization solution for seeds:

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a) 30% (v/v) Clorex solution (or any other commercial bleach that contains hypochloride) with 0.05% (v/v) Tween 20 b) Sodiumhypochloride, HCl conc. (Be careful, use gloves and protective goggles) 5. Visualization of leaf venation patterns Clearing solution: 80 g chloral hydrate in 30 ml water Mounting solution: 80 g chloral hydrate, 20 ml glycerol, and 10 ml water 6. Imprinting of plant surface 2% (w/v) low-melt agarose containing 0.01% (w/v) bromophenol blue

3. Methods 3.1. Generation of Homozygous Lines

Before working with mutant lines generated by insertion or point mutation it is important to remove putative additional mutations from the line by repeated backcrossing to wild type. For this purpose the mutant line is used to pollinate a wild type of the same ecotype. The progeny is then tested for the presence of the mutation and is used again to pollinate wild type. This cycle can be repeated several times. The progeny is finally analyzed for the presence of the mutation and selfed. In the next generation homozygous lines can be selected, which can then be used for the phenotypic characterization (see Note 1). Once a phenotype has been described for a mutant line, it is important to confirm that the phenotype is genetically linked to the change in the genotype (see Note 2).

3.2. Surface Sterilization of Seeds

If seeds are to be germinated on medium containing plates they must be sterilized first. Two possible methods are as follows.

3.2.1. Liquid Sterilization

1. Seeds are placed for 10 min in a 30% (v/v) Clorex solution (or any other commercial bleach that contains hypochloride) with 0.05% (v/v) Tween 20 and agitated every 30 s. About 300 ml solution in a 1.5 ml reaction tube is suitable for 100 seeds. 2. After 10 min the seeds are briefly centrifuged to help settle the seeds and the solution is removed. The seeds are washed three times with sterile water. 3. Sterile 0.1% (w/v) agarose can be added to the seeds to facilitate distribution of the seeds on media plates with a pipette.

3.2.2. Vapor Sterilization

1. A maximum of 100 seeds is added to each reaction tube; otherwise, the penetration of the vapor is not guaranteed. 2. The reaction tubes are placed with their lid open in a desiccator together with a beaker containing 100 ml sodium

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hypochloride into which 5 ml HCl (conc.) is added dropwise (use gloves!). 3. After 5 h treatment the seeds are placed in a sterile hood to remove the fumes. The advantage of this method is that the seeds are dry and can be stored in a sterile way for further use. The dry seeds can be distributed on media plates by tapping the tube. 3.3. Stratification of Seeds

Stratification (break of seed dormancy) of imbibed seeds by cold (2–4 days at 4–10°C in darkness) improves coordinated germination, which is important for most phenotypic assays as it allows us to compare seedlings of the same age. Stratification is best performed after the seeds have been placed on media. To keep the plates dark they can be wrapped in aluminum foil.

3.4. Growth Conditions

Controlled growth conditions (light intensities and spectra, moisture and temperature) are important for reproducible results. Growth chambers are usually easier to control than greenhouses. However, be aware that conditions within a growth chamber can also be variable. White light should be maintained at around 100–300 µmol photons m−2 s−1, temperature between 18–22°C, and humidity at 50–70%. For most tests long-day conditions are recommended, with 16 h light and 8 h dark. Juvenile phenotypes are best observed when the plants are raised in tissue culture, because all organs (esp. roots) are easily accessible for analysis under the microscope. Adult plants are observed on soil as this allows them to develop more naturally. For growing plants on soil the best option is the use of multitrays with about 50 individual 5-cm-diameter pots. This allows the growth of several mutant lines and wild type on one tray, with each single plant in its own pot. Plants that are grown on soil can either directly be sown onto soil (3 seeds per pot, thinned out to one plant per pot upon germination) or pre-grown in tissue culture without antibiotics. Seedlings should be transferred from tissue culture to soil when they have 4–6 true leaves. To be able to observe the phenotypes of soil-grown plants no Aracons or anything similar should impede the development of the plant. To minimize any spurious differences between the mutant and wild type caused by different growth conditions, mutant and wild-type plants should be grown next to each other in the same growth facility. They should be maintained on the same media plate or, when in soil, on the same tray. A quick test for a mutant phenotype, to determine whether a mutant phenotype exists or not, can be carried out with 12 parallel plants. However, for confirmation of a phenotype the experiments have to be repeated at least three times with an adequate number of parallel plants to generate statistically significant data.

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3.5. Recording the Mutant Phenotype

A visual record of a mutant phenotype can be captured with a digital or a video camera fixed to a support. This allows us to take images of different plates or pots of a series from the same distance therefore with comparable sizes. Cold white light from the sides reduces shadows and is recommended because using a flash for taking photographs generates reflections on the media plates. To increase the contrast black felt can be put underneath the objects. If green plants are to be photographed a red cloth can enhance the contrast. A ruler placed next to the objects enables the evaluation of physical parameters. The images can be imported into software such as ImageJ (NIH; http://rsb.info. nih.gov/ij/), which displays either area statistics, line lengths, or angles. To measure length a “straight line,” a “segmented line,” or a “freehand line” can be used. The data can be processed in a program such as Microsoft Excel. The length of 1 cm of the ruler on the digital image can be used to calibrate the program or to calculate the actual lengths in Excel. All data should be statistically evaluated (mean value, standard deviation, t-Test, or similar) using Excel or similar programs.

3.6. Phenotypic Screens

To systematically screen many individual mutant seed lines whose function is unknown, a broad spectrum of environmental and developmental assays may be needed to uncover conditional phenotypes (see Note 3). The search for a phenotype can have major pitfalls and has to be conducted with care (see Note 4). As the environmental conditions in each laboratory vary, these assays have to be adapted to the environment in the greenhouse or growth chambers and to the ecotypes used. To observe as many developmental phenotypes as possible we devised the following sequence of experiments (see Fig. 2.1): 1. Place 30–50 sterilized seeds of one or more mutants or alleles and the corresponding WT on one Petri dish, which can be divided into squares or segments with a marker on the back lid (A). On a second plate place about 20 seeds in a line (B). Depending on the size of the plate 2–3 lines can be added, leaving space for 2 cm upward and 1 cm downward. After stratification move plate A to the white light (WL). The germination rate can be determined from this WL incubated plate (see Sect. 3.6.2). Once the seedlings on this plate A have produced the first pair of true leaves (5–7 days), transfer about 20 seedlings to another plate (plate C) forming a line on the top third of the plate. This plate is positioned vertically in the growth chamber to evaluate root growth (see Sect. 3.6.5). After 4–7 days the plate is rotated 90° to test for gravitropic responses (see Sect. 3.6.6). After stratification plate B is illuminated for 4 h with WL to induce germination and then returned to darkness, but at room temperature and in a in a vertical position.

3.6.1. Streamlining of a Phenotypic Screen

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Fig. 2.1. Schematic flow chart of a streamlined phenotypic screen for Arabidopsis mutants. A, B, and C are the different media plates used (see Sect. 3.6.1). WL, white light; D, darkness

Plate B is analyzed after 4 days for its hypocotyl elongation in darkness (see Sect. 3.6.4) and then placed into unilateral light to test for phototropism (see Sect. 3.6.7). 4. After 14 days (4–6 true leaves) the rest of the seedlings from plate A can be transferred to soil and the adult phenotype can be observed every 2–3 days until the seeds are harvested (see Sects. 3.6.8–3.6.20). 3.6.2. Germination Assay

1. To test germination efficiency seeds are placed on media plates. About 30–100 seeds per mutant line are optimal, which are evenly spaced (1 seed per 2 mm2) on the plate. Several lines can be placed on the same plate within different segments. 2. After stratification, plates are transferred to white light. Plates should be checked daily for germination (1–7 days) scoring both germinated and nongerminated seeds. This is done best with a stereo microscope with illumination from below. Non-germinated seeds will appear dark whereas empty seed coats appear translucent. A seed is scored as germinated as soon as the radicle has protruded and/or upon emergence of green cotyledons. The radicle of wild type usually appears two days after being transferred into white light. 3. Germination is best evaluated by plotting the % germination against “days after imbibition”.

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To test for dormancy the germination assay is in principle the same except that the germination of non-stratified seeds is evaluated. In this case the efficiency to break the dormancy in the presence of only water and light is tested without activating the cold-induced degradation of gibberellins (GA) (6,7). For all germination assays it is extremely important to use seeds that have been stored for the same time and that have been gathered from mother plants that have been grown under the same conditions. Freshly harvested seeds have a different germination behavior (esp. higher dark germination) than stored seeds, whereas longer storage reduces the overall germination efficiency. Germination is influenced by hormone levels e.g., abscisic acid (ABA) (inhibition) and GA (promoting), deficiencies during seed development, mobilization of stored proteins, fatty acid catabolism, but also by environmental factors such as light and temperature (8–11). 3.6.3. Cotyledon Number, Size, Shape, and Color

Cotyledons are best evaluated using a stereo microscope. Arabidopsis produces two cotyledons and one cotyledon is usually slightly bigger then the other due to positioning in the embryo. Cotyledons are smaller and more roundish than adult leaves and have fewer trichomes. Changes in the cotyledon shape or number are an indication of changes in developmental processes. Cotyledons should be photosynthetically active and thus appear green.

3.6.4. Hypocotyl Length

Hypocotyl elongation is mainly a process of cell expansion, which is regulated by photoreceptors (mainly phytochromes and cryptochromes) and hormones such as gibberellins, brassinosteroids and auxins. 1. To measure hypocotyl length place about 50 sterilized seeds per line on agar plates. Four to eight different lines can be placed on one big agar plate (of at least 13 cm diameter and 2 cm height); one of them should be WT control. Seeds can be plated relatively dense (1 seed in 1–2 mm2), but the cotyledons should not obstruct or shade each other. 2. After stratification place seeds for 2–6 h in white light and then incubate in the dark at 20°C over night. 3. Place the seedlings in the appropriate light conditions or keep them in darkness and incubate for 4–6 days. Hypocotyl elongation can be tested under different light qualities (white, red, blue, and far-red) and quantities (12). Best results are obtained using non-saturating light conditions. For white light, fluence rates of 0.5–5 mE photons m−2 s−1 are most effective. For a more detailed analysis a fluence response curve is appropriate. For this experiment parallel media plates with seedlings are incubated under increasing light intensities and hypocotyl length is determined. To generate

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different light qualities white light can be filtered through appropriate filters (blue light 400 nm, red light 660 nm, farred light 730 nm), or LEDs with the appropriate spectra can be used (see Note 5). 4. To measure the hypocotyl length the seedlings are stretched with the help of forceps longitudinally on an agar plate, and this can be photographed with a digital camera (see Sect. 3.5). The length of the hypocotyl is measured from the apical meristem to the beginning of the root, often characterized by the still attached seed coat (see Fig. 2.2a). Alternatively, seeds can be placed in a row on agar plates so that the plates can be incubated vertically and the seedlings can grow parallel to the agar. The disadvantage is that fewer lines can be placed on one plate, as 2 cm has to be allowed for upward growth and 1 cm for downward growth. Furthermore, not

Fig. 2.2. Wild-type phenotypes of Arabidopsis. (a) Red-light grown seedlings as an example for the hypocotyl measurement. Line represents measured length. (b) Seedlings were placed on the vertical plates with their root tips extending to the upper dot and left growing for 4 days. For root length measure the distance between the upper dot and the root tip (“L”). (c) Parameters for evaluation of leaf size. P, length of the petiole; L, length of the leaf blade, W, width of leaf blade. On the right side a shaded leaf is shown in comparison to a “normal” leaf (ecotype Col-0). (d) Schematic drawing of the branching pattern of Arabidopsis. A1, main inflorescence; B1, side branch from the rosette. Additional numbers indicate axillary branches of higher orders. (e) Detail of a node, with a cauline leaf and an axial branch (ecotype Col-0). Note that the next cauline leaf emerges in an angle of about 120°. (f) Detail of a flower and several buds (ecotype Col-0) (see Color Plates ).

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all seedlings will grow in a straight fashion, which makes the measurement difficult. 5. Hypocotyl length can be expressed in absolute numbers (mm) or as % elongation compared with dark-grown seedlings. The latter allows a better comparison between different experiments, but can only be used if the hypocotyl elongation in darkness is as that in wild type. 3.6.5. Root Development

The establishment of root architecture is a tightly regulated process. New cells for root growth originate from the distal tip and lateral roots are formed from the pericycle cells (13,14,15). Root branching is modulated by auxin, but other aspects of root architecture, such as root hair development, are independent of auxin action. Root hairs are cylindrical outgrowths from root epidermal cells and are formed in rows at specific locations (16). The number of lateral roots is dependent on the availability of nutrients in the media; therefore, the root architecture will vary in media with or without sucrose. Furthermore, when mineral elements are scarce, plants often allocate a greater proportion of their biomass to the root system (17). This acclimatory response is a consequence of metabolic changes in the shoot and an adjustment of transport to the root. When plants are N- or P-deficient, root growth is accelerated and an increase in lateral root formation can be observed. 1. Root apical growth is best evaluated when seedlings are grown vertically. 2. Five- to seven-day-old seedlings are transferred to plates, spaced at 5–10 mm along a line in the top third of the plate with enough room for the roots to grow downward, and with a control on the same plate. The tip of the root is marked on the back with a dot. 3. Seedlings are grown in a vertical position and the progress should be monitored at 24-h intervals for 3–5 days. Then the length of growth can be determined by photographic analysis (Fig. 2.2b). 4. The plates can be left in vertical position until day 7–10, when the number of lateral roots can be counted and root hairs visually analyzed using the stereo microscope.

3.6.6. Gravity Assays

Amyloplast movement along the gravity vector, within gravitysensing cells in the root and shoot, is the most likely trigger of subsequent intracellular signaling, in which differential auxin distribution within the sensing cells plays a role (18). 1. Gravity assays are performed on 5-day-old etiolated hypocotyls, which have been grown on vertical plates or on roots of 7- to 10-day-old plants grown on vertical plates.

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2. The root tips or the tip of the hypocotyl is marked with a dot on the back side of the plate. 3. The plates are turned for 90° and the changed orientation of the organs is measured after 5–15 h. The angle can either be directly measured from the back of the plate or from a digital image. 3.6.7. Phototrophic Assay

Phototropism, the directional curvature of organs in response to lateral differences in light, is mainly perceived by the photoreceptor phototropin, and also by phytochrome A (12, 19). Growth toward a source of light is called positive phototropism (e.g., tips of shoots), and that away from the source is termed negative phototropism (e.g., roots). Changes in cell elongation rates across the bending organ, established through a lateral gradient of auxin across the organ, can lead to these phototropic curvatures. 1. Seedlings are pre-grown for 4 days on vertical plates in the dark to generate elongated hypocotyls. 2. The tips of the hypocotyls are marked with a dot on the back side of the plate. 3. Plants are subjected to unilateral light, preferentially blue light (0.1 µE photons m−2 s−1), for 2–8 h. The angle of the tropic re-orientation of etiolated hypocotyls is determined from digital images taken of the plants on the plate.

3.6.8. Physical Parameters of Rosette Leaves

Plants exhibit plasticity in leaf shape and structure, allowing them to optimize photosynthetic efficiency. In Arabidopsis several types of leaves develop differentially, according to light intensity and quality. Lateral leaf expansion is controlled not only by regulation of the cell cycle, but also by meristematic activities that determine the dorso-ventral orientation of leaf primordia (20, 21). Reduced light intensity inhibits leaf lamina expansion, while enhancing petiole elongation (22). This phenomenon is a component of the so-called shade-avoidance syndrome. Under low light, Arabidopsis develops “shade leaves” with only one layer of palisade tissue, whereas under high light, it develops “sun leaves” that have nearly two complete layers of palisade tissue (23). The photoreceptor phytochrome B deficient mutant (phyB), for example, displays a typical constitutive shade-avoidance syndrome. The leaves from well-developed rosettes of similar age and equivalent developmental stage (between 2- and 4-weeks-old plants) are compared to evaluate their physical parameters (see Note 6). 1. Leaves are removed from the plant and scanned in a flatbed scanner. 2. Parameters for evaluation from the digital image include (Fig. 2.2c)

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– Leaf area (mm2) – Length of the petiole – Length of the leaf blade – Width of leaf blade 3. To compare and quantify the data the following ratios can be calculated: – Leaf blade length/petiole length – Blade length/width 3.6.9. Leaf Vein Pattern

In Arabidopsis, leaf venation patterning is an early, progressive, and hierarchical process (24). The venation pattern of cotyledons is simple, while true leaves have more branch points per lamina area. 1. To visualize leaf venation patterns excised leaves are submerged and kept overnight in a clearing solution (80 g chloral hydrate in 30 ml water) until the tissue becomes transparent. 2. Whole leaves are mounted on slides in a solution of 80 g chloral hydrate, 20 ml glycerol, and 10 ml water. 3. Transmitted-light digital photographs are analyzed according to the numbers of branch points.

3.6.10. Epidermal Cell Shape and Guard Cell Position

The contours of individual cells, including guard cells, are easily distinguishable and allow the measurement of cell size, shape, and the positioning of guard cells. Stomata are not found adjacent to each other but with intermittent cells (25). To analyze the structure of leaf surface an imprint can be generated (26). 1. A drop of 2% (w/v) low-melt agarose containing 0.01% (w/v) bromophenol blue is pre-warmed at 50°C and placed on a glass slide. 2. The leaf is gently pressed into the gel. 3. After the gel solidifies, the leaf is peeled of and the gel is dried for another 10 min. 4. The gel cast can be analyzed without a cover glass under a differential phase contrast microscope.

3.6.11. Trichomes (Leaf Hairs)

Trichomes are best evaluated under the microscope. Here the form (number of branches) and the distribution are important. Changes in the form of the trichomes are often an indication of changes in the cytoskeleton, whereas changes in the development and numbers of branches of trichomes are often related to aberrations in the cell cycle (16, 27). The presence of trichomes on the adaxial or abaxial side of the leaves and especially cauline leaves has also been reported to be correlated to reproductive development and gibberellin levels. Juvenile leaves lack trichomes on their abaxial (lower) surface,

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mature leaves have trichomes on both surfaces, and cauline leaves may have few or no trichomes on their adaxial (upper) surface (28). 3.6.12. Leaf Color

Leaves should appear green and be able to perform photosynthesis. If the leaves are either albinotic, pale green, or variegated, they are most probably blocked at some stage of chloroplast biogenesis and/or photosynthesis. It is necessary to include sucrose in the medium to maintain most pale mutants. A rough estimate for chloroplast integrity can be obtained from chlorophyll levels. 1. Take leaf discs (1 cm diameter) from leaves, avoiding major veins to maintain consistent sampling. At least three parallel assays should be performed. 2. Weight leaf discs. 3. Extract chlorophyll by adding 1 ml of 80% acetone to each leaf disc cut into small pieces. 4. Place the test tubes in a dark place to avoid degradation of chlorophyll and incubate over night. If the extraction is not complete grind the leaf tissue with sand. 5. After centrifugation the pellet should appear white; otherwise, re-extract the tissue. 6. Measure absorbance of the supernatant at 663 and 645 nm against 80% acetone. Quantify chlorophyll for each sample using the following standards (29): Chlorophyll a (µg/ml) = 12.7 × E663 − 2.69 × E645 Chlorophyll b (µg/ml) = 22.9 × E645 − 4.68 × E663 Express on a fresh-weight basis. The total chlorophyll content can be calculated by the following formula: Chlorophyll a + b (µg/ml) = 20.2 × E645 + 8.02 × E663 The ratio of chlorophyll a to chlorophyll b in wild type is around 2.8–3. Mutants that are strongly impaired in photosynthesis can be detected because the absorbed light cannot be utilized and is emitted as red fluorescence light (30). Consequently, these mutants can be scored for by using a long-wavelength UV lamp or a source of blue light. 1. Place plants in the dark for 15 min. 2. Illuminate plants with a UV hand-lamp (365 nm). 3. Chlorophyll fluorescence can be detected by its pinkish color (see Note 7).

3.6.13. Number of Leaves

Arabidopsis has three leaf types: cotyledons, rosette leaves, and cauline leaves. Leaf number can be determined from adult stage plants grown under the same conditions. Plants have different leaf numbers when grown under short or long days, or under

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different temperatures (31). The ratio between total leaf number/ cauline leaf number or just absolute number of leaves 20 days after flowering can be compared. Remember to include the senescent and dried up leaves in the count. 3.6.14. Flowering Time

Primary environmental factors such as day length and temperature control flowering time in plants (32–35). In Arabidopsis thaliana, a number of signaling pathways that control flowering time have been identified: light signal transduction pathway (phytochrome, cryptochromes), vernalization pathway, and hormones such as gibberellins or ethylene play a role. Therefore, many photoreceptor, light signaling, and photoperiod mutants are also affected in flowering time. As a starting point flowering time in long days should be analyzed. Plants are grown in soil and at bolting (the first visible appearance of the inflorescence stem) the number of rosette leaves and the number of days until bolting is counted. Although it has recently become more common to use only the number of leaves to determine flowering time, including the number of days gives a better evaluation. In some cases counting leaves is difficult, if you do not want to destroy the plant, as senescent or rotten leaves are difficult to see underneath the rosette. In a second assay the flowering time under short days (8 h light/16 h dark) can be tested in a similar fashion. Flowering time is usually much prolonged under short-day conditions compared with long day. Because even a short flash of light during the dark period can alter the flowering time, it is essential that the plants be kept in total darkness during the dark period.

3.6.15. Stem Length

A number of factors can lead to a change in the Arabidopsis stature. Plants that possess reduced levels of gibberellin are dwarves whereas elevated gibberellin levels lead to an elongated spindly phenotype (36). Nonetheless, a dwarfish phenotype can also be observed in plants lacking essential nutrients, plants under constant stress, or under constant challenge by an overactive defense system. Plant size can also depend on the control of cell size, which is often related to the ploidy level of a cell. The overall length of the flowering plant can be measured with a ruler. To compare plants they should be of the same age or at the end of their life cycle, so that no further growth is to be expected. Additionally, the length of the main inflorescence stem can be measured. If the overall plant length is longer than the main bolt, then apical dominance is impaired. This usually leads to bushier plants (see Sect. 3.6.16).

3.6.16. Branching Pattern

In Arabidopsis, as in all seed plants, the primary axis of growth is developed during embryonic development. During postembryonic shoot development, side shoots are established from

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lateral meristems in the axils of leaves (37, 38) . Both the initiation of axillary meristems and the regulation of bud outgrowth are tightly regulated. Suppression of bud outgrowth is due to an inhibitory effect of the primary shoot apex, namely apical dominance (39). Auxin produced in the main shoot tip plays a major part in repressing bud outgrowth whereas cytokinin is considered to be an activator of shoot development. Other unknown hormones may also play a role in these processes. In Arabidopsis axillary branches can derive from the rosette, which develop after the main inflorescence stem has bolted or from the main inflorescence stem (Fig. 2.2d). These side branches can develop additional side branches of a higher order. Outgrowth of a second axillary meristem in the leaf axils is not usual (Fig. 2.2e). The bushiness of a plant can be illustrated by a digital image of the adult plant and evaluated by counting the number of side branches of each order, separately for the main inflorescence (Fig. 2.2d, A1) and the side stems from the rosette (Fig. 2.2d, B1). To uncouple the outgrowth of the lateral shoots during vegetative development (B1) from the apical dominance, adult plants can be decapitated 5 days after their bolting and the appearance of new shoots from the rosette can be counted after 10 days. 3.6.17. Phyllotaxis

In the growing plant shoot, new leaf and flower primordia emerge at well-defined positions, resulting in regular patterns (phyllotaxis). Whereas in Arabidopsis the cotyledons are opposed to each other with an angle of about 180° (decussate leaf position), the subsequent nodes (leaves and flowers) are in a spiral pattern (single primordia that are created sequentially) (40, 41). Changes in the phyllotaxis are easiest to be seen when the position of the cauline leaves or the siliques are observed (Fig. 2.2e). The subsequent node should be in a 120° angle to the previous one.

3.6.18. Flower and Fruit Morphology

Wild-type Arabidopsis flowers have, from the outside to the inside, 4 sepals (green), 4 petals (white), 6 stamens, and 2 carpels, which fuse to form the central pistil (Fig. 2.2f). To check the phenotype of flowers a stereo microscope should be used. The flower can be dissected with thin forceps. The presence of all organs in their correct size, shape, and number should be compared with a wild-type flower (42, 43).

3.6.19. Silique Morphology

The Arabidopsis fruit is a silique or pod (43). The length of the siliques can be measured with a ruler. The number of seeds within each silique can be counted. To determine the weight of a large number of seeds, use the weight of a pre-counted number of seeds (e.g., 25). Smaller siliques usually suggest reduced fertility. Small siliques with no seeds often imply a problem with the fertilization during the flower stage. By prying open the siliques aborted ovules should be visible.

Phenotyping of Arabidopsis Developmental

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3.6.20. Seed Morphology

After harvesting of the seeds several parameters can be observed: Seed size, color, and surface. Cytokinin is one of the factors that influences seed size (44), whereas deficiencies in anthocyanin biosynthesis lead to light brown or yellow seeds. If insufficient storage material was deposited, the seeds show wrinkles. The fertility of the seeds can then be tested in a germination assay (see Sect. 3.6.2).

3.6.21. Progression of Senescence or Leaf Browning

Senescence in green plants is a complex and highly regulated process that occurs as part of plant development or can be prematurely induced by stress or darkness (45, 46). (a) Whole plant senescence: 1. Grow plants for 3 weeks under long-day conditions (4–6 plants per pot). 2. Transfer plants to the dark for 6 days by wrapping the plants in aluminum foil. Determine chlorophyll content during this time every second day (see Sect. 3.6.12). 3. After growth in the dark, transfer plants back to longday conditions for 8 days. 4. Calculate survival rates after 8 days in WL. (b) Senescence in detached leaves: 1. Detach fully expanded leaves from 3-week-old plants and place them in the dark in Petri dishes with wet tissue paper. 2. Take images of leaves after 0, 3, and 4 days of dark treatment. 3. Furthermore, measure and calculate chlorophyll content according to the fresh weight (see Sect. 3.6.12).

4. Notes 1. The penetrance, especially of developmental phenotypes, is not always complete; therefore, it is important to work with homozygous lines to avoid confusion with a segregating population. 2. There are several ways to test whether the insertion in the gene of interest is the cause of the phenotype: (1) By crossing the homozygous insertion line to wild type and looking for co-segregation of the phenotype with the insertion or point mutation. If the mutation is the cause of the phenotype, they have to co-segregate. However, this test cannot exclude the possibility that the phenotype is caused by a second mutation that is closely linked to the insertion. (2) If two or more mutations in the same gene show a similar

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phenotype, the chances are very high that this phenotype is caused by this gene. Therefore, it is highly recommended to work, whenever possible, with several different alleles. (3) Complementation of plants homozygous for insertion with the wild-type version of the gene of interest. If the phenotype reverts to wild type, the mutation in the gene of interest must have been the cause of the phenotype. For this test the choice of the promoter under which the transgene is placed may be vital. The expression under its own promoter is the best, but the promoter fragment chosen might not carry all the regulatory signals. The CaMV 35S promoter might lead to ectopic expression and overexpression phenotypes, as it is a very strong promoter. 3. If no phenotype can be detected during normal growth, the plants can be challenged with stressors and hormones, while their phenotypic behavior is observed (47). 4. Observed phenotypes, especially pleiotropic ones, may arise for different reasons, which may or may not relate to the underlying defect studied. On the other hand, the observation of one phenotype and the resulting overlooking of other phenotypes can lead to wrong conclusions. It might be helpful to correlate the expression analysis from publicly available microarray data (48, 49) with the phenotype, to see if the gene is expressed under the developmental stage or in the tissue that shows a defect.Phenotypes might not be visible if similar genes are still expressed, whose proteins can substitute for the missing protein. This is often the case with protein families. In this case only the loss of both genes (double mutants) might yield a phenotype. 5. Note that it is important for the far-red light to exclude light below 700 nm. To reduce the (white) light intensity move the light source further away or cover the plates with sheets of white tissue paper. 6. Leaves within a rosette can be counted from the oldest to the youngest. It is helpful to always compare leaves of a similar stage, such as the fourth or sixth leaf appearing at bolting. 7. Note that some secondary metabolites can fluorescence in a strong reddish color.

Acknowledgments I would like to thank Dario Leister for his support and critical reading of the manuscript. I am grateful to Randy Foster for critical discussions and help with the manuscript. I would also like

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to thank numerous colleagues with whom I discussed different techniques. This work was supported by grants from the Deutsche Forschungsgemeinschaft (DFG, BO 1146/3; Arabidopsis Functional Genomics Network, AFGN, BO 1146/4) to C.B. References 1. The Arabidopsis Genome Initiative (AGI) (2000) Analysis of the genome sequence of the flowering plant Arabidopsis thaliana. Nature 408, 796–815. 2. Alonso, J.M., and Ecker, J.R. (2006) Moving forward in reverse: Genetic technologies to enable genome-wide phenomic screens in Arabidopsis. Nature Rev. Genet. 7, 524–536. 3. Østergaard, L., and Yanofsky, M.F. (2004) Establishing gene function by mutagenesis in Arabidopsis thaliana. Plant J. 39, 682–696. 4. Henikoff, S., Bradley, J.T., and Comai, L. (2004) TILLING. Traditional Mutagenesis Meets Functional Genomics. Plant Physiol. 135, 630–636. 5. Murashige, T., and Skoog, F. (1962) A revised medium for rapid growth and bioassays with tobacco tissue cultures. Physiol. Plant. 15, 473–497. 6. Gubler, F., Millar, A.A., and Jacobsen, J.V. (2005) Dormancy release, ABA and preharvest sprouting. Curr. Opin. Plant Biol. 8, 183–187. 7. Koornneef, M., Bentsink, L., and Hilhorst, H. (2002) Seed dormancy and germination. Curr. Opin. Plant Biol. 5, 33–36. 8. Razem, F.A., Baron, K., and Hill, R.D. (2006) Turning on gibberellin and abscisic acid signaling. Curr. Opin. Plant Biol. 9, 454–459. 9. Penfield, S., Graham, S., Graham, I.A. (2005) Storage reserve mobilization in germinating oilseeds: Arabidopsis as a model system. Biochem. Soc. Trans. 33, 380–383. 10. Footitt, S., Slocombe, S.P., Larner, V., Kurup, S., Wu, Y., Larson, T., Graham, I., Baker, A., and Holdsworth, M. (2002) Control of germination and lipid mobilization by COMATOSE, the Arabidopsis homologue of human ALDP. EMBO J. 21, 2912–2922. 11. Vicente-Carbajosa, J., and Carbonero, P. (2005) Seed maturation: developing an intrusive phase to accomplish a quiescent state. Int. J. Dev. Biol. 49, 645–651. 12. Fankhauser, C., and Casal, J.J. (2004) Phenotypic characterization of a photomorphogenic mutant. Plant J. 39, 747–760.

13. Casimiro, I., Beeckman, T., Graham, N., Bhalerao, R., Zhang, H., Casero, P., Sandberg, G. and Bennett, M.J. (2003) Dissecting Arabidopsis lateral root development. Trends Plant Sci. 8, 165–171. 14. Benfey, P.N., and Scheres, B. (2000) Root development. Curr. Biol. 10, R813–815. 15. Fukaki, H., Okushima, Y., and Tasaka, M. (2007) Auxin-mediated lateral root formation in higher plants. Int. Rev. Cytol. 256, 111–137. 16. Schellmann, S., Hulskamp, M., and Uhrig, J. (2007) Epidermal pattern formation in the root and shoot of Arabidopsis. Biochem. Soc. Trans. 35, 146–148. 17. Hermans, C., Hammond, J.P., White, P.J., and Verbruggen, N. (2006) How do plants respond to nutrient shortage by biomass allocation? Trends Plant Sci. 11, 610–617. 18. Morita, M.T., and Tasaka, M. (2004) Gravity sensing and signaling. Curr. Opin. Plant Biol. 7, 712–718. 19. Stone, B.B., Esmon, C.A., and Liscum, E. (2005) Phototropins, other photoreceptors, and associated signaling: The lead and supporting cast in the control of plant movement responses. Curr. Top. Dev. Biol. 66, 215–238. 20. Tsukaya, H. (2005) Leaf shape: genetic controls and environmental factors. Int. J. Dev. Biol. 49, 547–555. 21. Tsukaya, H. (2006) Mechanism of leafshape determination. Annu. Rev. Plant Biol. 57, 477–496. 22. Tsukaya, H., Kozuka, T., and Kim, G.T. (2002) Genetic control of petiole length in Arabidopsis thaliana. Plant Cell Physiol. 43, 1221–1228. 23. Kim, G.T., Yano, S., Kozuka, T., and Tsukaya, H. (2005) Photomorphogenesis of leaves: shade-avoidance and differentiation of sun and shade leaves. Photochem. Photobiol. Sci. 4, 770–774. 24. Candela, H., Martinez-Laborda, A., and Micol, J.L. (1999) Venation pattern formation in Arabidopsis thaliana vegetative leaves. Dev. Biol. 205, 205–216. 25. Nadeau, J.A., and Sack, F.D. (2002) Control of stomatal distribution on the Arabidopsis leaf surface. Science 296, 1697–1700.

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26. Horiguchi, G., Fujikura, U., Ferjani, A., Ishikawa, N., and Tsukaya, H. (2006) Largescale histological analysis of leaf mutants using two simple leaf observation methods: Identification of novel genetic pathways governing the size and shape of leaves. Plant J. 48, 638–644. 27. Schellmann, S., and Hulskamp, M. (2005) Epidermal differentiation: trichomes in Arabidopsis as a model system. Int. J. Dev. Biol. 49, 579–584. 28. Chien, J.C., and Sussex, I.M. (1996) Differential regulation of trichome formation on the adaxial and abaxial leaf surfaces by gibberellins and photoperiod in Arabidopsis thaliana (L.) Heynh. Plant Physiol. 111, 1321–1328. 29. Arnon, D.I. (1949) Copper enzymes in isolated chloroplasts. Polyphenoloxidase in Beta vulgaris. Plant Physiol. 24, 1–15. 30. Miles, D. (1982) The use of mutations to probe photosynthesis in higher plants. In: Edelman, M., Hallick, R.B., and Chua, N.H., (eds.) Methods in Chloroplast Molecular Biology. Elsevier Biomedical Press, New York, pp. 75–107. 31. Lempe, J., Balasubramanian, S., Sureshkumar, S., Singh, A., Schmid, M., and Weigel, D. (2005) Diversity of flowering responses in wild Arabidopsis thaliana strains. PLoS Genet. 1, 109–118. 32. Baurle, I. and Dean, C. (2006) The timing of developmental transitions in plants. Cell 125, 655–664. 33. Ausin, I., Alonso-Blanco, C., and MartinezZapater, J.M. (2005) Environmental regulation of flowering. Int. J. Dev. Biol. 49, 689–705. 34. Boss, P.K., Bastow, R.M., Mylne, J.S., and Dean, C. (2004) Multiple pathways in the decision to flower: enabling, promoting, and resetting. Plant Cell 16(Suppl.), S18–S31. 35. Corbesier, L., and Coupland, G. (2006) The quest for florigen: A review of recent progress. J. Exp. Bot. 57, 3395–3403. 36. Fleet, C.M. and Sun, T.P. (2005) A DELLAcate balance: the role of gibberellin in plant morphogenesis. Curr. Opin. Plant Biol. 8, 77–85.

37. Schmitz, G. and Theres, K. (2005) Shoot and inflorescence branching. Curr. Opin. Plant Biol. 8, 506–511. 38. McSteen, P., and Leyser, O. (2005) Shoot branching. Annu. Rev. Plant Biol. 56, 353–374. 39. Leyser, O. (2005) The fall and rise of apical dominance. Curr. Opin. Genet. Dev. 15, 468–471. 40. Reinhardt, D. (2005) Phyllotaxis--a new chapter in an old tale about beauty and magic numbers. Curr. Opin. Plant Biol. 8, 487–493. 41. Reinhardt, D. (2005) Regulation of phyllotaxis. Int. J. Dev. Biol. 49, 539–546. 42. Blazquez, M.A., Ferrandiz, C., Madueno, F., and Parcy, F. (2006) How floral meristems are built. Plant Mol. Biol. 60, 855–870. 43. Robles, P., and Pelaz, S. (2005) Flower and fruit development in Arabidopsis thaliana. Int. J. Dev. Biol. 49, 633–643. 44. Riefler, M., Novak, O., Strnad, M., and Schmulling, T. (2006) Arabidopsis cytokinin receptor mutants reveal functions in shoot growth, leaf senescence, seed size, germination, root development, and cytokinin metabolism. Plant Cell 18, 40–54. 45. Buchanan-Wollaston, V., Earl, S., Harrison, E., Mathas, E., Navabpour, S., Page, T., and Pink, D. (2003) The molecular analysis of leaf senescence - a genomics approach. Plant Biotechnol. J. 1, 3–22. 46. Lim, P.O., and Nam, H.G. (2005) The molecular and genetic control of leaf senescence and longevity in Arabidopsis. Curr. Top. Dev. Biol. 67, 49–83. 47. Bolle, C. (2008) Phenotyping of abioticresponses and hormone treatments in Arabidopsis. 48. Zimmermann, P., Hirsch-Hoffmann, M., Hennig, L. and Gruissem, W. (2004) GENEVESTIGATOR. Arabidopsis microarray database and analysis toolbox. Plant Physiol. 136, 2621–2632. 49. Toufighi, K., Brady, S.M., Austin, R., Ly, E., and Provart, N.J. (2005) The botany array resource: e-Northerns, expression angling, and promoter analyses. Plant J. 43, 153–163.

Chapter 3 Phenotyping of Abiotic Responses and Hormone Treatments in Arabidopsis Cordelia Bolle Abstract The disruption or modulation of signal transduction pathways does not always lead to drastic changes in plant growth and development. Therefore, many loss- or gain-of-function lines do not exhibit an obvious phenotype under normal greenhouse conditions. To be able to assign biological functions to these genes, the mutants need to be evaluated with a broad spectrum of assays to uncover conditional phenotypes. Here we provide an overview on how to evaluate plants in their development and their response to abiotic factors such as light, hormones, and different stressors. The assessment of the behavior of a plant under these conditions can be used to correlate a biological role with a genotype. This phenotypic analysis can be used for profiling of mutants. Key words: Phenotype, reverse genetics, signaling, abiotic stress, hormones, mutant analysis.

1. Introduction As reverse genetics projects become more and more popular due to the availability of collections of gene-specific knock-out mutants, the need to find phenotypes in these lines arises. Only a minority of mutants presents easily detectable phenotypic variations; therefore, a close observation of developmental deficiencies (1) and a broad spectrum of environmental and abiotic assays may be needed to uncover conditional phenotypes. Plants have a great potential to adapt their growth and development to changing environmental conditions. This phenotypic plasticity is also what makes changes in development in loss-of-function

T. Pfannschmidt (ed.), Plant Signal Transduction, vol. 479 Humana Press, a part of Springer Science + Business Media, LLC 2009 DOI: 10.1007/978-1-59745-289-2_3

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lines or gain-of-function lines difficult to detect. One option is to challenge plants with conditions that bring them to their extent of adaptability, evaluating developmental phases that are most vulnerable to stress, because under these conditions phenotypes might be more obvious. The growth and differentiation of plants are continuously adjusted to a multitude of environmental factors. Among the many abiotic factors are light, water, temperature, gravity, wind, oxygen, nutrients, and chemicals. Biotic factors are other organisms involved in symbiotic, pathogenic, or herbivorous interactions with plants. All of these environmental factors are recognized by plants, and have been identified for many receptors. In order to ensure adaptation to the environment, signals generated following perception need to be integrated by crosstalk between different signaling pathways. Hormones, light, and abiotic stresses trigger signal transduction cascades that control growth, development, and survival of plants, intersecting and influencing one another. Here we present protocols to use plants’ responses to abiotic factors such as light, hormones, and different stressors to assess alterations from wild type development. The assessment of the behavior of a plant under these conditions can be used to correlate a biological role with a genotype. This phenotypic analysis can be used for the profiling of mutants and thereby assigning then to specific signaling pathways. For a more detailed analysis of the loss-of-function and gain-of function lines the expression of well-described marker genes specific for different pathways can be analyzed by using the above described conditions.

2. Materials 1. Plates for tissue culture: Round, sterile plastic Petri dishes with a diameter of 145 mm are best for most experiments; otherwise, small Petri dishes can also be used. Some producers add a grid on the bottom lid, which helps with orientation and alignment. 2. Media plates: Half-strength (0.5X) Murashige and Skoog (MS) medium (2) is autoclaved together with 0.8% (w/v) agar at 121°C for 15 min and a magnetic stirring bar. This medium contains usually no sucrose as sucrose interferes with many signal transduction pathways. For root length assays performed in darkness add 1% sucrose to media. The medium is cooled down to 60°C and non-autoclaveable additives can be added (see Table 3.1 and Note 1). The medium should be stirred well before it is poured into the Petri dishes. After the plates have solidified and have been dried of excessive condensated

Synthetic cytokinin Osmotic stress Jasmonate

Kinetin

Mannitola

MeJa

Methyl-jasmonate

Indole-3-butyric acid

IBA

Auxin

Auxin

Indole-3 acetic acid

IAA

Brassinosteroid biosynthesis inhibitor

Gibberellin

(15)

Brz220

Brassinosteroid

GA4

Brassinolide

BL

Synthetic cytokinin

Gibberellin

Benzyladenine

BA

Ethylene inhibitor

GA3

Aminoethoxyvinyl-glycine

AVG

Ethylene inhibitor

ABA biosynthesis inhibitor

Silver nitrate

AgN03a

Ethylene precursor

Fluridone

1-Aminocyclopropane1-carboxylic acid

ACC

ABA

Cytokinin

Abscisic acid

ABA

synthetic Auxin

Function/pathways

cis-zeatin

2,4-Dichloro-phenoxyacetic acid

2,4-D

Chemical component

Table 3.1 Hormones and other chemicals added to the media for assays

EtOH

Water

1 N NaOH

EtOH or 1 N NaOH

EtOH

EtOH

EtOH

EtOH

1 N NaOH or DMSO

DMSO or 80% EtOH

DMSO or 80% EtOH

1 N NaOH or DMSO

Water

Water

EtOH

1 N NaOH or methanol

water (sodium salt), otherwise: 1 N NaOH

Dissolve inc

(continued)

500 mM in water −20°C

Directly added

100 mM in water −20°C

100 mM in water −20°C

100 mM in water −20°C

100 mM in water −20°C

100 mM in water −20°C

100 mM in water −20°C

100 mM in water −20°C

100 mM in water −20°C

100 mM in water −20°C

100 mM in water −20°C

100 mM in water −20°C

Add directly

100 mM in water −20°C

10 mM in water −20°C

1 mM in water 4°C

Stock solution storage

Responses to Hormones and Abiotic Stresses 37

Paclobutrazol

Polyethyleneglycol

PAC

PEG 6000 or 8000

Auxin transport inhibitor

TIBA Gibberellin biosynthesis inhibitor

Osmotic stress

Sucrosea

Uniconazole

Osmotic stress

Sorbitola

Auxin transport inhibitor

Salt stress

Water

1 N NaOH

Water

Water

Dissolved in autoclaved 0.5× MS media without agar sterilized by filter (0.22 µm)b

Water

EtOH DMSO

Water

1 N NaOH

Water

Dissolve inc

100 mM in water −20°C

100 mM in water −20°C

Directly added

Directly added

Double of desired final concentration

100 mM in water −20°C

100 mM in water −20/4°C

Directly added

100 mM in water −20/4°C

100 mM in water −20°C

Stock solution storage

b

Can be autoclaved in the media, all other additives have to be added after the media has cooled down to about 60°C. 0.5X MS media plates are covered with a PEG solution for 24 h to allow diffusion of the PEG into the media. For a media plate with 20 ml of solid media, 20 ml of PEG solution with the double concentration of the desired final concentration on the plate is added. After 24 h, the PEG solution is poured off and the plates are ready for use. c Can vary depending on the producer.

a

Osmotic stress

Naphthylphthalamic acid

NPA

2,3,5-Triiodobenzoic acid

Gibberellin biosynthesis inhibitor

Sodium chloride

NaCl2a

Synthetic auxin

1-Naphthaleneacetic acid

NAA

Oxidative stress

Function/pathways

Paraquat

Chemical component

Methyl-viologen

Table 3.1 (continued)

38 Bolle

Responses to Hormones and Abiotic Stresses

39

water they can be stored for up to 1 month at 4–10°C (without additives up to 2–3 months). Before using plates stored in the cold, they should be dried again in a sterile hood to remove excess water. For vertical plates increasing the concentration of agarose in the media to 1% (w/v) is advisable. 3. Sealing the plates: Bandage tape such as Leukopor or Micropore should be used to close the lids of the Petri dishes because in contrast to parafilm it allows gas exchange. 4. Sterilization solution for seeds: a. 30% (v/v) Clorex solution (or any other commercial bleach that contains hypochloride) with 0.05% (v/v) Tween 20 b. Sodiumhypochloride, HCl conc. (Be careful, use gloves, and protective goggles)

3. Methods 3.1. Generation of Homozygous Lines

Before working with mutants lines generated by insertion or point mutation, it is important to remove putative additional mutations from the line by repeated backcrossing to wild type. For this purpose the mutant line is used to pollinate a wild type of the same ecotype. The progeny is then tested for the presence of the mutation and is used again to pollinate the wild type. This cycle can be repeated for several times. The progeny is finally analyzed for the presence of the mutation and selfed. In the next generation homozygous lines can be selected, which can then be used for phenotypic characterization. For methods to confirm that the observed phenotype is correlated to a specific gene, see notes in (1).

3.2. Surface Sterilization of Seeds

If seeds are to be germinated on medium containing plates they must be sterilized first. Two possible methods are as follows.

3.2.1. Liquid Sterilization

1. Seeds are placed for 10 min in a 30% (v/v) Clorex solution (or any other commercial bleach that contains hypochloride) with 0.05% (w/v) Tween 20 and agitated every 30 s. About 300 µl solution in a 1.5-ml reaction tube is suitable for 100 seeds. 2. After 10 min the seeds are briefly centrifuged to help settle them and the solution is removed. The seeds are washed three times with sterile water. 3. Sterile 0.1% (w/v) agarose can be added to the seeds to facilitate distribution of the seeds on media plates with a pipette.

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3.2.2. Vapor Sterilization

1. A maximum of 100 seeds is added to each reaction tube; otherwise, the penetration of the vapor is not guaranteed. 2. The reaction tubes are placed with their lid open in an desiccator together with a beaker containing 100 ml sodium hypochloride into which 5 ml HCl conc. is added drop wise (use gloves!). 3. After 5 h treatment the seeds are placed in a sterile hood to remove the fumes. The advantage of this method is that the seeds are dry and can be stored in a sterile way for further use. The dry seeds can be distributed on media plates by tapping the tube.

3.3. Stratification of Seeds

Stratification (break of seed dormancy) of imbibed seeds by cold (2–4 days at 4–10°C in darkness) improves coordinated germination, which is important for most phenotypic assays as it allows comparing seedlings of the same age. Stratification is best performed after the seeds have been placed on media. To keep the plates dark they can be wrapped in aluminum foil.

3.4. Growth Conditions

Controlled growth conditions (light intensities and spectra, moisture and temperature) are important for reproducible results. Growth chambers are usually easier to control than greenhouses. However, be aware that conditions within a growth chamber can also be variable. White light should be maintained at around 100–300 µmol photons m−2 s−1, temperature between 18–22°C and humidity at 50–70%. For most tests long-day conditions are recommended, with 16 h light and 8 h dark. Most assays are performed in tissue culture, when all organs (especially roots) are easily accessible for analysis under the microscope and additives can be added in a controlled way. Plants that are grown on soil can either directly be sown onto soil (3 seeds per pot, thinned out to one plant per pot upon germination) or pre-grown in tissue culture without antibiotics. Seedlings should be transferred from tissue culture to soil when they have 4–6 true leaves. To minimize any spurious differences between the mutant and wild type caused by different growth conditions, mutant and wild type plants should be grown next to each other in the same growth facility. They should be maintained on the same media plate or, when in soil, on the same tray. A quick test for a mutant phenotype can be carried out with 12 parallel plants. However, for the confirmation of a phenotype the experiments have to be repeated at least three times with an adequate number of parallel plants to generate statistically significant data.

3.5. Recording the Mutant Phenotype

A visual record of a mutant phenotype can be captured with a digital or a video camera fixed to a support. This allows us to take images of different plates or pots of a series from the same distance therefore with comparable sizes. Cold white light from

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the sides reduces shadows and is necessary because a flash cannot be used to photograph media plates (reflections). To heighten the contrast black felt can be put underneath the objects. If green plants are to be photographed a red cloth can enhance the contrast. A ruler placed next to the objects enables the evaluation of physical parameters. The images can be imported into software such as ImageJ (NIH; http://rsb.info.nih.gov/ij/) which displays either area statistics, line lengths or angles. To measure length a “straight line,” a “segmented line,” or a “freehand line” can be used. The data can be processed in a program such as Microsoft Excel. The length of 1 cm of the ruler on the digital image can be used to calibrate the program or to calculate the actual lengths in Excel. All data should be statistically evaluated (mean value, standard deviation, t-Test, or similar) using Excel or similar programs. 3.6. Screen for a Phenotype Under Different Abiotic Conditions (see Note 2) 3.6.1. Germination Assay

1. To test germination efficiency seeds are plated on media plates. About 30–100 seeds per mutant line are optimal, which are evenly spaced (1 seed per 2 mm2) on the plate (see Note 3). Several lines can be placed on the same plate within different segments. 2. After stratification, plates are transferred to white light (16 h light/8 h dark; 21°C). Plates should be checked daily for germination (1–7 days) scoring both germinated and nongerminated seeds. This is done best with a stereo microscope with illumination from below. Non-germinated seeds will appear dark whereas empty seed coats appear translucent. A seed is scored as germinated as soon as the radicle has protruded and/or upon emergence of green cotyledons. 3. Germination is best evaluated by calculating germination efficiency without the respective additive, which is then assumed as 100% germination. Germination on the different concentrations of the additive can then be calculated as % germination.

3.6.2. Hypocotyl Length Assay

1. To measure hypocotyl length plate about 50 sterilized seeds per line on agar plates. About 4–8 different lines can be placed on one big plate, and one of them should be WT control. Seeds can be plated relatively dense (1 seed in 1–2 mm2), but the cotyledons should not obstruct or shade each other (see Note 4). 2. After stratification place seeds for 2–6 h in white light and then incubate in the dark (20°C) overnight. 3. Place the seedlings in the appropriate light conditions or keep them in the darkness and incubate for 4–6 days. 4. To measure the hypocotyl length the seedlings are stretched with the help of forceps longitudinally on a new agar plate,

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and this can be photographed with a digital camera (see Sect. 3.5). The length of the hypocotyl is measured from the apical meristem to the beginning of the root, often characterized by the still attached seed coat. 5. Hypocotyl length can be expressed in absolute numbers (mm) or as % elongation compared to control seedlings (either dark grown seedlings or seedlings on media without additives). The latter allows a better comparison between different experiments. 3.6.3. Root Growth Assay

1. Root apical growth is best evaluated when seedlings are grown vertically. 2. Five to seven-day-old light-grown seedlings are transferred to new plates, spaced at 5–10 mm along a line in the top third of the plate with enough room for the roots to grow downward, and with a control on the same plate. The tip of the root is marked on the back with a dot. 3. Seedlings are grown in a vertical position and the progress should be monitored at 24 h-intervals for 5 days; then the length of growth can be determined by photographic analysis (see Sect. 3.5). Root length can be expressed as absolute length or root growth inhibition. In the latter case root length of the control plants is set as 100% and those under varying conditions are expressed as % of the control. 4. The plates can be left in vertical position until day 7–10, when the number of lateral roots can be observed using the stereo microscope. The emerged lateral roots can be counted and the total length of lateral roots (cm) per seedling can be calculated.

3.6.4. Simultaneous Root Growth Assay on Several Conditions

Many of the conditions presented here can be assayed with the help of the root growth assay (see Sect. 3.6.3). Therefore, for a first, simpler screen 5-day-old seedlings can be transferred as described above to a variety of plates with different additives (see Table 3.2). About 12 seedlings per line are enough for an initial evaluation. In parallel to the root length, the number of lateral roots and hypocotyl elongation can be noted.

3.7. Influence of Hormones on Growth (see Note 5)

Abscisic Acid (ABA) is a sesquiterpenoid, similar to carotinoid, which is partially produced in plastids. Abiotic stresses, such as low water potential and salt stress, that cause water loss or reduced water uptake, drought, cold, and wounding are a major stimulus in eliciting ABA accumulation (see Sect. 3.8). ABA stimulates the closure of stomata, influences root and shoot growth, accumulates during the maturation of seeds, and has an effect on induction and maintenance of dormancy (3, 4, 5). Fluridone can be used as ABA biosynthesis inhibitor, but it is

3.7.1. Abscisic Acid (ABA)

Responses to Hormones and Abiotic Stresses

43

Table 3.2 Abiotic additives for a quick root growth assay. If not mentioned differently incubation of the vertical plates is at 21°C Chemical com- Pathway ponent/condition

Concentrations

Incubation in the light (L) or dark (D)

-

Control

-

L+D

ABA

ABA

2.5/25 µM

D

IAA or NAA

Auxin

0.1/1 µM

L

BL

Brassinosteroid

1 nM/1 µM

L

Kinetin or BA

Cytokinin

0.1/1 µM

D

ACC

Ethylene precursor

0.5 /5 µM

D

MeJa

Jasmonate

0.1 / 10 µM

L

NaCl

Salt stress

50 /150 mM

L

Mannitol

Osmotic stress

200/500 mM

L

Glucose/sucrose Osmotic stress

4/6% (w/v)

L

Methylviologen/ Oxidative stress paraquat

0.1/ 1 µM

L

14°C

Cold

L

27°C

Heat

L

not very specific as it inhibits carotinoid biosynthesis, leading to albino plants. As controls ABA-biosynthesis mutants such as aba1 can be used (6). To test for responses to ABA the following assays can be performed: a. Germination assay (see Sect. 3.6.1) 1. Germinate seeds on media containing different concentrations of ABA (0, 0.5, 1, 1.5, 2, 2.5, or 5 µM). 2. Evaluate the germination rate after 5 days. On 0.5 µM ABA wild type (Col-0) usually develops 50% green cotyledons, whereas on 1 µM ABA 50% radicle emergences can be observed. b. Root growth assay (see Sect. 3.6.3) 1. Transfer 5-day-old plants to vertical plates. Recommended condition for phenotypic analysis: media containing different concentrations of ABA (0, 2.5, 5, 10, 25, or 50 µM) or Fluridone (0, 1, or 10 µM). 2. Incubate for 5 days in the dark to test for root growth. At 25 µM ABA WT root growth is usually 50% inhibited.

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3.7.2. Auxin

Auxins stimulate cell elongation, cell division, and lateral root development and mediate the tropistic response of bending in response to gravity and light (7–9). The auxin supplied from the apical bud suppresses growth of lateral buds and auxin in the meristems determines the positions of primordia (for assays see Ref. 1). Indole-3-acetic acid (IAA) is derived from the amino acid tryptophan and is the only naturally occurring auxin. There are many synthetic auxins that are usually more stable: 1-naphthaleneacetic acid (NAA), indole-3-butyric acid (IBA), and 2,4-dichlorophenoxyacetic acid (2,4-D), which is also used as a herbizide. IAA and NAA are mainly used for assays; although they are similar in structure, they have slightly divergent responses, with NAA being less effective. The most common auxin transport inhibitors are naphthylphthalamic acid (NPA) and 2,3,5-triiodobenzoic acid (TIBA), although also p-chlorophenoxyisobutyric acid (PCIB) and 9-hydroxyfluorene-9-carboxylic acid (HFCA) can also be used. As controls the tir1 mutant, an auxin receptor mutant, can be used (10, 11). The artificial DR5 promoter coupled to a GUS reporter gene is an easy way to detect auxin distribution within a plant (12). To test for responses to auxins the following assays can be performed: a. Root growth assay (see Sect. 3.6.3) 1. Transfer 5-day-old plants to vertical plates. Recommended condition for phenotypic analysis: media containing different concentrations of IAA or NAA (0 - 0.01 - 0.1 - 1 - 5 µM) or 2,4 D (0 - 5 - 15 - 30 - 60 nM), NPA (5 - 10 µM), or TIBA (40 µM). 2. Incubate for 5 days in the light to test for root growth and count the number of lateral roots under the stereo microscope. At 1 µM IAA wild type root growth is usually 50% inhibited. b. Hypocotyl length assay (see Sect. 3.6.2) 1. Germinate seeds on a plate in the dark for 5 days. Recommended condition for phenotypic analysis: media containing different concentrations of IAA or NAA (0 - 0.1 - 1 10 - 100 µM). 2. Measure hypocotyl length. About 50% inhibition for wild type lies between 1–10 µM IAA or NAA.

3.7.3. Brassinosteroids

Brassinolide (BL) is the most bioactive form of the growthpromoting plant steroids termed brassinosteroids (BRs). These hormones are mainly involved in resistance to drought and cold weather, leaf bending, cell elongation, and cell division (13, 14). They interact with other hormones in stem elongation (auxin and gibberellins) and inhibit root elongation as a consequence of BR-induced ethylene synthesis. Many different brassinosteroids

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have been found; for the assays epi-brassinolide is the most commonly used form. Brz220 is a brassinosteroid biosynthesis inhibitor (15). As a control BR-deficient mutants such as det2 and cpd can be used, which appear dwarfish and have short petioles (16, 17). To test for responses to BL the following assays can be performed: a. Root growth assay (see Sect. 3.6.3) 1. Transfer 5-day-old plants to vertical plates. Recommended condition for phenotypic analysis: media containing different concentrations of epi-brassinolide (0 - 1 nM - 10 nM - 0.1 µM -1 µM - 5 µM). 2. Incubate for 5 days in the light to test for root growth. b. Hypocotyl length assay (see Sect. 3.6.2) 1. Germinate seeds on a plate in the dark or light for 5 days. Recommended condition for phenotypic analysis: media containing different concentrations of epibrassinolide (0 - 1 nM - 10 nM - 0.1 µM -1 µM), Brz220 (0 - 0.1 - 0.5 - 1 - 3 µM). 2. Measure hypocotyl length. c. Assay for petiole length in liquid media 1. Insert 14 day-old-plants from tissue culture media into a culture vial with liquid 0.5X MS-medium (0.5 ml per plant) and place on shaker. 2. Add BL to final concentration of 1 or 5 µM; in a control assay no BL is added. 3. Measure the petiole length of the third and fourth leaf after 10 days.

3.7.4. Cytokinin

Cytokinins are compounds with a structure resembling adenine, which promote cell division, leaf expansion, shoot initiation and bud formation in tissue culture, and growth of lateral buds (release of apical dominance), but they inhibit growth of the primary root and root branching, germination, modulation of light signaling, and delay leaf senescence (dark-induced chlorophyll loss) (18–20). Cis-zeatin is the only natural cytokinin used, but synthetic forms such as kinetin, adenine sulfate, and benzyladenine (BA) are also recommended for the assays. To test for responses to cytokinin the following assays can be performed: a. Root growth assay (see Sect. 3.6.3) 1. Transfer 5-day-old plants to vertical plates. Recommended condition for phenotypic analysis: media containing different concentrations of Kinetin or BA (0 - 0.1 - 0.5 - 1 µM). 2. Incubate for 5 days in the dark to test for root growth and number of lateral roots.

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b. Hypocotyl length assay (see Sect. 3.6.2) 1. Germinate seeds on a plate in the dark for 7 days. Recommended condition for phenotypic analysis: media containing different concentrations of BA (0 - 1 - 3 µM). 2. Measure hypocotyl length. c. Chlorophyll retention in the dark 1. Detach fully expanded leaves from 3-week-old plants and place them in the dark in Petri dishes with wet tissue paper. Per line at least 5 leaves should be used. 2. Spray the leaves with different concentrations of BA (0 - 0.01 - 0.1 - 1 µM). 3. Measure chlorophyll content after 0, 3, 6, and 9 days of darkness and calculate according to the fresh weight: – Weigh each leaf –

Extract chlorophyll by adding 1 ml of 80% acetone to each leaf cut into small pieces.

–

Place the test tubes in a dark place to avoid degradation of chlorophyll and incubate overnight. If the extraction is not complete grind the leaf tissue with sand.

–

After centrifugation the pellet should appear white; otherwise, re-extract the tissue.

–

Measure the absorbance of the supernatant at 663 and 645 nm against 80% acetone.Quantify chlorophyll for each sample using the following standards (after 21): Chlorophyll a (µg/ml) = 12.7 × E663 − 2.69 × E645 Chlorophyll b (µg/ml) = 22.9 × E645 − 4.68 × E663 Express on a fresh-weight basis! The total chlorophyll content can be calculated by the following formula:

Chlorophyll a + b (µg/ml) = 20.2 × E645 + 8.02 × E663 The chlorophyll content at the start of the experiments can be taken as a reference and set at 100%. 3.7.5. Ethylene

Ethylene, unlike the rest of the plant hormone compounds, is a gaseous hormone. It stimulates the release of dormancy, shoot and root growth and differentiation (triple response), fruit ripening, and leaf and fruit abscission (22–24). The “triple response” is exhibited by dark-grown seedlings exposed to ethylene. In Arabidopsis it is characterized by exaggerated curvature of the apical hook, radial swelling of the hypocotyl, and inhibition of hypocotyl and root growth. For assays the application of the gas is more complicated; therefore, 1-aminocyclopropane-1-carboxylic acid (ACC), which is an ethylene precursor, is used in the media. As inhibitors AgN03

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and aminoethoxyvinylglycine (AVG) can be added. As a control mutants such as ein2, ein3, ein5, ein6, or etr1 can be used (25). To test for responses to ethylene the following assays can be performed: a. Hypocotyl hook curvature assay 1. Germinate seeds on a vertical plate in the dark for 5 days. Recommended condition for phenotypic analysis: media containing different concentrations of ACC (1 - 5 - 10 25 µM). 2. Take a digital image and measure the angle between the top of the hypocotyl and the tip of the cotyledons. b. Hypocotyl length assay (see Sect. 3.6.2) 1. Germinate seeds on a plate in the dark for 5 days. Recommended condition for phenotypic analysis: media containing different concentrations of ACC (1 - 5 - 10 - 25 µM). 2. Measure hypocotyl length. c. Root growth assay (see Sect. 3.6.3) 1. The 5-day-old plants are transferred to vertical plates. Recommended condition for phenotypic analysis: media containing different concentrations of ACC (0.1 - 0.5 - 5 µM) or AVG (2 - 5 µM) and AgN03 (20 - 100 µM). 2. Incubate for 5 days in the dark to test for root growth. 3.7.6. Gibberellin

Gibberellins are diterpenes and many different forms with an entgibberellane skeleton have been identified. They stimulate stem elongation by stimulating cell division and elongation, and stimulate flowering and break seed dormancy (26–29). The two most often used forms are GA3 and GA4. Paclobutrazol (PAC) and uniconazole can be used as inhibitors. As controls GA-Biosynthesis mutants such as ga1 can be used (30). To test for responses to GA the following assays can be performed: a. Germination assay (see Sect. 3.6.1) 1. Germinate seeds on media containing different concentrations of GA3/GA4 (0 - 0.5 - 2 - 5 - 10 µM) or PAC/ Unicazole (0 - 2 - 5 µM). 2. Evaluate germination after 5 days. On 0.5 µM GA4 50% radicle emergences can be observed in wild type (Col-0). b. Hypocotyl length assay (see Sect. 3.6.2) 1. Germinate seeds on a plate in the light for 5 days. Recommended condition for phenotypic analysis: media containing different concentrations of GA3 or GA4 (0 - 0.5 - 2 - 5 – 10 µM). 2. Measure hypocotyl length. c. Treatment of plants with GA 1. Grow plants on soil. 2. Spray plants twice a week with 100 µM GA3.

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3. Measure the size of the rosette (diameter) after 3 weeks. 4. Record flowering time (see 1). 3.7.7. Methyl Jasmonate

Jasmonate is derived from the lipoxygenase-dependent oxidation of linolenic acid. MeJA induces the plant defense pathway in several species and seems to be an important stress-signaling molecule in plants (31, 32). To test for responses to MeJA the following assays can be performed: a. Root growth assay (see Sect. 3.6.3) 1. Transfer 5-day-old plants to vertical plates. Recommended condition for phenotypic analysis: media containing different concentrations of MeJA (0 - 0.1 - 1 - 10 - 100 µM). 2. Incubate for 5 days in the light to test for root growth.

3.8. Influence of Stress and Other Environmental Factors 3.8.1. Drought

Sensing of water status differs from sensing of other stimuli in that there is no chemical ligand that can be sensed, but other factors, such as reduction of turgor and changes in cell volume or membrane strain are most probably the primary stimulus detected (33–36). Prevention of drought stress depends upon minimizing stomatal and cuticular water loss and maximizing water uptake (through root growth and osmotic adjustment). To test for responses to drought the following assays can be performed: a. Drought assay on soil 1. Grow plants in soil (same size of pots, same amount of soil) for 3 weeks. 2. Withhold water from the plants for 9–12 days. 3. Score the number of wilted plants and express as the percentage of the total plants. 4. After 12 days re-water all the pots simultaneously (with the same amount). 5. Score the number of recovered plants that fully regained turgor and resumed growth after an additional 4 days and express as the percentage of the total plants wilted. b. Stomata closure Plants that cannot close their stomata wilt much faster than others (37). The stomatal closure can be assayed by looking at the stomata on the lower (abaxial) side of the leaves and by scoring the stomata as closed, open, and intermediate in the microscope (see Note 6).

3.8.2. Cold Stress

Chilling (temperatures below optimal but above freezing) and freezing temperatures can also cause osmotic stress in addition to their direct effect on metabolism (34, 35, 38). Survival during freezing-induced stress may depend upon prevention or delay of ice nuclei formation. Pretreatment of plants with lower temperatures

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increases their resistance to freezing temperatures. What occurs during this cold acclimation must be responsible for the freezing tolerance; one change is the increase in cellular concentrations of “compatible” osmolytes, such as proline. To test for responses to cold and freezing the following assays can be performed: a. Root growth assay at 14oC (chill) (see Sect. 3.6.3) 1. Transfer 5-day-old plants to vertical plates. 2. Keep plants 14 days in the light to test for root growth at 14°C in comparison with plants grown at 21°C. Check for identical light conditions. 3. Measure root length. b. Freezing tolerance assay/ cold acclimation 1. Grow plants on soil for 3 weeks (long day 16 h light/8 h dark, 21°C). 2. Transfer one set of plants to 4°C for one week (same light conditions) for cold acclimation, whereas the other set remains at 21°C. 3. Subject both set of plants to freezing at −5 to −10°C for 5 h. 4. Transfer plants immediately to 4°C under white light and incubate overnight. 5. Place plants into 21°C on the next morning. 6. After 7 days score damage to the plants and % survival. 3.8.3. Heat Stress

Plants have both an inherent ability to survive exposure to temperatures above the optimal for growth (basal thermo-tolerance) and an ability to acquire tolerance to otherwise lethal heat stress (acquired thermo-tolerance) (39). Acquired thermo-tolerance is induced by a short acclimation period at moderately high (but survivable) temperatures or by treatment with other non-lethal stress before heat stress. Heat stress has a complex impact on cell function, suggesting that many processes are involved in thermo-tolerance. High temperatures are known to affect membrane-linked processes due to alterations in membrane fluidity and permeability. Enzyme function is also sensitive to changes in temperature. Heat-induced alterations in enzyme activity can lead to imbalance in metabolic pathways, or heat can cause complete enzyme inactivation due to protein denaturation. Membrane and protein damage lead to the production of active oxygen species that cause heat-induced oxidative stress and can also promote programmed cell death. To test for responses to heat and thermotolerance the following assays can be performed: a. Germination assay (see Sect. 3.6.1) 1. Place seeds on media and immediately heat to 45°C for 220 min. Avoid drying out of the media by wrapping the plates with parafilm. As a control no heat treatment is applied (21°C).

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2. Further incubate plates at 21°C in the light (16 h light/8 h dark). 3. Score germination 1 week after heat treatment. b. Root growth assay (see Sect. 3.6.3) 1. Transfer 5-day-old plants to vertical plates. 2. Transfer plants to media for 14 days in the light to test for root growth at 27°C in comparison with plants grown at 21°C. Check for identical light conditions. c. Growth test 1. Grow 3-day-old seedlings on agar plate up to 3 weeks, in growth chambers at 35°C compared with seedlings at 21°C. 2. Measure the size of the rosette (diameter) and petiole length after 3 weeks (see 1). d. Hypocotyl length assay/thermo-tolerance test (see Sect. 3.6.2) 1. Germinate seeds on a vertical plate in the dark for 3 days (21°C). All the Petri dishes should contain the same amount of medium. 2. Seal the plates with the seedlings with parafilm and submerge them in a water bath. Heat seedlings in a water bath at 44°C for 15–30 min or at 46–50°C for 5 min. Use at least 3 parallel probes (30 seedlings per line). As a control one set of plates is not heated. 3. Label the hypocotyl positions after this heat treatment. 4. Keep the plants growing vertically for 3 days at 21°C, and then measure the length of hypocotyl elongation. e. Heat shock 1. Grow seedlings for 5–7 days in a Petri dish (90 × 15 mm) with 10 ml of solid medium at 21°C. This allows fast temperature changes and enables comparable temperature changes in all plates. 2. Seal the plate with the seedlings with parafilm and submerge it in a water bath at 44–45°C for 45 min. Use at least 3 parallel probes (30 seedlings per line). As a control one set of plates is not heated. 3. Cool the plate down with fanning or in the water bath at 21°C and keep at 21°C in the light. 4. Evaluate the thermo-tolerance by survival rate. Photograph plants 6–11 days after the heat shock treatment. Determine survival of plants relative to the wild-type control 5 days after heat stress. WT does not survive this treatment. This treatment leads to progressive bleaching after the heat treatment, which suggests that the damage

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caused by heating may have been due to oxidative stress occurring during the recovery phase. f. Acquired thermo-tolerance 1. Grow seedlings for 5 days in a Petri dish (90 × 15 mm) with 10 ml of solid medium at 21°C. This allows fast temperature changes and enables comparable temperature changes in all plates. 2. Pretreat plants 5 days after germination with 1 h at 37–38°C, and recover them for 2 days at 21°C. 3. Seal the plate with the seedlings with parafilm and submerge it in a water bath at 44–45°C for 45–60 min. Use at least 3 parallel probes (30 seedlings per line). As a control one set of plates is not heated. 4. Cool the plate down with fanning or in a water bath of 21°C and keep at 21°C in the light. 5. Evaluate the thermo-tolerance by survival rate. Photograph plants 6–11 days after the heat shock treatment. Survival of plants relative to the wild-type control is determined 8 days after heat stress. WT survives this treatment. 3.8.4. Salt Stress (with NaCl)

Salinity interferes with plant growth as it leads to physiological drought (difficulty in absorbing water), unusually high osmotic pressure, and ion toxicity (high concentrations of potentially toxic salt ions) (34–36, 40). To test for responses to salt stress the following assays can be performed: a. Root growth assay (see Sect. 3.6.3) 1. Transfer 5-day-old plants to vertical plates. Recommended condition for phenotypic analysis: media containing different concentrations of NaCl2 (0 - 50 - 100 - 150 - 200 mM). 2. Incubate for 5 days in the light to test for root growth. b. Growth assay 1. Pregrow plants for 1 month in the same kind of plastic pots with the same amount of soil (e.g., 250 cm3 soil) under long-day conditions at 21°C. Use ten plants of each genotype and three replicates each. 2. Soak pots in 150 or 400 mM NaCl solution (e.g., 200 ml per pot) freshly prepared every 3 days, for 2 or 3 weeks. 3. Measure shoot fresh weight and length and silique number after salt treatment.

3.8.5. Osmotic Stress

Osmotic stressors are stimuli that indicate an increase or decrease in the concentration of solutes outside the organism or cell (34, 35). Drought, high salinity, and freezing also impose osmotic stress on plants. While maintaining a positive turgor pressure, plant cells usually adjust their osmotic potential to meet the requirement of the whole plant in balancing its water budget. Significant changes

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in water potentials in the environment can impose osmotic stress to plants, which disrupts normal cellular activities, interferes with nutrient availability, or even causes plant death. Changes include, for example, morphological and developmental changes (e.g., life cycle, inhibition of shoot growth, and enhancement of root growth), adjustment in ion transport (such as uptake, extrusion, and sequestration of ions), and metabolic changes (e.g., carbon metabolism, the synthesis of compatible solutes). Some of these responses are triggered by the primary osmotic stress signals, whereas others may result from secondary stresses/ signals caused by the primary signals. These secondary signals can be phytohormones (e.g., ABA, ethylene, see 4), reactive oxygen species, and intracellular second messengers (e.g., phospholipids). Osmotic stress can be mimicked in the laboratory by several chemical components, such as mannitol, sorbitol, sucrose, NaCl (see Sect. 3.8.4), and PEG. It has to be taken into consideration that mannitol, sorbitol, and sucrose can have additional signaling functions besides changing the osmotic balance. To test for responses to osmotic stress the following assays can be performed: a. Germination assay (see Sect. 3.6.1) 1. Germinate seeds on media containing different concentrations of mannitol (0 - 200 - 300 - 400 - 500 mM), sorbitol (0 - 100 - 200 - 300 - 400 mM), and glucose (0 - 2 - 4 - 6% (w/v)). 2. Evaluate germination after 5 days in the light. b. Root growth assay (see Sect. 3.6.3) 1. Transfer 5-day-old plants to vertical plates. Recommended condition for phenotypic analysis: media containing different concentrations of mannitol (0 - 100 - 200 - 300 - 400 500 mM), sorbitol (0 - 100 - 200 - 300 - 400 mM), glucose (0 - 2 - 4 - 6% (w/v)), PEG 6000 or 8000 (0 - 5 - 10 - 20% (w/v)). 2. Incubate for 5 days in the light to test for root growth and total lateral root length. c. Growth assays on plates 1. Germinate seeds on media containing different concentrations of sucrose (0 - 4 - 7% (w/v)), glucose (0 - 4 - 6% (w/v)), and mannitol (0 - 2 - 4 - 6 - 8 - 10% (w/v)). 2. Evaluate the phenotype of the seedlings after 5 days. On 6% glucose or 8% mannitol, WT appears white and anthocyan accumulates. d. Growth assay on soil 1. Pre-grow plants for 1 month in the same kind of plastic pots with the same amount of soil (e.g., 250 cm3 soil) under long-day conditions at 21°C. Use ten plants of each genotype and three replicates each. 2. Irrigate pots with 200 mM mannitol for 1 week.

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3. Irrigate pots for another 3 days with water for recovery. 4. Measure shoot fresh weight and length and silique number after treatment. 3.8.6. Oxidative Stress

Oxidative stress arises from an imbalance in the generation and removal of reactive oxygen species (ROS) such as hydrogen peroxide (H2O2) and has been shown to occur in plants exposed to high and low temperatures, particularly in combination with high light intensities, drought exposure to air pollutants (e.g., ozone or sulfur dioxide), and ultraviolet light (41). Herbicides such as paraquat can also lead to oxidative stress. To test for responses to oxidative stress the following assays can be performed: a. Germination assay (see Sect. 3.6.1) 1. Germinate seeds on media containing different concentrations of methylviologen (paraquat) (0 - 0.1 - 0.5 - 1 -2 µM). 2. Evaluate germination after 5 days. b. Root growth assay (see Sect. 3.6.3) 1. Transfer 5-day-old plants to vertical plates. Recommended condition for phenotypic analysis: media containing different concentrations of methylviologen (paraquat) (0 - 0.1 - 0.5 - 1 -2 µM). 2. Incubate for 5 days in the light to test for root growth.

3.8.7. Light

Light is one of the most important abiotic factors controlling plant growth and development, starting at germination and until the reproductive phase (42–44). Plants have developed sophisticated light-sensing mechanisms (photoreceptors) to monitor changes in light quality, intensity, direction, and periodicity (day length). To generate different light qualities white light can be filtered through appropriate filters (blue light 400 nm, red light 660 nm, far-red light 730 nm) or LEDs with the appropriate spectra can be used (see Note 7). a. Germination assay (see Sect. 3.6.1) 1. Place seeds on media and illuminate with a far-red light pulse to revert active (Pfr) into inactive phytochrome (Pr). After this light pulse, plates are directly transferred into darkness. 2. Keep one set of plates for 7 days in the dark as a control. In the dark only a low amount of seeds should germinate. If many seeds germinate, the seeds are too freshly harvested, light could still penetrate, or the dormancy of the seeds is impaired (see 1). Illuminate a second set of plates after 3 h imbibition with 15 min of red light (660 nm, 1 µmol photons m−2s−1) and transfer them again for 7 days into the dark. Phytochrome B is responsible for germination under these conditions.

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Illuminate a third set of plates after 48 h imbibition with 15 min of far-red light (730 nm, 1 µmol photons m−2s−1) and transfer for 7 days into the dark. Phytochrome A is responsible for germination under these conditions. It is absolutely necessary to avoid any contaminating light besides the indicated light pulses, as this will change the outcome of the experiment. 3. Evaluate germination rate after these treatments and place seedlings in white light. After 3 days count those seeds that still did not germinate and subtract them, as these are “dead” seeds. 4. Calculate germination as % germination. b. Hypocotyl length assay (see Sect. 3.6.2) 1. Place seeds on the medium, stratify, and induce germination by a light pulse. Keep plates overnight in the dark (21°C). 2. Keep one set of plates in darkness, and place the others in different light conditions (see Note 8): White light 0.5–10 µmol photons m−2s−1 Blue light 2.0–10 µmol photons m−2s−1 Red light 0.5–35 µmol photons m−2s−1 Far-red light 0.5–10 µmol photons m−2s−1 Col-0 hypocotyl length is between 1 and 2 mm under saturating conditions, whereas in darkness the hypocotyl is between 1.1 and 1.4 mm (depending on the time in darkness). Non-saturating conditions are best for this assay, which means that the hypocotyl of the wild-type seedlings should be between 4 and 7 mm for an intermediate lightpoint. In most cases it is recommendable to perform a fluence rate–response curve to characterize the mutant in a more detailed way. 3. Incubate seedlings for 4–5 days under these light conditions. 4. Measure hypocotyl length. For a fluence rate–response curve the hypocotyl length (either in mm or % of dark value) can be plotted against the fluence rate on the x-axis (in a logarithmic scale). At least 20 seedlings should be measured for each condition and the experiments repeated at least three times. The temperature under the different light conditions should be maintained at the same level (around 21°C). Nonuniform germination results in variable hypocotyl length, with later germinating seedlings being shorter in stature. c. De-etiolation assay (see Note 9) 1. Place seeds on the medium, stratify, and induce germination by a light pulse. Keep seeds for 4 days in the dark (21°C). Wild type is characterized by an elongated hypocotyl, apical hook, and folded cotyledons.

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2. Transfer plates to white light (50–100 µmol photons m−2s−1). 3. Observe the opening of the apical hook and unfolding of the cotyledons after 6–12 h. Mutants that are hypersensitive to light show an increased opening of the cotyledons, which can be measured as the angle between the two cotyledons’ tips. When the seedlings are stretched on an agar plate for taking images one has to ensure that the angle between the cotyledons is not disturbed. 4. Measure the greening of the cotyledons after 12 h. For this 20 seedlings are extracted together and the chlorophyll content is specified per seedling (see Sect. 3.7.4). d. Root growth assay for UV-B light (see Sect. 3.6.3) 1. Transfer 5-day-old plants to vertical plates. 2. Irradiate plants for 15 min with UV-B light. One possible source is the Stratalinker (Stratagene) with 1.0 × 105 µJ of UV light, or broadband UV-B lamps (e.g., Philips TL40W/12RS) 1.5–1.7 µmol photons m−2 s−1 filtered through a layer of 3-mm 305-nm cut-off filters (half-maximal transmission at 305 nm) or a hand-held UV-lamp (312 nm) (see Note 10). One set of plants is not irradiated as a control. 3. Incubate plants vertically in light for 5 days and measure root length. UV-B irradiation is not a mere stress signal but can also serve as an environmental stimulus to direct growth and development (46). This includes hypocotyl growth inhibition and flavonoid accumulation.

4. Notes

1. Many substances that are added to the media are not autoclaveable and therefore should be sterile-filtered. Furthermore, in many instanced the solvent is not water; therefore, the correct control should have the same amount of solvent as in the different conditions. 2. The challenge with different abiotic factors is mainly performed in tissue culture as the amount of the added substances is easily controlled and therefore dose–response curves are more reliable and repeatable. The assays described here are dependent on the surrounding conditions and have sometimes to be adapted. A phenotype can be either hypersensitive or hyposensitive to a certain condition; therefore, it is recommended to find the level in which the wild type is

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affected to 50%. This allows the observation of effects that are higher and lower. Furthermore, using at least three different conditions gives a better evaluation of the response, although one or two might be enough for a first assessment. 3. For all germination assays it is extremely important to use seeds that have been stored for the same time and that have been gathered from mother plants that have been grown under the same conditions. This is because the dormancy of seeds depends on the seed age as freshly harvested seeds have a different germination behavior (especially higher dark germination) than stored seeds, whereas longer storage reduces the overall germination efficiency. Germination is also influenced by the endogenous hormone levels and the nutrient level within the seed, both of which are influenced by the status of the mother plant. 4. Alternatively, seeds can be placed in a row on agar plates so that the plates can be incubated vertically and the seedlings can grow parallel to the agar. The disadvantage is that fewer lines can be placed on one plate because 2 cm has to be allowed for upward growth and 1 cm for downward growth. Furthermore, not all seedlings grow in a straight fashion, which makes the measurement more difficult. 5. Cross-talk between the various signaling pathways makes the identification of the biological role difficult even after the identification of a phenotype. For example, screens for abscisic acid signaling mutants have turned up ethylene, gibberellic acid, and sugar signaling response genes, providing evidence for genetic interactions between ABA signaling and other hormones. 6. Examining stomata closure should be performed quickly as the detachment of leaves from the plant and the microscope lighting can change the status. 7. It is important for the far-red light source to not produce light below 700 nm. To reduce the light intensity move the light source further away or cover the plates with sheets of white tissue paper or additional filters. Note that the usual spectroradiometers to measure light intensity are not able to detect far-red light. 8. Under far-red light phytochrome A is responsible for the de-etiolation process, under red light mainly phytochrome B, whereas under blue light cryptochrome 1 and 2 are the major photoreceptors. 9. Light inhibits hypocotyl elongation and promotes cotyledon unfolding. Additionally, cotyledon size and phototrophic growth are influenced by light (see Ref. 1, 45). Many hormones intersect with light signal transduction such as gibberellins,

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ethylene, auxin, cytokinin, and brassinosteroids and influence developmental programs. However, light is not only important during the seedling development but also during later stages of plant development. Parameters that can be evaluated in the adult plant are, for example, petiole elongation and flowering time (see Ref. 1). 10. Note that the usual spectroradiometers to measure light intensity are not able to detect UV light.

Acknowledgement I would like to thank Dario Leister for his support and critical reading of the manuscript. Furthermore, I am grateful to Randy Foster for critical discussions and help with the manuscript. I would also like to thank numerous colleagues with whom I discussed different techniques. This work was supported by grants from the Deutsche Forschungsgemeinschaft (DFG, BO 1146/3; Arabidopsis Functional Genomics Network, AFGN, BO 1146/4) to C.B. References 1. Bolle, C. (2008) Phenotyping of Arabidopsis mutants for developmental effects of gene deletions. 2. Murashige, T. and Skoog, F. (1962) A revised medium for rapid growth and bioassays with tobacco tissue cultures. Physiol. Plant. 15, 473–497. 3. Gubler, F., Millar, A. A., and Jacobsen, J. V., (2005) Dormancy release, ABA and preharvest sprouting. Curr. Opin. Plant Biol. 8, 183–187. 4. Christmann, A., Moes, D., Himmelbach, A., Yang, Y., Tang, Y., and Grill, E. (2006) Integration of abscisic acid signalling into plant responses. Plant Biol. (Stuttg) 8, 314–325. 5. Verslues, P. E. and Zhu, J. K. (2005) Before and beyond ABA: upstream sensing and internal signals that determine ABA accumulation and response under abiotic stress. Biochem. Soc. Trans. 33, 375–379. 6. Barrero, J. M., Piqueras, P., GonzalezGuzman, M., Serrano, R., Rodriguez, P. L., Ponce, M. R., and Micol, J.L. (2005) A mutational analysis of the ABA1 gene of Arabidopsis thaliana highlights the involvement of ABA in vegetative development. J. Exp. Bot. 56, 2071–2083.

7. Leyser, O. (2006) Dynamic integration of auxin transport and signalling. Curr. Biol. 16, R424–433. 8. Teale, W. D., Paponov, I. A., and Palme, K. (2006) Auxin in action: Signalling, transport and the control of plant growth and development. Nat. Rev. Mol. Cell Biol. 7, 847–859. 9. Woodward, A. W. and Bartel, B. (2005) Auxin: regulation, action, and interaction. Ann. Bot. (Lond) 95, 707–735. 10. Dharmasiri, N., Dharmasiri, S., and Estelle, M. (2005) The F-box protein TIR1 is an auxin receptor. Nature 435, 441–445. 11. Kepinski, S. and Leyser, O. (2005) The Arabidopsis F-box protein TIR1 is an auxin receptor. Nature 435, 446–451. 12. Ulmasov, T., Murfett, J., Hagen, G., and Guilfoyle, T. J. (1997) Aux/IAA proteins repress expression of reporter genes containing natural and highly active synthetic auxin response elements. Plant Cell 9, 1963–1971. 13. Bishop, G.J. and Koncz, C. (2002) Brassinosteroids and plant steroid hormone signaling. Plant Cell 14(Suppl.), S97–S110. 14. Wang, Z. Y., Wang, Q., Chong, K., Wang, F., Wang, L., Bai, M., and Jia, C. (2006)

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Bolle The brassinosteroid signal transduction pathway. Cell Res. 16, 427–434. Katsuhiko, S., Uzawa, J., Hana, S. J., Yoneyamac, K., Takeuchic, Y., Yoshida, S., Asami, T. (2002) Brz220 a novel brassinosteroid biosynthesis inhibitor: stereochemical structure–activity relationship. Tetrahedron: Asymmetry 13, 1875–1878. Bancos, S., Nomura, T., Sato, T., Molnar, G., Bishop, G. J., Koncz, C., Yokota, T., Nagy, F., and Szekeres, M. (2002) Regulation of transcript levels of the Arabidopsis cytochrome p450 genes involved in brassinosteroid biosynthesis. Plant Physiol. 130, 504–513. Fujioka, S., Li, J., Choi, Y. H., Seto, H., Takatsuto, S., Noguchi, T., Watanabe, T., Kuriyama, H., Yokota, T., Chory, J., and Sakurai, A. (1997) The Arabidopsis deetiolated2 mutant is blocked early in brassinosteroid biosynthesis. Plant Cell 9, 1951–1962. Ferreira, F. J. and Kieber, J. J. (2005) Cytokinin signaling. Curr. Opin. Plant Biol. 8, 518–525. Riefler, M., Novak, O., Strnad, M., and Schmülling, T. (2006) Arabidopsis cytokinin receptor mutants reveal functions in shoot growth, leaf senescence, seed size, germination, root development, and cytokinin metabolism. Plant Cell 18, 40–54. Werner, T., Motyka, V., Laucou, V., Smets, R., Van Onckelen, H., and Schmulling, T. (2003) Cytokinin-deficient transgenic Arabidopsis plants show multiple developmental alterations indicating opposite functions of cytokinins in the regulation of shoot and root meristem activity. Plant Cell 15, 2532–2550. Arnon, D.I. (1949) Copper enzymes in isolated chloroplasts. Polyphenoloxidase in Beta vulgaris. Plant Physiol. 24, 1–15. Chen, Y. F., Etheridge, N., and Schaller, G. E. (2005) Ethylene signal transduction. Ann. Bot. (Lond) 95, 901–915. De Paepe, A. and Straeten, D. Van der (2005) Ethylene biosynthesis and signaling: An overview. Vitam. Horm. 72, 399–430. Etheridge, N., Hall, B. P., and Schaller, G. E. (2006) Progress report: ethylene signaling and responses. Planta 223, 387–391. Roman, G., Lubarsky, B., Kieber, J. J., Rothenberg, M., and Ecker, J. R. (1995) Genetic analysis of ethylene signal transduction in Arabidopsis thaliana: Five novel mutant loci integrated into a stress response pathway. Genetics 139, 1393–1409. Sun, T. P. and Gubler, F. (2004) Molecular mechanism of gibberellin signaling in plants. Annu. Rev. Plant Biol. 55, 197–223.

27. Swain, S. M. and Singh, D. P. (2005) Tall tales from sly dwarves: novel functions of gibberellins in plant development. Trends Plant Sci. 10, 123–129. 28. Thomas, S. G. and Sun, T. P. (2004) Update on gibberellin signaling. A tale of the tall and the short. Plant Physiol. 135, 668–676 29. Griffiths, J., Murase, K., Rieu, I., Zentella, R., Zhang, Z. L., Powers, S.J., Gong, F., Phillips, A. L., Hedden, P., Sun, T. P., and Thomas, S. G. (2006) Genetic characterization and functional analysis of the GID1 gibberellin receptors in Arabidopsis. Plant Cell 18, 3399–3414. 30. Sun, T. P. and Kamiya, Y. (1994) The Arabidopsis GA1 locus encodes the cyclase ent-kaurene synthetase A of gibberellin biosynthesis. Plant Cell 6, 1509–1518. 31. Liechti, R. and Farmer, E. E. (2003) The jasmonate biochemical pathway. Sci. STKE 2003, CM18. 32. Turner, J. G., Ellis, C., and Devoto, A. (2002) The jasmonate signal pathway. Plant Cell 14(Suppl.), S153–S164. 33. Valliyodan, B. and Nguyen, H. T. (2006) Understanding regulatory networks and engineering for enhanced drought tolerance in plants. Curr. Opin. Plant Biol. 9, 189– 195. 34. Mahajan, S. and Tuteja, N. (2005) Cold, salinity and drought stresses: an overview. Arch. Biochem. Biophys. 444, 139–158. 35. Xiong, L., Schumaker, K. S., and Zhu, J. K. (2002) Cell signaling during cold, drought, and salt stress. Plant Cell 14(Suppl.), S165– S183. 36. Zhu, J. K. (2002) Salt and drought stress signal transduction in plants. Annu. Rev. Plant Biol. 53, 247–273. 37. Roelfsema, M. R. and Hedrich, R. (2005) In the light of stomatal opening: new insights into ‘the Watergate’. New Phytol. 167, 665–691. 38. Thomashow, M. F. (1999) PLANT COLD ACCLIMATION: Freezing Tolerance Genes and Regulatory Mechanisms. Annu. Rev. Plant Physiol. Plant Mol. Biol. 50, 571–599. 39. Baniwal, S. K., Bharti, K., Chan, K. Y., Fauth, M., Ganguli, A., Kotak, S., Mishra, S. K., Nover, L., Port, M., Scharf, K. D., Tripp, J., Weber, C., Zielinski, D., and von KoskullDöring, P. (2004) Heat stress response in plants: a complex game with chaperones and more than twenty heat stress transcription factors. J. Biosci. 29, 471–487. 40. Chinnusamy, V., Zhu, J., and Zhu, J. K. (2006) Salt stress signaling and mechanisms of plant salt tolerance. Genet. Eng. (N Y) 27, 141–177.

Responses to Hormones and Abiotic Stresses 41. Apel, K. and Hirt, H. (2004) Reactive oxygen species: metabolism, oxidative stress, and signal transduction. Annu. Rev. Plant Biol. 55, 373–399. 42. Franklin, K. A., Larner, V. S., and Whitelam, G. C. (2005) The signal transducing photoreceptors of plants. Int. J. Dev. Biol. 49, 653–664. 43. Chen, M., Chory, J., and Fankhauser, C. (2004) Light signal transduction in higher plants. Annu. Rev. Genet. 38, 87–117.

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44. Sullivan, J. A. and Deng, X. W. (2003) From seed to seed: The role of photoreceptors in Arabidopsis development. Dev. Biol. 260, 289–297. 45. Fankhauser, C. and Casal, J. J. (2004) Phenotypic characterization of a photomorphogenic mutant. Plant J. 39, 747–760. 46. Ulm, R. and Nagy, F. (2005) Signalling and gene regulation in response to ultraviolet light. Curr. Opin. Plant Biol. 8, 477–482.

Chapter 4 Accurate Real-Time Reverse Transcription Quantitative PCR Marco Klatte and Petra Bauer Abstract Within the last few years real-time quantitative PCR has become the method of choice for the accurate quantification of mRNA levels. Compared to previous methods the sensitivity of real-time quantitative PCR improved to the detection limit of up to one single molecule per reaction tube. However, the improved sensitivity leads also to higher demands regarding experimental design. Here we describe an approved protocol to establish mRNA quantification by real-time RT qPCR in a straightforward manner. Key words: Real-time reverse transcription quantitative PCR, mRNA, absolute quantification, relative quantification, SYBR green I, housekeeping genes.

1. Introduction The quantification of mRNA levels is a major tool to investigate the response to environmental changes or the impact of mutations in plants. Nowadays high-throughput gene-chip experiments produce huge amounts of data covering the whole transcriptome of an organism. Gene-chip-based expression studies are a powerful tool to obtain an overview of regulatory networks and to identify unknown members of a regulatory network. However, confirmation of the observed expression patterns by more precise methods like real-time reverse transcription qPCR is still mandatory. Due to its high sensitivity, reverse transcription polymerase chain reaction (RT-PCR) is the method of choice to quantify low levels of mRNA (1). In the past, semi-quantitative RT-PCR, based on end-point detection of the PCR product, was widely used. The PCR reaction was stopped at a cycle number where

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the PCR reaction was assumed to be in the exponential phase and the amount of PCR product already detectable. After gel electrophoresis and southern blot hybridization, the signal intensities between two or more samples were used to determine differences in the initial amount of mRNA. In contrast to end-point detection, real-time reverse transcription quantitative PCR (real-time RT qPCR) is based on the detection of fluorescent marker molecules emitting light proportional to the amount of PCR product formed during the PCR reaction (2). By this, the fluorescence can be monitored and precisely quantified in real time during each cycle of the PCR reaction. The first significant increase in fluorescence, named threshold cycle (Ct) or crossing point (Cp), correlates directly with the initial amount of target DNA in the reaction tube. The early detection of PCR products results in a highly dynamic range in real-time RT qPCR: differences of up to 107 in initial template amount can be quantified compared with 103 in conventional RT-PCR (2, 3). Further, the use of mass standards in real-time RT qPCR enables absolute quantification and thus the quantitative comparison of expression levels of different genes, an aspect that cannot be addressed by conventional end-point qPCR. Here, we describe a straightforward and affordable reverse transcription based real-time RT qPCR protocol using the SYBR green I detection system. Our starting material is total RNA from various plant tissues (successfully tested with roots, rosette leaves, flowers and siliques) reverse transcribed to cDNA with M-MuLV reverse transcriptase using oligo-dT primer. To ensure reliability and reproducibility several precautions are necessary, which we describe briefly. Further, details about the theory of real-time RT qPCR and the requirements necessary to obtain reliable real-time RT qPCR results are reviewed elsewhere (4, 5, 6, 7). Plant material should be harvested and deep frozen as soon as possible after the plants are removed from controlled environmental conditions. Storage of plants on the bench or other delays before harvest should be avoided because the mRNA levels might change quickly because of environmental changes. To circumvent possible variations based on the circadian rhythm, plants need to be harvested always at the same daytime. Low RNA quality may compromise the results of downstream applications, which are labour and cost intensive. RNA should be ideally free of RNases, proteins and genomic DNA as well as any substances that can inhibit downstream enzymatic reactions. RNA quality should be assessed either by gel electrophoresis or by using a commercially available device like Bioanalyzer 2100 (Agilent) or Experion (BioRad). If RNA concentration is determined by UV/VIS spectrophotometer, ensure that no traces of genomic DNA compromise the measurement. A DNase I digest

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before RNA measurement might be advisable. Include multiple wavelengths in these measurements: an A260/A280 ratio between 1.8 and 2.0 is usually considered to be an indicator of good RNA quality. However, one might keep in mind that pH and ionic strength may lower the absorption ratio significantly even if the RNA is pure (8). A very important aspect in quantification of mRNA is the normalization of variations in the amount of initial starting material (4, 9). Internal control genes, often called “housekeeping genes”, are assumed to be constitutively expressed in the tissues and at the physiological conditions under investigation. Differences in the expression levels of an internal control gene in a set of samples are interpreted as differences in the amount of starting material. Therefore, these differences are suitable to normalize variations in the amount of starting material in the individual samples. The importance of proper internal control gene selection is often neglected. Usually only a single internal control gene like ACTIN2 (At3g18780) in Arabidopsis is used without any statistical confidence that this gene is a true constitutively expressed control gene under the conditions observed. A robust strategy to identify the most stably expressed internal control genes has been described recently (10). We recommend selecting two to three putative internal control genes from pathways independent from each other. Our internal control genes are elongation factor 1B alpha-subunit 2 (eEF1Balpha2, At5g19510) and ubiquitin-specific protease 6 (UBP6, At1g51710); for primer sequences refer to Ref. (11). Candidate control genes can be identified using gene-chip expression data, which are easily available through genevestigator (12) on the Internet. The performance of the selected control genes can be validated statistically e.g., by the tool geNorm (13) based on the formulas described in (10). A sophisticated experimental design is the prerequisite for successful high-throughput real-time RT qPCR. Diluting and aliquoting the cDNA before the experiment covers several aspects of accuracy. First, pipetting 10 µl of 1:10 diluted cDNA samples instead of 1 µl undiluted cDNA leads to an enormous improvement of pipetting accuracy. Hence, technical repetitions might even be reduced from three to two replicates per cDNA sample. Second, cDNA stored in aliquots will thaw only once directly before the PCR and therefore will not be subjected to harmful freeze–thaw cycles. Finally yet importantly, using aliquots from a single initial cDNA dilution guarantees a consistent cDNA composition. The expression data of large amounts of candidate genes derived from the same cDNA remain comparable. Real-time RT qPCR derived expression data can be presented either as absolute or relative expression values. In many cases the presentation of relative expression data is sufficient, representing the up- or down-regulation of gene expression in fold-changes

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compared with a control sample. However, sometimes it is desirable to know the absolute number of transcript per sample, e.g., to point out which member of a gene family is the predominant gene in a given tissue. Although there are existing methods to calculate reliable relative expression data without performing standard curve analysis (14), we recommend including mass standards in each PCR. The generation of relative expression data from absolute expression data is far less challenging when compared with the evaluation using Ct values (14). Further, mass standards are very useful when optimizing PCR conditions and verifying PCR quality.

2. Materials 2.1. PCR Establishment and Setup

All real-time RT qPCR experiments are performed by using the following consumables. 1. Enzyme for real-time RT qPCR: Ex Taq R-PCR Version 2.1 (TaKaRa, RR031) 2. SYBR green I nucleic acid gel stain (Roche Diagnostics, 1988131), diluted 1:200 in fresh DMSO (Sigma) and stored in aliquots at −20°C as main stock. The main stock is further diluted 1:10 with TE buffer (pH 7.5, see Note 1) and stored in 0.5 ml aliquots ready to use for real-time RT qPCR for long-term storage at −20°C. SYBR green I is stable up to three weeks at 4°C (see Note 1) when kept light protected. SYBR green I is a mutagen. Handle it with care. 3. PCR grade water (RNase and DNase free, Sigma W4502) is aliquoted to 50 ml Falcon tubes under sterile conditions and stored at −20°C. 4. Store PCR standards and cDNA in 8-tube PCR strips easy to reopen (Kisker Biotech, G002-A). 5. Sealing tape for 96-well PCR plates: iCycler iQ Optical Tape (Bio-Rad, 2239444). 6. All experiments are performed using the real-time PCR cycler Mx3000P and the MxPro operating Software, (version 3.20, Stratagene). 7. Primer 3 Software: http://frodo.wi.mit.edu/cgi-bin/ primer3/primer3_www.cgi 8. Amplification of PCR standards from clones or cDNA: Ex Taq Hot Start Version (TaKaRa, RR006). 9. Gel extraction: E.Z.N.A. Gel extraction kit (Omega Bio-Tek)

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10. Quantification of PCR products using gel electrophoresis: MassRuler DNA Ladder Mix (Fermentas, SM0403) 11. Primer: HPLC purified (Metabion) 2.2. Sample Preparation

1. Invisorb Spin Plant RNA Mini Kit (Invitek) 2. 3 M Na acetate, pH 5.2 (made with DEPC treated water) 3. 70% Ethanol (made with DEPC treated water) 4. DNase I (Fermentas #EN0521) 5. cDNA synthesis kit (Fermentas #K1622)

3. Methods The methods described below outline (1) the establishment and optimization of real-time RT qPCR, (2) sample preparation, (3) experimental setup and (4) data analysis. 3.1. PCR Establishment and Setup

3.1.1. Primer Design

The establishment of a robust real-time RT qPCR protocol is described in Sects. 3.1.1–3.1.3. This includes the description of primer design for real-qPCR, PCR standard generation and the optimization of primer concentrations. 1. Primers for real-time RT qPCR should be designed e.g., by using publicly available software like Primer 3 (15) with special attention to specificity. In our case the primers are 17–23 bases long, have a GC content of 40–60% and a Tm value of 54–56°C (2–4°C lower than the annealing temperature in the PCR program). The lengths of the amplicons are 80–150 bps; however, amplicons of up to 400 bps should work with SYBR green I. Because of the use of oligo-dT primers in cDNA synthesis, the primer annealing sites should be in proximity to the 3′ end of the gene (close to the poly-A tail of the mRNA). It is helpful to design primer pairs surrounding one intron or covering exon–exon junctions. If this is not possible include a primer pair that amplifies only genomic DNA but no cDNA will be able to subtract these values from the expression values if contaminations by genomic DNA are detected. Primer sequence specificity should be confirmed by blastn searches against available databases, e.g., the AGI transcript (-introns, + UTRs) and the AGI whole genome database of Arabidopsis thaliana available on the TAIR website (16). Primers should be ordered in HPLC purified quality to avoid contaminations with slightly shorter sequences. To facilitate downstream applications the primer concentrations should be adjusted to e.g., 100 µM.

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2. For generation of standards for real-time RT qPCR, another set of primers is needed. This primer set should amplify approx. 1 kb of the gene of interest including the target sequence for the real-time RT qPCR. The template for standard amplification can be cDNA; however, in cases of very closely related gene sequences it might be better to use individual cDNA or BAC clones to ensure uniformity of the amplified PCR standards. Some laboratories use cDNA clones directly as standards for real-time RT qPCR. The possible secondary structures of circular plasmid DNA might compromise PCR efficiency. To ensure consistency of amplification we prefer to use linear DNA as standard for real-time RT qPCR. 3.1.2. Generation of PCR Standards

Repeating freeze–thaw cycles are a major reason for degradation of the cDNA template and standards. To ensure a constant quality (and comparability) of standards in all real-time RT qPCR experiments, aliquots of each standard dilution are prepared. The aliquots are stored in PCR strips to ease future pipetting during PCR setup by using a multichannel pipette. 1. Perform a PCR with 30–35 cycles and separate PCR fragments on a 1% agarose gel. It is advisable to make three 25 µl PCR reactions in parallel to get sufficient amounts of PCR product. 2. Cut out the band and extract the DNA, for example, with the E.Z.N.A. Gel Extraction Kit (Omega Bio-Tek, Inc.) according to the manufacturer’s protocol. 3. Quantify and verify the purity of the extracted PCR product by UV-spectroscopy (A260 and A280). Use different dilutions (e.g., 10–50 fold) and ensure that A260 is in the linear range between OD = 0.05 and OD = 1. 4. Confirm the PCR product quantification by gel electrophoresis. Load three different dilutions (e.g., 1:1, 1:5, and 1:10) of the purified PCR product and two different amounts of the mass ruler (e.g., 5 and 10 µl) on the gel. By using 1.5% agarose gels and low voltage the sharpness of the bands will be enhanced. Stain the gel after electrophoresis to ensure equal staining. 5. Determine the molecular weight of the PCR product by a DNA calculation tool (e.g., (17)) and adjust the concentration of purified PCR standard product to 109 molecules per micro liter. 6. Make a serial dilution to obtain 1 mL of each 107, 106, 105, 104, 103, and 102 molecules per 10 µl solution (see Note 2). 7. Aliquot 30 µl of each serial dilution to pre-labelled “easy to reopen” PCR strips (in the order 107–102; see Note 3).

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One of the critical issues, especially in SYBR green I based real-time RT qPCR, is the minimization of non-specific amplification products. Maximum sensitivity can be achieved by using the lowest concentration of primers without compromising the efficiency of PCR. The primer concentration should be 50–300 nM and adjusted separately for each primer. The primer titration for one primer set requires 15 PCR reactions including technical triplicates. 1. Prepare primer dilutions according to Table 4.1a. Primer dilutions are prepared in “easy to re-open” PCR strips. The mastermix with the template but without the primer is prepared in a 1.5-ml reaction tube according to Table 4.1b and further aliquoted to a PCR strip, 58 µl in each well. Avoid direct light because SYBR green I is light sensitive. 2. Transfer 10 µl diluted primer and 10 µl mastermix to the PCR plate by using a 10 µl multichannel pipette. See pipetting scheme in Fig. 4.1a. 3. Seal the PCR plate thoroughly with optical tape (see Note 4) and spin down droplets in a plate centrifuge. 4. Run the PCR according to the scheme in Fig. 4.2a. 5. Plot the Ct values against the combinations of primer concentrations as shown in the diagram in Fig. 4.1b. 6. Select the primer ratio with the lowest Ct and the lowest primer concentrations. Verify the presence of a single specific PCR product by melting-curve analysis (see Fig. 4.3c). 7. For future PCR reactions, make primer pre-mixes that are 5X concentrated. Make aliquots of 424 µl of the primer mixes and store at −20°C (see Note 5).

3.2. Sample Preparation

3.2.1. RNA Isolation

The preparation of samples is described in Sects. 3.2.1–3.2.2, including the isolation of RNA from plant material and the generation of cDNA samples. 1. Isolate total RNA from max. 70 mg plant material by using the Invisorb Spin Plant Mini RNA kit (Invitek, Germany) according to the manufacturer’s protocol (see Note 6). Grind the plant material before RNA isolation by freezing it in N2 in a 2-ml reaction tube and using a motorized metal homogenizer rather than using mortar and pestle to obtain a fine powder (see Note 7). 2. Elute RNA with 50 µl buffer EB. 3. Add 5 µl 3 M Na-acetate, pH 5.2 and 125 µl 99% ethanol and precipitate at least 1 h at −20°C. 4. Centrifuge for 15 min at maximum speed and 4°C, discard supernatant, and add 500 µl ice-cold 70% ethanol. 5. Centrifuge for 15 min at maximum speed and at 4°C; discard supernatant.

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Table 4.1 Composition of primer- and mastermixes for real-time PCR. (a) To optimize primer concentrations, different final concentrations (50–300 nM) of forward and reverse primer are combined. (b) The mastermix for primer optimization containing a constant amount of standard (c) The mastermix for one PCR plate in a standard real-time PCR assay (a) Primer mixes(primer matrix)

1

2

3

4

5

Primer 5′

4 µl

8 µl

12 µl

16 µl

24 µl

Primer 3′

24 µl

16 µl

12 µl

8 µl

4 µl

H 2O

12 µl

16 µl

16 µl

16 µl

12 µl

Primer ratio

1:6

1:2

1:1

2:1

6:1

Primer 5′:3′ [nM]

500:300 100:200 150:150 200:100 300:50

(b) Mastermix (primer matrix)

1x

18x

5x buffer

4 µl

72 µl

dNTP (10 mM)

0.4 µl

7.2 µl

MgCL2 (250 mM)

0.16 µl

2.9 µl

SYBR green

0.5 µl

9 µl

Standard 10 /1 µl

4 µl

18 µl

H2O

0.84 µl

69.1 µl

Ex Taq HS

0.1 µl

1.8 µl

(c) Mastermix (PCR plate)

1x

106x

5x buffer

4 µl

424 µl

dNTP (10 mM)

0.4 µl

42.4 µl

MgCL2 (250 mM)

0.16 µl

17 µl

SYBR green

0.5 µl

53 µl

Primer mix

1 µl

424 µl

H2O

3.84 µl

89.0 µl

Ex Taq HS

0.1 µl

10.6 µl

Mix 1 mM primer stocks with H2O [µl]

4

6. Centrifuge for 1 min at maximum speed and 4°C, remove supernatant with fresh pipette tip, and air dry the pellet (see Note 8). 7. Resolve RNA in 20 µl buffer EB. 8. Dilute 2 µl RNA in 98 µl water and measure RNA quantity and quality at 260/280 nm.

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Fig. 4.1. Experimental design of primer optimization. (a) Pipetting scheme. By using PCR strips and a multichannel pipette, the pipetting time and accuracy are optimized and the risk of contamination minimized. Please refer to Table 4.1a and b for detailed composition of primer- and mastermix. (b) Plot of the Ct values against the primer combinations. In this example, primer set 1 performs best with both primers at 150 nM concentration. Primer set 2 performs best with 200 nM forward and 100 nM reverse primer (see arrows).

3.2.2. DNase Treatment and cDNA Synthesis

RNA isolated with the Invisorb Spin Plant RNA Mini Kit usually contains little or no traces of genomic DNA. To ensure that only cDNA is present in real-time RT qPCR analysis we perform DNase I digestion of the RNA samples before cDNA synthesis by default. Mastermixes are prepared for all steps. 1. Mix 1 µl 10X DNase I buffer and 1 µl Ribolock RNase inhibitor with 1 µg total RNA and adjust the volume to 10 µl with water in a 200 µl PCR tube. Include one nontemplate control (NTC). 2. Incubate for 45 min at 37°C in a PCR block; cool down to 12°C. 3. Add 2 µl of a 1:1 mixture of 25 mM EDTA and 100 µM oligo(dT)18 primer. 4. Incubate for 10 min at 65°C, and cool down to 12°C (see Note 9). 5. Add 8 µl of a mixture containing 4 µl 5X RT-buffer, 2 µl dNTP, 1 µl RNase inhibitor, and 1 µl reverse transcriptase. 6. Incubate for 2 h at 42°C followed by a deactivation step for 10 min at 70°C and subsequent cooling to 4°C in a PCR block. 7. Add 180 µl PCR grade water to each cDNA sample and mix by pipetting up and down (see Note 10). 8. Store cDNA at −20°C.

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Fig. 4.2. (A) Scheme of the PCR program. (B) Pipetting scheme of a standard real-time PCR assay. (1) Transfer 10 µl mastermix (see Table 4.1c) with a multichannel pipette to each well, aliquot mastermix to 8-tube PCR strip before (129 µl per tube in the strip). (2) Transfer 10 µl cDNA to the rows B–H. (3) Transfer 10 µl PCR standard to the wells A1–A6 and A7–A12. Especially when pipetting PCR standards ensure that you do not move with the pipette tips over other wells. NTC = non template control from cDNA synthesis.

3.3. Experimental Setup

The experimental setup presented here aims at maximizing accuracy and economical efficiency. Usually the first row (A1–A12) of a 96-well PCR plate is reserved for the PCR standards. The remaining 84 wells are left to analyse 26 cDNA samples (plus non-template and water control) when performing technical triplicates. This number can be raised to 40 cDNA samples when performing technical duplicates. We always try to fill up the entire plate to achieve the best ratio between PCR standards and cDNA samples. 1. Dilute the cDNA 1:10 a second time with PCR grade water. Prepare aliquots of 70 µl into 7-tube (cut) PCR strips, each single tube containing another individual cDNA sample.

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Fig. 4.3. Verfication of real-time RT qPCR quality and integrity. (A) Amplification plots of PCR standards in a range of 107 to 102 template molecules per reaction tube. Each reaction was performed in duplicate. The Ct values of technical replicates are almost identical, showing high reproducibility of the experimental data. (B) Standard curve based on the amplification of standards in (a). The correlation coefficient (R2 = 1) and the PCR efficiency (102.7%) are usually calculated by the operating software of the real-time RT qPCR machine. (C) Melting-curve analysis of three different samples. circle: correct PCR product with Tm = 81°C, square: PCR artefact, presumably primer dimers, with Tm = 74°C. triangle: the correct PCR product and primer dimers were formed.

Each PCR strip contains the cDNA necessary for two PCR plates (genes to be analysed), which will be pipetted in parallel. For backup reasons we recommend the preparation of a minimum of 2 aliquots of 70 µl more than needed. Store the cDNA aliquots at −20°C.

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2. To save time, prepare two PCR plates in parallel, e.g., for analysis of two genes. Thaw the components for the PCR as well as one set of cDNA strips and the PCR standards on ice. 3. Vortex the PCR strips containing cDNA and standards after thawing and spin them down. 4. Prepare the mastermixes according to Table 4.1c (one mastermix per gene). Avoid direct light as SYBR green I is light sensitive. Mix the mastermixes and prepare aliquots of 129 µl in each well of an 8-tube PCR strip. 5. Transfer the two mastermixes to the two 96-well PCR plates (one plate per gene) by using a 10 µl multichannel pipette and fine pipette tips as indicated in Fig. 4.2b. 6. Turn the PCR plate by 180° (see Note 11). 7. Transfer the cDNA samples to the 96-well PCR plates as indicated in Fig. 4.2b. 8. Transfer the PCR standards to the 96-well PCR plates as indicated in Fig. 4.2b. 9. Seal the PCR plate as previously described and spin down in a plate centrifuge (see Note 4). 10. Run the PCR according to the PCR program indicated in Fig. 4.2a. Store the second PCR plate light protected at 4°C while the first PCR plate is in the PCR machine. One single PCR run lasts approx. 95 min in the Stratagene Mx3000P PCR cycler with the program indicated in Fig. 4.2a. 3.4. Data Analysis

The analysis of real-time RT qPCR data is described in Sects. 3.4.1–3.4.2. This includes the verification of PCR quality and integrity and the data evaluation and graphical presentation.

3.4.1. Verification of PCR Quality and Integrity

For each real-time RT qPCR experiment the quality and integrity of each individual PCR reaction must be verified before data analysis. 1. Always test the reproducibility of technical repetitions. The validation of reproducibility should be done with care. Samples with little amounts of template DNA tend to show higher deviations because the Ct values of these samples lie at the end of the exponential phase of PCR. However, in contrast to deviations of highly concentrated samples, deviations within replicates of low concentrated samples have only a little impact on the absolute expression level. Overall, we usually consider differences of max. 1 Ct to be acceptable. Samples with higher deviations are excluded from further data analysis. Pipetting 10 µl template DNA improves pipetting accuracy significantly and consequently deviations can be expected to be much lower than 1 Ct (see Fig. 4.3a).

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2. Always determine the PCR efficiency. Real-time RT qPCR is based on the assumption of exact doubling of DNA during each PCR cycle. However, PCR efficiency can be compromised by PCR inhibitors, sub-optimal primer design, or degraded components in the PCR reaction mixture. PCR efficiency can be verified by analyzing the standard curve (see Fig. 4.3b). The slope of the standard curve should be between −3.2 and −3.5, representing a PCR efficiency of approx. 95–105% (efficiency = 1 + 10(−1/slope)) (7). cDNA samples can contain PCR inhibitors not present in the PCR standards. Therefore, it is necessary to test the PCR efficiency at least once with serial dilutions of cDNA samples. Identical slopes of the amplification plots (see Fig. 4.3a) of mass standards and cDNA samples indicate similar PCR efficiencies. Depending on the cDNA synthesis system used a cDNA dilution of 1:5 to 1:10 before PCR was shown to be necessary for elimination of inhibitory effects on the PCR reaction (18). PCR efficiencies higher than 110% or lower than 90% indicate the necessity of additional PCR optimization steps (see Note 12). 3. Validate the PCR specificity. This is determined by meltingcurve analysis and is based on the specific melting temperature (Tm) of amplification products. During melting-curve analysis the fluorescence of each sample is continuously monitored. The PCR block is slowly heated from a temperature below to a temperature above the Tm of the amplification products (e.g., 55–95°C). Loss of the helical structure of a PCR product at its Tm leads to the release of intercalated fluorescent dye and by this, to an overall decrease of fluorescence in the reaction tube. Usually the decrease in fluorescence is displayed as the first negative derivative (−dF/dT) of the melting curve, leading to sharp peaks determining the precise Tm of each amplification product. Because the Tm depends on the size and GC content of the PCR product, melting-curve analysis is a powerful tool to identify amplified products and distinguish them from primer dimers and other amplification artefacts, usually having a Tm different from the Tm of the desired PCR product. In Fig. 4.3c three different melting curves are shown. While the peak height (=change of fluorescence) in the sample with the correct PCR products (circles) is relatively high, PCR samples containing artefacts usually show lower peak heights. In cases of co-amplification of correct PCR product, e.g., primer dimers, the fluorescence data collected at 72°C during real-time RT qPCR cannot be used for quantification of correct PCR product. If additional optimization steps are necessary, it is possible to include an additional temperature step after the elongation step in each PCR cycle, e.g., 20 s at 78°C, to measure the fluorescence of correct PCR product exclusively. However, because the PCR efficiency might be

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compromised if PCR artefacts are formed, it is desirable to optimize the PCR in such a way that PCR artefacts are not present (see Note 12). 3.4.2. Data Evaluation and Graphical Presentation

The operating software of modern real-time RT qPCR cyclers usually offers comfortable opportunities to analyse and display the expression data. Here we describe a manual method of data analysis independent from the operating software. In Fig. 4.4 a concrete step-by-step scheme of data evaluation is presented. See (11) for the biological background. 1. Export the expression data from the operating software to a spreadsheet PC program like Microsoft Excel. 2. Isolate sample names and absolute expression values. 3. Subtract possible expression values of water controls from each individual sample (Fig. 4.4a). For genes where the primer design does not allow to discriminate cDNA from genomic DNA, values from a primer set specific for genomic DNA should be subtracted here.

Fig. 4.4. Data processing and analysis of absolute expression values extracted from the operating software of the realtime RT qPCR machine. For better understanding the formulas used in the spreadsheet program are set in quotation marks. (a) Subtraction of possible water contaminations from the expression data. If necessary, contaminations in the non-template control (NTC) or contaminations by genomic DNA must also be subtracted (not shown). (b) Determination of mean normalization factors using two internal control genes EF1Balpha2 (EF) and UBP6. (c) Normalization of the expression values of the remaining genes by multiplication with the mean normalization factor. (d) Bar diagram showing the normalized absolute expression values. A logarithmic ordinate is useful in displaying large differences in expression levels. (e) Alternative data presentation showing the regulation of the genes in fold changes relative to the control sample.

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4. Determine normalization factors by analyzing the absolute expression data of each internal control gene (Fig. 4.4b). Divide the absolute expression values of a control sample (e.g., Wild type, root, standard condition) by the absolute expression of each individual sample. The mean normalization factors obtained are used to multiply the expression values of all other genes. Truly constitutively expressed internal control genes will yield very similar normalization factors with a low deviation. Perform statistical analysis at this place to test if the internal control genes used are indeed constitutively expressed. 5. For each gene, normalize the absolute expression data of each individual sample with the corresponding mean normalization factor (Fig. 4.4c). The normalized data can be displayed directly in a bar diagram; use logarithmic ordinate if necessary (Fig. 4.4d). Another possibility is the presentation with bars relative to a control sample to emphasize fold changes in the regulation of the genes. For this, divide the absolute normalized expression data from each sample by the absolute normalized expression data of the control sample (Fig. 4.4e).

4. Notes 1. Degradation products of SYBR green I are inhibitors of PCR. SYBR green I is stable at 4°C for approx. 3 weeks in a pH range of 7.5–8.3. As the pH of TE buffer rises when stored in the cold it is important to avoid using the common TE buffer with pH 8.0. The pH at 4°C will rise up to approx. 8.5 leading to fast degradation of SYBR green I (18). 2. Perform the dilution in 1:10 steps, e.g., 100 µl DNA solution to 900 µl PCR grade water, to maximize the accuracy of the dilution series. 3. A practical way to maintain PCR mass standard stocks is to prepare a certain amount of PCR strips in advance. For example, 16 PCR strips with six tubes each will fit a common 96-well PCR rack. Choose PCR racks with lids to be able to store them in stacks for storage at −20°C. 4. It is important to seal the tape very tightly on the plate. Otherwise, the sample can evaporate during PCR. The paper covering the sticky side of the optical tape can ideally be used to wrap around the thumb allowing one to strike tightly over the optical tape without causing scratches. Do not vortex the sealed PCR plate because it is not possible to spin down droplets sticking on the inner side of the sealing tape.

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5. Aliquots of ready-to-use primer mixes and cDNA are stored at −20°C. Harmful freeze–thaw cycles are avoided and the risk of contaminations is minimized. By this, a consistent quality of primer mixes is achieved. 6. We use 70 mg plant material instead of the 100 mg recommended by the manufacturer of the plant RNA extraction kit. In our experience, especially leaf samples tend to clog in the column leading to low yields of RNA. Extracting RNA from less plant material results in higher RNA yield. 7. Very fine tissue powder is a prerequisite for a good yield with any RNA extraction method. When using a preformed metal stick, alternate grinding and mixing is essential to gain homogeneous fine powder. Consequent N2 cooling of the samples and the metal stick are necessary to avoid loss of material. 8. Air dry the RNA pellet until no droplets of ethanol can be seen on the walls of the tube. Depending on the environment 30–60 min drying should be sufficient. Do not use a vacuum centrifuge because the RNA pellet might get too dry leading to low solubility of the RNA difficult. 9. The addition of EDTA and heating to 65°C deactivate DNase I in the sample. In parallel the heating step is used to denature secondary structures of mRNA. When cooling down, oligo(dT)18 primer can anneal before cDNA synthesis. 10. The dilution of the cDNA after cDNA synthesis depends on the cDNA synthesis system used and is performed to exclude inhibitory effects on the PCR reaction. According to Ref. (18) a 1:10 dilution is necessary with the Fermentas cDNA synthesis system, as used in our experiments. 11. When pipetting small volumes such as 10 µl to the PCR plate, touching the inner wall of the wells with the pipette tip can hardly be avoided. The droplet would stick on the pipette tip instead of falling to the PCR tube. To avoid possible contaminations and loss of volume, the 96-well PCR plate is turned horizontally by 180° before pipetting the cDNA. By this, the opposite side of the inner wall of the well will be touched when pipetting the cDNA. 12. Add DMSO (2–10% final) to minimize the formation of primer dimers and to lower PCR efficiencies higher than 110%. At PCR efficiencies below 90% the addition of glycerol (5–10% final) might improve the PCR reaction. Be aware that low PCR efficiencies can also be an indicator of PCR inhibitors present in the mastermix or the cDNA.

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References 1. Weis, J. H., Tan, S. S., Martin, B. K., and Wittwer, C. T. (1992) Detection of rare mRNAs via quantitative RT-PCR. Trends Genet. 8, 263–264. 2. Higuchi, R., Fockler, C., Dollinger, G. and Watson, R. (1993) Kinetic PCR analysis: realtime monitoring of DNA amplification reactions. Biotechnology (N Y) 11, 1026–1030. 3. M. Tevfik Dorak - Real-Time PCR. http:// dorakmt.tripod.com/genetics/realtime.html. 4. Bustin, S. (2002) Quantification of mRNA using real-time reverse transcription PCR (RT-PCR): trends and problems. J. Mol. Endocrin. 29, 23–39. 5. Gachon, C., Mingam, A. and Charrier, B. (2004) Real-time PCR: what relevance to plant studies? J. Exp. Bot. 55, 1445–1454. 6. Bustin, S. A., Benes, V., Nolan, T., and Pfaffl, M. W. (2005) Quantitative real-time RT-PCR-a perspective. J. Mol. Endocrinol. 34, 597–601. 7. Nolan, T., Hands, R. E., and Bustin, S. A. (2006) Quantification of mRNA using real-time RT-PCR. Nature Protocols 1, 1559–1582. 8. Wilfinger, W. W., Mackey, K., and Chomczynski, P. (1997) Effect of pH and ionic strength on the spectrophotometric assessment of nucleic acid purity. BioTechniques 22, 474–476. 9. Pfaffl, M. W. (2001) A new mathematical model for relative quantification in real-time RT-PCR. Nucleic Acids Res. 29, e45. 10. Vandesompele, J., De Preter, K., Pattyn, F., Poppe, B., Van Roy, N., De Paepe, A.,

11.

12.

13. 14.

15.

16. 17. 18.

and Speleman, F. (2002) Accurate normalization of real-time quantitative RT-PCR data by geometric averaging of multiple internal control genes. Genome Biol. 3, RESEARCH0034. Wang, H.-Y., Klatte, M., Jakoby, M., Bäumlein, H., Weisshaar, B., and Bauer, P. (2007) Iron deficiency-mediated stress regulation of four subgroup Ib BHLH genes in Arabidopsis thaliana. Planta. 226(4), 897–908. Zimmermann, P., Hirsch-Hoffmann, M., Hennig, L. and Gruissem, W. (2004) GENEVESTIGATOR. Arabidopsis microarray database and analysis toolbox. Plant Physiol. 136, 2621–2632. geNorm. http://medgen.ugent.be/jvdesomp/genorm/. Pfaffl, M. W., Horgan, G. W., and Dempfle, L. (2002) Relative expression software tool (REST) for group-wise comparison and statistical analysis of relative expression results in real-time PCR. Nucleic Acids Res. 30, e36. Rozen, S. and Skaletsky, H. (2000) Primer3 on the WWW for general users and for biologist programmers. Methods Mol. Biol. 132, 365–386. TAIR - The Arabidopsis Information Resource. http://www.arabidopsis.org/. IDT - Oligo Analyzer. http://www.idtdna. com/analyzer/Applications/OligoAnalyzer/. Karsai, A., Muller, S., Platz, S. and Hauser, M. (2002) Evaluation of a homemade SYBR green I reaction mixture for real-time PCR quantification of gene expression. BioTechniques 32, 790–792.

Chapter 5 Probing Spatio-Temporal Intracellular Calcium Variations in Plants Axel Mithöfer, Christian Mazars, and Massimo E. Maffei Abstract Calcium (Ca2+) ions act as intracellular second messengers in many different signalling processes in plant cells and thus contribute to the amplification step of the signalling pathway and the specificity of the adaptative response. Dynamics of calcium described as spatial and temporal changes of the Ca2+ concentrations either in the cytosol and/or in other compartments of the plant cell are now accepted to generate “calcium signatures”, which might be responsible for the initiation of specific downstream events leading to the mounting of an appropriate response. To identify and elucidate the properties of such signatures, highly sensitive and specific methods have been developed and are used to measure and monitor variations in intracellular Ca2+ concentrations. Two of these methods, namely bio-luminescence and fluorescence in combination with confocal laser scanning microscopy, are presented. Key words: Aequorin, calcium-sensitive dye, confocal laser scanning microscopy, intracellular calcium concentrations, Arabidopsis, tobacco.

1. Introduction Like in animal cells, free intracellular [Ca2+] variations in plant cells are key signals in many regulatory functions, playing a major role in mediating various endogenous and exogenous signals to cellular responses. Thus, [Ca2+] changes have been described during growth and differentiation processes, mitosis, cytosolic streaming, stomata regulation, induction of defence responses, and stress adaptation (1,2). It is now well established that intracellular [Ca2+] variations take place as “calcium signatures” rather than as uniformly shaped changes (3–5). Calcium signatures have been correlated with final responses on the basis of their obvious parameters such as the shape, the amplitude, the duration, T. Pfannschmidt (ed.), Plant Signal Transduction, vol. 479 Humana Press, a part of Springer Science + Business Media, LLC 2009 DOI: 10.1007/978-1-59745-289-2_5

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the compartmentation and, in the case of oscillations, the frequency of the signals (5). To measure transient intracellular [Ca2+] changes in plant cells, two main approaches are often used based on fluorescent and bio-luminescent techniques. Fluorescent techniques use either chameleon probes, which are genetically encoded calcium indicators based on derivatives of green fluorescent protein (GFP) (not described here) or low affinity fluorescence indicators, whereas bio-luminescent techniques mainly use the aequorin technique based on a Ca2+-sensitive protein constitutively expressed in plant cells. Here we focus on the aequorin technique and the use of fluorescent calcium indicators through confocal laser scanning microscopy. Aequorin is a Ca2+-binding photoprotein composed of an apoprotein (apoaequorin), which has an approximate molecular weight of 22 kDa and a prosthetic group, a luciferin molecule, coelenterazine (Mr 432). In the presence of molecular oxygen, the functional holoprotein, aequorin, reconstitutes spontaneously. The protein contains three EF-hand Ca2+-binding sites. When these sites are occupied by Ca2+, aequorin undergoes a conformational change and behaves as an oxygenase that converts coelenterazine into excited coelenteramide, which is released together with carbon dioxide. When the excited coelenteramide relaxes to its ground state, blue light (λ = 469 nm) is emitted (6). This emitted light can be easily detected with a luminometer and correlates with the particular [Ca2+]. Confocal microscopy (CM) is an optical sectioning method used to reduce the image blur that is caused by inclusion of light from outside the plane-of-focus in a cross-section of a thick sample. In CM, the path of out-of-focus light is physically blocked before detection (7). As a comparatively non-destructive imaging technique, confocal laser scanning microscopy (CLSM) has a number of distinct advantages over alternative imaging modalities; primarily CLSM facilitates the in situ characterization of the 3D architecture of tissue microstructure (8). Several papers have addressed the benefits that CM affords during analysis of the spatial properties of intracellular [Ca2+] signals (9–12). However, it is generally difficult to measure [Ca2+] in a non-invasive method and without artefacts. It is also particularly tricky to measure [Ca2+] in a physiologically relevant context, which allows one to compare results obtained in the laboratory with the physiological status of plants in the field. With the emergence of technologies like the transgenic aequorin system or CLSM in combination with sensitive fluorescent probes for localizing intracellular calcium, it will be possible to decipher the earliest signals in the calcium signal transduction pathways and further understand the regulatory role of calcium in cell physiology either in cell division and differentiation, or responses to biotic and abiotic challenges.

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2 Materials 1. 1 l Erlenmeyer culture flasks (VWR, Darmstadt, Germany).

2.1 Measuring Free Calcium Variations in Plant Cells or Plant Seedlings Using the Recombinant Aequorin Technology

2. Linsmaier and Skoog medium with macro- and microelements including vitamins (L0230 DUCHEFA: http:// www.duchefa.com;Haarlem, the Netherlands).

2.1.1 Tobacco BY-2 Cell Culture

5. Ultra High Quality (UHQ) water.

2.1.2 Arabidopsis thaliana Seed Sterilization and Germination

1. Cell culture dishes 120 × 120 mm (VWR, Darmstadt, Germany).

3. Dichlorophenoxy-acetic acid (2,4D) (Sigma, Munich, Germany). 4. Cellulose taps type38 (VWR, Darmstadt, Germany).

2. Cell culture dishes 35 mm (VWR, Darmstadt, Germany). 3. 2 mL microtubes (Sarstedt, Nürnbrecht, Germany). 4. Polyamide open mesh fabrics (SEFAR NITEX mesh opening 37 µm, (Sefar, http://www.sefar.com;Thal, Switzerland). 5. Murashige and Skoog Medium (Sigma, Munich, Germany). 6. Sucrose (Euromedex, Souffelweyersheim, France). 7. Agar (Europio, Courtaboeuf Cedex B, France). 8. Stock solution of 9.6% active chlorine Javel extract (Javel Oxena, Portes-Les-Valences, France). 9. Ethanol 99%. 10. Seeds of Arabidopsis thaliana plants, ecotype RLD1, constitutively expressing a 35S-aequorin construct (13) (see Note 1). 11. UHQ sterile water.

2.1.3 Calcium Measurement

Sirius single tube luminometer (Berthold Technologies, Thoiry, France) with a sample drawer accepting cell culture dishes (35 mm). 1. 50 ml BD Falcon conical tubes (BD Biosciences, Heidelberg, Germany). 2. Polystyrene tubes (3 ml, 11 × 55 mm). 3. Native coelenterazine (Interchim, Mannheim, Germany) as 5 mM stock solution in methanol and stored frozen at −20°C as 10 µl aliquots in 0.2 ml PCR microtubes. 4. Lysis solution: 100 mM CaCl2 in 10% ethanol supplemented with 2% Nonidet P40 (v/v). 5. Minimal medium (MM): 2 mM Mes-KOH pH 5.8, 175 mM mannitol, 0.5 mM K2SO4, 0.5 mM CaCl2. 6. Reconstitution buffer (RB): 10 mM Tris-HCl pH 7.4, 5 mM EDTA, 0.5 M NaCl, 5 mM β-mercaptoethanol, 0.1% gelatine. 7. Dilution buffer (DB): 200 mM Tris-HCl pH 7.0, 5 mM EDTA.

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2.2 Confocal Laser Scanning Microscopy 2.2.1 Microscopy Equipment

The following settings refer to the CLSM Leica TCS SP2 mounted on a Leica DM6000 microscope (Leica Microsystems, Milano, Italy). The microscope has three lasers: Ar (458, 476, 488, 514 nm), Ar/Kr (568, 647 nm) and He/Ne (543, 633 nm). It is equipped with programmable filters, an acousticoptical (AOTF) filter for each wavelength, dicroic mirrors, 4 photomultipliers (3 for fluorescence and 1 for transmitted light) and spectral scanning. Other features are FRET and FRAP. Objectives used are Dry 10x and 20x and immersion (water) 40x and 63x. Scanning modes are 2D, 3D and 4D. Output images and movies formats are TIFF and AVI, respectively. The typical microscope settings for the localization of calcium with the dye calcium orange (see Sect. 2.2.2 for dye specifications) are given in Table 5.1.

Table 5.1 Logical size

Physical length

Physical origin

X: 1024 Pixel

750.00 µm

0.00 m

Y: 1024 Pixel

750.00 µm

0.00 m

Z: 16 Pixel

−149.94 µm

20.00 mm

Voxel information Grey resolution

8 bit

Range begin

0

Range end

255

Type

Intensity

Scanner settings ScanMode

xyz

Pinhole [m]

80.12 µm

StepSize

10.00 µm

Voxel-width

732.42 nm

Voxel-height

732.42 nm

Voxel-depth

10.00 µm

Zoom

1

Scan-direction

Uni (continued)

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Table 5.1 (continued) Logical size

Physical length

Physical origin

Y-scan-direction

Uni

Sequential scan mode

None

Frame-accumulation

1

Frame-average

1

Line-average

4

Section Z

16

Hardware settings Visible AOTF (488)

67.00%

Visible AOTF (543)

100.00%

PMT 2 (Offset)

−78.4

PMT 2 (Voltage)

740.3 V

PMT 3 (Offset)

−18.4

PMT 3 (Voltage)

483.3 V

PMT Trans (Offset)

2.7

PMT Trans (Voltage)

147.7 V

Beam Exp.

3

Excitation beam splitter

FW DD 488/543

TLD_Settings

100

Inverse flag topo

1

Scan field rotation

−0.04°

Z position

−20.35 nm

Scan speed

400 Hz

SP Mirror 2 (left)

574 nm

SP Mirror 2 (right)

580 nm

SP Mirror 2 (stain)

0

Objective

HC PL FLUOTAR 20.0x0.50 DRY

Numerical aperture (Obj.)

0.5

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2.2.2 Calcium Dyes

Among the several calcium indicators the following have been mostly used in plant science: 1. FLUO-3. The most important properties of Fluo-3 are an absorption spectrum compatible with excitation at 488 nm by argon-ion laser sources and a very large fluorescence intensity increase in response to Ca2+ binding. 2. FLUO-4. Fluo-4 and its esterified form Fluo4-AM have a visible wavelength excitation (compatible with argon-ion laser sources) and a large fluorescence increase upon binding Ca2+. 3. FLUO-4FF, FLUO-5F, FLUO-5N. These are analogues of Fluo-4 with lower Ca2+-binding affinity, making them suitable for detecting intracellular calcium levels in the 1 µM to 1 mM range that would saturate the response of Fluo-3 and Fluo-4. 4. FLUO-4 DEXTRANS. These are Fluo-4 coupled to a biologically inert dextran carrier (molecular weight = 10,000), providing a new and potentially valuable tool for measuring Ca2+ transients. 5. CALCIUM GREEN-1. It is structurally similar to Fluo-3, but is more fluorescent at low calcium concentrations, facilitating the determination of base line Ca2+ levels and increasing the visibility of resting cells. 6. CALCIUM GREEN-2. It has two fluorescent reporter groups, which are believed to quench one another in the absence of calcium, and it undergoes a much larger increase in fluorescence emission upon calcium binding than does Calcium Green-1. Its lower affinity for calcium makes it particularly suitable for measuring relatively high spikes of calcium, up to 25 µM. 7. CALCIUM ORANGE and CALCIUM CRIMSON. These are spectrally similar to tetramethylrhodamine and Texas Red. The long-wavelength spectral characteristics of these indicators allow them to be used in combination with fluorescein and ultraviolet excitable dyes. 8. INDO-1 or FURA-2. These are “dual-emission” and “dualexcitation” types of calcium dyes, respectively. To use either of these indicators, however, appropriate modifications of standard CLSM need to be made, for instance, a high-power argon-ion laser is required to obtain ultraviolet (UV) excitation, and compensatory changes along the optical path must be incorporated to deal with the lens aberrations and reduced signal throughputs that are associated with UV illuminations. At low concentrations of the indicator, use of the 340/380 nm excitation ratio for Fura-2 or the 405/485 nm emission ratio for Indo-1 allows accurate measurements of the intracellular Ca2+ concentration.

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Measurements of Indo-1 and Fura-2 fluorescence can usually be made over a period of an hour without significant loss of fluorescence resulting from either leakage or bleaching. In addition, Fura-2 and Indo-1 are bright enough to permit measurements at intracellular concentrations of dye unlikely to cause significant Ca2+ buffering or damping of Ca2+ transients. 9. RHODAMINE-BASED INDICATORS. Rhod-2 has fluorescence excitation and emission maxima at 552 and 581 nm, respectively. Variants with longer-wave length excitation and emission (X-Rhod-1) and lower Ca2+-binding affinity (Rhod-5N, Rhod-FF, etc.) have been developed (i.e., at Molecular Probes). Rhod-2 is used as a selective indicator for mitochondrial Ca2+ in most eukaryotic cells.

3 Methods 3.1 Measuring Free Calcium Variations in Plant Cells or Plant Seedlings Using the Recombinant Aequorin Technology 3.1.1 Tobacco BY-2 Cell Culture

1. Prepare Linsmaier and Skoog (LS) growth medium with 4.3 g l−1 of the LS salts supplemented with 30 g l−1 of sucrose and 1 mg mL−1 2,4D in UHQ water. 2. Adjust to pH 5.8 with 1 M KOH. 3. Pour 200 ml of medium in 1-l culture Erlenmeyer flasks and autoclave for 20 min at 120°C. 4. Inoculate the sterile medium with 2% of a 14-day-old cell culture of BY-2 tobacco cells (The packed cell volume is between 60 and 70%). 5. Grow the cells in the dark at 24°C under constant agitation on a rotary shaker (130 rpm).

3.1.2 Arabidopsis thaliana Seed Sterilization

1. Use a laminar flow hood. 2. Prepare a working solution of Javel (2.4% active chlorine) by diluting 4 times the stock solution with UHQ water (v/v). 3. Prepare the disinfection solution (Sol A) by diluting 6 times the working solution of Javel in ethanol 99%. 4. Put 50 to 100 µl of Arabidopsis seeds in a 2-ml microtube and add 1 ml Sol A, and vortex the tube every 2 to 3 min during an incubation time not exceeding 10 min. 5. After seed decantation, remove the supernatant carefully. 6. Rinse the seeds with 1 ml ethanol 99%, discard the supernatant, and repeat once (see Note 2). 7. Keep the tube open and dry off the excess of ethanol for a maximum of 2 h in the laminar flow hood.

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3.1.3 Arabidopsis thaliana Germination and Growth

1. Prepare a half-strength medium with Murashige and Skoog medium (2.2 g l−1) containing 0.8% agar and 1% sucrose. Sterilize and pour 50 ml of this sterile medium in each cell culture dish with appropriate antibiotic if necessary. 2. When the medium is cool enough, layer on the top of the solidified medium, a sterile 120 × 120 mm Nytex fabric (mesh opening 37 µm) prepared beforehand and sterilized. 3. Prepare a 120 × 120 mm piece of white paper and draw two black lines at, respectively, 40 and 80 mm from the top. Place this paper mask beneath the culture dish where the seeds have to be layered. 4. Distribute the sterile seeds along the two lines. 5. Close the Petri dish with the cover and seal it with parafilm; then place it in the fridge for 48 h to synchronize the germination before placing it for 8 to 10 days in a vertical position in the growth chamber at 23°C, with a 16 h photoperiod and 40% humidity.

3.1.4 Calcium Measurements with Tobacco Cells

1. Harvest the cells from a cell suspension in the exponential growth phase and centrifuge in a 50-ml BD Falcon tube for 5 min at 100 g. Measure the packed cell volume. 2. Remove the supernatant and wash the cells once with two volumes of fresh complete or minimal medium (MM). 3. Resuspend the cells in fresh culture medium or MM, to obtain a 20% packed cell volume. 4. Incubate in the dark 5 to 7 ml of the cell suspension in a 50-ml BD Falcon conical tube with 2.5 µM coelenterazine under constant agitation (130 rpm) for a minimum of 3 h (see Note 3). 5. Using a pre-cut 200-µl pipette tip, remove 50–100 µl of cells pre-incubated with coelenterazine and pour them in the bottom of luminometer tubes. 6. Add the compound to be tested in the minimal volume (1–10% v/v) to prevent osmolarity disturbance or solvent side effect (see Note 4). Homogenate by quickly rotating the tube, place it in the luminometer chamber, and close the chamber to start light measurements. 7. At the end of the chosen duration time for measurement, automatically inject 300 µl of lysis buffer to completely discharge the remaining aequorin.

3.1.5 Calcium Measurements with Seedlings

1. At time minus 15 h before calcium measurements, prepare a 2.5 µM coelentarazine solution in UHQ water from a stock solution.

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2. Distribute 250–500 µl of this solution in the 35-mm culture dish or in the wells of a 24-well microplate for more convenience if several samples have to be tested. 3. Harvest 8- to 10-day-old seedlings from the culture dish and place the roots of bunches of 3–5 seedlings in the wells containing the diluted coelenterazine. Incubate overnight in the dark at room temperature (see Note 5). 4. At time 0, remove the coelentarazine solution and replace it with UHQ water. 5. Test the calcium response of each group of seedlings by placing them in a 35-mm culture dish containing 1 ml of UHQ water placed in the open drawer (chamber) of the luminometer. 6. Remove the water by tilting the luminometer to 45° from the horizontal position and add an equal amount of testing solution (solution containing the compound to be tested for inducing calcium responses) on the opposite side of the roots in the 35-mm culture dish (see Note 6). 7. Close the measuring chamber; the photon counting will start. This counting corresponds to the basal photon emission before any treatment (control). At a chosen time (generally 1–2 min) when the photon baseline is stable tilt back the luminometer to the horizontal position to allow the testing solution to reach the roots of the seedlings. 8. When the calcium response has reached the initial baseline level, open the measuring chamber, tilt the luminometer again, remove the testing solution, and replace it by slowly adding 1 ml of lysis solution. 9. Close the chamber by pushing back the drawer, tilt back the luminometer to the horizontal position to allow the lysis solution to reach the seedlings, and discharge the free aequorin thanks to the calcium-enriched medium. 3.1.6 Control of Putative Side Effects of Chemicals on Aequorin Activity

1. Extract the apoaequorin by grinding in liquid nitrogen a known amount of plant cells or seedlings constitutively expressing aequorin. 2. Resuspend the powder in minimum volume of RB. 3. Collect the supernatant obtained after centrifugation at 12,000 g for 10 min. 4. Reconstitute aequorin by adding coelenterazine (2.5 µM) in the dark for 2 h. 5. Dilute a known volume of the supernatant in 0.5 ml of DB in the presence or absence of the compound that has to be checked in a luminometer tube.

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6. Insert the tube in the luminometer and inject 0.5 ml of 50 mM CaCl2. Compare the luminescence response in the presence and absence of the compound (see Note 7). 3.1.7 Calcium Calibration

1. Transfer all the light values using Excel software in a working sheet. 2. Open a calibration sheet where the formula will be pasted in for calibrating with reference to the RLU values sheet (see Note 8).

3.2 Confocal Laser Scanning Microscopy

3.2.1 Perfusion of Dyes into Leaves and CLSM Observation

Unlike “normal” microscopy, only a spot of a minimum size in the specimen is illuminated in CM, and to enable an image to be formed from this, the specimen must be scanned. The modern CLSM running in fluorescent mode uses a laser light source to excite a fluorescing contrasting agent within an imaged sample. An illumination light is launched from a gas or solid-state laser of one specific or several wavelengths and is subsequently filtered to produce the wavelengths required. The image of the spot is directed through a pinhole stop in an intermediate image plane. The smaller the pinhole, the less stray light or fluorescence from out-of-focus areas within the specimen reaches the detector. It is thus possible to obtain optical sections of the specimen and reconstruct its 3D structure, and vertical “virtual” optical sections are then reconstructed from the resulting 3D blocks. As a result, only light from the focal plane can reach the detector (a photomultiplier). All other (out-of-focus) planes are blocked out. This results in an “optical section”. The images are stored electronically and displayed on a monitor. A series of optical sections can be recorded by moving a motor a slight distance along the z-axis each time an image has been recorded. Such a z-series permits the electronic reconstruction of the three-dimensional structure using suitable computer programs. 1. Dyes, such as Fluo-3 AM (acetoxy-methyl ester of Fluo-3), are purchased in vials containing the molecule as a stock solution in DMSO (see Note 9). 2. Dilute in 50 mM MES buffer, pH 6.0, with the addition of 0.5 mM calcium sulphate and 2.5 µM DCMU (which stops photosynthetic electron flow) to reach the concentration of 5 µM. This resulting solution is used for an initial treatment of leaves. 3. Use leaves attached to the plant. In order to perfuse the dye insert the leaf blade between two glass cover slips. A drop (about 20 µl) of 5 µM Fluo-3 AM solution is applied and covered with another glass slide. 4. The leaf is gently fixed over a glass slide.

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5. Thirty min after treatment with Fluo-3 AM, the leaf is fixed on a confocal scanning laser microscope stative without separating the leaf from the plant. 6. Measurements are taken in intact leaves both in the presence and absence of exogenous calcium. 7. The microscope is operated with a Krypton/Argon laser at 488 and 568 nm wavelengths: the first wavelength excitates the Fluo-3 dye, resulting in emission of green light and the second mostly excitates chloroplasts, which emit red fluorescence. 8. Images generated by the CLSM software are analysed using the image software (see Note 10).

4 Notes

1. Any line of A. thaliana or tobacco or any other plant constitutively expressing apoaequorin can be similarly used. 2. Rinsing in ethanol should be as brief as possible; just vortex the tube, wait for decantation of seeds and discard the supernatant. At the second rinse, as much as possible ethanol should be eliminated and the seeds should be dispersed on the wall of the tube to get a rapid drying. 3. A full aequorin reconstitution takes approximately 3 h; however, incubation can be longer (i.e., overnight). A 20% packed cell volume appears to be, in our hands, a good cell/ volume ratio to be easily pipetted with pre-cut 200 µl pipette tips and to give detectable luminescence responses. 4. If compounds are added solubilized in a solvent such as ethanol or DMSO, we noticed that up to 1% (v/v) ethanol or DMSO has no significant effect on luminescence responses. If compounds have already been added diluted in growth or minimal medium, higher volumes can be injected through the luminometer injector but controls with similar volumes of medium without drugs or with similar amounts of solvent have to be performed. 5. Measuring calcium response using three seedlings gives a strong calcium response easily detected with the luminometer. The response obtained is equivalent to an averaged response that would have been obtained from three independent measurements of light collected from a single seedling. During incubation of seedlings with coelenterazine, do not incubate more than five seedlings together since the

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light emitted could saturate the detector of the luminometer in particular at the step of total aequorin discharge. In addition, if several measurements are necessary prepare sets of three to five seedlings but not more; otherwise, the roots interact with each other during the incubation period with coelenterazine and it is quite impossible then to separate the seedlings without injuring the roots or partially discharging the aequorin through the mechanical shock. A 24-well flat bottom microplate could be appropriated to incubate 24 lots of seedlings in a minimum volume of coelentarazine. 6. This step is important because all the roots of the seedlings have to be arranged on one side of the plate (upper part of the dish on opposite side of the medium after tilting the luminometer) to completely separate the roots from the medium upon tilting the luminometer. This procedure will avoid any untimely contact of the roots with the testing solution before luminescence counting i.e., before closing the drawer and starting the photon counting. 7. This experiment should be done when a new chemical (drugs etc.) is tested since it could directly act on aequorin protein and modify its calcium binding properties giving rise to artefacts. The volume of supernatant diluted in DB should be adjusted to get a significant luminescence response without saturating the photon detector. 8. The emitted light was calibrated into Ca2+ concentrations by a method based on the calibration curve of Allen and coworkers (14): [Ca

2+

1/3 1/3 1/3 ] = {(L 0 / L max ) + [KTR(L 0/ L max ) ] -1}/{KR(L0 / L max ) ]}

L0 is the luminescence intensity per second and Lmax is the total amount of luminescence present in the entire sample over the course of the experiment. [Ca2+] is the calculated Ca2+ concentration, KR is the dissociation constant for the first Ca2+ ion to bind, and KTR is the binding constant of the second Ca2+ ion to bind to aequorin. The luminescence data were determined using the KR and KTR values of 7 × 106 M−1 and 118, respectively, using native coelenterazine and the specific aequorin isoform that we have used in these experiments. If the seedlings express the nucleoplasmin–aequorin construct (15) used to measure nuclear calcium variations, the KR and KTR values of 2 × 106 M−1 and 55, respectively, as calculated (16), should be used. Note also that in each experiment, the concentration of reconstituted aequorin should not be limiting i.e., strong values of RLU should be obtained at the end of the experiment after discharging the remaining aequorin under any of the experimental conditions.

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9. In order to conduct successful calcium imaging studies by CLSM, the dye-loaded cells must remain physiologically viable during imaging. Although a number of fluorescent Ca2+-sensitive small molecule dyes are commercially available, these are all anionic at physiological pH and do not diffuse into cells at any appreciable rate unless extracellular pH is lowered. A wide range of Ca2+ dyes are available, but the majority of cytoplasmic measurements have used the single-wavelength Ca2+ indicators Fluo-3 and Calcium Green-1 (Molecular Probes, Eugene, OR, U.S.A.), notably for confocal measurements. The long-wavelength calcium indicators Calcium Green, Oregon Green 488 BAPTA, Calcium Yellow, Calcium Orange, and Calcium Crimson are visible light-excitable probes derived from fluorescein, Oregon Green 488 dye, lucifer yellow, tetramethyl rhodamine, and Texas Red, respectively. Companies that sell these products (e.g., Molecular Probes) prepare these indicators in the water-soluble potassium salt form, cell-permeant acetoxymethyl (AM) ester form, and in some cases as compartmentalization-resistant dextran conjugates. Upon binding to calcium, these indicators exhibit an increase in fluorescence emission intensity with little shift in wavelength. The spectral characteristics of the long-wavelength indicators have three major advantages: (a) their emissions are in a spectrum range where cellular autofluorescence and scattering backgrounds are often very low; (b) the energy of the excitation light is low, reducing the potential for cellular photodamage; (c) the wavelengths required for optimal excitation are compatible with those produced by laser-based instrumentation, such as CLSM. 10. In situ calibrations are performed by exposing loaded cells to controlled Ca2+ buffers in the presence of ionophores such as A-23187 (A1493), 4-bromo A-23187 (B1494) and ionomycin (I24222). Alternatively, cell permeabilization agents such as digitonin or Triton X-100 can be used to expose the indicator to the controlled Ca2+ levels of the extracellular medium. To determine either the free calcium concentration of a solution or the Kd of a single-wavelength calcium indicator, the following equation is used: ⎡ F − Fmin ⎤ [Ca 2+ ]free = K d ⎢ ⎥ ⎣ Fmax − F ⎦ F is the fluorescence of the indicator at experimental calcium levels, Fmin is the fluorescence in the absence of calcium and Fmax is the fluorescence of the calcium-saturated probe. The dissociation constant (Kd) is a measure of the affinity

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of the probe for calcium. Calibration of the Ca2+ indicator is an essential component of calcium measurements; general reviews of the use of these indicators include those by Ref. (11). The standard curve obtained from the calibration can then be used to convert fluorescence measurements obtained from experimental samples into estimates of free Ca2+ concentration. Loading has been proved to be possible with all forms of dyes, and both ratiometric and non-ratiometric methods reported relative changes in [Ca2+]. However, in general only the ratiometric methods allow good spatial definition of the cytosolic [Ca2+] gradient. References 1. Hepler, P. K., and Wayne, R. O. (1985) Calcium and plant development. Ann. Rev. Plant Physiol. 36, 397–439. 2. Sanders, D., Brownlee, C., and Harper, J. F. (1999) Communication with Calcium. Plant Cell 11, 691–706. 3. Mithöfer, A., Ebel, J., Bhagwat, A. A., Boller, T., and Neuhaus-Url, G. (1999) Transgenic aequorin monitors cytosolic calcium transients in soybean cells challenged with β-glucan or chitin elicitors. Planta 207, 566–574. 4. Walter, A., Mazars, C., Maitrejean, M., Hopke, J., Ranjeva, R., et al. (2007) Structural requirements of jasmonates and synthetic mimetics as inducers of Ca2+ signals in the nucleus and the cytosol of plant cells. Angew. Chem. – Int. Ed. 46, 4783–4785. 5. Allen, G. J., Chu, S. P., Harrington, C. L., Schumacher, K., Hoffmann, T., et al. (2001) A defined range of guard cell calcium oscillation parameters encodes stomatal movements. Nature 411, 1053–1057. 6. Kendall, J., and Badminton, M. N. (1998) Aequorea victoria bioluminescence moves into an exciting new era. Trends Biotechnol. 16, 216–224. 7. Reddy, G. V., Gordon, S. P., and Meyerowitz, E. M. (2007) Unravelling developmental dynamics: transient intervention and live imaging in plants. Nature Rev. Mol. Cell Biol. 8, 491–501. 8. Foldes-Papp, Z., Demel, U., and Tilz, G. P. (2003) Laser scanning confocal fluorescence microscopy: an overview. Int. Immunopharmacol. 3, 1715–1729.

9. Fricker, M., Runions, J., and Moore, I. (2006) Quantitative fluorescence microscopy: From art to science. Ann. Rev. Plant Biol. 57, 79–107. 10. Stricker, S. A., and Whitaker, M. (1999) Confocal laser scanning microscopy of calcium dynamics in living cells. Microsc. Res. Tech. 46, 356–369. 11. Takahashi, A., Camacho, P., Lechleiter, J. D., and Herman, B. (1999) Measurement of intracellular calcium. Physiol. Rev. 79, 1089–1125. 12. Tsien, R. Y., Miyawaki, A., Llopis, J., Griesbeck, O., Zacharias, D., et al. (1999) New molecular sensors and coupling mechanisms for calcium signals. FASEB J. 13, A1514. 13. Knight, M. R., Campbell, A. K., Smith, S. M., and Trewavas, A. J. (1991) Transgenic plant aequorin reports the effects of touch and cold-shock and elicitors on cytoplasmic calcium. Nature 352, 524–526. 14. Allen, D. G., Blinks, J. R., and Prendergast, F. G. (1977) Aequorin luminescence: relation of light emission to calcium concentration – a calcium-independent component. Science 195, 996–998. 15. Badminton, M. N., Kendall, J. M., SalaNewby, G., and Campbell, A. K. (1995) Nucleoplasmin-targeted aequorin provides evidence for a nuclear calcium barrier. Exp. Cell Res. 261, 236–243. 16. van der Luit, A. H., Olivari, C., Haley, A., Knight, M. R., and Trewavas, A. J. (1999) Distinct calcium signaling pathways regulate calmodulin gene expression in tobacco. Plant Physiol. 121, 705–714.

Chapter 6 Dynamic Redox Measurements with Redox-Sensitive GFP in Plants by Confocal Laser Scanning Microscopy Andreas J. Meyer and Thorsten Brach Abstract Continuous control of metabolism and development is a key feature of life and is of particular importance under stress conditions. While under normal conditions most cellular compartments maintain a reducing environment, the cellular redox state can be influenced by external factors. Redox changes might in turn be employed as part of a signalling cascade leading to molecular responses to adverse situations. To enable dynamic measurements of the cellular redox poise in vivo, reduction-oxidation sensitive GFP (roGFP) can be expressed in plant cells and observed by confocal microscopy. When imaged by confocal microscopy this probe exhibits significant opposing shifts in the fluorescence intensities excited at 488 and 405 nm upon formation of an intramolecular disulfide bridge, which enables ratiometric analysis. The formation of the disulfide bridge is directly responsive to the redox state of the glutathione redox buffer within the subcellular compartment to which roGFP is targeted. Key words: Redox-sensitive GFP·, Arabidopsis·, glutathione·, glutaredoxin·, GSH·, GSSG·, ratiometric imaging.

1. Introduction Cellular redox homeostasis is a key feature of all living organisms. Cells maintain a highly reducing internal milieu in most subcellular compartments like the cytosol, the plastids and the mitochondria. Metabolic functions of enzymes in these compartments depend on reducing conditions and it has been known for long that certain metabolic conditions lead to changes in the redox state and hence the activity of certain enzymes. The best described examples are thioredoxin-dependent physiological processes that can be switched off and on depending on whether the plant is exposed to light or not (1–4). In this way deviations

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from the steady state redox poise are exploited for signalling purposes. Similarly, abiotic and biotic stress factors often trigger increased concentrations of reactive oxygen species (ROS) and thus a partial oxidation of the cellular milieu (5,6). Cells are capable of sensing this oxidation and then respond appropriately to the respective stress factor. However, analysis of the underlying signalling mechanisms so for has been hampered because changes in the cellular redox poise could not be measured dynamically. The recent development of redox-sensitive GFP (roGFP) now offers a possibility to overcome the limitations of earlier static measurements of ROS (7). roGFP contains two cysteine residues engineered into the 11-stranded β-barrel such that they are located on two adjacent β-strands. Formation of a disulfide bridge between these two cysteines leads to slight conformational changes around the chromophor that cause distinct changes in the spectral properties (8,9). In consequence, the excitation spectrum of roGFP exhibits two excitation peaks with opposite redoxdependent behaviour, which enables ratiometric measurements. In vitro, roGFP is responsive to changes in the ambient redox potential, but these changes are slow. In contrast to the in vitro situation, the sensor responds much faster when expressed in Arabidopsis cells and it has been shown that this faster response is dependent on the presence of glutaredoxins (GRXs) capable of interacting with roGFP (10). GRXs mediate equilibration of roGFP with the redox potential of the cellular glutathione redox buffer. Thus, changes in the glutathione redox potential are immediately transduced to roGFP (10). Because GRXs can reversibly oxidize and reduce the artificial target roGFP, imaging of roGFP fluorescence for the first time allows dynamic monitoring of distinct changes in cellular redox poise. 1.1. Properties of roGFP

The chromophor of GFP is formed through intramolecular cyclization of the three amino acids Ser65-Tyr66-Gly67. Subsequent oxidation leads to formation of a conjugated system of Π-electrons capable of absorbing and emitting visible light. Wild-type GFP has two excitation peaks that reflect different protonation states of the phenol residue of Tyr66 within the chromophor. The first peak with a maximum at 395 nm corresponds to the protonated, neutral form of the chromophor (A-band) and the second peak with a maximum at 475 nm corresponds to the de-protonated, anionic form (B-band) (11). Although wild-type GFP contains two cysteine residues (Cys48 and Cys70), the molecule is not sensitive to changes in the redox conditions in the surrounding medium. Substitution of two surface exposed amino acid residues, located on the adjacent antiparallel β-strands 7 and 10 of the GFP barrel with cysteines, renders the GFP molecule redox-dependent (8). To avoid undesirable redox reactions the surface exposed Cys48 was replaced by serine, a mutation that has no impact on the fluorescent properties of the chromophor. Formation of a disulfide

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bridge between the two engineered cysteines results in small but significant movements on the part of both the main chain and side chains in its vicinity, resulting in a change in the chromophor environment (8). Hanson et al. supposed that these changes in the chromophor environment perturb the equilibrium between neutral and anionic forms of the chromophor (8). 1.2. Variants of roGFP

Different versions of roGFP have been described (8,9). The version roGFP1 is derived from the original GFP sequence and still contains the chromophor Ser65-Tyr66-Gly67. The version roGFP2 differs only in that Ser65 is replaced by Thr65. This mutation leads to a major shift in the excitation properties with the main excitation peak shifting from 395 to 490 nm. In addition the S65T mutation renders roGFP2 slightly pH-sensitive, leading to a quenched fluorescence emission at low pH. The ratiometric properties in general, however, are unaffected ((12), Meyer and Brach, unpublished data). The pKa of roGFP2 also depends slightly on the degree of oxidation with a pKa of 5.6 in the fully reduced form and a pKa of 6.0 in the fully oxidized state (8) (see Note 1). The midpoint redox potential of thiols is dependent on the degree of protonation and will become more negative with increasing pH. Although this midpoint potential shift would ideally depend on the pH according to the Nernst equation, a slight deviation was observed for roGFP2 due to titratable amino acid residues close to the cysteine residues (8). However, as long as the pH is constant this pH-dependence is not relevant for quantitative analysis of the measured fluorescence ratios. Several further changes in amino acids close to the reactive cysteines have been described (13,14). Introduction of basic amino acids in positions around the cysteines causes the pKa of the cysteines to increase and hence renders the redox potential slightly less negative. The additional amino acid modifications had some minor effects on the equilibrium constant for the equilibration of roGFP with the redox pairs in the bulk solvent. Despite being less reducing, roGFP2 (E’0 = −280 mV) was generally oxidized (somewhat) faster than roGFP1 (E’0 = −291 mV), indicating that kinetic aspects play an important role in determining the speed of equilibration. These differences as well as the changes due to introduction of basic amino acids around the roGFP cysteines mentioned here are only of minor importance when roGFPs are expressed in Arabidopsis. In this case roGFP directly interacts with glutaredoxins (GRXs), which are present in large numbers in Arabidopsis. This interaction makes the equilibration of roGFP with glutathione as a redox buffer several orders of magnitude faster than in the absence of GRXs and also several orders of magnitude faster than roGFP equilibration with other redox pairs. In the presence of catalysing GRXs, roGFP is consequently a specific in vivo redox sensor of the glutathione redox potential (10). The first roGFP versions generated were developed in a wildtype GFP background with the codon usage of Aequorea vic-

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toria. While such proteins can be expressed in bacteria, plants and the model plant Arabidopsis in particular recognize a cryptic intron leading to aberrant mRNA processing (15). For expression in plants it is thus important to use codons that avoids artificial splicing. This can be achieved with constructs adapted to the human codon usage (16). 1.3. Imaging roGFP Fluorescence

The redox-dependent fluorescence of roGFPs can be imaged on a conventional epifluorescence microscope equipped with a fast filter changer and a cooled CCD camera (8,9,17). Epifluorescence imaging, however, suffers from strong out-of-focus blur and thus is not useful to obtain roGFP fluorescence along the z-axis in thick specimen. In addition, autofluorescence from phenolic compounds in the cell walls and secondary metabolites in the vacuole can be a major problem in plant specimen. Particularly after excitation with UV-light this autofluorescence may obscure the redox-dependent signal from roGFP. Moreover, roGFPs can be specifically targeted to all different subcellular compartments, which also emphasizes the demand for high-resolution imaging. This demand for highresolution imaging is fulfilled by confocal laser scanning microscopy (CLSM), and with the broad availability of reasonably priced UV laser diodes for excitation at 405 nm, it is possible to do confocal ratiometric imaging of roGFPs. In contrast to epifluorescence microscopy, for which the excitation wavelengths can be chosen fully flexibly from the continuous spectrum of the excitation source, CLSM is restricted to a few discrete laser wavelengths. For ratiometric redox imaging with roGFPs on most configured instruments, this will be 405 and 488 nm. This restriction has some implications on the choice of the most suitable roGFP version. roGFP1 exhibits excitation maxima at 400 and 475 nm with the A-band (400 nm) being the more prominent one. In addition to the change in the relative intensity of the two excitation peaks, the B-peak in roGFP2 is also shifted by about 15 nm to a maximum of 490 nm (9). The spectral shift means that roGFP2 can be excited at both laser wavelengths almost on its excitation peak, whereas excitation of roGFP1 would be off the peaks. This results in a larger dynamic range for roGFP2 when the fluorophors are imaged by confocal microscopy. The described protocols, however, apply to both versions of roGFP.

2. Materials 2.1. Expression and Purification of Recombinant roGFP

1. Expression vector pET-30a(+) (Novagen, Darmstadt, Germany) for expression of roGFP with an N-terminal His-tag. 2. Escherichia coli HMS174.

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3. LB medium with 50 mg/l kanamycin. 4. About 1 mM isopropyl β-D-1-thiogalactopyranoside (IPTG, Duchefa, Haarlem, the Netherlands). 5. Sonificator. 6. Centrifuge. 7. Ni-NTA column: flush a HiTrap chelating HP column (GE Healthcare, München, Germany) with 50 mM NiCl2 solution at a flow rate of 1 ml/min to load the column with Ni2+. 8. Binding buffer: 50 mM Tris-HCl, pH 8.0, 250 mM NaCl, 1 mM phenylmethanesulphonylfluoride (PMSF, Serva, Heidelberg, Germany). 9. Washing buffer: 50 mM Tris-HCl, pH 8.0, 250 mM NaCl, 50 mM imidazole (Sigma, Deisenhofen, Germany). 10. Elution buffer: 50 mM Tris-HCl, pH 8.0, 250 mM NaCl, 250 mM imidazole (Sigma). 11. About 10 mM EDTA. 12. About 0.02% NaN3 for column storage. 2.2. In Vitro Characterization of roGFP Fluorescence

1. Fluorimeter. 2. Two Quartz cuvettes. 3. About 10 mM dithiothreitol (DTTred) for reduction and 10 mM trans-4,5-dihydroxy-1,2-dithiane (DTTox, Sigma) for oxidation of roGFP. 4. About 10 mM H2O2. 5. Fluorescence platereader capable of dual excitation, with filter sets for excitation of roGFP at both excitation peaks and an emission filter for GFP fluorescence. 6. Multi-well plates with a clear bottom for fluorescence measurements (Greiner Bio-One, Frickenhausen, Germany). 7. Phosphate buffer: 100 mM K2HPO4/KH2PO4, pH 7.0, 1 mM EDTA; store at room temperature. 8. Desalting columns (Zeba Desalt Spin Columns, Pierce, Rockford, IL)

2.3. Transient Expression of roGFP in Tobacco Leaves

1. Tobacco (Nicotiana tabacum L. or N. benthamiana Domin.) plants grown on soil for about 6 weeks. 2. Agrobacterium tumefaciens strain harbouring a binary vector, with roGFP behind a strong promoter such as the CaMV 35S promoter. If appropriate, a targeting sequence for different organelles can be included. 3. LB medium with the appropriate antibiotic resistance selection drug for the roGFP construct carried by these bacteria.

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4. Infiltration medium: 5% (w/v) sucrose, 10 mM MgCl2 (see Note 2). 2.4. Stable Expression of roGFP in Arabidopsis thaliana Plants with Different Genetic Background

1. Wild-type or mutant plants that are grown under long-day conditions to induce flowering. 2. Agrobacterium tumefaciens strain harbouring a binary vector with roGFP behind a strong promoter such as the CaMV 35S promoter. If appropriate, a targeting sequence for different organelles can be included. 3. Dipping medium: 5% (w/v) sucrose, 0.02% (v/v) Silwet-77. 4. Agar plates (1%) with Murashige and Skoog (MS) medium (Duchefa) containing the appropriate selection antibiotic and 250 mg/l cefotaxime (Duchefa). 5. Fluorescence stereomicroscope with appropriate filter combinations for observation of roGFP1 or roGFP2 (see Note 3).

2.5. Ratiometric Confocal Imaging of roGFP Fluorescence

1. Agar plates (1%) with MS medium containing the appropriate selection antibiotic. 2. Surface sterilizing solutions: 6% (w/v) sodium hypochlorite (NaOCl) with 0.02% (v/v) Tween-20, 70% ethanol. 3. Sterile distilled water. 4. Confocal microscope with excitation wavelengths at 405 and 488 nm. 5. Glass microscope slides. 6. Microscope coverslips (22 × 40 × 0.17 mm) (Assistent, Karl Hecht, Sondheim, Germany) 7. Insulation tape (150 µm thick). 8. Propidium iodide (PI, Molecular Probes/Invitrogen, Eugene, OR): prepare 5 mM stock solution in water and freeze in single use (50 µl) aliquots at −20°C. 9. Nutrient medium: prepare 1 M KNO3, 1 M KH2PO4, pH 5.6, 1 M MgSO4, 1 M Ca(NO3)2 and 20 mM Fe-EDTA. Store these stocks at 4°C. For experiments prepare nutrient medium consisting of 5 mM KNO3, 2.5 mM KH2PO4, pH 5.6, 2 mM MgSO4, 2 mM Ca(NO3)2 and 0.05 mM FeEDTA. Adjust pH to 5.8 with KOH and autoclave. 10. Perfusion chamber fed by gravity flow.

2.6. Calibration of roGFP Fluorescence

1. Confocal microscope with excitation wavelengths at 405 and 488 nm. 2. Microwell plate with a coverslip bottom for microscopic applications (see Note 4). 3. Dithiothreitol (DTTred) for reduction and trans-4,5-dihydroxy-1,2-dithiane (DTTox, Sigma) for oxidation of roGFP.

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3. Methods Recombinant roGFP isolated from bacteria under aerobic conditions is always fully oxidized after purification. The reduction of roGFP by DTT can be followed on a fluorimeter or on a plate reader. To obtain reduced roGFP, the protein can be incubated with 2 to 10 mM DTT for 10 min. In order to remove free DTT from the medium, the roGFP solution can be rapidly desalted on spin columns. The following procedures for confocal imaging of roGFP fluorescence are written for applications based on a Zeiss LSM510 Meta equipped with a UV-laser diode (Point Source i-flex 2000, 25 mW), an Argon laser (LASOS LGK 7812 ML-4, 30 mW) and a green Helium-Neon laser (LASOS LGK 7786 P, 1 mW). All applications can be easily adapted to any other confocal microscope equipped with equivalent laser light sources. 3.1. Expression and Purification of Recombinant roGFP

1. Clone the roGFP sequence with NcoI and NotI into the expression vector pET30a(+) generating an N-terminal Histag. 2. Transfect E. coli HMS174 with the expression vector and select for kanamycin resistance. 3. Grow liquid cultures and induce expression of His:roGFP with 1 mM IPTG. 4. Harvest cells after 20 h growth at room temperature to prevent the formation of inclusion bodies. 5. Centrifuge cells for 10 min at 12,100 g and 4°C. 6. Resuspend the pellet in 10 ml binding buffer and destroy cells by sonification for 5 min. 7. Remove cell debris by centrifugation for 15 min at 27,200 g and 4°C. 8. Filter the supernatant through a 0.45-µm filter to obtain a sterile crude extract. 9. Rinse the Ni2+-charged HiTrap chelating HP column with binding buffer for 10 min at a flow rate of 1 ml/min. 10. Apply the crude extract to the column using a pump for 30 min. 11. Rinse with washing buffer for 5 min to remove weakly bound proteins. 12. Elute roGFP protein by flushing the column with elution buffer for 5 min. 13. Rinse column with 10 mM EDTA to remove Ni2+ from the column. 14. Store column loaded with 0.02% NaN3 at 4°C.

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15. The amount of eluted protein can be determined by a Bradford assay. 16. Protein can be stored at −80°C. 3.2. In Vitro Characterization of roGFP Fluorescence

1. Determine the spectral properties on a fluorimeter. About 1 µM of roGFP (see Note 5) in 100 mM potassium phosphate buffer pH 7.0 is placed in a cuvette and excitation spectra are collected with emission at 510 nm (Fig. 6.1). 2. Measure the excitation spectrum of fully oxidized roGFP. To oxidize roGFP, add 10 mM DTTox or 10 mM H2O2 to the buffer (see Note 6). 3. Measure the excitation spectrum of fully reduced roGFP. To fully reduce roGFP, add 10 mM fresh DTTred to the buffer and incubate for 15 min.

3.3. Transient Expression of roGFP in Tobacco Leaves

1. Inoculate 4 ml of LB medium containing the appropriate selection antibiotic with a single Agrobacterium colony carrying the vector for roGFP and grow for 24 h at 28°C and shaking at 200 rpm. 2. Pellet 2 ml bacteria by centrifugation at 16,000 g and discard the supernatant. 3. Wash the bacteria twice with 1 ml of infiltration medium at 4°C and pellet in between. 4. Resuspend the pelleted bacteria in infiltration medium to OD600 of 0.2–0.6. 5. Pressure infiltrate leaves of well-watered tobacco plants using a plastic syringe without a needle.

Fig. 6.1. Excitation spectra of roGFP2. Purified recombinant protein was incubated in phosphate buffer in the presence of 1 mM DDTox or DTTred. (□) fully reduced roGFP2; (○) fully oxidized roGFP2. Emission was always measured at lem = 510 nm.

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6. Mark the infiltrated leaf area with a black marker pen. 7. Place plants in a growth room under normal growth conditions 8. At appropriate times (see Note 7) excise 1 cm2 leaf pieces from the infiltrated leaf area and mount the piece in a drop of 50 µM PI solution on a glass microscope slide. Cover with a coverslip. 9. Examine roGFP expression in epifluorescence mode first before continuing with confocal imaging for ratiometric redox measurements (see Sect. 3.5). 3.4. Stable Expression of roGFP in Arabidopsis Plants with Different Genetic Background

1. Grow a 4 ml overnight starter culture of Agrobacterium harbouring the desired roGFP construct. 2. Inoculate 300 ml LB medium with this starter culture and grow for ~3 h at 28°C and shaking at 200 rpm to an OD600 of 0.8–1.0. 3. Centrifuge for 15 min at 12,100 g and 4°C. 4. Resuspend the pelleted cells in dipping medium to an OD600 of 0.8–1.0. 5. Dip Arabidopsis flower stalks for 15 s and leave in humid conditions under low light for 24 h. 6. Grow plants for seed production. 7. Collect seeds and store in a dry place at room temperature. 8. Screen transformed seeds on agar plates with appropriate selection antibiotics and verify roGFP expression in resistant plants under a fluorescence stereomicroscope. 9. Grow roGFP expressing plants and produce T2 seeds.

3.5. Ratiometric Confocal Imaging of roGFP Fluorescence

1. For growth of sterile Arabidopsis seedlings perform sterilization and plating of seeds in a laminar flow hood. Place 40–60 transgenic Arabidopsis seeds in a 1.5-mL Eppendorf tube and sterilize by brief incubation with 1 ml 6% NaOCl supplemented with 0.02% Tween-20 and subsequent washing with 1 ml 70% ethanol at room temperature. 2. Wash the seeds twice with 1 ml sterile water, and transfer seeds with a thin pipette to agar plates containing the respective antibiotic for selection of the transgene and 250 mg/l cefotaxime for inhibition of bacterial growth. Alternatively, seeds can be placed on sterile filter paper first, and then transferred to the agar plates with a needle. Wetting the needle tip makes transfer of single seeds easier. About 20–30 seeds can be placed in rows on a single plate. 3. Close the agar plates with micropore tape to allow gas exchange. 4. Place plates for 2 days at 4°C in the dark to ensure even germination.

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5. After stratification place the agar plates in vertical orientation in a growth chamber at 21°C for germination. 6. Pre-screen seedlings under a fluorescence stereomicroscope for high and even roGFP expression. 7. Mount samples expressing roGFP in 50 µM PI (see Note 8) on glass slides for CLSM. For tobacco leaves transiently expressing roGFP, excise 1 cm2 omitting the larger veins. Arabidopsis seedlings germinate after 3 days and can be used at any time after germination. For investigation of redox effects in primary roots 5-day-old seedlings are most useful (see Note 9). 8. Cover with a coverslip and fix the coverslip to the slide. This may be of particular importance for imaging on an inverted stage (see Notes 10 and 11). 9. Observe the sample for green fluorescence in epifluorescence mode using 490 nm excitation (can be obtained with an FITC filter set or a GFP filter with 470/20 nm excitation and 505–530 nm emission). 10. For tobacco leaves choose cells with intermediate or stronger fluorescence (see Note 12). 11. Switch on the 405-nm laser diode, the Ar-laser for excitation at 488 nm and the HeNe-laser for excitation at 543 nm. Set the output of the Ar-laser to 50% of its maximum value (see Note 13). 12. Start configuration of the scanning protocol. Dual excitation of roGFP requires imaging in two separate tracks because emission for both excitation wavelengths is green and needs to be detected on the same channel. To guarantee correct ratiometric analysis of the two images, the images need to be taken in line mode with laser switching after each line. In track 1 the sample is excited with 488 nm for the anionic form of roGFP and 543 nm for PI. In addition, a transmission image can be collected. In track 2 the sample is excited with 405 nm for the neutral form of roGFP. 13. Start the scanning process with only excitation at 488 and 543 nm (Track 1) and ‘fast’ scanning. While looking at the scanned sample reduce the transmission of the acoustooptical tunable filters (AOTF) to the minimum level that allows clear detection of cells. Adjust the gain settings for the photomultiplier tube (PMT) and the amplifier offset to achieve an optimal signal-to-noise ratio. For careful adjustment of images use the image range indicator palette in which black pixels are displayed in blue and white pixels in red. To prevent artefacts, the gain should be adjusted to a value such that only a few red pixels appear in the image.

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The amplifier offset should be adjusted in a way that only a few blue pixels appear in the image and that non-cellular regions appear in grey values just above zero. Also, adjust the pinhole for a suitable depth of the image. The fluorescence intensity of the image generated with 488 nm excitation should be in the upper part of the full dynamic range of the PMT. 14. Activate track 2 and adjust the AOTF to generate a sufficiently bright image (see Note 14). The intensity should be adjusted to the lower part of the full dynamic range of the detector. 15. Place a fully oxidized sample on the microscope (i.e., a sample in 10 mM H2O2) and repeat Steps 13 and 14. For the fully oxidized sample the 488-nm image should be at the lower end of the intensity range and the 405-nm excited image should be significantly brighter than in the reduced sample (see Note 15). 16. Start collecting images with 12 bit in a 512x 512-pixel frame and averaging over 4 lines (see Note 16). 17. To avoid using regions of the images where the signal strength was below that known to give reliable ratio results in the evaluation, such pixels need to be masked out through a thresholding approach. Calculate the image ratio by dividing the images acquired with 405 nm excitation by those collected with 488 nm excitation on a pixel-by-pixel basis (see Note 17). 18. Map the resultant ratio image to a pseudo-colour Look Up Table, which is adjusted such that ratios indicating oxidized roGFP are displayed in red and ratios indicating reduced roGFP are shown in blue. Background regions without roGFP expression are black. 19. Collected images for raw data, PI labelling as proof of cell viability and calculated ratio images can be assembled in Adobe Photoshop (Fig. 6.2). 3.6. Calibration of roGFP Fluorescence

1. Set up fresh solutions of DTTred (10 mM) and DTTox (10 mM) in phosphate buffer, pH 7.0. Degas the solutions and flush with N2 to reduce air oxidation. 2. Set up a calibration series with 1 µM roGFP in DTT buffer solutions with known redox potentials. The midpoint redox potential E’0 of DTT at pH 7.0 is −323 mV (8). For full reduction of roGFP use only DTTred and for full oxidation use only DTTox. 3. Place a drop of the roGFP solutions and a background control without roGFP in a multi-well plate with a coverslip bottom and incubate for 30 min in a N2 atmosphere.

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Fig. 6.2. Ratiometric imaging of Arabidopsis roots expressing roGFP2. (a) Maximum projections of serial optical sections through an Arabidopsis root. Images were collected with a 10x lens before and after addition of 10 mM H2O2. Images were collected with a grey resolution of 4096 levels (12-bit digital range). Ratio images were calculated and mapped to a pseudo-colour Look Up Table. The mean ratio for the control image was 0.27 ± 0.08 and for the H2O2 treated root 1.06 ± 0.48. Scale bars = 50 µm. (b) Fluorescence intensity distribution for control roots containing fully reduced roGFP and (c) roots treated with 10 mM H2O2 so that the roGFP is fully oxidized. Histograms for 488 nm excitation are shown in the lower panel and histograms for 405 nm excitation in the upper panel (see Color Plates ).

4. Place the multi-well plate on the microscope stage and image the fluorescence in each well with exactly the same instrument settings as used before for the in vivo measurements (see Note 18). 5. The degree of oxidation (OxDroGFP) can be calculated according to the following equation: 6. OxDroGFP =

(I 488red

R − R red / I 488ox )(R ox − R) + (R − R red )

7. In this equation R is the actual 405/488 nm ratio, Rred is the 405/488 nm ratio for fully reduced roGFP and Rox is the ratio for fully oxidized roGFP. The factor I488red/I488ox refers to the fluorescence intensities measured with excitation at 488 nm for fully oxidized and fully reduced roGFP (see Note 19). 8. Once the degree of oxidation has been determined it can directly be converted into the actual redox potential according to the following equation: ′ − 9. EroGFP = E0roGFP

RT 1 − OxDroGFP In zF OxDroGFP

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4. Notes 1. Because roGFP1 is less pH-sensitive than roGFP2, roGFP1 has been suggested to be the first choice for redox imaging (8). While this might hold true when imaging is done with an imaging system in which the choice of excitation wavelengths is fully flexible, CLSM imaging allows only a few distinct wavelengths to be used. With the excitation wavelengths 405 and 488 nm, roGFP2 offers a dynamic range of about 5-fold whereas roGFP1 shows only a 2- to 3-fold ratio change between fully reduced and fully oxidized protein. Provided that no major pH-shifts occur, roGFP2 is thus more suitable for CLSM imaging because of its significantly larger dynamic range. 2. Acetosyringone (3′,5′-dimethoxy-4′-hydroxy acetophenone; Aldrich) might be added to a final concentration of 100 µM to facilitate better infection. 3. For roGFP2 a standard GFP filter combination with excitation at 470/40 nm and emission at 525/50 nm works fine. For roGFP1 the excitation filter should be replaced with a filter of 400/40 nm or similar. 4. For upright microscopes silicone gaskets can be fixed to a slide so that small volumes of roGFP solution can be placed in the no-leak wells. 5. The exact total amount is not critical as the ratiometric analysis is independent of the protein concentration. 6. Longer incubation in H2O2 may destroy the protein, resulting in loss of overall fluorescence. The relative intensities of the two excitation peaks, however, remain unaffected and still indicate fully oxidized roGFP. 7. The expression needs to be monitored for up to 5 days to identify the time point with the best expression and correct targeting of roGFP. Especially when roGFP is meant to be targeted to organelles, some residual protein might be left in the cytosol, thus obscuring the redox signal from the organelles. 8. For extended time lapse imaging use a nutrient medium without PI as this will very slowly leak into the cells. For control of root viability, the PI solution can be perfused at the end of the experiment. PI does not label shoot tissues as efficiently as roots due to restricted penetration. 9. Roots can be used at a later stage, but the transfer of roots to microscope slides is more difficult with increasing root length. Alternatively, for imaging with an inverted microscope it is possible to grow roots in coverslip-based chambers (Nunc).

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A small amount of nutrient medium with agar is placed into these chambers and sterilized seeds are placed on the surface. Roots will grow down through the agar and further along the coverslip at the bottom. Roots grown in this way are ideally suited for extended time-lapse imaging. The chamber also allows treatment of seedlings from the open top. 10. To avoid root squashing, insulation tape with a thickness of 150 µm can be used as a spacer between the slide and the coverslip. 11. To treat seedlings with different nutrient solutions, inhibitors of distinct metabolic pathways or toxic compounds, seedlings are best placed in a perfusion chamber with a coverslip bottom. To avoid movement of the seedlings in the perfusion flow, it is necessary to clamp the seedling to the coverslip with a grid. 12. Transgenic Arabidopsis seedlings can be screened for strong and even fluorescence on a fluorescence stereomicroscope before transfer of seedlings to a slide for CLSM. 13. Lasers need at least 30 min to warm up and stabilize. 14. The images are scanned in line switching mode. Because of the rapid line switching the PMT cannot be re-adjusted while switching from one excitation wavelength to another. In this case the gain and offset of the PMT and the pinhole size cannot be adjusted and thus the only option to modify the intensity of the image generated with 405 nm excitation is to alter the intensity of the excitation light. 15. Steps 13 and 14 must be iterated to find ideal settings for ratio images using the maximum achievable dynamic range. 16. z-Stacks can be collected if this is applicable. For collection of z-stacks care needs to be taken to ensure that extended imaging of one area of the tissue does not cause artificial oxidation of the probe due to the imaging. Thus, laser power should be reduced to the absolute minimum required to produce a clear image with a sufficient signal-to-noise ratio. 17. Larger numbers of images can be ratioed automatically by custom written software in ImageJ (version 1.37v, Wayne Rasband, NIH, USA) or MatLab (The MathWorks Inc.). In this case a new output Look Up Table (LUT) to colour code the ratio images can also be applied automatically. 18. If the fluorescence of the roGFP solution is much brighter than the fluorescence measured in vivo, the concentration of roGFP may need to be reduced to avoid exceeding the dynamic range used. 19. If the isosbestic point could be used as a reference point for excitation, the coefficient I488red/I488ox would become 1, and thus the equation would simplify to OxDroGFP = (R − Rred)/ (Rox − Rred).

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Acknowledgements We thank James Remington (University of Oregon) for providing the roGFP constructs. This work was financially supported by a grant from the University of Heidelberg and the Deutsche Forschungsgemeinschaft (DFG) (Grant ME1367/3–2). Thorsten Brach was supported by the Schmeil-Stiftung. References 1. Schürmann, P. and Jacquot, J. -P. (2000) Plant thioredoxin systems revisited. Annu. Rev. Plant Physiol. Plant Mol. Biol. 51, 371–400. 2. Buchanan, B. B. and Balmer, Y. (2005) Redox regulation: a broadening horizon. Annu. Rev. Plant Biol. 56, 187–220. 3. Buchanan, B. B. (1980) Role of light in the regulation of chloroplast enzymes. Annu. Rev. Plant Physiol. 31, 341–374. 4. Scheibe, R. (1991) Redox-modulation of chloroplast enzymes: a common principle for individual control. Plant Physiol. 96, 1–3. 5. Foyer, C. H. and Noctor, G. (2005) Redox homeostasis and antioxidant signaling: a metabolic interface between stress perception and physiological responses. Plant Cell 17, 1866–1875. 6. Mittler, R., Vanderauwera, S., Gollery, M., and Van Breusegem, F. (2004) Reactive oxygen gene network of plants. Trends Plant Sci. 9, 490–498. 7. Meyer, A. J. and Fricker, M. D. (2008) Imaging thiol-based redox processes in live cells. In: Sulfur Metabolism in Phototrophic Organisms, R. Hell, C. Dahl, D. Knaff, and T. Leustek, Eds., Dordrecht: Springer, pp. 483–501. 8. Hanson, G. T., Aggeler, R., Oglesbee, D., Cannon, M., Capaldi, R. A., Tsien, R. Y., and Remington, S. J. (2004) Investigating mitochondrial redox potential with redoxsensitive green fluorescent protein indicators. J. Biol. Chem. 279, 13044–13053. 9. Dooley, C. T., Dore, T. M., Hanson, G. T., Jackson, W. C., Remington, S. J., and Tsien, R. Y. (2004) Imaging dynamic redox changes in mammalian cells with green fluorescent protein indicators. J. Biol. Chem. 279, 22284–22293.

10. Meyer, A. J., Brach, T., Marty, L., Kreye, S., Rouhier, N., Jacquot, J.-P., and Hell, R. (2007) Redox-sensitive GFP in Arabidopsis thaliana is a quantitative biosensor for the redox potential of the cellular glutathione redox buffer. Plant J., 52, 973–986. 11. Elsliger, M., Wachter, R., Hanson, G., Kallio, K., and Remington, S. (1999) Structural and spectral response of green fluorescent protein variants to changes in pH. Biochemistry 38, 5296–5301. 12. Remington, S., and Hanson, G. T. (2006) Oxidation reduction sensitive green fluorescent protein variants. Patent US 7,015,310 B2. 13. Hansen, R., Østergaard, H., and Winther, J. (2005) Increasing the reactivity of an artificial dithiol-disulfide pair through modification of the electrostatic milieu. Biochemistry 44, 5899–5906. 14. Cannon, M. B. and Remington, S. J. (2006) Re-engineering redox-sensitive green fluorescent protein for improved response rate. Protein Sci. 15, 45–57. 15. Haseloff, J., Siemering, K. R., Prasher, D. C., and Hodge, S. (1997) Removal of a cryptic intron and subcellular localization of green fluorescent protein are required to mark transgenic Arabidopsis plants brightly. Proc. Natl. Acad. Sci. U S A 94, 2122–2127. 16. Chiu, W., Niwa, Y., Zeng, W., Hirano, T., Kobayashi, H., and Sheen, J. (1996) Engineered GFP as a vital reporter in plants. Curr. Biol. 6, 325–330. 17. Jiang, K., Schwarzer, C., Lally, E., Zhang, S., Ruzin, S., Machen, T., Remington, S. J., and Feldman, L. (2006) Expression and characterization of a redox-sensing green fluorescent protein (reduction-oxidationsensitive green fluorescent protein) in Arabidopsis. Plant Physiol. 141, 397–403.

Chapter 7 Imaging of Reactive Oxygen Species In Vivo Steven M. Driever, Michael J. Fryer, Philip M. Mullineaux and Neil R. Baker Abstract Reactive oxygen species (ROS) are involved in many signalling pathways and numerous stress responses in plants. Consequently, it is important to be able to identify and localize ROS in vivo to evaluate their roles in signalling. A number of probes that have a high affinity for specific ROS and that are effectively taken up by cells and tissues are commercially available. Applications to intact leaves of singlet oxygen sensor green (SOSG), nitroblue tetrazolium (NBT), di-amino benzidine (DAB) and Amplex Red to detect singlet oxygen, superoxide and hydrogen peroxide are described. Imaging of the probes in the cells and tissues of leaves allows sites of ROS production to be identified. Key words: Amplex Red, di-amino benzidine, hydrogen peroxide, imaging, nitroblue tetrazolium, reactive oxygen species, singlet oxygen, singlet oxygen sensor green, superoxide.

1. Introduction In biological systems reactive oxygen species (ROS), such as singlet oxygen (1O2), superoxide (O2−) and hydrogen peroxide (H2O2), are potentially harmful molecules that can cause severe perturbations of function. The production of ROS in plants is widespread in nearly all cell types and tissues. Plants have evolved an extensive system of antioxidants to scavenge these ROS. In recent years it has been shown that in situations where the production of ROS does not exceed the scavenging capacity of the antioxidant system, ROS are involved in sensing and signal transduction pathways in response to a wide range of environmental and biological stresses (1,2). To examine the generation and

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roles of ROS in living tissues, techniques are required that do not perturb the systems severely but indicate the presence and sites of production of ROS. One of the problems with detecting ROS in biological systems is that these molecules can have very short lifetimes. Consequently, probes to detect ROS should be introduced near to the production site so that they can react with ROS directly upon their production. Furthermore, because of the ubiquitous and high-affinity scavenging systems in tissues, probes to detect ROS must have a high affinity for the ROS to compete with the ROS scavenging systems in the tissues. Here we describe how four commercially available probes can be used for the spatial and temporal detection of 1O2, O2− and H2O2 in intact leaves. Feeding these probes into leaves and imaging the leaves allows for the detection of ROS in different cell and tissue types. The techniques described are applicable to leaves of most higher plants, but similar techniques can be used on roots of higher plants and with algae and plant cell cultures.

2. Materials 2.1. Plant Material

Plants of Arabidopsis thaliana are grown from seed in Levington F2+S compost to mature rosette stage under controlled environmental conditions (photosynthetically active photon flux density of 150 µmol m−2 s−1 during an 8-h photoperiod at 22°C and ca. 50% relative humidity). Although leaves of Arabidopsis thaliana are used in the experiments shown here, the methods reported have been successfully applied to mature leaves of Zea mays and Phaseolus vulgaris.

2.2. Singlet Oxygen Sensor Green

Singlet oxygen sensor green (SOSG) is a stable fluorescein derivative compound that is highly specific for 1O2, and exhibits weak blue fluorescence with excitation peaks at 372 and 393 nm and emission peaks at 395 and 416 nm. On reaction with 1O2, it exhibits a much greater fluorescence with excitation and emission peaks at 504 and 525 nm, characteristics that are used to identify 1 O2 (3). Commercially available aliquots of SOSG (Invitrogen, Paisley, UK) containing 100 µg are dissolved in 33 µl methanol and diluted with 50 mM sodium phosphate buffer pH 7.4 to make a stock solution of 0.5 mM SOSG. Further dilution of this stock solution to 260 µM is required to produce a solution suitable for feeding to leaves. Solutions should be prepared fresh for each set of experiments.

2.3. Nitroblue Tetrazolium

Nitroblue tetrazolium (NBT; 2,2′-bis(4-nitrophenyl)-5,5′diphenyl-3,3′-(3,3′-dimethoxy-4,4′-diphenylene)ditetrazolium

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chloride; Sigma-Aldrich, Gillingham, UK) is a yellow compound that is reduced by O2− to form an insoluble blue formazan deposit (4–6), which can be imaged in leaves after extraction of chlorophyll (7). A solution of 6 mM NBT is prepared for feeding to leaves (see Note 1). 2.4 Di-amino Benzidine

Di-amino benzidine (DAB; 3,3′,4,4′-biphenyltetramine; SigmaAldrich, Gillingham, UK) reacts with H2O2 to form brown polymerization products, which can be detected in leaves after chlorophyll extraction (7–9). DAB is dissolved in water by reducing the pH to 3.6 using HCl. A solution of 23 mM DAB is prepared for feeding to leaves. DAB degrades in light and should be kept in a dark bottle (see Note 2).

2.5 Amplex Red

The colourless probe Amplex Red (10-acetyl-3,7-dihydroxyphenoxazine; Invitrogen Paysley, UK) reacts with H2O2 in the presence of peroxidase and forms resorufin (7-hydroxy-3H-phenoxazin-3-one) (10,11). Resorufin is fluorescent with excitation and emission peaks at 570 and 590 nm, respectively (11). Amplex Red is prepared in 50 µl aliquots of 10 mM in dimethylsulfoxide, which are then diluted into 50mM sodium phosphate buffer pH 7.5 to give a final concentration of 2 mM Amplex Red for feeding to leaves. This feeding solution should be prepared fresh for each experiment. The Amplex Red solutions for experiments on leaves do not require the addition of horseradish peroxidase since leaves contain high levels of peroxidases. Amplex Red will degrade in high light. Consequently, when illuminating leaves containing Amplex Red a wavelength of light that is not strongly absorbed by Amplex Red should be used. This is achieved by passing the light through a “Rose Pink” filter (Lee Filters, Andover, Hampshire, UK) (see Note 3).

3. Methods Leaves are fed with specific ROS probes through their transpiration stream and then exposed to a light treatment. After light treatment, chlorophyll is extracted from leaves fed with NBT or DAB. Leaves fed with SOSG or Amplex Red do not require chlorophyll extraction. Images are then taken either using bright field or fluorescence emission. 3.1. Feeding of Probes and Light Treatment

1. Cut leaves from the plant at the petiole under water with a sharp blade and then cut a second time under water to ensure an airlock at the cut surface does not occur. Airlocks can prevent effective uptake of probes into the transpiration stream.

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2. Keeping a droplet of water on the tip of the petiole of the leaf by holding the leaf with the petiole lowermost, transfer the leaf to a feeding solution containing a specific probe. 3. Place part of the leaf in a leaf cuvette (e.g., CIRAS-2, PP Systems, Hitchin, UK; LI-6400, LI-COR, Lincoln, Nebraska, USA) in which light, temperature and humidity can be controlled. Place the cuvette over a region of the leaf remote from the cut petiole to avoid interference of ROS production from the wound response associated with cutting the petiole. 4. Cover areas of the leaf, apart from the region enclosed in the cuvette but including the petiole, with aluminium foil to exclude light. Also, keep the feeding solution in the dark by covering with foil. For the duration of the feeding, expose the leaf area in the cuvette to a photosynthetic photon flux density (PPFD) of 50 µmol m−2 s−1 to minimize the possibility of light-induced production of ROS. Use an air flow through the cuvette of 500 ml min−1 and maintain the CO2 concentration in the cuvette at 400 ppm. Control the humidity in the chamber to give a vapour pressure deficit (VPD) of 1 kPa. 5. Feed probes to the leaf for up to 2.5 h to ensure that they are taken up and distributed throughout the entire leaf. The probe feeding time may need to be modified depending upon plant species, growth conditions, leaf age and environmental conditions during feeding. It is essential that leaves do not lose turgor during the feeding procedure. 6. After feeding the probe, expose the leaf area inside the cuvette to a given light treatment. 3.2. Chlorophyll Extraction

Chlorophyll extraction is required only for leaves fed with NBT or DAB. 1. After a given light treatment, immerse leaves in a solution of ethanol, lactic acid and glycerol (4:1:1 by volume) and place in a water bath at 80°C for 25 min. 2. Remove the leaves and place them on filter paper to remove the excess solution before imaging.

3.3. Bright Field Imaging

After the light treatment place leaves infiltrated with NBT or DAB on a flat bed scanner (e.g., Hewlett Packard Precision Scan) and image ensuring even illumination of the samples. Take a 32-bit colour image of each leaf at a resolution of 300 dpi (Fig. 7.1).

3.4. Fluorescence Imaging

Fluorescent products obtained when SOSG and Amplex Red react with 1O2 and H2O2, respectively, are imaged using a Peltier-cooled charge coupled device (CCD) camera (Wright Instruments Ltd.,

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Fig. 7.1. Imaging of hydrogen peroxide (a) and superoxide (b) production in leaves using DAB and NBT, respectively. Colour images are shown of Arabidopsis leaves infiltrated with DAB (a) and NBT (b). The lower half of each leaf was kept darkened as a control (Control). The upper halves of the leaves were exposed to a PPFD of 350 mmol m−2 s−1 for 60 min. Chlorophyll was extracted from the leaves before imaging. Marked superoxide production is observed in response to wounding in the control area of (b) (Reproduced from Ref. (7) with permission) (see Color Plates ).

Middlesex, UK) with a custom built light-emitting diode (LED) lighting system (Technologica Ltd, Colchester, UK). LED lighting provides a constant and homogenous excitation light with a maximum at 466 nm over an area of 9 by 12 cm. Detection of specific fluorescence emissions is achieved by placing optical band pass filters in front of the lens of the CCD camera. Band pass filters with maxima for wavelengths of 525 and 590 nm (Edmund Optics Inc., York, UK) were used for detection of reaction products of SOSG and Amplex Red, respectively. The camera is controlled by FluorImager V1.01 software (Technologica Ltd, Colchester, UK), which was purposely designed for image acquisition (576 by 384 pixels) and for controlling exposure time with this setup. The acquired images are processed using ImageJ software (12). 1. Place leaves infiltrated with SOSG or Amplex Red under the CCD camera fitted with a band pass filter for their respective fluorescence emission wavelength. 2. Illuminate leaves while images are taken by the camera using shutter times selected to obtain images with suitable contrast and brightness. Keep the shutter times and illumination the same for all leaves that are to be compared.

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A

B

C

Stressed

Control

-DCMU

+ DCMU

increasing photon emission

Fig. 7.2. Imaging singlet oxygen production in leaves. False colour images of Arabidopsis leaves infiltrated with SOSG and exposed to a PPFD of 350 µmol m−2 s−1 PPFD for 60 min, a light treatment that would not be expected to produce significant amounts of singlet oxygen in the absence of an inhibitor of photosystem II electron transport. (a) Control leaf was fed with SOSG only. (b) Leaf was infiltrated and fed with SOSG. A solution of 200 µM 3-(3′,4′-dichlorophenyl)1,1-dimethylurea (DCMU) was painted onto the upper leaf surface before exposure to light. DCMU inhibits photosystem II electron transport and stimulates 1O2 production. A wound response of singlet oxygen production is evident at the cut end of the petiole of the control leaf (a); however, there is little light-induced singlet oxygen produced in the leaf blade. (c) Imaging of hydrogen peroxide production in a leaf using Amplex Red. False colour image of fluorescence emission from an Arabidopsis leaf after infiltration with Amplex Red. The upper half of the leaf was exposed to a PPFD of 350 µmol m−2 s−1 for 60 min (Stressed). The lower half of the leaf was kept in the dark (Control). Marked hydrogen peroxide production is observed in response to wounding in the control area around the veins and petiole (see Color Plates ).

3. Copy the images to ImageJ and adjust brightness and contrast for similar grey value scales in 8-bit mode. A false colour scale can be applied to highlight differences in fluorescence emission intensities between leaves. Images of leaves infiltrated with SOSG and Amplex Red are shown in Fig. 7.2 (see Note 4).

4. Notes 1. NBT is difficult to dissolve, especially at the concentrations used here. It is therefore advisable to use NBT of at least 98%

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purity. When diluting in water, shake vigorously for 30 min or until all NBT is dissolved. Alternatively, a small amount of methanol can be used to dissolve the NBT before diluting into water to produce the feeding solution concentration. NBT in solution can be kept for several days at 7°C and it does not degrade significantly in the light. 2. DAB is difficult to dissolve in water unless the pH is reduced. To dissolve DAB, prepare a solution of water by adding HCl until pH 3.6 is reached and then add the DAB. Shake the solution until all the DAB is dissolved. The pH of the solution can be adjusted upwards, but some of the DAB will precipitate. It is important to note that DAB is photodegradable and should be kept protected from light. DAB in solution can be kept at 7°C for a maximum of two days; however, it is advisable to prepare a fresh solution for each set of experiments. Take care when handling DAB because it is toxic; wear suitable protective clothing. Preparation of DAB solutions should be done in a fumehood. Dispose of DAB solutions according to your local regulations. 3. Amplex Red is photodegradable and this must be prevented during light treatments by using the “Rose Pink” filter to apply light. This filter blocks wavelengths between 500 and 570 nm, which are wavelengths absorbed strongly by Amplex Red. 4. Image processing is only intended to highlight and normalize images taken at different times and from different samples. All manipulations of images are done in 8-bit grey scale mode. Depending on your system, other types of images may be acquired that need different ways of processing. A simple and easy way to adjust the scales of different grey scale images is using an imaging program such as ImageJ. ImageJ is a free java-based program, which can be downloaded from http://rsb.info.nih.gov/ ij/download.html.On this website documentation and plug-ins can be found for this program.

Acknowledgements This work was supported by funds from the Biotechnology and Biological Sciences Research Council, the Natural Environment Research Council and the University of Essex.

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References 1. Foyer, C.H. and Noctor, G. (2003) Redox sensing and signalling associated with reactive oxygen species in chloroplasts, peroxisomes and mitochondria. Physiol. Plant. 119, 355–364. 2. Apel, K and . Hirt, H. (2004) Reactive oxygen species: Metabolism, oxidative stress, and signal transduction. Ann. Rev. Plant Biol. 55, 373–399. 3. Molecular Probes (2004) Product Information. http://probes.invitrogen.com/ media/pis/mp36002.pdf?id=mp36002 4. Flohe, L and . Otting, F. (1984) Superoxide dismutase assays. Methods Enzymol. 105, 93–104. 5. Bielski, B.H.J., Shiue, G.G and ., Bajuk, S. (1980) Reduction of nitro blue tetrazolium by CO2- and O2- radicals. J. Phys. Chem. 84, 830–833. 6. Beyer, W.F. and Fridovich, I. (1987) Assaying for superoxide dismutase activity: some large consequences of minor changes in conditions. Anal. Biochem. 161, 559–566. 7. Fryer, M.J., Oxborough, K., Mullineaux, P.M. and , Baker, N.R. (2002) Imaging of photo-oxidative stress responses in leaves. J. Exp. Bot. 53, 1249–1254.

8. Thordal-Christensen, H., Zhang, Z., Wei, Y. and , Collinge, D.B. (1997) Subcellular localisation of H2O2 in plants. H2O2 accumulation in papillae and hypersensitive response during the barley-powdery mildew interaction. Plant J. 11, 1187–1194. 9. Orozoco-Cardenas, M. and Ryan, C.A. (1999) Hydrogen peroxide is generated systemically in plant leaves by wounding and systemin via the octadecanoid pathway. Proc. Natl. Acad. Sci. USA 96, 6553–6557. 10. Mohanty, J.G., Jaffe, J.S., Schulman, E.S., and Raible, D.G. (1997) A highly sensitive fluorescent micro-assay of H2O2 release from activated human leukocytes using a dihydroxyphenoxazine derivative. J. Immunol. Methods 202, 133–141. 11. Zhou, M., Diwu, Z., Panchuk-Voloshina, N., and Haugland, R.P. (1997) A stable nonfluorescent derivative of resorufin for the fluorometric determination of trace hydrogen peroxide: applications in detecting the activity of phagocyte NADPH oxidase and other oxidases. Anal. Biochem. 253, 162–168. 12. Abramoff, M.D., Magalhaes, P.J., and Ram, S.J. (2004) Image processing with ImageJ. Biophotonics Int.July, 36–42.

Chapter 8 Identification of Thioredoxin Targeted Proteins Using Thioredoxin Single-Cysteine Mutant-Immobilized Resin Ken Motohashi, Patrick G. N. Romano, and Toru Hisabori Abstract Thioredoxins (Trx) are a ubiquitous family of proteins that modulate the enzymatic activity of their substrate proteins by redox regulation. This is achieved by reduction of a disulfide bond within their target proteins. A conserved pair of cysteine residues in Trx is required for catalysis of the dithiol–disulfide exchange with their target proteins. A single-cysteine mutant capable of forming a stable mixed disulfide bond with target proteins was immobilized on resin and used to capture potential target proteins. By using this method, a number of previously unidentified Trx-target protein candidates were captured from various organisms and organelles. Following the development of this technique, more than one hundred proteins have been reported as potential Trx targets, allowing significant progress to be made in our knowledge and understanding of Trx-target proteins. Key words: Cysteine, disulfide, di-thiol, redox regulation, thioredoxin, thioredoxin affinity chromatography.

1. Introduction Thioredoxin (Trx) is a small ubiquitous protein, which works as a redox equivalent transducer in cells. This important protein was first identified in Escherichia coli (E. coli) as a hydrogen donor to ribonucleotide reductase (1). In higher plant chloroplasts, Trx regulates the activity of various proteins, including Calvin cycle enzymes, through disulfide bond reduction of the target proteins (2–4). Until 2000, knowledge of these target proteins was very limited; though many proteins were known to contain potential target disulfide bonds, only nine chloroplast enzymes had been identified as Trx-target proteins in higher plants (5). Because the T. Pfannschmidt (ed.), Plant Signal Transduction, vol. 479 Humana Press, a part of Springer Science + Business Media, LLC 2009 DOI: 10.1007/978-1-59745-289-2_8

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transfer of reducing equivalents is linked to photosynthetic electron transfer and chloroplast enzyme activity is upregulated in the light, we postulated that there must be a significant number of Trx-target proteins within this complex organelle. This observation prompted us to develop the so-called Trx-affinity chromatography, an effective screening method that allowed us to successfully identify many Trx-target candidate proteins (6–9). Kick-started by the accumulation of genomic information from various organisms, this proteomics-based strategy has become a popular and efficient method for Trx-target protein isolation. Since 2001, this strategy has led to the development of several Trx-target protein screening protocols (6, 10–14), allowing significant progress to be made in our understanding of redox regulation by Trx (15–17). Two stromal thioredoxin isoforms, Trx-f and Trx-m, in higher plant chloroplasts are extensively characterized and are known to regulate the activity of a number of thiol enzymes. These Trxs have two highly conserved cysteine residues located within an active cysteine motif (Fig. 8.1). By substitution of the internal Cys (Fig. 8.1b) with a Ser residue (WCGPC to WCGPS), a mutant Trx is prepared, which can be bound to an immobilized resin matrix and used as a “bait” by allowing it to form stable disulfide intermediates with target proteins (Fig. 8.2). Upon incubation of a protein mixture containing Trx-target proteins with the resin described above, Trx-target candidate proteins that form stable disulfide bonds with the single-cysteine mutant Trx can be separated by centrifugation. Trx-target candidate proteins are then eluted from the resin by reduction of the mixed disulfide bond by dithiothreitol (DTT) (Fig. 8.3). The method described here to capture the Trx-targeted protein is based on the interaction between the reactive cysteine of Trx and the disulfide bond on the target proteins. To assure specific interaction between Trx and the target protein, it is necessary to examine their specific binding by using a control protein under the experimental conditions used. For this purpose, two different cysteine mutants of Trx, such as a Trx-h1 (CS)mutant (39WCGPC43 to 39WCGPS43) and a Trx-h1 (SC)-mutant (39WCGPC43 to 39WSGPC43), must be useful (7). Another possible control is activated thiol Sepharose 4B, which is an artificial thiol-reactive resin (see Sect. 3.2 and Fig. 8.4). Plant genome analysis revealed the existence of multiple Trx genes and those for Trx-like proteins (18). In order to understand the function of these Trx isoforms and Trx-like proteins, preparation and immobilization of a single-cysteine mutant of these proteins on a resin matrix represents a useful way of screening for target proteins (9, 19, 20). Here we describe the screening protocol for the identification of target proteins of HCF164, a Trx-like protein located in the

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Fig. 8.1. Amino acid sequence alignment and three-dimensional protein structure of plant Trxs. (a) The amino acid sequences of Trxs were aligned by ClustalW. Identical amino acid residues within the three aligned Trxs were reversed. AtTrxh1: Arabidopsis Trx-h1, SpTrxf: spinach Trx-f, SpTrxm: spinach Trx-m. Black triangles show the N-terminal position of recombinant mature form of Trx-f and Trx-m. Trx-f was expressed from the indicated position (black triangle) using Met as an initial position. Trx-m was expressed from the indicated position (black triangle) with the start Met in front of Lys. Conserved reactive cysteine motif in Trx-family proteins are indicated by a hatched bar. The internal Cys replaced with Ser in Trx-mutant is indicated by an asterisk. (b) Three-dimensional protein structure of spinach Trx-m (PDB code 1FB0). Two-conserved cysteine residues at the reactive cysteine motif are indicated with a CPK model.

thylakoid lumen, as the basis for the application of this method to other Trx-family proteins. HCF164 is a thylakoid bound Trxlike protein, which contains a membrane-spanning domain and a thioredoxin-like CXXC motif in the N- and C-terminal regions,

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Fig. 8.2. Strategy used to capture Trx-target proteins using single-cysteine mutant of Trx. The internal cysteine at the reactive cysteine motif of Trx is substituted with serine (WCGPC to WCGPS) and Trx-mutant proteins are immobilized on the resin. The target proteins can form a mixed-disulfide intermediate with Trx-mutant-immobilized resin and are eluted by reduction.

Fig. 8.3. Compositions of Trx-target candidate proteins in spinach chloroplast stroma captured by Trx-f (left panel ) and Trx-m (right panel ) single-cysteine mutant-immobilized resin. The Trx-targeted proteins were captured by Trx-mutant-immobilized resin, extensively washed, and finally eluted with DTT by reduction of the mixed disulfide bond. The proteins in the final washing step (NaCl-wash) and DTT eluted fraction (DTT-elute) were separated by SDS-PAGE. Black triangles show the Trx-mutants released from the resin.

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Fig. 8.4. Specificity and efficiency of Trx single-cysteine mutant-immobilized resin. The protein composition of the eluates from three different resins is analyzed by two-dimensional gel electrophoresis. (a) About 100 mg of whole cell lysate proteins from dark grown Arabidopsis. (b) Proteins captured by activated thiol Sepharose 4B (80 nmol of activated thiol groups, equal mol of immobilized Trx-h1 (1 mg)). (c) Proteins captured by Trx-h1 (CS)-mutant-immobilized resin (1 mg of Trx), and (d) Proteins captured by Trx-h1 (SC)-mutant-immobilized resin (1 mg of Trx). Activated thiol Sepharose 4B is a thiolreactive resin and initially appeared to react with SH-residues in proteins. However, the results showed that this resin could not trap the majority of Trx-target proteins, suggesting that, in addition to the highly reactive external cysteine residue, the three-dimensional environment surrounding the reactive cysteines of Trx is an important element in the recognition of the target proteins. In contrast to Trx-h1 (CS), SC mutant, which lacks the external cysteine residue, reacted poorly with the target proteins (Reproduced from Ref. (7) with permission from the Japanese Society of Plant Physiologist).

respectively (21). The thioredoxin-like domain of HCF164 extends into the thylakoid lumen (9), where the HCF164 appears to exercise its function (see Sect. 3.3 and Fig. 8.5).

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Fig. 8.5. Screening of the target candidate proteins from Arabidopsis thylakoids by HCF164sol single-cysteine mutant-immobilized resin. In order to screen the target protein candidates of HCF164, we prepared a recombinant Trx-like domain of HCF164 (designated as HCF164sol), in which the N-terminal membrane-spanning region of HCF164 had been deleted (9). (a) The HCF164sol single-cysteine mutant-immobilized resin was used to screen for interacting proteins in a mixture containing Arabidopsis thylakoids solubilized with 1% n-octyl-β-D-glucoside (OG). Thylakoids, thylakoids from Arabidopsis leaves; OG-solubilized, n-octyl-β-D-glucoside solubilized thylakoids from Arabidopsis leaves; Flow-through, flow-through fraction after incubation with HCF164sol-immobilized resin. (b) Elution profile of Trx-target candidate proteins captured by the HCF164sol (CS)mutant. Final-wash, final washed fraction of HCF164sol-immobilized resin; DTT-elute, eluted proteins by DTT from HCF164sol-immobilized resin (Reproduced from Ref. (9) with permission from the American Society for Biochemistry and Molecular Biology).

2. Materials 2.1. Screening of TrxTarget Proteins from Stroma Fraction of Spinach Chloroplasts

1. LB medium (1 l): tryptone 10 g, yeast extract 5 g, NaCl 10 g, agar 15 g 2. 2xYT liquid medium (1 l): tryptone 16 g, yeast extract 10 g, NaCl 5 g 3. French press (5501-M, Ohtake Works, Tokyo) 4. QAE-toyopearl 550c (Tosoh) 5. SP-toyopearl 650M (Tosoh) 6. DEAE-toyopearl 650M (Tosoh) 7. Butyl-toyopearl 650M (Tosoh)

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8. Trx-f buffer 1: 25 mM Tris-HCl (pH 8.1) containing 0.5 mM DTT 9. Trx-f buffer 2: 25 mM MES-NaOH (pH 6.1) containing 0.5 mM DTT 10. Trx-m buffer: 25 mM Tris-HCl (pH 7.5) containing 0.5 mM DTT 11. CNBr-activated Sepharose-4B (GE Healthcare) 12. Coupling buffer: 0.1 M NaHCO3 (pH 8.3) containing 0.5 M NaCl 13. Blocking buffer: 0.1 M Tris-HCl (pH 8.0) 14. Percoll (GE Healthcare) 15. Washing buffer 1: 0.1 M acetate buffer (pH 4.0) containing 0.5 M NaCl 16. Washing buffer 2: 0.1 M Tris-HCl (pH 8.0) containing 0.5 M NaCl 17. Washing buffer 3: 50 mM Tris-HCl (pH 8.0) 18. Chloroplast disrupting buffer: 50 mM Tricine-KOH (pH 8.0) 2.2. Binding Specificity of Trx-Single-Cysteine MutantImmobilized Resin

1. Murashige and Skoog medium (SIGMA) 2. Protease inhibitor cocktail for plant cell extracts (SIGMA) 3. Homogenizing buffer: 50 mM Tricine-KOH (pH 8.0), 400 mM sucrose, 50 mM NaCl, 2% (v/v) protease inhibitor cocktail for plant cell extracts 4. Washing buffer 4: 20 mM Tris-HCl (pH 7.5), 200 mM NaCl 5. Activated thiol Sepharose 4B (GE Healthcare) 6. Trichloroacetic acid (20% (w/v) solution) 7. IPG buffer (GE Healthcare) 8. IPG Strips (pH 4–7, 7cm) (GE Healthcare) 9. Multiphor II system (GE Healthcare) 10. Sample buffer 1: 8 M urea, 2% (w/v) CHAPS, 40 mM DTT, 0.5% (w/v) IPG buffer, and 0.001% (w/v) bromophenol blue 11. Sample buffer 2: 50 mM Tris-HCl, pH 6.8, 6 M urea, 30% (v/v) glycerol, 16 mM DTT, and 1% (w/v) SDS 12. Sample buffer 3: 50 mM Tris-HCl, pH 6.8, 6 M urea, 30% (v/v) glycerol, 240 mM iodoacetamide, and 1% (w/v) SDS 13. Rehydration solution: 8 M urea, 2% CHAPS, 10 mM DTT, 2% IPG buffer, and 0.001% bromophenol blue

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2.3. Screening of Target Proteins for Trx-Related Proteins

1. Arabidopsis thylakoids preparation buffer: 330 mM sorbitol, 1 mM MgCl2, and 50 mM HEPES-NaOH (pH 7.6) 2. Chloroplast disrupting buffer: 50 mM Tricine-KOH (pH 8.0) 3. Arabidopsis thylakoids washing buffer: 10 mM sodiumdihydrogen pyrophosphate-NaOH (pH 7.8) 4. Solubilization buffer: 50 mM Tricine-KOH (pH 8.0) containing 2 mM EDTA and 1% n-octyl-β-D-glucoside

3. Methods 3.1. Screening of TrxTarget Proteins from Stroma Fraction of Spinach Chloroplasts

1. Spinach Trx-f (C40S) and Trx-m (C41S) are purified by similar protocols for wild-type Spinach Trx-f (9) and Trx-m (22) as described, though a minor modification is required for the single-cysteine mutant.

3.1.1. Preparation of Spinach Trx-f (C40S)

1. Recombinant spinach Trx-f (C40S) is expressed in E. coli (23). The Trx-f (C40S) expression plasmid is transformed to E. coli BL21(DE3) cells, and the transformed cells are grown on an LB plate containing ampicillin (50 µg/ml). 2. Inoculate a colony incubated for 12 h at 37°C to 10 ml 2x YT liquid medium containing 50 µg/ml ampicillin. Transfer the small culture at mid log phase to 2 L of 2x YT liquid medium containing 100 µg/ml ampicillin. 3. Grow E. coli cells in the culture of 2 l until OD600 = 0.8, and induce expression of the recombinant protein by adding isopropyl-β-D(−)-thiogalactopyranoside (final concentrations of 0.5 mM). Grow E. coli cells for 3 h and harvest them by centrifugation. 4. Suspend E. coli cells in Trx-f buffer 1, and disrupt them by a French press at 4°C. 5. Centrifuge disrupted cells at 100,000× g for 40 min, and apply the supernatant to a QAE-TOYOPEARL 550c column (Tosoh, Tokyo), which is equilibrated with Trx-f buffer 1 at 4°C in advance. Wash the column with Trx-f buffer 1 and elute with a 0–300 mM linear gradient of NaCl in Trx-f buffer 1 at 5 ml per each fraction. 6. Detect fractions containing Trx-f (C40S) by SDS-PAGE (15%), and extensively dialyze with a dialysis membrane (MWCO 6–8,000) against Trx-f buffer 2. Apply Trx-f (C40S) fractions to an SP-TOYOPEARL 650M column (Tosoh, Tokyo) equilibrated with Trx-f buffer 2. Wash the column

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with Trx-f buffer 2, elute with a 0–250 mM linear gradient of NaCl in Trx-f buffer 2, and fractionate every 5 ml. 7. Add glycerol (final concentration: 20%) to the purified Trx-f (C40S) protein preparation, and store it at −80°C. 3.1.2. Preparation of Spinach Trx-m (C41S)

1. Recombinant spinach Trx-m (C41S) is expressed in E. coli BL21 (DE3) (23), as described in Sect. 3.1.1. 2. Suspend E. coli cells in Trx-m buffer, disrupt cells by a French press and centrifuge at 100,000× g for 40 min at 4°C. 3. Apply the supernatant to a DEAE-TOYOPEARL 650M column (Tosoh, Tokyo) equilibrated with Trx-m buffer at 4°C. Wash the column with Trx-m buffer and elute with a 0–150 mM linear gradient of NaCl in Trx-m buffer, and fractionate every 5 ml. 4. Collect the peak fraction containing Trx-m, and add solid ammonium sulfate to a final concentration of 1.6 M. 5. Apply the solution to a BUTYL-TOYOPEARL 650M column (Tosoh, Tokyo) equilibrated with Trx-m buffer containing 1.6 M ammonium sulfate. Wash the column with Trx-m buffer containing 1.6 M ammonium sulfate, elute with a 1.6–0 M inverse gradient of ammonium sulfate in Trx-m buffer, and fractionate every 5 ml. 6. Add glycerol (final concentration: 20%) to the purified Trx-m (C41S) protein preparation, and store the preparation at −80°C.

3.1.3. Preparation of TrxImmobilized Resin

1. Dialyze the purified Trxs (Trx-f (C40S) and Trx-m (C41S)) extensively against coupling buffer (see Note 1). 2. Adjust the protein concentration of Trx (Trx-f (C40S) or Trx-m (C41S)) to 1 mg/ml. 3. Incubate CNBr-activated Sepharose-4B powder (150 mg) in 1 mM HCl for 15 min, to preserve the activity of the reactive groups on the resin. 4. Wash the CNBr-activated Sepharose-4B with coupling buffer to remove HCl three times. 5. For coupling, mix Trx solution containing 1 mg Trx (Trx-f (C40S) or Trx-m (C41S)) with CNBr-activated Sepharose4B, and incubate the mixture for 1 h at room temperature with occasional gentle stirring in a plastic tube (15 ml). 6. Remove excess uncoupled-Trx by washing with coupling buffer as described in Step 4. 7. In order to quench any remaining active groups, add blocking buffer, and incubate the resin for 2 h at room temperature (see Note 2).

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8. Wash the resin with 5 volumes of washing buffer 1 at least three times, and then wash with washing buffer 2 at least three times. 3.1.4. Screening of Trx Target Proteins

1. Prepare spinach chloroplast stroma lysate (10 mg protein) by following the standard protocol as described (6). 2. Prepare chloroplasts from spinach leaves and purify them by a 40–70% (v/v) discontinuous Percoll gradient. After centrifugation at 4,500× g for 5 min, collect the lower green band between the 40 and 70% layers as the intact chloroplast. 3. Wash this preparation subsequently by centrifugation to remove Percoll, suspend chloroplasts in disrupting buffer at a chlorophyll concentration of 1 mg/mL, and incubate for 15 min at 4°C. 4. Obtain the stroma lysate protein as the supernatant of this suspension after centrifugation for 1 h at 100,000× g. 5. Incubate chloroplast stroma lysate aliquots with Trx mutant (1 mg)-immobilized resin for 60 min at room temperature in a plastic tube (50 ml). 6. Wash the resin with washing buffer 3 by centrifugation, to remove nonspecifically bound proteins. Wash the resin further with washing buffer 3 containing 200 mM NaCl. Repeat this step until the absorbance of the washing solution at 280 nm approaches zero (see Note 3). 7. To elute the Trx-targeted proteins, finally incubate the resin with washing buffer 3 containing 200 mM NaCl and 10 mM DTT for 30 min at 25°C, and obtain the aliquot containing the captured proteins by separation of the resin by centrifugation (800× g, 3 min, at 4°C). 8. In order to determine the nature of the proteins obtained, separate these by SDS-PAGE (15%) (Fig. 8.3), followed by N-terminal amino acid sequencing, time of flight mass spectrometry, or other methods for protein identification.

3.2. Binding Specificity of Trx-Single-Cysteine Mutant-Immobilized Resin 3.2.1. Preparation of Trx-h1 Immobilized Resin and Activated Thiol Sepharose 4B

1. Arabidopsis Trx-h1 (CS) and Trx-h1 (SC) are purified as described (7). The recombinant Trx-h1 (CS) and Trx-h1 (SC) are expressed in E. coli BL21(DE3) and purified using the same method as Trx-m (C41S), as described in Sect. 3.1.2. 2. Trx-h1 mutants (80 nmol) are immobilized to the resin as spinach Trx-f and Trx-m (Sect. 3.1.3). 3. Activated thiol Sepharose 4B is prepared as follows. Swell freeze-dried activated thiol Sepharose 4B (20 mg, 80 nmol of activated thiol groups) and wash it in distilled water. Incubate

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the activated thiol Sepharose 4B slurry with washing buffer 3 containing 1 mM DTT for 15 min at room temperature to expose active SH-groups, and wash it three times with washing buffer 2. 3.2.2. Trapping of Thiol-Reactive Proteins in Arabidopsis Lysate by Trx-h1 Immobilized Resin and Activated Thiol Sepharose 4B

1. Surface sterilize 80 mg of Arabidopsis ecotype Columbia seeds with 5% (w/v) sodium hypochlorite, rinse and saw it in 2 l of Murashige and Skoog medium (24) containing 2% (w/v) sucrose, and place it in a shaking incubator (100 rpm) at 23°C for 3 weeks in the dark. White root hairs are predominantly obtained and this is useful to minimize the contamination of the cell lysate preparation by chloroplasts. 2. Homogenize Arabidopsis tissues in homogenizing buffer at 4°C using a Waring blendor. 3. Pass the homogenate through four layers of gauze and centrifuge it at 7,000× g for 30 min at 4°C. The supernatant is then further centrifuged at 100,000× g for 50 min at 4°C. Use the supernatant as the cell lysate. 4. Incubate the cell lysate containing 30 mg protein with 0.5 ml of Trx-immobilized resin (80 nmol of Trx was immobilized) at room temperature for 1 h with gentle stirring in a plastic tube (50 ml). 5. Wash the resin with washing buffer 4 to remove nonspecifically bound proteins. A washing buffer 4 containing 0.1% (w/v) SDS can also be used to effectively reduce the amount of proteins that bind to the resin nonspecifically. Repeat each washing step until the absorbance of the washing solution at 280 nm approaches zero. 6. Incubate the resin in washing buffer 4 containing 0.1% SDS and 10 mM DTT for 1 h at room temperature to reduce the mixed-disulfide intermediates formed. 7. Separate the proteins from the resin by centrifugation (800× g for 3 min) and collect the supernatant containing Trx-target proteins by pipetting.

3.2.3. Separation of Eluted Proteins by 2-D Electrophoresis

1. A standard 2-D gel electrophoresis protocol should be applied to separate the proteins obtained from the Trximmobilized resin. First precipitate the eluted proteins with tri-chloro-acetic acid (final concentration: 10% (w/v)), wash with acetone, and suspend it in sample buffer 1. 2. Perform isoelectric focusing by using the Multiphor II system (GE Healthcare) according to the manufacturer’s instructions. IPG strips (pH 4–7, 7 cm) are rehydrated with a rehydration solution. 3. Carry out isoelectric focusing at 0–300 V linear gradient for 10 min, 300–3,500 V linear gradient for 3.5 h, and 3,500 V

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for 3 h. After electrophoresis, equilibrate the gel strips with sample buffer 2, and then incubate with sample buffer 3 (see Note 4). 4. For second dimensional separation, vertical gradient slab SDS-PAGE (7.5–15% (w/v) gradient gel, BioCraft, Tokyo, Japan) is used. 5. After electrophoresis, visualize the separated proteins by Coomassie Brilliant Blue R-250 staining (Fig. 8.4). 3.3. Screening of Target Proteins for Trx-Related Proteins 3.3.1. Preparation of Arabidopsis Thylakoids

1. Arabidopsis thylakoids are prepared by following the protocol as described (9). 2. Harvest Arabidopsis (ecotype Columbia) rosette leaves from 4- to 5-weeks-old plants and rapidly homogenize them in a Waring blendor in ice-cold Arabidopsis thylakoid preparation buffer containing 5 mM sodium ascorbate, 0.05% BSA, 2 mM EDTA, and 1 mM MnCl2, filtered through Miracloth (Calbiochem), and centrifuge the suspension at 2,000× g for 5 min. 3. Resuspend the resulting chloroplasts in Arabidopsis thylakoid preparation buffer containing 5 mM sodium ascorbate, and centrifuge it at 1,300× g for 5 min. 4. Resuspend chloroplasts with Arabidopsis thylakoid preparation buffer containing 2 mM sodium ascorbate, 1 mM MnCl2, 2 mM EDTA, 2 mM NaNO3, 5 mM NaHCO3, 0.5 mM K2HPO4, and 5 mM sodium diphosphate and centrifuge at 1,300× g for 5 min. 5. Resuspend the precipitated chloroplasts in chloroplast disrupting buffer containing 2 mM EDTA and 1% protease inhibitor mixture for plant cell extracts (Sigma) to a final concentration of 0.5 mg/ml chlorophyll. 6. Centrifuge the broken chloroplast suspension at 7,500× g for 10 min at 4°C and wash twice with Arabidopsis thylakoid washing buffer to remove peripheral proteins. 7. Arabidopsis thylakoids are obtained as a precipitate of this suspension following centrifugation for 10 min at 7,500× g, for screening of HCF164 target proteins.

3.3.2. Screening of HCF164 Target Proteins

1. Incubate Arabidopsis thylakoids (20 mg chlorophyll) in 50 ml solubilization buffer containing 1% protease inhibitor cocktail for plant cell extracts (Sigma) for 60 min at 4°C with gentle mixing, and centrifuge at 140,000× g for 30 min. The solubilized thylakoid protein fraction is consequently obtained as a supernatant. 2. Incubate this detergent-solubilized fraction with the HCF164sol (CS)-mutant-immobilized resin for 60 min at room temperature in a plastic tube (50 ml).

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3. Wash the resin to remove nonspecifically bound proteins using solubilization buffer containing 500 mM NaCl, and solubilization buffer containing 500 mM NaCl and 0.1% SDS. 4. Repeat the washing step until the absorbance of the washing solution at 280 nm approaches zero. Finally, suspend the resin in solubilization buffer containing 0.1% SDS and 20 mM DTT, and incubate for 60 min to reduce the mixeddisulfide intermediates on the resin. The solution containing Trx-target proteins is separated from the resin by centrifugation (800× g for 3 min) and the supernatant is collected by pipetting. 5. Separate the eluted proteins by SDS-PAGE (15% (w/v)) and stain with Coomassie Brilliant Blue R-250 (Fig. 8.5). 6. Determine each of the stained protein bands by N-terminal amino acid analysis and by peptide mass fingerprint analysis using matrix-assisted laser desorption ionization time-offlight mass spectrometry (6–9).

4. Notes 1. When Tris-buffer remains in the sample solution, the Trx coupling yield is significantly decreased since the Tris-base blocks the active groups on CNBr-activated Sepharose4B. Therefore, purified Trx should be extensively dialyzed against coupling buffer to remove Tris or other amino-residues in the buffer. 2. In Sect. 3.1.3, after Trx coupling, the protein concentrations of the supernatant of the coupling solution should be determined to estimate Trx-coupling yield to the resin. Under the conditions shown in this protocol, the protein concentration of the supernatant is almost zero. 3. When the nonspecifically bound proteins are not completely removed from the resin by the final washing step (Sect. 3.1.4), a further washing step is required. In such a case, a washing step using 0.1% SDS after the 200 mM NaCl washing step is effective. After washing buffer 3 containing 200 mM NaCl is removed, washing buffer 3 containing 200 mM NaCl and 0.1% SDS should be added to the precipitated resin and gently stirred. Centrifuge at 800× g for 1 min and remove the supernatant. Repeat this washing step twice. The resin is finally incubated in washing buffer 3 containing 200 mM NaCl, 0.1% SDS and 10 mM DTT to elute the captured

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proteins. After centrifugation the supernatant is transferred to a new tube. Eluates are analyzed by SDS-PAGE, identified by N-terminal amino acid sequencer and peptide mass finger printing method. 4. The reduced alkylation of target proteins by iodoacetamide protects further modification of cysteine residues.

Acknowledgements We thank Fumie Koyama for technical assistance in sample preparation.

References

1. Laurent, T. C., Moore, E. C., and Reichard, P. (1964) Enzymatic Synthesis of Deoxyribonucleotides. IV. Isolation and characterization of thioredoxin, the hydrogen donor from Escherichia coli B. J. Biol. Chem. 239, 3436–3444. 2. Buchanan, B. B. (1991) Regulation of CO2 assimilation in oxygenic photosynthesis: the ferredoxin/thioredoxin system. Perspective on its discovery, present status, and future development. Arch. Biochem. Biophys. 288, 1–9. 3. Schurmann, P. (1995) Ferredoxin: thioredoxin system. Methods Enzymol. 252, 274–283. 4. Jacquot, J. P., Lancelin, J. M., and Meyer, Y. (1997) Thioredoxins: structure and function in plant cells. New Phytol. 136, 543–570. 5. Schurmann, P., and Jacquot, J. P. (2000) Plant Thioredoxin Systems Revisited. Annu. Rev. Plant Physiol. Plant Mol. Biol. 51, 371–400. 6. Motohashi, K., Kondoh, A., Stumpp, M. T., and Hisabori, T. (2001) Comprehensive survey of proteins targeted by chloroplast thioredoxin. Proc. Natl. Acad. Sci. U S A 98, 11224–11229. 7. Yamazaki, D., Motohashi, K., Kasama, T., Hara, Y., and Hisabori, T. (2004) Target proteins of the cytosolic thioredoxins in Arabidopsis thaliana. Plant Cell Physiol. 45, 18–27.

8. Hosoya-Matsuda, N., Motohashi, K., Yoshimura, H., Nozaki, A., Inoue, K., Ohmori, M., and Hisabori, T. (2005) Antioxidative stress system in cyanobacteria. Significance of type II peroxiredoxin and the role of 1-Cys peroxiredoxin in Synechocystis sp. strain PCC 6803. J. Biol. Chem. 280, 840–846. 9. Motohashi, K., and Hisabori, T. (2006) HCF164 receives reducing equivalents from stromal thioredoxin across the thylakoid membrane and mediates reduction of target proteins in the thylakoid lumen. J. Biol. Chem. 281, 35039–35047. 10. Yano, H., Wong, J. H., Lee, Y. M., Cho, M. J., and Buchanan, B. B. (2001) A strategy for the identification of proteins targeted by thioredoxin. Proc. Natl. Acad. Sci. U S A 98, 4794–4799. 11. Balmer, Y., Koller, A., Val, G. D., Schurmann, P., and Buchanan, B. B. (2004) Proteomics uncovers proteins interacting electrostatically with thioredoxin in chloroplasts. Photosynth Res. 79, 275–280. 12. Kadokura, H., Tian, H., Zander, T., Bardwell, J. C., and Beckwith, J. (2004) Snapshots of DsbA in action: detection of proteins in the process of oxidative folding. Science 303, 534–537. 13. Kumar, J. K., Tabor, S., and Richardson, C. C. (2004) Proteomic analysis of thioredoxin-

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targeted proteins in Escherichia coli. Proc. Natl. Acad. Sci. U S A 101, 3759–3764. Marchand, C., Le Marechal, P., Meyer, Y., Miginiac-Maslow, M., Issakidis-Bourguet, E., and Decottignies, P. (2004) New targets of Arabidopsis thioredoxins revealed by proteomic analysis. Proteomics 4, 2696–2706. Buchanan, B. B., and Balmer, Y. (2005) Redox regulation: a broadening horizon. Annu. Rev. Plant Biol. 56, 187–220. Hisabori, T., Hara, S., Fujii, T., Yamazaki, D., Hosoya-Matsuda, N., and Motohashi, K. (2005) Thioredoxin affinity chromatography: a useful method for further understanding the thioredoxin network. J. Exp. Bot. 56, 1463–1468. Hisabori, T., Motohashi, K., HosoyaMatsuda, N., Ueoka-Nakanishi, H., and Romano, P. G. (2007) Towards a functional dissection of thioredoxin networks in plant cells. Photochem. Photobiol. 83, 145–151. Meyer, Y., Reichheld, J. P., and Vignols, F. (2005) Thioredoxins in Arabidopsis and other plants. Photosynth. Res. 86, 419–433. Broin, M., Cuine, S., Eymery, F., and Rey, P. (2002) The plastidic 2-cysteine peroxiredoxin is a target for a thioredoxin involved in the protection of the photosynthetic apparatus against oxidative damage. Plant Cell 14, 1417–1432.

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20. Rey, P., Cuine, S., Eymery, F., Garin, J., Court, M., Jacquot, J. P., Rouhier, N., and Broin, M. (2005) Analysis of the proteins targeted by CDSP32, a plastidic thioredoxin participating in oxidative stress responses. Plant J 41, 31–42. 21. Lennartz, K., Plucken, H., Seidler, A., Westhoff, P., Bechtold, N., and Meierhoff, K. (2001) HCF164 encodes a thioredoxinlike protein involved in the biogenesis of the cytochrome b6f complex in Arabidopsis. Plant Cell 13, 2539–2551. 22. Motohashi, K., Koyama, F., Nakanishi, Y., Ueoka-Nakanishi, H., and Hisabori, T. (2003) Chloroplast cyclophilin is a target protein of thioredoxin. Thiol modulation of the peptidyl-prolyl cis-trans isomerase activity. J. Biol. Chem. 278, 31848– 31852. 23. Stumpp, M. T., Motohashi, K., and Hisabori, T. (1999) Chloroplast thioredoxin mutants without active-site cysteines facilitate the reduction of the regulatory disulphide bridge on the gamma-subunit of chloroplast ATP synthase. Biochem J. 341, 157–163. 24. Murashige, T., and Skoog, F. (1962) A revised medium for rapid growth and bio assays with tobacco tissue cultures. Physiol. Plantarum 15, 473–497.

Chapter 9 Determination of In Vivo Protein Phosphorylation in Photosynthetic Membranes Julia P. Vainonen, Alexander V. Vener, and Eva-Mari Aro Abstract Light- and redox-controlled reversible phosphorylation of thylakoid proteins regulates short- and longterm acclimation of plants to environmental cues. The major phosphoproteins in thylakoids belong to photosystem II and its light-harvesting antenna but phosphorylation of subunits of other thylakoid protein complexes has been detected as well. The detection methods include electrophoretic separation of proteins and detection of phosphoproteins with a phosphoaminoacid-specific antibody or phosphoprotein-specific dye. The use of mass spectrometry allows the identification of exact phosphorylation site(s) in the proteins. Various methods for detection of phosphoproteins in thylakoids are outlined including phosphopeptide preparation for mass spectrometric analyses and quantitative analysis of protein phosphorylation. Key words: Protein phosphorylation, phosphothreonine antibody, phosphospecific dye, mass spectrometry.

1. Introduction Protein phosphorylation plays a key role in many cellular processes including signal transduction and stress responses. Plants as sessile organisms are continuously exposed to abiotic and biotic stresses that endanger their survival. Therefore, the mechanisms of acclimation to environmental cues are of particular importance for plants. More than 5% of Arabidopsis thaliana genome (2% in human genome) encode for protein kinases and phosphatases regulating an extensive and dynamic phosphoprotein and signaling network (1, 2).

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Environmental factors such as light, temperature and nutrient availability exert a strong effect on the function of plant chloroplasts and modulate protein phosphorylation of the photosynthetic thylakoid membrane, which in turn induces signaling cascades for proper acclimation to the changed environment. Several proteins of the photosynthetic membranes of plant chloroplasts undergo reversible phosphorylation (3). Phosphorylation of thylakoid proteins is light- and temperature-dependent through redox changes in both the membrane and the stromal compartments of chloroplasts. These redox changes apparently induce a cascade of protein kinase and phosphatase reactions that upon chloroplast protein phosphorylation/dephosphorylation lead to acclimation of the photosynthetic apparatus to changes in the quantity and quality of light or other environmental parameters (4–7). The most extensively studied thylakoid phosphoproteins include the Lhcb1 and Lhcb2 polypeptides of the light harvesting complex of photosystem II (LHCII) (8–11). Such reversible phosphorylation is responsible for chloroplast state transitions that balance the distribution of light energy between the two photosystems upon changing illumination conditions. Also, the minor LHCII antenna protein CP29 (12, 13), photosystem II core subunits D1 and D2 (8, 9), CP43 (8, 9, 11), and PsbH (8, 9) undergo redox-dependent reversible phosphorylation and thereby regulate, for example, the light-dependent turnover of the PSII reaction centre protein D1. The membrane phosphoprotein TMP14 belongs to photosystem I (12, 14), and phosphorylation sites were also identified in the Rieske Fe-S protein of cytochrome b6f complex (11). Several different experimental strategies can be employed to study the dynamics of protein phosphorylation in thylakoid membranes. Here we describe the commonly used methods for detection and identification of in vivo phosphorylation of thylakoid membrane proteins including immunoblotting with p-Thr antibody, fluorescence staining with Pro-Q Diamond phosphoprotein stain, and mass spectrometry-based approaches. Mass spectrometry also allows the mapping of the exact phosphorylation site within the protein sequence.

2. Materials 2.1. Preparation of Thylakoid Membranes

1. Homogenizer (Ultra Turrax T5FU) (IKA-Labortechnik Staufen, Germany) 2. Miracloth (Calbiochem)

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3. Buffer 1:50 mM Hepes-NaOH, pH 7.5, 300 mM sorbitol, 5 mM MgCl2, 1 mM EDTA, 4 mM sodium ascorbate, 10 mM NaF, and 1% bovine serum albumin (BSA) 4. Buffer 2: 10 mM Hepes-NaOH, pH 7.5, 5 mM sorbitol, 5 mM MgCl2, and 10 mM NaF 5. Buffer 3: 10 mM Hepes-NaOH, pH 7.5, 100 mM sorbitol, 5 mM NaCl, 10 mM MgCl2, and 10 mM NaF 6. Buffer 4: 50 mM Tricine, pH 7.8, 330 mM sorbitol, 1 mM EDTA, 10 mM KCl, 10 mM NaF, 0.15% BSA, 4 mM sodium ascorbate, and 7 mM L-cysteine 7. Buffer 5: 10 mM Tricine, pH 7.8, 5 mM MgCl2, and 10 mM NaF 8. Buffer 6: 50 mM Tricine, pH 7.8, 100 mM sorbitol, 5 mM MgCl2, and 10 mM NaF 9. Porra acetone: 80% acetone buffered with 25 mM HepesKOH, pH 7.8 2.2. SDSPolyacrylamide Gel Electrophoresis (SDS-PAGE)

1. SDS-PAGE equipment 2. Laemmli solubilization buffer and electrophoresis buffer (15) 3. Separating buffer: 1.5 M Tris-HCl, pH 8.8 4. Stacking buffer: 0.5 M Tris-HCl, pH 6.8 5. Fifty percent acrylamide/1.33% bis solution and N,N,N,N′tetramethyl-ethylenediamine (TEMED, BioRad, Hercules, CA) 6. 20% SDS 7. Ammonium persulphate: prepare fresh 10% solution in water before use 8. Prestained molecular weight markers, broad range (New England Biolabs)

2.3. Western Blotting with P-Thr Antibody

1. A semidry transfer system 2. Whatman 3MM filter paper 3. Polyvinilidene fluoride (PVDF) membrane (Millipore) 4. Transfer buffer: 48 mM Tris-HCl, 39 mM glycine, 0.0375% (w/v) SDS, and 20% (v/v) methanol 5. BSA for blocking (A-7030 Sigma) 6. Fat-free milk for blocking 7. TBS: 20 mM Tris-HCl, pH 7.5, and 500 mM NaCl 8. TBS-T: TBS containing 0.05% Tween-20 9. Primary antibody: p-Thr polyclonal antibody (New England Biolabs)

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10. 10. Secondary antibody: antirabbit IgG conjugated to horseradish peroxidase 11. Enhanced chemiluminescent (ECL) reagents (Amersham) 2.4. Fluorescent Staining with Pro-Q Diamond

1. Pro-Q Diamond Phosphoprotein gel stain (Molecular Probes); store at 2–6°C, protected from light 2. Fix solution: 45% MeOH, 5% acetic acid 3. De-staining solution: 4% ACN/50 mM NaOAc, pH 4.0 4. SYPRO Ruby total protein stain (Molecular Probes); store at 2–6°C, protected from light 5. Wash solution: 10% methanol, 7% acetic acid 6. Rotary shaker 7. Visible-light-based scanner or trans-illuminator

2.5. Phosphopeptide Enrichment and Mass Spectrometry

1. Sequence-grade modified trypsin (Promega, Madison, WI) 2. Acetyl chloride 3. Anhydrous methanol (Aldrich) 4. Anhydrous deuterium-containing d-alcohol (Aldrich)

methanol:

d3-methyl

5. Chelating Sepharose Fast Flow (Pharmacia) 6. FeCl3 7. Acetonitrile (ACN) 8. Trifluoroacetic acid 9. Formic acid 10. ZipTip C18 (Millipore)

3. Methods The methods described below present three possible ways to determine in vivo thylakoid protein phosphorylation. The immunodetection with p-Thr antibody and staining with Pro-Q Diamond allow identification of the phosphoproteins and estimation of their relative amount. Mass spectrometry allows the identification of the exact phosphorylation sites in the proteins and the states of phosphorylation for individual proteins. These methods can be used for analysis of phosphorylated proteins in general with the exception of sample preparation before SDS-PAGE or trypsin digestion, which would require a specialized protocol.

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3.1. Detection of Thylakoid Phosphoproteins by P-Thr Antibodies

This method allows the detection of endogenous level of thylakoid protein phosphorylation under specific environmental conditions (Fig. 9.1). To avoid the dephosphorylation of thylakoid phosphoproteins during thylakoid isolation, 10 mM NaF should be added to all buffers used for thylakoid preparation (see Note 1). The control sample should be illuminated with far red (<700 nm) and red light (650 nm, above 150 µmol photon m−2 s−1) to completely dephosphorylate and phosphorylate the thylakoid proteins, respectively.

3.1.1. Isolation of Thylakoid Membranes after In Vivo Phosphorylation

The leaves should be frozen in liquid nitrogen directly after light treatment. BSA, sodium ascorbate, and NaF are added to the buffers before use. All procedures should be carried out at 4°C under dim light. 1. Homogenate 2 g of leaves in 10 ml of a grinding buffer (buffer 1) using a homogenizer/ultraturrax. 2. Filter the homogenate through one layer of Miracloth and centrifuge at 1500× g for 5 min. 3. Resuspend the chloroplast pellet in 5 ml of a lysis buffer (buffer 2) and centrifuge at 3000× g for 5 min. 4. Resuspend the thylakoid pellet in 5 ml of a storage buffer (buffer 3) and centrifuge at 3000× g for 5 min. 5. Finally resuspend the thylakoid pellet in a small volume (50– 100 µl) of storage buffer (buffer 3). 6. Save an aliquot for determination of chlorophyll concentration; freeze the rest of the thylakoids in liquid nitrogen and store at −70°C (see Note 2).

Fig. 9.1. Thylakoid protein phosphorylation in differentially light-treated leaves as demonstrated by immunoblotting with p-Thr antibodies. After dark adaptation the Arabidopsis leaves were illuminated for 3 h under different light intensities before isolation of thylakoid membranes. Various light conditions are indicated as D, darkness; LL, low light, 50 µmol photon m−2 s−1; GL, growth light, 120 µmol photon m−2 s−1; HL, high light, 600 µmol photon m−2 s−1.

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The concentration of chlorophyll a and b can be easily measured with a spectrophotometer based on the Beer-Lambert Law and the extinction coefficient for chlorophyll (16). 1. Resuspend 2–5 µl of the isolated thylakoids in 1 ml of Porra acetone. 2. Vortex the tube, and centrifuge it at 12,000× g for 5 min. 3. Measure the supernatant absorbance at 646.6 nm (A646.6) and 663.6 nm (A663.6) using the absorbance at 750 nm as a reference. 4. Calculate the chlorophyll concentration using the following equations: Chlorophyll a (µg/ml) = 12.25 (A663.6) − 2.55 (A646.6) Chlorophyll b (µg/ml) = 20.31 (A646.6) − 4.91 (A663.6) Total chlorophyll (µg/ml) = 17.76 (A646.6) + 7.34 (A663.6) 3.1.2. Separation of Thylakoid Proteins by SDS-PAGE and Transfer of Proteins to a PVDF Membrane

SDS-PAGE is run according to the Laemmli protocol (15). 1. Prepare 0.75- to 1.0-mm thick polyacrylamide gels (160 × 200 mm) using 15% acrylamide with 6 M urea in the separation gel. 2. Solubilize thylakoid proteins in a Laemmli sample buffer by incubation for 15 min at 55°C. 3. Load the thylakoid sample corresponding to 0.75 µg of chlorophyll in each well. 4. Run the gels at 10 mA/gel for 16 h (see Note 3). 5. Transfer the proteins to a PVDF membrane (Immobilon P) according to the manufacturer’s instructions.

3.1.3. Visualization of Proteins Using P-Thr Antibody

1. Block the membrane with 5% BSA (fatty-acid free) in TBS, for 1.5 h. 2. Incubate the membrane in primary antibody (dilution 1:3000 in 1% BSA prepared in TBS-T) overnight. 3. Wash the membrane five times for 5 min with TBS-T. 4. Incubate the membrane with secondary antibody (dilution 1:10,000 in 1% BSA prepared in TBS-T) for 2 h. 5. Wash the membrane five times for 5 min with TBS-T and two times for 5 min with TBS (see Note 4). 6. Detect phosphoproteins using a chemiluminescence kit according to the manufacturer’s instructions. 7. Scan the blots and compare intensities with controls included in the gel side by side with the samples (see Note 5).

3.2. Immunodetection with Protein-Specific Antibody

The phosphorylated and non-phosphorylated forms of some thylakoid proteins can be separated by SDS-PAGE because of retarded mobility of the former in the gel (see Note 6). To make the quantitative analyses, it is necessary to verify the equal cross-reaction

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of the protein-specific antibody with two forms of the protein. Successful separation of phosphorylated and non-phosphorylated forms of PSII D1 protein can be done using the following conditions: 1. Prepare and load gels as described in Sect. 3.1.2. 2. Run the gels at 30 mA/gel for 16 h. 3. Transfer the proteins to a PVDF membrane (Immobilon P) according to the manufacturer’s instructions. 4. Block the membrane with 5% fat-free milk in TBS for 1.5 h. 5. Incubate the membrane with primary D1 antibody (dilution 1:8,000, 1% fat-free milk in TBS-T) overnight. 6. Continue as described in Sect. 3.1.3 beginning at Step 3. 3.3. Detection of Thylakoid Phosphoproteins by Pro-Q Diamond Stain

This method allows direct, in-gel detection of phosphoproteins (Fig. 9.2) employing a very simple experimental protocol. The sensitivity of staining correlates with the number of phosphorylated residues present in the protein. Pro-Q Diamond is most useful in combination with SYPRO Ruby protein stain, which allows detecting the phosphorylated proteins and total protein (SYPRO Ruby) in the same gel. These reagents are fully compatible with mass spectrometry. 1. Prepare gels and solubilize thylakoid proteins as described in Sect. 3.1.2. 2. Load the thylakoid sample corresponding to 2 µg of chlorophyll in each well. 3. Run gels as described in Sect. 3.1.2. 4. Fix the gels overnight with fix solution.

Fig. 9.2. Thylakoid protein phosphorylation in differentially light-treated leaves as detected by Pro-Q Diamond staining. After dark adaptation the Arabidopsis leaves were illuminated for 3 h under different light intensities before isolation of thylakoid membranes. Various light conditions are indicated as D, darkness; LL, low light, 50 µmol photon m−2 s−1; GL, growth light, 120 µmol photon m−2 s−1; HL, high light, 600 µmol photon m−2 s−1.

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5. Wash the gels 3 times for 15 min with 200 ml of ddH2O. 6 Incubate gels in 50 ml of Pro-Q Diamond solution for 3 h in darkness. 7. De-stain gels by incubation three times for 60 min with 100 ml of de-staining solution in darkness (see Note 7). 8. Rinse the gel two times for 5 min with ddH2O. 9. Visualize thylakoid phosphoproteins using a visible-lightbased scanner, a visible-light trans-illuminator, or a 300 nm trans-illuminator (see Note 8). 10. Rinse the gel two times for 5 min with ddH2O. 11. Incubate the gel in 50 ml of SYPRO Ruby overnight in darkness. 12. Wash the gel in 100 ml of wash solution for 30 min in darkness. 13. Rinse the gel two times for 5 min with ddH2O. 14. Visualize the thylakoid proteins on UV or blue-light transilluminator. 3.4. Mass Spectrometric Analyses of Thylakoid Phosphoproteins

The methods described below outline the preparation of the surface-exposed peptides from thylakoid membranes, peptide esterification and enrichment of the phosphopeptides by immobilized metal affinity chromatography (IMAC), identification and sequencing of phosphopeptides by electrospray ionization mass spectrometry (ESI-MS), and quantitative analysis of protein phosphorylation using liquid chromatography coupled to mass spectrometry or stable isotope labeling.

3.4.1. Isolation of the Surface-Exposed Peptides from Thylakoid Membranes

1. Homogenize 2 g of Arabidopsis leaves with a blender in 35 ml of ice-cold preparation buffer (Buffer 4).

3.4.1.1. Preparation of Thylakoid Membranes

2. Filtrate the suspension through four layers of Miracloth and centrifuge at 1,000× g for 5 min. 3. Resuspend the pellet in 10 ml of preparation buffer and centrifuge at 3,000× g for 5 min. 4. Resuspend the pellet in 10 ml of lysis buffer (Buffer 5) and homogenize five times in a potter grinder. 5. Centrifuge at 6,000× g for 5 min. 6. Wash thylakoids once with 10 ml of washing buffer (Buffer 6) and twice with 10 ml of 25 mM ammonium bicarbonate (NH4HCO3)/10 mM NaF (see Note 9). Centrifuge the thylakoids after each resuspension at 6,000× g for 5 min. 7. Resuspend the thylakoids in 25 mM NH4HCO3/10mM NaF to a final concentration of 3 mg of chlorophyll per milli liter.

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1. Incubate the thylakoid suspension with sequence-grade modified trypsin (Promega) in the ratio of 5 µg trypsin per mg of chlorophyll at 22°C for 3 h (see Note 10). 2. Freeze the sample in liquid nitrogen, thaw, and centrifuge the digestion product at 15,000× g for 20 min. 3. Transfer the soluble peptides in the supernatant into a new tube, resuspend the pellet in the same volume of ddH2O, and centrifuge at 15,000× g for 20 min. 4. Pool the supernatants and centrifuge at 100,000× g for 20 min. The supernatant containing released peptides can be used either for direct mass spectrometric analyses or for phosphopeptide enrichment by IMAC. The released peptides could be methyl-esterified for blocking of peptide carboxylic groups, thus preventing unspecific binding of these groups to immobilized metal ions.

3.4.2. Esterification of Isolated Peptides

1. Lyophilize the peptides to dryness (see Note 11). 2. Add four times 20 µl of acetyl chloride to 0.5 ml of anhydrous methanol dropwise with stirring. The mixture is explosive; the procedure should be carried out under the hood (see Note 12). 3. Add the reagent to lyophilized peptides (250 µl per 50 mg of chlorophyll used for trypsin digestion) and incubate at room temperature for 2 h with occasional vortexing. 4. Lyophilize the modified peptides to dryness. The methyl-esterification of peptides gives an increment in peptide mass of 14 Da per modification. This mass increment should be taken into account when the sample is analyzed by mass spectrometry.

3.4.3. Enrichment of Phosphopeptides by IMAC

Enrichment of phosphopeptides from the thylakoid peptide mixture prepared as described in Sects. 3.3.1 and 3.3.2 allows their successful identification and sequencing by MS. Phosphopeptides are enriched by affinity chromatography using columns charged with Fe(III) (see Note 13). 1. Pack a micro-column in a gel loader tip with 3–4 µl of chelating Sepharose Fast Flow beads (7.5 µl of 50% slurry). 2. Wash the column with 40 µl of 0.1% acetic acid and charge the column with 80 µl of 100 mM FeCl3. 3. Wash the unbound salts two times with 20 µl of 0.1% acetic acid. 4. Reconstitute the peptides in 10 µl of H2O/ACN/MeOH (1:1:1) and load the peptides onto the column.

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5. Wash the column with 40 µl of 0.1% acetic acid, 40 µl of 0.1% acetic acid/20% ACN and, finally, with 40 µl of 20% ACN. 6. Elute the phosphopeptides with 40 µl of 20 mM Na2HPO4 (pH 9.0)/20% ACN (see Note 14). The phosphopeptides enriched by IMAC could be directly analyzed by HPLC-MS. For ESI-MS without HPLC separation, the sample should be desalted using ZipTip C18. 1. Pre-wet ZipTip C18 with 50% ACN in water. 2. Equilibrate ZipTip C18 with 0.1% trifluoroacetic acid in water. 3. Load the peptides by pipetting 10–20 µl of sample. 4. Wash the ZipTip with 0.1% trifluoroacetic acid in water. 5. Elute the peptides with 10 µl of 50% ACN in water. 3.4.4. Identification and Sequencing of Phosphopeptides Using ESI-MS

Several instrumental techniques could be used for specific identification of phosphopeptides in complex peptide mixtures. Precursor-ion scanning in negative mode ESI-MS allows detection of phosphopeptides that produce the diagnostic fragment ions with m/z -79 (PO3−) and -97 (H2PO4−). This type of experiment is preferable for specific detection of phosphopeptides because it simplifies the spectrum of masses for phosphorylated peptides among all non-phosphorylated peptides. Neutral loss scanning using a tandem mass spectrometer in positive mode exploits the specific loss of a neutral phosphate group (m/z 98 for singly charged ions, m/z 49 for doubly charged ions) from phosphoserine and phosphothreonine containing peptides. However, this method gives a number of false positives and there is a need to know the charge of the ion losing phosphate. The positive mode of ESI-MS is used for phosphopeptide fragmentation and sequencing. To increase the positive ionization of peptides the peptide solution is acidified by addition of formic acid to a concentration of 1–2%. Complete fragmentation of the phosphorylated peptides requires clean selection of their molecular ions for fragmentation, which is not usually possible from the total mixture of thylakoid peptides. The mixtures of phosphopeptides enriched by IMAC as described in Sect. 3.3.3 are suitable for direct tandem MS and sequencing. The nano-electrospray capillaries are loaded with 2 µl of peptide solution in 50% ACN/1% formic acid in water and MS fullscan spectra are recorded. These spectra are then analyzed for the presence of the peptide ions. The candidate ions are then selected and subjected to collision-induced dissociation with the instrument settings recommended by the vendor of the particular mass spectrometer. Phosphopeptides undergo characteristic neutral loss of phosphoric acid (m/z 98) during fragmentation, which allows distinguishing them from non-phosphorylated ones. Interpretation

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of the product ion spectra provides both identification of phosphopeptide sequence and determination of the modified residues. 3.4.5. Relative Quantification of the Phosphorylation Level

Changes in thylakoid protein phosphorylation can be quantitatively measured by labeling of the control and experimental samples with stable isotopes. The peptides isolated from control and experimental thylakoid samples as described in Sect. 3.3.1 can be differentially labeled during the esterification step before IMAC enrichment. 1. Prepare 2N methanolic HCl by dropwise addition of 80 µl of acetyl chloride to 0.5 ml of anhydrous methanol with stirring. 2. Prepare 2N methanolic DCl by dropwise addition of 80 µl of acetyl chloride to 0.5 ml of anhydrous d3-methyl d-alcohol (deuterium-containing methanol) with stirring. 3. Add 250 µl methanolic HCl and DCl to lyophilized peptides isolated from 50 µg of chlorophyll from control and experimental samples, respectively (see Note 15). 4. Incubate the reaction mixture at room temperature for 2.5 h. 5. Mix the modified peptides 1:1 and lyophilize the solvents. The differentially labeled peptides are subjected to IMAC enrichment as described in Sect. 3.3.3 and analyzed by ESI-MS in positive mode. The peptide esterification with light- and heavyisotope gives the mass increment of 14 and 17 Da, respectively. Signals for methyl esters of phosphopeptides appear as doublets separated by n(3 Da)/z, where n is the number of carboxylic groups in the peptide and z is the charge in the peptide (Fig. 9.3). The difference in intensity of the two signals in each doublet reflects the ratio in phosphorylation of a particular site between the control and experimental sample.

4 Notes 1. 10 mM NaF is added to all preparation buffers to unspecifically inhibit all protein phosphatase activities and, thus to avoid dephosphorylation of thylakoid phosphoproteins. Omit NaF in case the phosphorylated thylakoids are used to study the dephosphorylation process. 2. Freezing and storage of thylakoids at −20°C may lead to irreversible aggregation of proteins. 3. Do not run gels containing urea in a cold room (4°C) because of urea precipitation. Alternatively, use thermo-stated equipment.

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Fig. 9.3. Differential and stable isotope labeling for relative quantification of proteins. Positive-mode MS spectrum zoomed in the region where D1 and D2 phosphopeptides are detected. Sites of esterification (-Me) are marked in the corresponding peptide sequences. (a) Light-isotope labeled peptides from the control sample were mixed 1:1 with heavy-isotope-labeled peptides from the experimental sample. (b) Reverse labeling was performed. Reduced phosphorylation of D1 and D2 proteins was detected in the experimental sample.

4. Use TBS in the two last washing steps because Tween decreases the intensity of the chemiluminescence. 5. The relative quantification of protein spots can be performed with a FluorChem 8000 image analyzer (Alpha Innotech Corp.). The intensity of spots should be compared with control samples illuminated with far red (<700 nm), where the thylakoid proteins are dephosphorylated, and red light (650 nm, above 150 µmol photon m−2 s−1) where the phosphorylation level reaches the maximum. 6. The difference in electrophoretic mobility of phosphorylated and non-phosphorylated forms does not occur for every phosphoprotein. 7. Pro-Q Diamond is a reversible dye, and therefore a short de-staining time will result in a strong background while a long de-staining time will decrease the number of detected phosphoproteins.

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8. Use of a 300-nm trans-illuminator will lead to reduced sensitivity. 9. The washes of thylakoid membranes with 25 mM NH4HCO3 increase the access of trypsin to the surface-exposed regions of thylakoid membrane proteins because of de-stacking of thylakoids. The resulting peptide supernatant is compatible with direct analyses by MS. 10. Do not perform proteolytic treatment of thylakoids at 37°C because some of the thylakoid phosphoproteins are rapidly dephosphorylated by heat-shock-activated membrane protein phosphatases (17). 11. The peptides should be lyophilized completely because any traces of water will decrease the efficiency of peptide esterification. 12. 2N methanolic HCl is explosive upon contact with water. Do not throw used tips into open waste boxes; let the excess of the reagent evaporate under the hood. 13. Ga (III) ions can be used for phosphopeptide enrichment by IMAC (18). Phosphopeptides can also be enriched by using titanium dioxide (TiO2) columns (11). 14. The elution of phosphopeptides can be done stepwise with 4 × 10 µl of 20 mM Na2HPO4/20% ACN. In this case the majority of phosphopeptides will be in the second and third elutes. 15. Perform the reverse labeling of peptides as an internal control, i.e., label the control sample with a heavy isotope and the experimental sample with a light isotope. The ratio in peak intensities between the experimental and control sample should be independent on the way of labeling.

Acknowledgements The work is supported by Academy of Finland, The Kone Foundation, and The Swedish Research Council for Environment, Agriculture and Spatial Planning.

References 1. The Arabidopsis Genome Initiative (2000) Analysis of the genome sequence of the flowering plant Arabidopsis thaliana. Nature 408, 796–815.

2. Venter, J.C., Adams, M.D., Myers, E.W., Li, P.W., Mural, R.J., Sutton, G.G et-al.., (2001) The sequence of the human genome. Science 291, 1304–1351.

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3. Bennett, J. (1991) Protein phosphorylation in green plant chloroplasts. Annu. Rev. Plant Physiol. Plant Mol. Biol. 42, 281–311. 4. Allen, J.F. (1992) Protein phosphorylation in regulation of photosynthesis. Biochim. Biophys. Acta 1098, 275–335. 5. Rintamäki, E., Salonen, M., Suoranta, U.M., Carlberg, I., Andersson, B., Aro, E.-M. (1997) Phosphorylation of light-harvesting complex II and photosystem II core proteins shows different irradiance-dependent regulation in vivo. Application of phosphothreonine antibodies to analysis of thylakoid phosphoproteins. J. Biol. Chem. 272, 30476–30482. 6. Aro, E.-M. and Ohad, I. (2003) Redox regulation of thylakoid protein phosphorylation. Antioxid Redox Signal. 5, 55–67. 7. Pursiheimo, S., Martinsuo, P., Rintamäki, E. and Aro, E.-M. (2003) Photosystem II protein phosphorylation follows four different regulatory patterns induced by environmental cues. Plant Cell Environ. 26, 1995–2003. 8. Michel, H., Griffin, P.R., Shabanowitz, J., Hunt, D.F., and Bennett, J. (1991) Tandem mass spectrometry identifies sites of three post-translational modifications of spinach light-harvesting chlorophyll protein II. Proteolytic cleavage, acetylation, and phosphorylation. J. Biol. Chem. 266, 17584–17591. 9. Vener, A.V., Harms, A., Sussman, M.R., and Vierstra, R.D. (2001) Mass spectrometric resolution of reversible protein phosphorylation in photosynthetic membranes of Arabidopsis thaliana. J. Biol. Chem. 276, 6959–6966. 10. Vainonen, J.P., Hansson, M., and Vener, A.V. (2005) STN8 protein kinase in Arabidopsis thaliana is specific in phosphorylation of photosystem II core proteins. J. Biol. Chem. 280, 33679–33686.

11. Rinalducci, S., Larsen, M.R., Mohammed, S., and Zolla, L. (2006) Novel protein phosphorylation site identification in spinach stroma membranes by titanium dioxide micro-columns and tandem mass spectrometry. J. Proteome Res. 5, 973–982. 12. Hansson, M. and Vener, A.V. (2003) Identification of three previously unknown in vivo phosphorylation sites in thylakoid membranes of Arabidopsis thaliana . Mol. Cell. Proteome. 2, 550–559. 13. Bergantino, E., Dainese, P., Cerovic, Z., Sechi, S., Bassi, R. (1995) A post-translational modification of the photosystem II subunit CP29 protects maize from cold stress. J. Biol. Chem. 270, 8474–8481. 14. Krouchtchova, A., Hansson, M., Paakkarinen, V., Vainonen, J.P., Zhang, S., Jensen, P.E., et al. (2005) A previously found thylakoid membrane protein of 14 kDa (TMP14) is a novel subunit of plant photosystem I and is designed PSI-P. FEBS Lett. 579, 4808–4812. 15 . Laemmli , U.K. (1970) Cleavage of structural proteins during assembly of head of bacteriophage-T4 . Nature 227 , 680 – 685 . 16. Porra, R.J. (2002) The chequered history of the development and use of simultaneous equations for the accurate determination of chlorophylls a and b. Photosynth Res. 73, 149–156. 17. Rokka, A., Aro, E.M., Herrmann, R.G., Andersson, B., and Vener, A.V. (2000) Dephosphorylation of photosystem II reaction center proteins in plant photosynthetic membranes as an immediate response to abrupt elevation of temperature. Plant Physiol. 123, 1525–1536. 18. Posewitz, M.C. and Tempst, P. (1999) Immobilized gallium (III) affinity chromatography of phosphopeptides. Anal. Chem. 71, 2883–2892.

Chapter 10 Examining Protein Stability and Its Relevance for Plant Growth and Development Claus Schwechheimer, Björn C. Willige, Melina Zourelidou, and Esther M. N. Dohmann Abstract Eukaryotes control many aspects of growth and development such as cell cycle progression and gene expression through the selective degradation of regulatory proteins by way of the 26S proteasome. Generally, proteasomal degradation requires the poly-ubiquitylation of degradation targets by E1 ubiquitin activating enzymes, E2 ubiquitin conjugating enzymes, and E3 ubiquitin ligases. Specificity is brought to the process by E3 ubiquitin ligases, which engage in direct interactions with the degradation substrate to bring it into the proximity of the E2 enzyme. The abundance of genes encoding E3 ligase subunits in plant genomes invites the hypothesis that protein degradation plays an important role in the control of many plant growth processes, and it is therefore not surprising that proteasomal degradation has already been implicated in several important response pathways. However, most of the genes with a predicted role in the ubiquitin-proteasome pathway still remain to be characterized and the identity of their degradation substrates needs to be revealed. In this chapter, we give an overview of the ubiquitin-proteasome system and the pathway proteins that have been examined in Arabidopsis to date. We review the methods required to identify and characterize the proteins that play a role in protein degradation or that are the target for proteasomal degradation. Key words: E3 ubiquitin ligases, proteasome, protein degradation, reporter protein, ubiquitin.

1. Introduction – The Ubiquitin– Proteasome System

Eukaryotes control many aspects of growth and development such as cell cycle progression and gene expression through the selective degradation of regulatory proteins with the ubiquitin–proteasome system (1–4). The components of the ubiquitin–proteasome system are E1 ubiquitin activating enzymes, E2 ubiquitin conjugating

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Fig. 10.1. Overview of the principal steps of protein poly-ubiquitylation and protein degradation by the 26S proteasome. UBI, ubiquitin; SUB, substrate; E1, E1 ubiquitin activating enzyme; E2, E2 ubiquitin conjugating enzyme; E3, E3 ubiquitin ligase.

enzymes, and E3 ubiquitin ligases, which together promote the formation of poly-ubiquitin chains on lysine residues of a given degradation substrate and the 26S proteasome, which recognizes, de-ubiquitylates, unfolds, and degrades the poly-ubiquitylated proteins (see Fig. 10.1). E3 ubiquitin ligases (complexes) interact with the degradation substrate and bring the substrate into the vicinity of the E2 enzyme; thereby they confer specificity to the degradation process. E3 ligases form a large and diverse protein family with distinct substrate specificities in eukaryotes and many of these protein families are conserved in plants (4, 5). 1.1. E3 Ubiquitin Ligases – The Specificity Components of Ubiquitylation

The E3 ligases can be subdivided into those that function in the context of protein complexes and those that function (seemingly) as monomers (see Fig. 10.2). SKP1/CDC53/F-box domain protein (SCF) complexes are a predominant E3 subfamily in Arabidopsis (6, 7). SCF complexes are composed of a cullin1 subunit (CDC53 in yeast), RING-box protein1 (RBX1), the adaptor subunit SKP1 (in Arabidopsis ASK), and an interchangeable F-box domain protein (FBP), the latter functioning as the degradation substrate receptor subunit. Analyses of the Arabidopsis genome sequence predict more than 694 FBPs in Arabidopsis (6). Although several cases have been reported where FBPs engage in protein complexes other than SCF-type complexes (8), these cases are

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Fig. 10.2. E3 ubiquitin ligases are the specificity components of the ubiquitin–proteasome system since they specifically recognize the degradation substrate. The three types of cullin-RING ligases (CRLs) conserved in plants are depicted here. Each of these protein complexes requires the conjugation of the ubiquitin-related protein NEDD8 for (full) activity and can associate with an interchangeable degradation substrate receptor subunit.

comparatively rare, and it can be anticipated that most of the Arabidopsis FBPs engage in SCF-type E3 ligase complexes, where they confer distinct degradation substrate specificities. While the biological role of the vast majority of these FBPs and SCF complexes remains to be determined, the general importance of FBPs and SCF complexes for plant development is underlined by the fact that several of the already understood FBPs are implicated in important developmental processes such as auxin (SCFTIR1/AFBs) (9–13), ethylene (SCFEBF1/2) (14–16), and gibberellic acid (SCFSLY1/SNE ) response (17–21), light-regulated development (SCFEID1) (22, 23), and cell cycle control (SCFSKP2) (24). A complete list of known FBP functions is provided (see Table 10.1). In addition to SCF-type E3 ligases, other E3 complexes with similar protein composition have been reported, and together with SCF-type E3s these form the cullin-RING ligase (CRL) superfamily (see Fig. 10.2). Similarly to SCF-complexes, these CRL complexes contain a cullin subunit (cullin3 in the case of BTB/POZ complexes and cullin4 in the case of DCX complexes) and RBX1 as well as an (adaptor and an) interchangeable substrate receptor subunit (25–29) (see Table 10.1). Besides these CRLs, which in Arabidopsis may amount to close to one thousand different E3 complexes, several other E3 ligases are known (4). The anaphase promoting complex/cyclosome (APC), which has been predominantly implicated in the degradation of mitotic cell cycle regulators, represents another multimeric E3 ligase complex with some similarity to CRLs (30–36) (see Table 10.1). Similar to the CRLs, APC contains a cullin-related (APC2) and an RBX1related (APC11) subunit; however, the subunit composition and the regulation of APC is very different from that described for the other CRLs, e.g. APC2 unlike cullin1, cullin3, and cullin4 is not modified by neddylation (see below). Besides CRLs and APC, apparent monomeric E3 ligases such as HECT-domain (37–39), RING-domain (40–43), and U-box E3s (44, 45) exist in plants,

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Table 10.1 List of Arabidopsis mutants with defects in E3 ubiquitin ligase (subunits) Name

Mutant name

Biochemical function

Biological function

Reference

Cullin ring ligases (CRLs) – general components CULLIN1 (AtCUL1)

atcul1

cullin1 homologue Embryogenesis

auxinresistant6 (axr6)

CULLIN3a/CULLIN3b

Auxin response

(109–111)

Cytokinin response

(112)

Jasmonate response

(110, 112, 113)

Floral development

(110, 112, 114)

Photomorphogenesis

(110, 112)

cul3a/cul3b cullin3 homologue Embryogenesis cul3a

RBX1 homologue

ARABIDOPSIS SKP1 (ASK1) ask1

ASK2

(27)

cullin4 homologue Photomorphogenesis

RBX1

ask2

(25, 115)

cullin3 homologue Far-red light response (26)

cul3a/cul3b cullin3 homologue CULLIN4a/CULLIN4b

(108)

SKP1 homologue

SKP1 homologue

(29)

Organ formation

(29)

Cell cycle

(116)

Auxin response

(75, 117)

Jasmonate response

(75)

Cold response

(75)

Floral development

(114)

Embryogenesis

(118)

Male meiosis

(119)

Floral development

(114, 120)

Embryogenesis

(118)

Floral development

(114)

SGT1A

sgt1a

SGT1 homologue

Auxin response

(121)

ENHANCER OF tir1–1 AUXIN RESISTANCE (ETA1)/SGT1B

sgt1b/eta3/sgt1b

SGT1 homologue

Auxin response

(121, 122)

Jasmonate response

(122)

Plant defence

(123) (continued)

Table 10.1 (continued)

Name

Mutant name

Biochemical function

Biological function

Reference

Anaphase promoting complex (APC) APC2

apc2

APC2 homologue

Female gametogenesis (34)

APC6/CDC16

nomega

APC6 homologue

Female gametogenesis (33)

HOBBIT/CDC27

hobbit

CDC27 homologue

Cell cycle progression

ABI3-INTERACTING PRO- aip2 TEIN2 (AIP2)

RING-finger protein

Abscisic acid response (78)

AtCHIP

U-box protein

Abscisic acid response (45)

F-box protein

Auxin response

(98)

AUXIN SIGNALING F-BOX afb5 PROTEIN5 (AFB5)

F-box protein

Auxin response

(121)

ARABILLO-1

arabillo-1

F-box protein

Lateral root development

(124)

ARABILLO-2

arabillo-2

F-box protein

Lateral root development

(124)

ARM REPEAT PROTEIN INTERACTING WITH ABF2 (ARIA)

aria

BTB/POZ protein Abscisic acid response (125)

BIG BROTHER (BB)

bb

RING-finger protein

BLADE-ON-PETIOLE1 (BOP1)

bop1

BTB/POZ protein Leaf morphogenesis

CEGENDUO (CEG)

ceg

F-box protein

Lateral root formation (127)

CONSTITUTIVE PHOTOMORPHOGENIC1 (COP1)

cop1

RING-finger protein

Photomorphogenesis

(128, 129)

CORONATINE INSENSITIVE1 (COI1)

coi1

F-box protein

Jasmonate signalling

(129)

DE-ETIOLATED1 (DET1)

det1

Photomorphogenesis

(28, 29, 130, 131)

Photomorphogenesis

(22, 23)

(36)

Substrate receptor subunits

AUXIN RECEPTOR F-BOX PROTEIN1–3 (AFB1–3)

afb1–3

Organ size

(107) (126)

EMPFINDLICHER IM eid1 DUNKELROTEN LICHT (EID1)

F-box protein

ETHYLENE OVERPRODUCTION1 (ETO1)

eto1

BTB/POZ protein Ethylene Biosynthesis

(132)

ETO-LIKE1 (EOL1), EOL2

eol1, eol2

BTB/POZ protein Ethylene biosynthesis

(132) (continued)

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Table 10.1 (continued)

Name

Mutant name

Biochemical function

Biological function

Reference

F-BOX PROTEIN7 (FBP7)

fbp7

F-box protein

Translation

(133)

FLAVIN BINDING, KELCH fkf1 REPEAT, F-BOX (FKF1)

F-box protein

Circadian clock

(134–136)

HISTONE MONOUBIQUI- hub1 TINATION1 (HUB1)

RING-finger protein

Leaf and root growth

(137)

HISTONE MONOUBIQUI- hub2 TINATION2 (HUB2)

RING-finger protein

Leaf and root growth

(137)

HIGH EXPRESSION OF OSMOTICALLY SENSITIVE GENE (HOS1)

hos1

RING-finger protein

Cold response

(106, 138)

KAKTUS/HECT-CONTAINING UBIQUITIN PROTEIN LIGASE3 (UPL3)

kak/upl3

HECT-domain domain

Trichome development

(38, 39)

KEEP ON GOING (KEG)

keg

RING-finger protein

Abscisic acid

(41)

LOV KELCH PROTEIN2 (LKP2)

lkp2

F-box protein

Circadian rhythm

(139)

NON-EXPRESSOR OF PATHOGENESIS RELATED GENES1 (NPR1/

npr1

BTB/POZ protein Plant defence

NON-INDUCIBLE IMMUNITY1 (NIM1)

nim1

BTB/POZ protein Plant defence Shoot lateral branching

(140)

ORESARA9 (ORE9)/MORE ore9/max2 AXILLARY BRANCHES2 (MAX2)

F-box protein

(141–143)

PROTEOLYSIS1 (PRT1)

prt1

RING-finger protein

RPM1-INTERACTING PROTEIN2 (RIN2)

rin2

RING-finger protein

Hypersensitive response

(103)

RPM1-INTERACTING PROTEIN2 (RIN3)

rin3

RING-finger protein

Hypersensitive response

(103)

SLEEPY1 (SLY1)

sly1

F-box protein

Gibberellin response

(17, 19, 21)

SNEEZY (SNE)

F-box protein

Gibberellin response

(18)

SINA ARABIDOPSIS THAL- sinat5 IANA HOMOLOGUE5 (SINAT5)

RING-finger protein

Auxin response

(105)

(144)

(continued)

Examining protein stability

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Table 10.1 (continued) Name

Mutant name

SKP2A/SKP2B

Biological function

Reference

skp2a/skp2b F-box protein

Cell cycle

(24, 83)

SUPPRESSOR OF nim1 (SON1)

son1

F-box protein

Induced defence response

(145)

SUPPRESSOR OF phytochromeA1–5 (SPA1–5)

spa1 – spa5

WD40 repeat protein

Photomorphogenesis

(146– 148)

TRANSPORT INHIBITOR RESISTANT1 (TIR1)

tir1

F-box protein

Auxin response

(10, 98, 149)

TUBBY-LIKE PROTEIN9 (TLP9)

tlp9

F-box protein

Abscisic acid response (150)

UNUSUAL FLORAL ORGANS (UFO)

ufo

F-box protein

Floral development

(114, 120, 151)

VIER F-BOX PROTEIN1 (VFB1), VFB2 – VFB4

vfb1, vfb2, F-box protein vfb3, vfb4

Root and lateral root growth

(152)

XBAT32

xba32

RING-finger protein

Lateral root development

(153)

RING-finger protein

Abscisic acid response (154)

F-box protein

Circadian clock

XERICO

ZEITLUPE(ZTL)/ LOV KELCH PROTEIN 1(LKP1)

ztl/lkp1

Biochemical function

(155)

but only a few of these monomeric E3s have been characterized at the biological level to date (see Table 10.1). 1.2.The 26S Proteasome

The 26S proteasome is a highly conserved multi-protein complex that is composed of the 20S central cylinder and two 19S cap complexes, which are in turn composed of two sub-complexes, the ‘lid’ and the ‘base’, that associate in an ATP-dependent manner with the 20S proteasome (46, 47). Poly-ubiquitylated degradation substrates are recognized and de-ubiquitylated by the 19S ‘lid’, unfolded by the 19S ‘base’, and degraded by the protease subunits of the 20S ‘core complex’. Just like proteasome mutants from other eukaryotes, Arabidopsis mutants deficient in 26S proteasome function are defective in cell cycle progression and these mutants arrest growth at the zygotic stage. However, several viable weak mutants of specific 26S proteasome subunits with a presumably weak defect in 26S proteasome activity have been described (see Table 10.2) (48–53).

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Table 10.2 List of Arabidopsis mutants with defects in general components of the ubiquitin–proteasome pathway Name

Mutant name

Biological Function

Degradation substrate

Reference

20S Proteasome ‘core complex’ PAD1

pad1

Leaf polarity

(48)

PBE1

pbe1

Leaf polarity

(48)

19S Regulatory particle ‘base’ RPN1a

rpn1a

Embryo development

CYCLIN B1

(50)

Leaf polarity

(48)

Unknown

(50)

RPN1b

rpn1b

RPT2a

haltered root (hlr/ Meristem rpt2a) maintenance

RPT4a

rpt4a

Leaf polarity

(48)

RPT5a

rpt2a

Leaf polarity

(48)

RPN8a

asymmetric leaves enhancer3 (ae3)

Leaf polarity

(48)

RPN8b

rpn8b

RPN9a

rpn9a

Leaf polarity

(48)

RPN12A

rpn12a

Cytokinin response

(52)

rpn12a

Auxin response

(52)

AUXIN-RESISTANT1 (AXR1)

axr1

Auxin response

(156, 157)

E1-CONJUGATING ENZYME RELATED1 (ECR1)

ecr1

Auxin response

(158)

Hormone responses, meristem function

(29, 159)

Leaf venation

(160)

Photomorphogenesis

(161)

IAA17/ AXR3

(48, 53)

19S Regulatory particle ‘lid’

(48)

Neddylation

CULLIN-ASSOCIATED cand1 NEDDYLATION DISSOCIATED1 (CAND1) hve/cand1 COP9 Signalosome (De-neddylation) COP9 SIGNALOSOME SUBUNIT1 (CSN1)/FUSCA6 (FUS6)

csn1/fus6

(continued)

Examining protein stability

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Table 10.2 (continued) Name

CSN2/FUS12

Mutant name

csn2/fus12

CSN3/FUS11/CONSTITU- csn3/fus11/cop13 TIVE PHOTOMORPHOGENIC13 (COP13)

Biological Function

Degradation substrate

Reference

Jasmonate response

(62)

Photomorphogenesis

(162)

Photomorphogenesis

General growth and morphogenesis

(163)

CSN4/FUS4/COP8

csn4/fus4/cop8

Photomorphogenesis

(164)

CSN5A/ARABIDOPSIS JAB1 HOMOLOG1 (AJH1)

csn5a

Photomorphogenesis

(60, 75, 165)

General growth and morphogenesis

(165)

Auxin response

(60, 64, 75)

Jasmonate response

(75)

Photomorphogenesis

(60, 165)

Auxin response

(60)

Photomorphogenesis

(166)

General growth and morphogenesis

(167)

CSN5B/AJH2

CSN6A

csn5b

csn6a

CSN6B

csn6b

Photomorphogenesis

(166)

CSN7/FUSCA5 (FUS5)

csn7/fus5

Photomorphogenesis

(168)

CSN8/FUS7/FUS8/COP9

csn8/fus7/fus8/ cop9

Photomorphogenesis

(56, 169)

cop10

Photomorphogenesis

(170, 171)

cin4

Hormone responses

(172)

Auxin response

(75)

Jasmonate response

(75)

Cold response

(75)

E2 Ubiquitin-conjugating enzymes CONSTITUTIVE PHOTOMORPHOGENIC10 (COP10)

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1.3. NEDD8 Conjugation/ De-Conjugation and the COP9 Signalosome

The COP9 signalosome (CSN) is another protein complex that has been implicated in the regulation of the ubiquitin– proteasome system (54) . The fact that this evolutionarily conserved protein complex was first identified in plants based on constitutively photormorphogenic ( cop ) Arabidopsis mutants has attracted much attention in the plant community and beyond (see Table 10.2 ) (55– 58) . CSN is an eightsubunit complex and each of its eight subunits is homologous to one of eight subunits of the 19S ‘lid’ (59) . CSN interacts directly with CRLs – but presumably not with APC or other types of E3 ligases and is required for proper CRL E3 function (60– 65) . The mode of CSN action with regard to CRL function has been a matter of debate over the past years. CSN has been implicated in CRL subunit stability as well as CRL assembly (66– 74) . Critical to CSN function is its ability to remove the ubiquitin-related NEDD8/RUB1 modification from the cullin subunit of CRLs (de-neddylation) (64, 75) . A recent biochemical study from the mammalian system with the degradation substrate p27 KIP1 and the FBP SKP2 suggests that the availability of the degradation substrate and its association with the cognate FBP lead to the formation of a specific SCF complex and that this complex is stabilized by the conjugation of NEDD8 to cullin (neddylation) (66) . In the absence of a degradation substrate, e.g. when all molecules of a specific degradation substrate have been degraded, CSN-dependent cullin de-neddylation will destabilize the specific E3 CRL, and in this way CSN may allow us to maintain a free pool of cullin-RBX1 heterodimer that can then engage in de novo E3 ligase formation. In the context of these observations, the fact that CSN function is required for E3 ligase activity may then be interpreted such that the de novo formation of specific CRLs is impaired in the absence of CSN-dependent de-neddylation.

1.4. Degradation Substrates of the Ubiquitin-Proteasome System

While the proteins of the ubiquitylation machinery, the 26S proteasome and CSN, can now be identified based on sequence homologies to characterized components from other eukaryotes or based on the presence of specific protein domains (3), the identification of the respective degradation substrates is not trivial. In a few cases, notably in the case of the cyclin cell cycle regulators, protein instability was predicted and confirmed in analogy to the behaviour of functionally homologous proteins from other eukaryotes (76, 77). In general, however, there are no reliable prediction criteria for protein instability, and findings about the instability of plant proteins as well as about the biological significance of this instability are normally predicted based on genetic data or based on the interaction with components of the ubiquitin–proteasome system such as E3 ligase subunits (see Table 10.3).

Examining protein stability

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Table 10.3 List of Arabidopsis proteins targeted by the ubiquitin–proteasome system Name

Mutant

Biological function

E3

Reference

ABSCISIC ACID INSENSITIVE3 (ABI3)

abi3

Abscisic acid signalling

AIP2

(78)

ABSCISIC ACID INSENSITIVE5 (ABI5)

abi5

Abscisic acid signalling

KEG

(41)

CASEIN KINASE II REGULATORY SUBUNIT 4 (CKB4)

Circadian clock

Unknown

(173)

ClpP4

Chloroplast function

AtCHIP

(174)

CYCLIND3;1

Cell cycle

Unknown

(77)

CYCLING DOF FACTOR1 (CDF1)

Circadian rhythms

FKF1

(135)

DIMERIZATION PARTNER PROTEIN (DPB)

Cell cycle

SCFSKP2A

(83)

E2Fc

Cell cycle

SCFSKP2A

(24)

ETHYLENE OVERPRODUCTION2 (ETO2)

eto2

Ethylene biosynthesis

ETO1

(132)

ETHYLENE OVERPRODUCTION3 (ETO3)

eto3

Ethylene biosynthesis

ETO1

(132)

FAR-RED ELONGATED HYPOCOTYL1 (FHY1)

fhy1

Photomorphogenesis

Unknown

(80)

Chromatin modification

HUB1

(137)

HISTONE2B NAM/CUC PROTEIN (NAC1)

nac1

Auxin signalling

SINAT5

(105)

PHYTOCHROME A (PhyA)

phyA

Photomorphogenesis

COP1

(128)

PHYTOCHROME INTERACTING pif1 FACTOR1 (PIF1)

Photomorphogenesis

Unknown

(81)

PHYTOCHROME INTERACTING pif3 FACTOR3 (PIF3)

Photomorphogenesis

Unknown

(82)

PROTEIN PHOSPHATASE2A (PP2A)

rcn1

Abscisic acid response

AtCHIP

(174)

ZEITLUPE (ZTL)/LOV KELCH PROTEIN1 (LKP1)

ztl

Circadian clock

Unknown

(175)

2. Testing Protein Stability 2.1. General Considerations

In the simplest case, the abundance of protein in a given sample is the result of its de novo synthesis via transcription and translation and its turnover by proteasome-dependent or proteasome-independ-

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ent protein degradation. Thus, increases or decreases in protein abundance may be altered by an increase or decrease in its transcription, its translation, and/or its turnover. Thus, decreases in protein abundance can only be correlated with increased protein turnover when transcription and translation rates are unaffected. Consequently, only a careful examination of at least these three processes will allow to identify the different process(es) that regulate protein abundance in a given experiment, and still the relative contribution of each individual event to the overall change may be hard to interpret. Alternatively, much of the complexity can be taken from such an analysis when the turnover of a (putative) degradation substrate is uncoupled from transcription and translation, e.g. through the use of the protein biosynthesis inhibitor cycloheximide (50–100 µM), which blocks protein synthesis rapidly and efficiently (for examples see (78, 79)) (see Fig. 10.3a). Following cycloheximide treatment, the abundance of the protein of interest can be followed over time using degradation substrate-specific antibodies or suitable reporter fusion constructs (see below) in the absence or presence of the degradation-inducing stimulus (e.g. phytohormones, light, temperature). In cases where protein instability is observed, the next question should be whether this turnover is mediated by the ubiquitin–proteasome system or by other proteolytic pathways. This can be addressed through the use of 26S proteasome-specific inhibitors whose application should lead to the stabilization of the otherwise unstable degradation substrate if the turnover is mediated by the 26S proteasome. The most widely used inhibitor is MG132 (Z-Leu-Leu-Leu-CHO; for examples see (78, 80–82)), but also MG115 (Z-Leu-LeuNorvalinal-CHO; (81)), ALLN (N-Acetyl-Leu-Leu-Nle-CHO; (80)), and epoxomycin (83) have been employed as proteasome inhibitors in plant studies. 2.2. Following Protein Abundance Using Antibodies, Fusion Tags, and Reporter Proteins

The availability of a specific antibody allows examining protein abundance and degradation in various contexts such as different growth conditions or different mutant backgrounds. Therefore, an antibody is the most versatile means for protein analysis. However, unstable proteins are often – and in fact almost by definition – low abundant proteins and it is therefore often difficult to generate specific antibodies that identify the protein of interest with high affinity. For this reason, many researchers prefer to generate transgenic constructs for the expression of fusion proteins that express the protein under investigation as a fusion to an epitope tag (MYC, HA, FLAG etc.) or to a reporter protein, such as green fluorescent protein (GFP) and other fluorescent proteins, β-glucuronidase (GUS; (12)) or luciferase (LUC; (64, 81, 84)). An increasing range of fusion vectors are available in the

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plant community for each of these tags and reporters (85–87) and highly specific epitope tag- or reporter-specific antibodies are commercially available. However, when using fusion proteins to examine protein instability great attention should be paid to a number of experimental parameters to guarantee that the observations are biologically relevant. First, protein function could be affected when a protein is expressed as a fusion protein and the fusion protein may behave differently from the wild type protein with respect to protein turnover. For this reason, the fusion protein should be tested for functionality by complementation of a mutant phenotype (if available). This is particularly important because the fusion protein may have lost the affinity for its cognate E3 ligase or may localize to a different subcellular compartment from the wild type protein or its cognate E3 ligase. Second, attention should be paid to the expression level of the fusion protein. When choosing high-level expression systems, the amount of degradation substrate generated may by far exceed the turnover rate that can be promoted by the specific E3 ligase. Therefore, gene expression under the control of the endogenous promoter or in the genomic context provides the most significant data. Alternatively, protein expression can be fine-tuned using inducible gene expression systems (88, 89). An interesting example from the literature is the use of a heat-shock promoter driven AUX/IAA gene expression system that was used to follow protein degradation in response to auxin (12). Third, each of the commonly used reporter proteins has its individual advantages and also disadvantages. The GUS reporter protein allows easy quantification of reporter protein activity. However, GUS is inherently stable, has a long half-life, and tends to form aggregates, which may be resistant to ubiquitin-mediated degradation (90). GFP, its derivatives, and other fluorescent proteins allow for the non-invasive in vivo detection of reporter activity at the cellular and subcellular level. However, the reporters’ activities are often difficult to quantify because of photobleaching effects and the proteins’ slow and temperature-dependent folding (90–92). Finally, LUC offers the advantage of being comparatively unstable and of allowing for protein detection with a high sensitivity in a quantitative manner (87, 90). However, an often overlooked disadvantage when trying to quantify LUC activity as a means for fusion protein degradation resides in the fact that LUC activity is lost after the LUC reaction has taken place because the reaction catabolite blocks the active site of the enzyme (90, 93). Hence, the kinetics of protein degradation or accumulation can only be assessed by repeated LUC measurements from different samples processed in parallel.

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3. Interactions Between Unstable Proteins and E3 Ubiquitin Ligases 3.1 Molecular Approaches

3.2. Genetic Interactions

Once it has been established that a protein of interest is degraded via the ubiquitin–proteasome system, it may be interesting to identify the E3 ligase that promotes its degradation or inversely to identify the substrate(s) of a given E3. Only in a few cases it has so far been possible to identify the E3 ligase and its degradation substrate by screening yeast two-hybrid cDNA expression libraries to identify interacting proteins (14, 15, 94). It is often hypothesized that the reason for this resides in the fact that the degradation substrate may need to be post-translationally modified, e.g. by phosphorylation, to gain affinity for the E3, and that such a post-translational modification does not occur in the heterologous yeast host. Based on the presently published interaction data, it appears that yeast two-hybrid interaction screens are less suitable for the identification of F-box protein–substrate interactions but more suitable for the identification of RING-finger–substrate interactions (see Table 10.3). Once it has been established that a given protein is subject to proteasomal degradation, the biological relevance of this degradation for the biological process can be tested genetically. Mutations that fail to perceive the destabilizing signal or that fail to interact with the E3 ligase may be available or may be generated. Such mutations will consequently lead to a stabilization of the protein under investigation (see Fig. 10.3a for an example). Should protein degradation be important for the biological process under investigation, plants expressing a stabilized mutant form of the protein or plants overexpressing the protein under investigation may have the phenotype of a mutant with a defect in the respective degradation machinery (79). Conversely, the contribution of the accumulation of the degradation target for a given E3 ligase phenotype may be assessed by introducing loss-of-function alleles of the degradation target into the mutant background to prevent degradation target accumulation. If degradation target accumulation is responsible for the mutant phenotype, a suppression of the E3 ligase phenotype would be expected (see Fig. 10.3b and c for an example). Auxin signalling and gibberellic acid signalling are two predominant examples in the plant literature where such mutations have been identified. In auxin response, gain-of-function mutations have been described for a number of so-called AUX/IAA repressor proteins (reviewed in Ref. (95)). These mutants are defective in auxinregulated growth processes such as embryogenesis, root elongation growth, lateral root formation, and gravitropic responses. Since the expression of the AUX/IAA gain-of-function variants under the

Fig. 10.3. Biochemical and genetic analyses to examine protein instability. (a) Immunoblot to monitor the turnover of the DELLA protein RGA, a degradation target of the F-box protein SLY1, by endogeneous gibberellic acid (-GA) and following giberellic acid treatment (+GA). In both cases protein synthesis was blocked with 50 µM cycloheximide. RGA degradation is blocked in the sly1-10 F-box protein mutant and in the gid1a-c gibberellic acid receptor mutant. (b) Dwarfism of the gid1a-c mutant can be suppressed by introducing loss-of-function alleles of RGA (rga-24) and its close homologue GAI (gai-t6). Please note that the allele dosage-dependent genetic suppression. (c) The same genetic suppression is observed when introducing the rga-24 and gai-t6 mutations into the sly1-10 background. Interestingly, although both mutants are blocked in DELLA protein degradation and although the genetic interactions indicate that DELLA protein accumulation is responsible for the observed phenotypes, the severity of the mutant phenotypes is strikingly different, a finding that is indicative of the existence of mechanisms other than protein degradation that regulate DELLA protein activity.

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control of the promoter of another AUX/IAA gene induces the growth defects that are typical for gain-of-function mutations of the AUX/IAA gene whose promoter is used rather than that of the AUX/IAA gene, it appears that the 29 members of the AUX/IAA protein family have largely redundant function (96). The functional redundancy between the AUX/IAA proteins is also underlined by the observation that loss-of-function mutants for individual AUX/ IAA genes have only mild phenotypes, if any (97). In contrast, each of the gain-of-function mutants carries a mutation in the so-called domain ll of these mutant proteins (95). Domain II mutant AUX/ IAAs fail to interact in an auxin-dependent manner with the cognate F-box protein subunit TRANSPORT INHIBITOR RESPONSE1 (TIR1) or any of its close homologues AUXIN BINDING F-BOX1 – 3 (AFB1-3) (10, 98). Domain II of the AUX/IAAs is sufficient to mediate auxin-dependent protein degradation; thus it serves as a degron that can be used to destabilize fusion proteins in response to the auxin stimulus (12, 84, 99, 100). In gibberellic acid response, the gai gain-of-function mutant expresses a stabilized variant of the repressor GAI, which is resistant to GA-induced proteolysis (101). The gai mutant expresses a mutant protein with a 17 amino acid deletion of the DELLA domain. Just like the ga1-3 GA biosynthesis mutants or the gid1a-c GA receptor mutants, the GA insensitive gai mutant seedlings are dark-green lateflowering dwarfs (Fig. 10.3) (79, 101, 102). Growth de-repression, thus normal plant growth, requires the degradation of GAI and its close homologue RGA and is dependent on GA (biosynthesis), GA perception by the GID1 receptors, and the activity of the F-box protein SLY1 (17, 21). GAI and RGA are not degraded in gid1a-c and sly1 mutants (see Fig. 10.3a). The finding that the dwarfism of these mutants as well as the mutants’ other phenotypes can be almost completely suppressed by introducing loss-of-function alleles of GAI and RGA indicates that GAI and RGA are predominant growth repressors in these mutants (see Fig. 10.3b and c) (17, 21, 79, 102). In the context of the genetic studies on GAI it is also interesting to note that the gai suppressor mutation gar2-1 was identified to be a sly1 gain-offunction allele, which expresses a mutant sly1 protein with increased sensitivity towards the DELLA proteins (17, 21). Thus, the block of degradation imposed by loss of the DELLA domain in the gai mutant is partially overcome by the increased GA-independent affinity of the mutant sly1 F-box protein towards its degradation target. 3.3. Ubiquitylation Assays

Biochemical proof for the interaction of a given E3 ligase and a degradation substrate can be brought about by in vitro ubiquitylation experiments. The fact that several E3 ligases, notably the widely studied SCF complexes, function as protein complexes has rendered their examination in in vitro studies difficult, and to date there is no report of an in vitro ubiquitylation experiment using a recombinant reassembled plant E3 complex. In recent years, how-

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ever, several laboratories have been successful in performing in vitro ubiquitylation experiments with monomeric E3s such as RINGfinger and U-box proteins. Typically, commercially available yeast or rabbit E1 enzyme and purified recombinant Arabidopsis E2 enzymes (AtUBC1, AtUBCH5b, AtUBC8, AtUBC9, AtUBC10 (40, 41, 78, 103–107)) are used for these studies together with ubiquitin, which can be purchased from several suppliers, and a substrate protein, if available. In cases where no substrate is available for the ubiquitylation reaction, auto-ubiquitylation of the E3 is commonly observed and used as a proof that the candidate E3 ligase has indeed ubiquitin ligase activity.

4. Final Remarks Much of this chapter is concerned with the E1, E2, E3 cascade and the degradation substrates as the major players of the ubiquitin-proteasome system. It should be noted that the vast knowledge about the regulation of proteasomal degradation from yeast and mammalian systems indicates that proteasomal protein degradation is regulated at multiple other levels e.g. by phosphorylation, de-phosphorylation, and de-ubiquitylation (3). To date only limited information is available in the plant literature that allows judging the importance of this regulation in proteolysisdependent processes in plants. However, it can be anticipated that these mechanisms play a major role in most proteolysisdependent processes but are to be discovered. References 1. Hershko, A., and Ciechanover, A. (1998) The ubiquitin system. Annu. Rev. Biochem. 67, 425–479. 2. Reinstein, E., and Ciechanover, A. (2006) Protein degradation and human diseases: the ubiquitin connection. Ann. Intern. Med. 145, 676–684. 3. Bachmair, A., Novatchkova, M., Potuschak, T., and Eisenhaber, F. (2001) Ubiquitylation in plants: a post-genomic look at a posttranslational modification. Trends Plant Sci. 6, 463–470. 4. Schwechheimer, C., and Villalobos, L. I. (2004) Cullin-containing E3 ubiquitin ligases in plant development. Curr. Opin. Plant Biol. 7, 677–686. 5. Willems, A. R., Schwab, M., and Tyers, M. (2004) A hitchhiker’s guide to the cullin ubiquitin ligases: SCF and its kin. Biochim. Biophys. Acta 1695, 133–170.

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negative effects on development. Plant Cell 16, 2984–3001. Gusmaroli, G., Figueroa, P., Serino, G., and Deng, X. W. (2007) Role of the MPN subunits in COP9 signalosome assembly and activity, and their regulatory interaction with Arabidopsis Cullin3-based E3 ligases. Plant Cell 19, 564–581. Peng, Z., Serino, G., and Deng, X. -W. (2001) Molecular characterization of subunit 6 of the COP9 signalosome and its role in multifaceted development processes in Arabidopsis. Plant Cell 13, 2393–2407. Karniol, B., Malec, P., and Chamovitz, D. A. (1999) Arabidopsis FUSCA5 encodes a novel phosphoprotein that is a component of the COP9 complex. Plant Cell 11, 839–848. Wei, N., and Deng, X. -W. (1992) COP9: A new genetic locus involved in light-regulated development and gene expression in Arabidopsis. Plant Cell 4, 1507–1518. Suzuki, G., Yanagawa, Y., Kwok, S., Matsui, M., and Deng, X. W. (2002) Arabidopsis COP10 is an ubiquitin-conjugating enzyme variant that acts together with COP1 and the COP9 signalosome in repressing photomorphogenesis. Genes Dev. 16, 554–559. Yanagawa, Y., Sullivan, J. A., Komatsu, S., Gusmaroli, G., Suzuki, G., Yin, J., Ishibashi, T., Saijo, Y., Rubio, V., Kimura, S., Wang, J., and Deng, X. W. (2004) Arabidopsis COP10 forms a complex with DDB1 and DET1 in vivo and enhances the activity of ubiquitin conjugating enzymes. Genes Dev. 18, 2172–2181. Vogel, J. P., Schuerman, P., Woeste, K., Brandstatter, I., and Kieber, J. J. (1998) Isolation and characterization of Arabidopsis mutants defective in the induction of ethylene biosynthesis by cytokinin. Genetics 149, 417–427. Perales, M., Portoles, S., and Mas, P. (2006) The proteasome-dependent degradation of CKB4 is regulated by the Arabidopsis biological clock. Plant J. 46, 849–860. Luo, J., Shen, G., Yan, J., He, C., and Zhang, H. (2006) AtCHIP functions as an E3 ubiquitin ligase of protein phosphatase 2A subunits and alters plant response to abscisic acid treatment. Plant J. 46, 649–657. Kim, W. Y., Geng, R., and Somers, D. E. (2003) Circadian phase-specific degradation of the F-box protein ZTL is mediated by the proteasome. Proc. Natl. Acad. Sci. USA 100, 4933–4938.

Chapter 11 Probing Circadian Rhythms in Chlamydomonas reinhardtii by Functional Proteomics Volker Wagner and Maria Mittag Abstract In the unicellular flagellated green alga Chlamydomonas reinhardtii several processes are regulated by the circadian clock. To study circadian controlled processes, the cell’s clock is synchronized in a 12 h light–12 h dark cycle (LD12:12) before the cells are released into constant conditions of dim light and temperature. Under these free-running conditions circadian rhythms will continue with a period of about 24 h and cells can be harvested during specific time-points of subjective day and night. These cells were then used for isolating basic proteins by heparin-affinity chromatography, separating them on twodimensional PAGE and comparing the amount of their expression at four different time-points of subjective day and night. Among 230 proteins, we could find two proteins whose expression level changed more than fourfold throughout the circadian cycle. These proteins were identified as a protein disulfide isomerase (PDI)-like protein and a tetratricopeptide repeat (TPR) protein by liquid-chromatographyelectrospray ionization mass spectrometry (LC-ESI-MS). Key words: Chlamydomonas reinhardtii, circadian rhythms, functional proteomics, protein disulfide isomerase, tetratricopeptide repeat protein.

1. Introduction The unicellular green alga Chlamydomonas reinhardtii serves as a model for a wide range of biological processes (1, 2). Some of them, like photosynthesis and nitrogen metabolism, are common to plant cells while others like the structure and composition of flagella and basal bodies are similar to animal cells and highly relevant for the understanding of several human diseases. Still others, such as sensing and acclimation to environmental factors

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or the control of biological processes by the circadian clock, are fundamental to all eukaryotes (3). In recent years, specific cellular processes and cell “compartments” of C. reinhardtii have been investigated by applying proteomic strategies. This was possible because of two main reasons: (1) The alga can be easily and quickly grown in large amounts (4) and therefore, biochemical purification procedures of certain sub-proteomes can be well established. (2) Genome sequences from all three genetic compartments of C. reinhardtii (nucleus, mitochondria, and chloroplast) are available as well as more than 200,000 ESTs, which have been assembled into 10,000 contigs, representing unique cDNAs (5). In C. reinhardtii several processes including phototaxis, chemotaxis, and others have been shown to be controlled by the circadian clock (summarized in Ref. 6). They reach their maximal activities during the subjective day (e.g., phototaxis) or night (e.g., chemotaxis). A key property of circadian rhythms is that they persist under constant conditions of light and temperature with a period of about 24 h. Thus, cells whose clock has been synchronized under a LD 12:12 cycle continue with the rhythm if the cells are released under the free-running conditions of constant dim light (LL) or darkness (DD) and temperature. Using these conditions, rhythmic changes can be correlated to the control of an endogenous circadian clock rather than representing processes that are purely activated or inactivated by light or darkness. In the current approach we have applied functional proteomics to find novel components of circadian pathways by taking cell samples from four different time-points of the circadian cycle and analyzing the expression pattern of basic proteins that were enriched by heparin-affinity chromatography (7). In contrast to the often-applied micro- or macroarray studies to analyze circadian changes in the transcript level, such an approach will directly focus on the expression level of the protein. Thus, circadian proteins encoded by mRNAs that are constantly present during the day–night cycle but that are being translated in a circadian manner can be also depicted. Further, circadian fluctuating mRNAs that do not always result in circadian expression of proteins are automatically neglected by this way (summarized in Ref. 3). Application of heparin-affinity chromatography enriches basic proteins. About 230 proteins were detectable on 2-DE PAGE from this sub-proteome (7). The quantification procedure from the proteins expressed at specific circadian time-points involved a normalization procedure in which the volume of each spot was divided by the total volume of all non-saturated spots. Normalized protein spots, which have shown circadian changes with an amplitude of variance ³4 at the four different LL time-points

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Fig. 11.1. 2-DE of heparin bound proteins. (a) Heparin bound proteins were enriched from C. reinhardtii crude extracts, which were prepared from cells harvested at four LL time points (LL25, LL29, LL37, LL41) and applied to 2-DE. Proteins were visualized by silver staining. The four gel pictures represent the area where two protein spots (138 and 163) were identified, which change their expression pattern in a circadian manner with an amplitude ³4 in four independent datasets. The corresponding protein of spot 138 has an IEP of 9.15 and an average molecular weight of 36.2 kDa. Its amount peaks in the subjective night (LL41). The other circadian-expressed protein (spot 163) has an apparent IEP of 8.13 and an average molecular weight of 29.5 kDa. Its expression increases from the subjective day to the subjective night with a maximum of expression at LL41. Both proteins show a rhythm in their amount under conditions of constant dim light and temperature demonstrating that their expression is controlled by the circadian clock. The corresponding proteins from spot 138 and 163 were cut out of the gels and tryptic digested. LC-ESI-MS analysis identified spot 138 as a tetratricopeptide repeat protein and spot 163 as a protein disulfide isomerase. (b) Average strength of expression on the basis of normalized spot volume (method A) for gel spots 138 and 163 from four datasets using ImageMaster 2D software. Quantification was carried out as described under Methods. (Reprinted by permission of Federation of the European Biochemical Societies from Ref. (7)).

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(LL25 and 29 representing the subjective day; LL37 and 41 representing the subjective night) within four independent datasets, were further examined by mass spectrometry (LC-ESI-MS). Application of this method (7) led to the identification of a circadian-expressed PDI-like protein whose maximal expression occurred during the subjective night (LL41) and its lowest expression was at early subjective day (LL25, Fig. 11.1a). In addition, a circadian-expressed TPR-protein was identified. Its maximal expression also occurred at LL41 while its minimum was during the middle of the subjective day (LL29, Fig. 11.1a).

2. Materials 2.1. Cell Culture and Preparation of Crude Protein Extracts

1. C. reinhardtii wild type strain SAG 73.72 was obtained from the Culture Collection of Algae at the University of Göttingen, Germany (Sammlung von Algenkulturen der Universität Göttingen: SAG). 2. Chemicals and solutions for the preparation of high-saltacetate (HSA) medium: (a) Yeast extract (ROTH, no. 2363.3) (b) 10 X HSA buffer: 6.2 mM K2HPO4, 6.8 mM KH2PO4, 9.3 mM NH4Cl, 15 mM Na-acetate, pH: 6.8. (c) Trace element solution: The following amounts are needed for 1 l: 22 g ZnSO4 · 7H2O, 11.4 g H3BO4, 5.06 g MnCl2 · 4H2O, 5 g FeSO4 · 7H2O, 1.61 g CoCl2 · 6H2O, 1.57 g CuSO4 · 5H2O, 1.1 g (NH4)6Mo7O24 · 4H2O, 50 g Na2-EDTA (see Note 1). (d) 81 mM stock solution MgSO4 · 7H2O (e) 100 mM stock solution CaCl2 · 2H2O 3. Protein extraction buffer: 10 mM sodium phosphate buffer, pH 7.0, 14 mM dithiothreitol (DTT), and 4% (v/v) Complete protease inhibitor cocktail (ROCHE Molecular Biochemicals).

2.2. Heparin-Affinity Chromatography

1. FPLC-Buffers: Starting buffer: 10 mM sodium phosphate buffer, pH 7.0, 2 mM DTT. Washing buffer: 150 mM NaCl, 10 mM sodium phosphate buffer pH 7.0, 2 mM DTT. Elution buffer: 10 mM sodium phosphate buffer, pH 7.0, 2 mM DTT, 1 M NaCl. 2. HiTrap Heparin HP column, (Amersham Biosciences, now part of GE Healthcare).

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1. Dialysis buffer: 10 mM sodium phosphate, pH 7.0, 2 mM DTT. 2. QuickSep micro dialyzing capsules (ROTH). 3. Dialysis membrane: ZelluTrans MWCO: 8000–10,000 (ROTH)

2.4. Trichloroacetic Acid (TCA)/Acetone Precipitation

2.5. Two-Dimensional Polyacrylamide Gel Electrophoresis (2-DE PAGE)

Ten percent TCA dissolved in acetone containing 0.1% DTT and acetone containing 0.1% DTT, both kept at −20°C.

1. Ettan Dalt six electrophoresis unit and Ettan Dalt gel caster (Amersham Biosciences, now part of GE Healthcare). 2. Ettan IPGphor electrophoresis unit and IPG (immobilized pH gradient) strip holder (Amersham Biosciences, now part of GE Healthcare). 3. 2-DE lysis buffer: 9 M urea, 2% (w/v) 3-[(3-cholamidopropyl) dimethylammonio]-1-propanesulfonate (CHAPS), 1% DTT, 0.8% (v/v) ampholyte pH 3–10. 4. IPG strip (pH 3–10, 24 cm separation distance, manufactured by Amersham Biosciences, now part of GE Healthcare). 5. IPG rehydration solution: 8 M urea, 0.5% (w/v) CHAPS, 20 mM DTT, 10% (w/v) glycerol, 0.2% (v/v) ampholyte pH 3–10). 6. IPG equilibration solution 1: 6 M urea, 50 mM Tris-HCl, pH 8.8, 2% (w/v) sodium dodecyl sulfate (SDS), 30% (w/v) glycerol, 1% (w/v) DTT. 7. IPG equilibration solution 2: 6 M urea, 50 mM Tris-HCl, pH 8.8, 2% (w/v) SDS, 30% (w/v) glycerol, 4% (w/v) iodoacetamide, 0.03% (w/v) bromophenol blue. 8. Highly liquid paraffin (MERCK, no. 1.07174) for covering the IPG strips. 9. Acrylamide/piperazinediacrylamide (PDA) solution (30:0.27): 375 ml of a 40% (w/v) acrylamide stock solution from Bio-Rad, 1.35 g of PDA (Bio-Rad), 125 ml of water. Filter this solution through a 0.45-µm sterile filter and store it protected from light at 4°C. 10. Gel buffer: 1.5 M Tris-HCl pH 8.8 11. 5% (w/v) sodium thiosulfate stock solution. 12. N,N,N,N’-tetramethyl-ethylenediamine (TEMED, Sigma)

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13. Ammonium persulfate (APS): prepare 10% solution in water and immediately freeze in single aliquots (200 µl) at −20°C for further use. 14. Water-saturated isobutanol: Shake equal volumes of water and isobutanol in a glass bottle and allow separating. Use the top layer. Store at RT. 15. Electrode buffer: 0.1% SDS, 25 mM Tris, 0.192 M glycine. 16. 0.5% (w/v) agarose in electrode buffer for fixation of the IPG strip. 2.6. Silver Staining and Detection of Spots

1. Fixation solution: 40% (v/v) ethanol, 10% acetic acid. 2. Incubation solution: 30% (v/v) ethanol, 0.2% (w/v) sodium thiosulfate, 0.5 M sodium acetate. 3. Silver solution: 0.1% (w/v) silver nitrate, 0.01% (v/v) formaldehyde. 4. Developing solution: 2.5% (w/v) Na2CO3 (water free), 0.01% (v/v) formaldehyde. 5. Stop solution: 0.05 M disodium EDTA. 6. Duoscan f40 scanner from Agfa. 7. ImageMaster-2D v4.01 software from Amersham Biosciences (now part of GE Healthcare).

2.7. Capillary Liquid ChromatographyElectrospray Ionization (LC-ESI)-MS Analysis

1. Silver spot destaining solution: a 1:1 mixture of 30 mM potassium ferricyanide and 100 mM sodium thiosulfate (see Note 2). 2. Trypsin stock solution: 500 ng trypsin/µl dissolving buffer, both provided by PROMEGA. 3. Trypsin digest solution: 60 ng/µl in 50 mM NH4HCO3, stored at 4°C. 4. HPLC system: Dionex Ultimate HPLC pump with nanoflow setup, PepMap C18 reversed-phase capillary HPLC column and Dionex Famos autosampler with 1 µl injection possibility, directly connected to the MS. 5. LC-Packings PepMap C18 column (75 µm [inner diameter] by 150 mm) with a 3-µm particle size and a 100-Å-pore diameter (NAN-75-15-03-PM). 6. Mass spectrometer: 3D-Ion trap mass spectrometer (LCQ Deca XP, Thermo Electron Corp, San Jose, CA) with electrospray ionization. 7. Fused silica needle (75-µm inner diameter, a length of 12 cm, and a 30-µm aperture (FS 360-75-30; New Objective, Woburn, Mass.). 8. All instruments were controlled through Xcalibur System software (Thermo Electron Corp, San Jose, CA).

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9. HPLC aqueous phase A: 5% (v/v) acetonitrile and 0.1% formic acid. HPLC aqueous phase B: 80% (v/v) acetonitrile and 0.1% formic acid (see Note 3). 10. HPLC sample tubes: Polypropylene caps (P/N 160135) and vials (P/N 163974) from LC Packings.

3. Methods The green alga C. reinhardtii can be efficiently grown in a highsalt-acetate medium (4). To check for circadian expression, we transferred cells that had been exposed to a light–dark cycle to constant conditions of dim light and temperature. Under these free-running conditions, circadian rhythms will continue (8) and cells can be harvested at specific time-points representing the subjective day or night. By applying heparin-affinity chromatography, we enriched basic proteins. However, any other sub-proteome can be also isolated and used for quantitative comparisons by 2-DE along with the normalization procedure described here. Methods for efficient separation of basic proteins on 2-DE have been established along with immobilized pH gradients (9, 10). Procedures of functional proteomics based on silver staining and destaining procedures (11, 12) of 2-DE separated proteins have already been applied for C. reinhardtii to identify thylakoid membrane proteins by LC-ESI-MS (13, 14). For quantification and comparison of the amount of the proteins visualized by silver-stained 2-DE gels, we used a normalization procedure (7). Among ca. 230 protein spots, two proteins whose expression changed more than fourfold during the circadian cycle could be depicted. Identification of the tryptic digested peptides from these proteins by LC-ESI-MS analysis was based on the use of a nano-HPLC (Ultimate, Dionex) coupled to an ESI-MS with a 3D ion trap (LCQ, Thermo Electron Corp., San Jose, CA) along with a software relying on the Sequest algorithm (15, 16, 17) and C. reinhardtii databases (7). However, peptide identification can be also carried out by other LC-ESI-MS or yet other mass spectrometry units and other available software programs. 3.1. Cell Culture

1. For the cultivation of C. reinhardtii use a high-salt-acetate (HSA) medium that should be prepared in the following way: 900 ml of distilled water is filled in a Fernbach flask equipped with a stirring bar and supplemented with 100 ml 10 X HSA buffer. Subsequently, add 1 ml of the MgSO4· 7H2O stock solution, 1 ml of the CaCl2 · 2H2O stock solution, 1 ml of the trace element solution, and 3 g of yeast extract. After all components are mixed, seal the flask with

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a batting plug and autoclave it for 40 min at 120°C. Before inoculating the cells, the solution should cool down to RT. 2. C. reinhardtii wild-type strain SAG 73.72 is grown in HSA medium under a 12-h light/12-h dark cycle by constant stirring with a light intensity of 71 µE/m2 s at 24°C and then put under constant conditions of dim light [LL: (15 µE/m2 s)] before harvesting. The beginning of the dim light period is defined as time zero (LL0). The numbers (LL25, LL29, LL37, LL41) indicate how many hours the cells have been kept under LL conditions. 3. Harvest cells by centrifugation (4°C, 5 min, 8000× g) at different circadian times corresponding to early subjective day (LL25), the middle of the subjective day (LL29), the beginning of the subjective night, (LL37) and the middle of the subjective night (LL41). Freeze cells in liquid nitrogen and store them at −80°C (see Note 4). 3.2. Preparation of Crude Extracts

1. Fill 2/3 of a 15-ml Falcon tube with glass beads (diameter: 0.25–0.30 mm). 2. Wash the glass beads two times with protein extraction buffer. 3. Resuspend the cells in a small amount of extraction buffer and apply them to the glass beads. 4. Lyse the cells by vortexing (highest speed) five times for 1 min. Place the tube on ice for 1 min after each vortexing step. 5. Remove intact cells by centrifugation at 3000× g for 15 min at 4°C. 6. Remove the cell debris from the supernatant by centrifugation at 90,000× g for 1 h at 4°C. 7. Take the supernatant (protein crude extract) and determine the protein concentration using the Bio-Rad Protein Assay (referring to the manual from Bio-Rad).

3.3. Heparin-Affinity Chromatography

1. Apply 3 mg of the soluble protein (S 90000) to an FPLC (fast performance liquid chromatography) heparin-affinity chromatography HiTrap-column at a flow rate of 1 ml/min. 2. After sample application, wash the column with five volumes of starting buffer and five volumes of washing buffer to elute weakly bound proteins from the column. 3. Elute the proteins with a high affinity to heparin with 1 ml elution buffer.

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1. Prepare a 1-in2. dialysis membrane and incubate it in dialysis buffer for 5 min. 2. Fill the sample into a 1 ml micro dialyzing capsule and center the dialysis membrane over the well of the capsule. Push the collar over the membrane until it meets the flange, locking the membrane in place. 3. Place the loaded micro dialyzing capsules in a stirred beaker. 4. Adjust the stir bar speed so that the capsule tumbles continuously. 5. Dialyze the high-salt eluted proteins for 3 h against a 500fold volume of dialysis buffer at 4°C. 6. After the dialysis, determine the protein concentration using the Bio-Rad Protein assay. Refer to the manual from BioRad for the procedure. 7. Mix one volume of the dialyzed sample with four volumes of 10% TCA dissolved in acetone (−20°C) containing 0.1% DTT and keep it at −20°C overnight. 8. Following a centrifugation step (35,000× g, 30 min, 4°C), discard the supernatant and resuspend the pellet in acetone containing 0.1% DTT. 9. After 1 h incubation at −20°C, centrifuge the sample again (35,000× g, 30 min, 4°C). Discard the supernatant and dry the pellet under vacuum.

3.5.Two-Dimensional SDS-PAGE for Efficient Separation of Basic Proteins

These instructions assume the use of an EttanIPGphor electrophoresis unit for the first dimension and an Ettan Dalt six electrophoresis unit including the Ettan Dalt gel caster for the second dimension (see Sect. 2.5). 1. Solubilize the protein pellet for 2 h at RT in 2-DE lysis buffer. 2. Centrifuge the resuspended sample for 30 min at 26,800× g. 3. Fill 450 µl hydration solution into the IPG strip holder and place the IPG strip on it with the gel side down. Cover the strip carefully with paraffin. The rehydration of the IPG strip takes minimum 12 h, but can also be done longer. 4. Remove the rehydrated strip from the strip holder, discard any excessive rehydration solution, and apply 150 µg of the resuspended sample into the lateral sample application well of the strip holder near the anode. 5. Place an extra paper strip (e.g., blotting paper for Western blots) soaked with 15 mM DTT on the surface alongside the cathodic end of the IPG strip, before you put the rehydrated IPG strip again with the gel side down into the strip holder, and cover it with paraffin.

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6. Start the isoelectric focusing with the following IPGphor program: 200 V (1 h), 500 V (1 h), 1000 V (1 h), linear gradient from 1000 to 8000 V (1 h), and 8000 V (6 h). The current should be limited to 50 µA/IPG strip. 7. In the meanwhile, prepare a 10% polyacrylamide/PDA gel using the acrylamide/PDA (30:0.27) stock solution. The Ettan Dalt gel caster allows the preparation of six gels (25-cm wide, 21-cm high, 1-mm thickness). The following instructions refer to the preparation of four gels. Place the four assembled gels and the two spacer plates that were provided with the Ettan equipment into the gel caster. 8. Mix 88 ml Tris-HCl, pH 8.8 from the 1.5 M stock solution with 140 ml 30:0.27 acrylamide/PDA solution, add 1.4 ml of a 5% stock solution of sodium thiosulfate, and fill it up with water to 350 ml. Use a stirring bar for mixing the substances and add 133 µl TEMED and 1.3 ml of 10% APS to start the polymerization process. 9. Pour the solution gently into the gel caster. The liquid should reach a line just 1 cm beyond the top of the glass plates. Carefully overlay the gel with water-saturated isobutanol. Polymerization should be finished in 1 h. 10. After the end of the IPGphor program, remove the strip from the strip holder (see Note 5) and incubate it for 15 min in IPG equilibration solution 1 and 15 min in IPG equilibration solution 2. 11. Place the strips with the plastic sheet side to the higher glass plate and carefully push it between the plates directly on the top of each gel. 12. Prepare a small plastic spacer normally used for mini gels and place it at one side of the gel in a vertical position between the glass plates where no IPG strip is present. This will provide a well for the molecular weight marker. 13. Heat up the 0.5% agarose in electrode buffer to 40–50°C. Fix the spacer and the strip on the gel with this solution. Fill also the space between the glass plates at the lower part of the gel with agarose, because otherwise air can be trapped there during electrophoresis. 14. Place the gels and the spacer plates into the electrophoresis unit and fill it up (upper and lower buffer reservoir) with 5 l of electrode buffer. 15. Apply 20 µl of the molecular weight marker (Broad Range from Bio-Rad) to the well after removing the spacer and run the electrophoresis at 4°C and 2.5 Watt per gel overnight in the Ettan Dalt six electrophoresis unit (~12 h). 16. After electrophoresis, stain the gel with silver according to the protocol given below.

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1. Before removing one of the glass plates check carefully to which plate the gel is attached and remove the other one. Cut away the agarose containing the IPG strip and mark the gel at one corner to recognize the orientation of the pH gradient. Carefully place the gel in a tray of the appropriate size. All incubation steps are facilitated by gentle shaking. 2. Incubate the gel for 30 min in fixation solution. 3. Incubate the gel for 30 min up to overnight in the incubation solution. 4. Wash the gel three times for 10 min with distilled water. 5. Soak the gel for 40 min in the silver nitrate solution. 6. After a short wash with 50 ml distilled water, add 50 ml of developing solution. Gently shake it for a few seconds and remove the solution again. 7. Now place the gel in the developing solution and incubate it for 5–10 min. The exact developing time depends on the amount of protein and the thickness of the gel and must be optimized for each experimental purpose (see Note 6). 8. Stop the staining process by removing the developing solution and rinse the gel for 15 min with the stop solution. 9. Place the gel in distilled water to remove the EDTA.

3.7. Comparative Quantitative Analysis of Proteins from Four 2-DE Gels per Time-Point

The analyses of the stained 2-DE gels including the quantitative analysis of the spot volume of all proteins were carried out with the ImageMaster-2D software. This procedure can be adapted to any other quantitative analysis software systems. 1. At first, normalize all protein spots in each gel by the following procedures: Divide the volume of each spot by the total volume of all non-saturated spots in the gel (normalization method A) or by the total volume of all spots having a spot volume less than 200,000 (normalization method B). Volume is defined as the sum of the intensities of every pixel (gray level) in the spot. As suggested by the ImageMaster2D manual, multiply the resulting values by a scaling factor representing the total area of all non-saturated spots in the gel (method A) or the total area of all spots with a volume <200,000 (method B) (see Note 7). 2. Now you can compare the expression ratios of any protein you are interested in. It is possible to determine the average percentage and the standard error of means (SEM) of the normalized spot volumes from the datasets for a selected protein. In our experiments, protein spots having a normalized volume, which show an amplitude of variance ³4 during the chosen circadian cycle (LL25, LL29, LL37, LL41) within the four independent experiments, were further considered. The highest volume of a circadian-expressed protein spot

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with regard to the different LL time points is set to 100% (see Fig. 11.1b). 3.8. Tryptic Digest of Proteins in Gel Spots

1. Excise the spot, place in an Eppendorf tube, and add 300– 500 µl/spot destaining solution (see Note 8). 2. Incubate it for 8 min at RT and discard supernatant. 3. Add 100 µl/spot destaining solution and incubate 1 min (minimum) at RT until the gel spot appears colorless and discard supernatant. 4. Wash four times with 1 ml distilled water (1 min/wash). 5. Incubate the gel in 0.5 ml acetonitrile/H2O-mixture (3/2) for 20 min, discard the supernatant, and dry the gel in the speed-vac for 15 min. 6. Then incubate (rehydrate) the gel piece with 50 µl 50 mM NH4HCO3 for 10–15 min and discard the supernatant. 7. Repeat Steps 5 and 6. 8. Incubate the gel in 0.5 ml acetonitrile/H2O-mixture (3/2) for 20 min, discard the supernatant, and dry the gel in the speed-vac for 15 min. 9. Add 10 µl trypsin digest solution to the dried gel piece and incubate for 30 min on ice. 10. Add 50 µl 50 mM NH4HCO3 and incubate for 12 h up to 16 h at 37°C. 11. Centrifuge shortly, remove supernatant (S1), and save it in an Eppendorf tube. 12. To elute the peptides from the gel, add 80–100 µl acetonitrile/H2O-mixture (3/2) and incubate for 2 h at RT with shaking. 13. Then transfer the acetonitrile supernatant (S2) into the Eppendorf tube that already contains the other supernatant (S1). 14. Dry the pellet in a speed vac for 2–3 h. This dried pellet can be stored at −80°C. 15. For the MS analysis, dissolve the dried pellet in 5 µl of the column buffer and transfer the sample into the HPLC sample tubes.

3.9. Capillary Liquid ChromatographyElectrospray Ionization (LC-ESI)-MS Analysis

The resulting peptides from the tryptic digest were separated by reversed phase high-pressure liquid chromatography (HPLC) and analyzed by LC-ESI-MS. These instructions assume the use of the specified systems. The HPLC part is easily adaptable for other HPLC systems, the MS part is specially described for an ion trap mass spectrometer with electrospray ionization. 1. The HPLC system is directly connected to the MS. All instruments are controlled by Xcalibur System software (Thermo Electron Corp, San Jose, CA).

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2. Deliver the aqueous phase (A) (0.1% (v/v) formic acid in 5:95 acetonitrile-water) and the organic phase (B) (0.1% (v/v) formic acid in 80:20 acetonitrile-water) by an Ultimate (LC-Packings) micropump as follows: 5% B in the first 8 min, 5–50% B from 8 to 38 min, 50–95% B from 38 to 39 min, 95% B from 39 to 49min, 95–5% B from 49 to 50 min, and 5% B from 50 to 75 min. 3. Split the solvents delivered by the micropump before reaching the column by a splitter at a flow ratio of 1:1250, with the postsplitter flow rate set at 250 nl/min. 4. Fractionate the peptide mixture on an LC-Packings PepMap C18 column with a 3 µm particle size and a 100-Å-pore diameter. 5. Provide the interface between the liquid phase and the gas phase by a fused silica needle that is mounted on a nanoelectrospray ionization source (New Objective). 6. Apply a voltage difference of +1.3 kV across the fused silica needle to accomplish atmospheric pressure electrospray ionization. The aperture of the fused silica needle is positioned 1–2 mm from the opening of the ion transfer capillary and slightly off axis to minimize the entrance of non-vaporized solvent into the LCQ Deca XP ion trap mass spectrometer. No sheath gas is used. 7. For collision induced dissociation, set the collision energy in the ion trap to 35% of 5 V. Run the instrument cycling between one full MS scan and MS/MS scans of the three most abundant ions. 8. Analyze the mass spectrograms by using the Bioworks software (version 3.1; see Note 9) from Thermo Electron Corp. (San Jose, Ca, USA) including the SEQUEST algorithm and the appropriate databases. They should be protein sequences in the fasta-file format. In our experiments, we used the C. reinhardtii EST and genomic databases. Consider only high scores (see Note 10) of peptides (Xcorr > 1.5 if the charge z of the peptide is 1; Xcorr > 2 if the charge z of the peptide is 2; Xcorr > 2.5 if the charge z of the peptide is 3). Do the searches for tryptic peptides allowing two missed cleavages of trypsin.

4. Notes 1. The trace element solution should be prepared as follows: Dissolve all salts except disodium EDTA in 75 ml distilled water. For the H3BO3 and the FeSO4 · 7H2O solutions it is necessary to heat them up. Add the disodium EDTA to 250 ml water and heat the solution until the EDTA is dissolved. Mix the hot FeSO4 · 7H2O solution with the hot EDTA

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solution. Combine all other salt solutions in a 1-l beaker, heat them up to 70°C, add the hot FeSO4-EDTA solution, and adjust the pH carefully to 6.5–6.8 using a 20% KOH solution at this high temperature. Fill the beaker up to 1 l and let it cool down. It should be a clear green-colored solution. Stir the covered beaker at RT for several days until the solution shows a purple color and then filtrate it through a sterile filter. The ready-to-use trace element solution should be stored in 1-ml aliquots at −20°C. 2. The working mixture of potassium ferricyanide and sodium thiosulfate is unstable and therefore should be prepared fresh for each reaction. 3. All solvents for the LC-ESI-MS analysis should be of LC grade. 4. Do not store the cells longer than 3 months at −80°C. 5. If necessary, the IPG strips can be stored without any loss of resolution after the isoelectric focusing is finished. Therefore, the strips should be placed with the gel side up on a plastic sheet that is then covered by a plastic bag, sealed, and stored at −80°C. 6. The detection limit for silver-stained spots is approximately 0.05–0.1 ng protein per spot. 7. All protein spot volumes of each gel were normalized according to method A and B as described in Sect. 3.7. In method A, the volume of each spot was divided by the total volume of all non-saturated spots. The determination of saturated spots with the ImageMaster-2D software saturation map feature gave the result that no saturated spots were present in the 16 analyzed 2-DE gels. Since some protein spots that have a spot volume of >200,000 appear close to saturation level, we also used a variation of method A, entitled method B, to check if such spot volumes have an influence on the results. In this case, the volume of each spot is divided by the total volume of all spots, which have a volume of less than 200,000. Since these types of normalization methods tend to produce extremely small values, the results are multiplied by a scaling factor. ImageMaster 2-DE suggests two options for the scaling factor to use. One option is to multiply by a constant factor. The other option, which we used, is to multiply by the total area of all the evaluated spots in the gel. Gels with more spots would have a higher total spot volume, usually resulting in a lower normalized volume; this method compensates for the differences in spot density. Please note that the results from method A and B turned out to be very similar (7). 8. Destaining of silver-stained gel spots is necessary as silver disturbs the mass spectrometric analysis.

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9. It should be noted that meanwhile the Bioworks software has been updated and version 3.1 may not be commercially available anymore. The current update is version 3.3. 10. The cross-correlation factor “Xcorr” of the SEQUEST-analysis of the MS spectra describes the cross-correlation between the experimentally measured MS/MS spectrum and an in situ generated MS/MS spectrum of candidate peptides in the databases.

Acknowledgments We appreciate the free delivery of information by the USA (Department of Energy) and Japanese genome projects of C. reinhardtii very much. Our work was supported by grants of the Deutsche Forschungsgemeinschaft (DFG). References 1. Grossman, A. R. (2005) Paths toward algal genomics. Plant Physiol. 137, 410–427. 2. Harris, E. H. (2001) Chlamydomonas as a model organism. Annu. Rev. Plant Physiol. Plant Mol. Biol. 52, 363–406. 3. Wagner, V., Gessner, G., and Mittag, M. (2005) Functional proteomics: a promising approach to find novel components of the circadian system. Chronobiol. Int. 22, 403– 415. 4. Harris, E. H. (1989) The Chlamydomonas Sourcebook. Academic Press, Inc. San Diego, CA. 5. Grossman, A. R., Harris, E. E., Hauser, C., Lefebvre, P. A., Martinez, D., Rokhsar, D., Shrager, J., Silflow, C. D., Stern, D., Vallon, O., and Zhang, Z. (2003) Chlamydomonas reinhardtii at the crossroads of genomics. Eukaryot. Cell 2, 1137–1150. 6. Mittag, M., Kiaulehn, S., and Johnson, C. H. (2005) The circadian clock in Chlamydomonas reinhardtii. What is for? What is similar to? Plant Physiol. 137, 399–409. 7. Wagner, V., Fiedler, M., Markert, C., Hippler, M., and Mittag, M. (2004) Functional proteomics of circadian expressed proteins from Chlamydomonas reinhardtii. FEBS Lett. 559, 129–135. 8. Mittag, M. (1996) Conserved circadian elements in phylogenetically diverse algae. Proc. Natl. Acad. Sci. USA 93, 14401–14404.

9. Görg, A., Obermaier, Ch., Boguth, G., Csordas, A., Diaz, J. -J., and Madjar, J. -J. (1997) Very alkaline immobilized pH gradients for two-dimensional electrophoresis of ribosomal and nuclear proteins. Electrophoresis 18, 328–337. 10. Görg, A., Boguth, G., Obermaier, Ch., Posch, A. and Weiss, W. (1995) Twodimensional polyacrylamide gel electrophoresis with immobilized pH gradients in the first dimension (IPG-Dalt): the state of the art and the controversy of vertical versus horizontal systems. Electrophoresis 16, 1079–1086. 11. Heukeshoven , J. and Dernick, R. (1988) Improved silver staining procedure for fast staining in PhastSystem Development Unit. I. Staining of sodium dodecyl sulfate gels. Electrophoresis 9, 28–32. 12. Gharahdaghi , F., Weinberg, C. R., Meagher, D. A., Imai, B. S., and Mische, S. M. (1999) Mass spectrometric identification of proteins from silver-stained polyacrylamide gel: A method for the removal of silver ions to enhance sensitivity. Electrophoresis 20, 601– 605. 13. Hippler, M., Klein, J., Fink, A., Allinger, T. and Hoerth, P. (2001) Towards functional proteomics of membrane protein complexes: analysis of thylakoid membranes from Chlamydomonas reinhardtii. Plant J. 28, 595–606.

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14. Stauber, E. J., Fink, A., Markert, C., Kruse, O., Johanningmeier, U. and Hippler, M. (2003) Proteomics of Chlamydomonas reinhardtii light-harvesting proteins. Eukaryot. Cell 2, 978–994. 15. Mortz, E., Vorm, O., Mann, M., and Roepstorff, P. (1994) Identification of proteins in polyacrylamide gels by mass spectrometric peptide mapping combined with database search. Biol. Mass. Spectrom. 23, 249–261.

16. Link , A. J., Eng, J., Schieltz, D. M., Carmack, E., Mize, G. J., Morris, D. R., Garvik, B. M., and Yates, J. R. 3rd. (1999) Direct analysis of protein complexes using mass spectrometry. Nat. Biotechnol. 17, 676–682. 17. Eng , J., McCormack, A. L., and Yates, J. R. (1994) An approach to correlate tandem mass spectral data of peptides with amino acid sequences in a protein database. J. Am. Soc. Mass. Spectrom. 5, 976–989.

Chapter 12 Bimolecular Fluorescence Complementation (BiFC) to Study Protein–Protein Interactions in Living Plant Cells Katia Schütze, Klaus Harter, and Christina Chaban Abstract Dynamic networks of protein–protein interactions regulate numerous cellular processes and determine the ability of cells to respond appropriately to environmental stimuli. However, the study of protein complex formation in living plant cells has remained experimentally difficult and time-consuming and requires sophisticated technical equipment. In this report, we describe a bimolecular fluorescence complementation (BiFC) technique for visualization of protein–protein interactions in plant cells. This approach is based on the formation of a fluorescent complex by two non-fluorescent fragments of the yellow fluorescent protein (YFP) brought together by the association of interacting proteins fused to these fragments. We present the BiFC vectors currently available for the transient and stable transformation of plant cells and provide a detailed protocol for the successful use of BiFC in plants. Key words: Protein–protein interaction, BiFC, YFP, tobacco infiltration, protoplast transfection, intracellular localization, bZIP transcription factors.

1. Introduction The identification and characterization of protein–protein interaction provide crucial information for understanding the molecular mechanisms underlying biological processes. Although numerous methods for the detection of interacting proteins have been developed, they often operate under non-native conditions or in non-plant systems. In vitro interaction assays using recombinant proteins, which are not folded properly, may result in the formation of non-specific aggregations or loss of interaction. Furthermore, co-immuno-precipitation experiments on the basis of plant

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extracts bear the risk that proteins that are normally localized in different cell compartments are observed to interact (1, 2). The yeast two-hybrid screen is widely used for the detection of interacting proteins in living cells, while retaining the native state of proteins. However, this heterologous host system suffers from significant false-positive and false-negative rates (1, 3, 4). The cloning of spontaneously fluorescent protein from the jellyfish Aequorea victoria opened a new era in the visualization of proteins in vivo. With the help of this approach several methods have been developed to analyse protein–protein interactions in the homologous host system. By monitoring the fluorescence resonance energy transfer (FRET) between colour variants of GFP fused to given proteins, molecular interactions have been successfully examined (reviewed in Ref. 5). Nevertheless, the necessity for the expression of high protein levels and for sophisticated equipment to determine small changes in the fluorescence restricts FRET-based methods to selected applications and experienced laboratories (the advantages and limitations of the method have been recently reviewed by Bhat et al., 6). Here, we describe the relatively fast and inexpensive method, bimolecular fluorescence complementation (BIFC), which was originally applied for protein–protein interaction studies in animal cell systems (7) and has since been successfully used in the plant field. It is based on the formation of a fluorescent complex by nonfluorescent fragments of the enhanced yellow fluorescent protein (YFP) when brought together by the interaction of two partners fused to these fragments (Fig. 12.1, 7–9). Besides the direct visualization of complex formation in living cells, this method also allows the detection of the intracellular location at which the protein association occurs (9, 10). However, it should be mentioned that sometimes very high expression of the YFP fragments may lead to the detection of unspecific fluorescence (9). This might be due to the intrinsic tendency of YFP fragments to form irreversible complexes (11). This phenomenon might also enhance sporadic false-positive signals. By removing the amino acids 153 to 155 in the N-terminal fragment of YFP by the introduction of a pre-mature stop codon

Fig. 12.1. Reconstitution of fluorescent YFP by its two non-fluorescent fragments mediated by protein–protein interaction. YFP fragments (YFPN, N-terminal fragment [amino acids 1–155]; YFPC, C-terminal fragment [amino acids 156–239]) are fused to the proteins of interest. Interaction of these proteins leads to the formation of a fluorescent complex by the fragments of the YFP.

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(153STOP), the stability of the fluorophore complex could be reduced, thus lowering the false-positive rate and probably making it possible to follow the dynamics of protein–protein interactions in vivo (C. Oecking, personal communication). In this report we present sets of plant-compatible BiFC vectors, which allow the examination of protein–protein interactions using either the N- or C-terminal fusion of YFP fragments in transiently or stably transformed plant systems. Vectors have been generated containing Gateway-compatible or classic multiple cloning sites. Using these vectors, we investigated dimerization between the subfamily C members of the Arabidopsis thaliana basic region-leucine zipper (bZIP) transcription factors (9). The detailed protocols for the transformation of Agrobacterium tumefaciens, the infiltration of Nicotiana benthamiana leaves and the transfection of protoplasts are described here. Depending on particular bZIP proteins studied, the interacting complexes were detected either in the nucleus or in the cytoplasm of the plant systems (Fig. 12.2). The differential intracellular location of bZIP factors was confirmed by an alternative method (12) and may contribute to the distinctive mechanisms of the regulation of bZIP factordependent transcription (13). Our study indicates that the BiFC technique represents a fast, efficient and convenient tool to investigate protein–protein interactions in living plant cells.

Fig. 12.2. BiFC visualization of AtbZIP63 and AtbZIP10 homodimerization in Arabidopsis protoplasts and Agrobacterium infiltrated tobacco leaves. (I) Epifluorescence and (II) bright field images of Arabidopsis protoplasts (upper row) and epidermal leaf cells (lower row) infiltrated with a mixture of Agrobacterium suspensions harbouring the constructs encoding the indicated fusion proteins. In both cell systems AtbZIP63 homodimers accumulate exclusively in the nucleus, whereas AtbZIP10 homodimers are found in the nucleus and the cytoplasm.

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Table 12.1 Vectors for transient protoplast transfection1 YFP-Fusion

Cloning

Vector

Type

Reference

C-terminal

Gateway

pUC-SPYNE GW

High copy

(9)

pUC-SPYCE GW

High copy

(9)

pUC-SPYNE

High copy

Oecking, unpublished

pUC-SPYNE

High copy

(9)

pUC-SPYCE

High copy

(9)

pUC-SPYNE

High copy

Oecking, unpublished

pE SPYNE

High copy, binary

(18)

pE SPYCE

High copy, binary

(18)

153STOPGW

MCS

153STOP

N-terminal

Gateway

1

A schematic sketch of vectors is shown in Fig. 12.3.

Table 12.2 Vectors for either tobacco leaf infiltration or stable transformation of plants1 YFP-Fusion Cloning

Vector

Type

Reference

C-terminal

pSPYNE-35S GW

Binary

Lahaye, unpublished

pSPYCE-35S GW

Binary

Lahaye, unpublished

pSPYNE-35S

Binary

(9)

pSPYCE-35S

Binary

(9)

pE SPYNE

High copy, binary

(18)

pE SPYCE

High copy, binary

(18)

Gateway

MCS

N-terminal

Gateway

1

A schematic sketch of vectors is shown in Fig. 12.3.

2. Materials 2.1. Plant BiFC Vectors

1. Wall digestion solution without enzymes: 8 mM CaCl2, 0.4 M mannitol, pH 5.5, filter sterile.

2.2. Protoplast Transfection

2. Wall digestion solution: 1% cellulase, 0.25% macerozym, 8 mM CaCl2, 0.4 M mannitol, pH 5.5, filter sterile.

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3. W5 solution: 154 mM NaCl, 125 mM CaCl2, 5 mM KCl, 5 mM glucose, pH 5.8–6.0, autoclave. 4. MMM solution: 15 mM MgCl2, 0.1% MES, 0.5 M mannitol, pH 5.8, autoclave. 5. PEG solution: 40% PEG 4000, 0.4 M mannitol, 0.1 M Ca(NO3)2, pH 8–9 (the pH needs 1–2 h to stabilize), autoclave. 6. Macro stock (1000 ml): 1.5 g NaH 2PO 4 · H 2O, 9.0 g CaCl 2 · 2H 2O, 25 g KNO 3, 2.5 g NH 4NO 3, 1.34 g (NH 4) 2SO 4, 2.5 g MgSO 4 · 7H 2O, add H 2O up to 1 l, autoclave. 7. Micro stock (100 ml): 75 mg KI, 300 mg H3BO3, 1 g MnSO4 7H 2O (0.6 g MnSO 4 * H 2O), 200 mg ZnSO 4 · 7H 2O, 25 mg Na2MoO4 · 2H2O, 2.5 mg CuSO4 · 5 H2O, 2.5 mg CoCl2 · 6H2O, add H2O up to 100 ml, filter sterile and freeze. 8. K3 solution (100 ml): 10 ml macro stock, 0.1 ml micro stock, 0.1 ml vitamin stock, 0.5 ml EDTA stock, 1 ml Caphosphate stock, 10 mg myo-inositol, 25 mg d(+)-xylose, 13.7 g sucrose, pH 5.6, filter sterile and freeze in 10 ml aliquots. 9. Vitamin stock (100 ml): 100 mg nicotinacid, 100 mg pyridoxin HCl, 1 g thiamin · HCl, add H2O up to 100 ml, filter sterile and freeze. 10. EDTA stock (1000 ml): 7.46 g EDTA dissolve in 300 ml H2O and cook, 5.56 g Fe*SO4 7H2O dissolve in 300 ml H2O and cook, mix and add H2O up to 1 l, autoclave and keep in the dark. 11. Ca-phosphate stock (200 ml): 1.26 g CaHPO4 · 2H2O dissolve in H2O, add H2O up to 200 ml, pH 3 with 25% HCl, autoclave and keep in the dark. 2.3. Transformation of Agrobacterium cells

Agrobacterium tumefaciens GV3101/pMP90, a strain based on the C58 and pTiC58 genotypes (14–16) that carries genes for resistance to gentamycin and rifampicin, was used in this study. An Agrobacterium strain containing the p19 protein of tomato bushy stunt virus was used to suppress gene silencing in transformed tobacco leaves (17). 1. YEB-medium (1000 ml): 5 g bactopeptone, 5 g beef extract, 1 g yeast extract, 5 g saccharose, 0,5 g MgSO4 · 7H2O and add 15 g agar for solid medium, autoclave. 2. TE buffer: 10 mM Tris-HCl, pH 7.5, 1 mM EDTA, autoclave.

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3. The antibiotics for the strain and for the plasmids were used at the following concentrations: rifampicin 100 µg/ml, gentamycin 40 µg/ml, kanamycin 25 µg/ml, carbenicillin 50 µg/ml. 2.4. Infiltration of Nicotiana benthamiana

1. AS-medium (100 ml): 1 ml 1 M MES-KOH, pH 5.6, 333 µl 3 M MgCl2, 100 µl 150 mM acetosyringon (in DMSO, stored in aliquots at −20°C); prepare fresh from stock solutions. 2. Sterile single-use syringes (1 ml)

2.5. SDS-PAGE and Western Analysis

1. Sample buffer: 8 M urea, 2% SDS, 0.1 M DTT, 20% glycerol, 0.1 M Tris-HCl (pH 6.8), 0.004% bromophenol blue (store frozen in aliquots). 2. Blocking buffer: TBS (50 mM Tris-HCl pH 7.4, 150 mM NaCl) containing 0.1% Tween-20 and 4% milk powder. 3. Primary antibodies: rat monoclonal anti-HA and mouse monoclonal anti-c-myc (Roche). 4. Secondary antibodies: anti-rat IgG alkaline phosphatase conjugate developed in goat (Sigma) and anti-mouse IgG alkaline phosphatase conjugate developed in goat (BIO-RAD) 5. AP-buffer: 100 mM Tris-HCl pH 9.5, 100 mM NaCl, 5 mM MgCl2 6. NBT-Solution: 50 mg/ml nitro-blue tetrazolium chloride (NBT) in 70% dimethylformamide, store in aliquots at −20°C. 7. BCIP-Solution: 50 mg/ml 5-bromo-4-chloro-3-indolylphosphate, disodium salt (BCIP) in H2O, store in aliquots at −20°C. 8. AP-staining solution: mix 66 µl of NBT-solution and 33 µl of BCIP solution in 10 ml AP-buffer.

3. Methods 3.1. Cloning into BiFC vectors

Different sets of vectors are available for N-terminal or C-terminal YFP fusions. Because the fusion can interfere with the interaction domains of proteins of interest, it is important to decide whether the N-terminal or C-terminal YFP fusion is more suitable for the study (see Note 1). For example, with some S-group bZIP transcription factors from Arabidopsis, the interaction could only be shown when the YFP fragments were fused to the N-terminus of the proteins (18). Depending on the cloning strategy (classic or Gateway) the cDNA can be cloned through restriction and ligation or through LR-recombination from the entry clone into the

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3.2. BiFC in Protoplasts 3.2.1. PEG Mediated Transfection of Arabidopsis thaliana Cell Suspension Protoplasts

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destination/expression clone. The presence and correct orientation of the insert should be confirmed with appropriate restriction enzymes and sequencing (compare Tables 12.1 and 12.2, see Fig. 12.3). 1. Use cells 3 days after sub-cultivation. 2. Collect the cells (2 × 10 ml culture) by centrifugation at 400× g for 5 min. 3. Wash once with 10 ml of wall digestion solution without enzymes, centrifuge at 100 × g for 5 min. 4. Resuspend each pellet in 7 ml of digestion solution and dispense each into a Petri dish. 5. Incubate at 26°C in the dark for 6 h in a shaker at 50 rpm. 6. Collect protoplasts in 2 tubes by centrifugation at 100× g for 5 min.

Fig. 12.3. Schematic representation of plant-compatible BiFC vectors. (a) Classic BiFC vectors with C-terminal fusion of the YPF fragments: pUC-SPYNE/ 35S-pSPYNE and pUC-SPYCE/35S-pSPYCE (9). 35S, 35S promoter of the cauliflower mosaic virus; MCS, multiple-cloning site. Unique restriction sites are illustrated with normal letters, others with italic letters. c-myc, c-myc affinity tag; HA, hemagglutinin affinity tag; YFPN, N-terminal fragment of YFP (amino acid 1–155); YFPC, C-terminal fragment of YFP (amino acid 156–239); NosT, terminator of the nos-gene. (b) Gateway compatible BiFC vectors with C-terminal fusion of the YPF fragments: pUC-SPYNE GW/35S-pSPYNE GW and pUC-SPYCE GW/35SpSPYCE GW. attR1-CmR-ccdB-attR2, Gateway conversion cassette (Invitrogen). (c) Gateway compatible BiFC vectors with N-terminal fusion of the YPF fragments: pE SPYNE and pE SPYCE (18).

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7. Wash once with 10 ml of wall digestion solution without enzymes. 8. Centrifuge at 100× g for 5 min. 9. Remove the supernatant and resuspend cells in the remaining solution, slowly add 10 ml of W5 solution, mix gently and centrifuge at 100× g for 5 min. 10. Resuspend pellets in 10 ml of W5 solution, take an aliquot for counting, store 20 min in the dark at 4°C and count protoplasts during this time, for example in a Fuchs-Rosenthal chamber. 11. Centrifuge at 100× g for 5 min, remove all the W5 solution and resuspend pellet in MMM solution to a density of 2 × 106 protoplasts/ml. 12. Redistribute samples of 250 µl into tubes, add 30 µg of plasmid DNA, (high DNA purity is required) very slowly add 250 µl PEG solution, mix gently and incubate 15–20 min. 13. Gradually add 10 ml of W5 solution; this needs to be done very slowly, to not disrupt the protoplasts. 14. Centrifuge at 100× g for 5 min, and remove the supernatant. 15. Resuspend the protoplasts in 2 ml of K3 solution, and incubate at 26°C in the dark. 3.2.2. Microscopy

Usually the expression of the fusion proteins is efficient 12–18 h after the transfection. Fluorescence was assayed with the Nikon eclipse 90i microscope and quantification of the fluorescence intensity was performed using the Metamorph software (Universal Imaging Corporation Downington, PA, USA). The optimal excitation wavelengths for YFP are in the range of 490–515 nm; the maximal emission intensity is observed in the range of 520–560 nm. Therefore, the YFP fluorescence might be easily visualized with a fluorescence microscope using an appropriate commercially available filter set.

3.3. BiFC in Nicotiana benthamiana

Two alternative protocols can be used for successful transformation of Agrobacterium tumefaciens cells. Electroporation (according to the method described by Mersereau et al. 19): 1. Grow Agrobacterium cells at 28°C to a OD600 of 1–1.5 and harvest the cells by centrifugation at 3,000× g for 5 min.

3.3.1. Transformation of Agrobacterium Cells

2. Wash the cells five times in sterile, cold water, centrifuge at 3,000× g for 5 min and remove the supernatant. 3. Resuspend the cells in 10% glycerol to a density of about 1010 cells/ml and store them frozen at −70°C. 4. Thaw the frozen cells in ice and add 0.5–1 µl of plasmid DNA to 50 µl of recipient cells and transfer the cells to a cold electroporation cuvette.

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5. Carry out the electroporation by applying a single electrical pulse of 2.5 kV and 400 Ω (e.g., Bio-Rad Gene Pulser). 6. After applying the pulse immediately suspend the cells in 1 ml of cold YEB medium and incubate for 1–2 h at 28°C. 7. Recover the cells by short centrifugation, resuspend in 0.1 ml YEB medium and spread on appropriate selective media. Let colonies grow for 2–3 days at 28°C. Chemical transformation (freeze–thaw method, according to Ref. 20): This method is simple, rapid and robust, although the cell transformation frequency is comparatively low. 1. Grow a single Agrobacterium colony in 5 ml of YEB (with antibiotics) medium overnight at 28°C. 2. Add 2 ml of the overnight culture to 50 ml YEB medium and let the culture grow to an OD600 of 0.5–0.7 at 28°C. 3. Centrifuge the cell suspension at 3,000× g for 15 min at 4°C, and discard the supernatant. 4. Wash the cells once in 5 ml pre-cooled TE buffer and finally resuspend the pellet in 5 ml of ice-cold fresh YEB medium. Use aliquots of 0.2 ml directly for transformation or freeze in liquid nitrogen and store at −70°C. 5. Add about 1–5 µg of plasmid DNA to the competent cells, incubate 5 min on ice and freeze the cells in liquid nitrogen (5 min). 6. Thaw the cells by incubating for 5 min at 37°C. 7. Add 1 ml of YEB medium and incubate for 2–4 h at 28°C with shaking. 8. Centrifuge for 30 s at 3,000× g, discard the supernatant, resuspend the cells in 0.1 ml YEB medium and spread the cells on a YEB agar plate containing an appropriate antibiotic selection. 9. Incubate the plate at 28°C. Transformed colonies should appear in 2–3 days. Colonies used for tobacco infiltration should be checked by colony PCR. 3.3.2. Infiltration of Nicotiana benthamiana (according to Ref. 21)

1. Distribute a single colony on a fresh plate and let it grow for 1–2 days at 28°C. 2. Inoculate the cells in 5 ml of YEB medium and incubate the culture overnight under shaking at 28°C. 3. Centrifuge the culture for 15 min at 4,000× g and resuspend pelleted cells in 1 ml of AS medium. Dilute the cells with AS medium to OD600 0.7–0.8 (about 1 ml per leaf). 4. Prepare working suspensions by mixing appropriate clones containing the BiFC constructs and the p19 plasmid at a 1:1:1 ratio (total volume of 3 ml per leaf); let them stand for 2–4 h.

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5. Give the excess of water to tobacco plants (see Note 2). It is convenient to use one plant for each variant. Infiltration of two or three leaves per plant with the same sample might be advisable especially when protein expression level is a problem. 6. Co-infiltrate mixed Agrobacterium strains into the abaxial air space of tobacco leaves (see Fig. 12.4). It is important that the total leaf air space is infiltrated. In the case of incomplete infiltration mark the remaining area with a pen because after a while the infiltrated area will not be distinguishable from that not infiltrated. 3.3.3. Microscopy

Observe the fluorescence in the epidermal cell layer of the lower leaf surface expressing the fusion proteins 1–3 days after infiltration (see Notes 3 and 4).

3.4. SDS-PAGE and Western Analysis

It is necessary to verify the expression of the BiFC fusion proteins in transfected protoplasts and Agrobacterium infiltrated tobacco leaf discs (see Note 5 and Fig. 12.5).

3.4.1. Sample Preparation from Protoplasts

1. Dilute 0.5 ml of protoplast suspension with 0.5 ml water in an Eppendorf tube and centrifuge at 5,000× g for 10 min to collect the protoplasts. 2. Resuspend the protoplasts in 50–100 µl hot SDS-sample buffer and denature for 5 min at 95°C. Shortly spin down cell debris and separate 15–20 µl of the supernatant by SDS-PAGE. Run two parallel gels for subsequent detection of fusion proteins by both anti-HA and anti-c-myc antibodies.

Fig. 12.4. Agrobacterium infiltration of Nicotiana benthamiana leaves. (a) A 5-week-old N. benthamiana plant that is appropriate to be infiltrated. The leaves at the optimal developmental stage for infiltration are indicated by arrows. (b) Infiltration of the Agrobacterium suspension into the abaxial air space of a tobacco leaf (see Color Plates ).

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Fig. 12.5. Western blot analysis of overexpressed proteins in Agrobacterium-infiltrated tobacco leaves. The expression of the proteins is demonstrated by immunodetection using alkaline phosphatase with (a) anti-c-myc antibodies for YFPN-fusion proteins and (b) anti-HA antibodies for YPFC-fusions. Total protein extracts from tobacco leaves infiltrated with an Agrobacterium suspension containing constructs encoding the following fusion proteins (1) AtbZIP10-YFPN/AtbZIP63-YFPC, (2) AtbZIP63-YFPN/AtbZIP9-YFPC, (3) AtbZIP9-YFPN/AtbZIP63-YFPC were separated on a 12.5% SDS-PAGE gel. The size of the marker proteins is shown on the left (see Color Plates ) . 3.4.2. Sample Preparation from Infiltrated Tobacco Leaves

1. Excise a leaf disc of about 40–50 mg, homogenize it in liquid nitrogen in an Eppendorf tube, add 150–200 µl of hot SDS-sample buffer and vortex. Incubate 5 min at 95°C. 2. Centrifuge 5 min at maximal speed to remove cell fragments. 3. Separate 15–20 µl of the supernatant by SDS-PAGE. Run two parallel gels for subsequent detection of fusion proteins by both anti-HA and anti-c-myc antibodies. SDS-PAGE and Western blot transfer are to be carried out using standard protocols.

3.4.3. Immunodetection of the Fusion Proteins

1. Incubate the membranes for 2–3 h at room temperature or overnight at 4°C in blocking buffer with continuous shaking. 2. Discard the blocking buffer and incubate the membranes for 2 h at room temperature with the primary anti-HA (1:800) or anti-cmyc (1:1000) antibodies in TBS containing 0.1% Tween-20. 3. Remove the primary antibody (can be reused several times) and wash 3 × 5 min with TBS-Tween. 4. Incubate the membrane for 1–2 h at room temperature with the secondary antibody (anti-rat-AP 1:7000 for anti-HA and anti-mouse-AP 1:3000 for anti-c-myc).

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5. Remove the secondary antibody (can be reused several times) and wash 3 × 10 min with TBS-Tween. 6. Equilibrate the membrane in AP-buffer for a short time. 7. Stain with fresh AP staining solution until the signal is clearly visible (15 min to overnight) – do not shake! To reduce background, keep the membranes at 4°C when a longer reaction time is necessary. 8. Stop the reaction by rinsing the membranes twice with water. Dry the membranes.

4. Concluding remarks Although the BiFC approach was launched not so long ago, it is becoming more and more widely used in the field of protein–protein interaction research. This can be attributed to its main advantages, which include the possibility to assay protein interaction directly in planta with comparative simplicity. Up to now, this assay has been primarily used to verify the interaction of particular proteins, although it is not limited to this. Other possible applications such as multiple protein–protein interaction studies by using multicolor BiFC (22) and identification of novel interactions by in planta screening are presently being developed and tested. The latter would be achievable by co-transfection of protoplasts with the constructs, one of which bears the protein of interest fused with N-terminus of YFP and the second a cDNA-library fused with C-terminus of YFP, and by subsequent sorting and analysis of positive, i.e., fluorescent protoplasts (K. Berendzen and K. Harter, unpublished).

5. Notes 1. The presence of signal peptides or transmembrane domains determines the protein terminus at which the YFP fragment must be fused. However, in the case of BiFC, additional protein features, namely their allosteric configuration and orientation during interaction, also play an important role. As these features are usually difficult to predict, the testing of all possible protein combinations is crucial. Moreover, the testing of the fusion proteins for their functional activity is recommended when feasible (e.g. by complementation of a mutant phenotype). 2. The developmental stage of the Nicotiana benthamiana plants is important for the success of this method. Good

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results were obtained with 4- to 6-week-old plants and a leaf size of about 5 cm2 (see Fig. 12.4). Older plants already developing flowers are less appropriate. 3. As mentioned in the introduction, the expression levels of investigated proteins are quite important. Usually, the highest rate of protein accumulation is observed on the second day after infiltration. Therefore, when the expression of the BiFC fusion proteins is low or the interaction study is performed with the shortened N-terminal YFP fragment (SPYNE153STOP), which by itself decreases the intensity of the fluorescence, the microscopic analysis of the tobacco leaves should be carried out on this day. In contrast, if too high expression of the fusion proteins leads to non-specific complex formation, it might be helpful to survey the leaves 24 h or 5 days after infiltration. Fluorescence is usually detectable for 2–3 days and may persist for up to 6 days. 4. It is not necessary to peel off the epidermis to monitor the fluorescence; however, it might improve the quality of the images. 5. In order to correctly interpret the obtained pictures it is absolutely necessary to analyse the protein expression level by means of western analysis as shown in Fig. 12.5. This is possible due to the presence of c-myc and HA epitopes in the expressed proteins fused with N-terminal and C-terminal YFP fragments, respectively.

Acknowledgments The authors gratefully acknowledge the support by Caterina Brancato, ZMBP, University of Tübingen, for the experiments. We also thank F. de Courcy for her support in proofreading the manuscript. This work was supported by an SFB 446 grant to K.H. References 1. Phizicky, E. M., and Fields, S. (1995) Proteinprotein interactions: methods for detection and analysis. Microbiol. Rev. 59, 94–123. 2. Howell, J. M., Winstone, T. L., Coorssen, J. R., and Turner, R. J. (2006) An evaluation of in vitro protein-protein interaction techniques: Assessing contaminating background proteins. Proteomics 6, 2050–2069. 3. Qi, Y., Ziv, B. -J., and Klein-Seetharaman, J. (2006) Evaluation of different biological data and computational classification methods for use in protein interaction prediction. Proteins 63, 490–500.

4. von Mering, C., Krause, R., Snel, B., Cornell, M., Oliver, S. G., Fields, S., and Bork, P. (2002) Comparative assessment of large scale data sets of protein–protein interactions. Nature 417, 399–403. 5. Lalonde, S., Ehrhardt, D. W., and Frommer, W. B. (2005) Shining light on signaling and metabolic networks by genetically encoded biosensors. Curr. Opin. Plant Biol. 6, 574–581. 6. Bhat, R. A., Lahaye, T., Panstruga, R. (2006) The visible touch: in planta visualization of protein-protein interactions by fluorophorebased methods. Plant Methods 2, 12.

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7. Hu, C. -D., Chinenov, Y., and Kerppola, T. (2002) Visualization of interactions among bZIP and Rel family proteins in living cells using bimolecular fluorescence complementation. Mol. Cell 9, 789–798. 8. Bracha-Drori, K., Shichrur, K., Katz, A., Oliva, M., Angelovici, R., Yalovsky, S., and Ohad, N. (2004) Detection of protein–protein interactions in plants using bimolecular fluorescence complementation. Plant J. 40, 419–427. 9. Walter, M., Chaban, C., Schütze, K., Batistic, O., Weckermann, K., Näke, C., Blazevic, D., Grefen, C., Schumacher, K., Oecking, C., Harter, K., and Kudla, J. (2004) Visualization of protein interactions in living plant cells using bimolecular fluorescence complementation. Plant J. 40, 428–438. 10. Hoff, B. and Kück, U. (2005) Use of bimolecular fluorescence complementation to demonstrate transcription factor interaction in nuclei of living cells from the filamentous fungus Acremonium chrysogenum. Curr. Genet. 47, 132–138. 11. Magliery, T. J., Wilson, C. G., Pan, W., Mishler, D., Ghosh, I., Hamilton, A. D., Regan, L. (2005) Detecting protein-protein interactions with a green fluorescent protein fragment reassembly trap: scope and mechanism. J. Am. Chem. Soc. 127, 146–157. 12. Näke, C. (2001) Charakterisierung von CPRF2-homologen bZIP-Proteinen aus Arabidopsis thaliana unter besonderer Berücksichtigung ihrer intrazellulären Verteilung. Inaugural Dissertation. Biologische Fakultät, Universität Freiburg, Germany. 13. Kaminaka, H., Nake, C., Epple, P., Dittgen, J., Schutze, K., Chaban, C., Holt, B. F. 3rd , Merkle, T., Schafer, E., Harter, K., and Dangl, J. L. (2006) bZIP10-LSD1 antagonism modulates basal defense and cell death in Arabidopsis following infection. EMBO J. 20, 4400–4411. 14. Bechthold, N., Ellis, J., and Pelletier, G. (1993) In planta Agrobacterium-mediated

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gene transfer by infiltration of adult Arabidopsis thaliana plants. CR. Acad. Sci. Paris Life Sci. 316, 1194–1199. Katavic, V., Haughn, G. W., Reed, D., Martin, M., and Kunst, L. (1994) In planta transformation of Arabidopsis thaliana. Mol. Gen. Gene.t 245, 363–370. Koncz, C., and Schell, J. (1986) The promoter of the TL-DNA gene 5 controls the tissuespecific expression of chimeric genes carried by a novel type of Agrobacterium binary vector. Mol. Gen. Genet. 204, 383–386. Voinnet, O., Rivas, S., Mestre, P., and Baulcombe, D. (2003) An enhanced transient expression system in plants based on suppression of gene silencing by the p19 protein of tomato bushy stunt virus. Plant J. 33, 949–956. Weltmeier, F., Ehlert, A., Mayer, C. S., Dietrich, K., Wang, X., Schütze, K., Alonso, R., Harter, K., Vicente-Carbajosa, J., and Dröge-Laser, W. (2006) Combinatorial control of Arabidopsis proline dehydrogenase transcription by specific heterodimerisation of bZIP transcription factors. EMBO J. 12, 3133–3143. Mersereau, M., Pazour, G. J., and Das, A. (1990) Efficient transformation of Agrobacterium tumefaciens by electroporation. Gene 31, 149–151. Höfger, R. and Willmitzer, L. (1988) Storage of competent cells for Agrobacterium transformation. Nucl. Acids Res. 16, 9877. Romeis, T., Ludwig, A. A., Martin, R., and Jones, J. D. G. (2001) Calcium-dependent protein kinases play an essential role in a plant defence response. EMBO J. 20, 5556–5567. Hu, C. D. and Kerppola, T. K. (2003) Simultaneous visualization of multiple protein interactions in living cells using multicolor fluorescence complementation analysis. Nat. Biotechnol. 21, 539–545.

Chapter 13 Fluorescence Cross-Correlation Spectroscopy of Plant Proteins Hideki Muto, Masataka Kinjo, and Kotaro T. Yamamoto Abstract Fluorescence cross-correlation spectroscopy (FCCS) is a technique that physically evaluates the molecular interaction between two fluorophore-tagged molecules such as proteins and oligonucleotides in a quantitative manner. Because it simply makes use of the coincidental movement of two molecules, it could avoid the complexity that sometimes occurs in other fluorescent techniques such as fluorescence resonance energy transfer. The present chapter describes procedures for FCCS of plant proteins expressed and measured in HeLa cells. Determination in plant cells is also mentioned briefly. Key words: Fluorescence cross-correlation spectroscopy, fluorescent proteins, molecular interaction, single-molecule measurements.

1. Introduction The fluorescence emitted from a small number of fluorophorecontaining molecules in a small confocal volume fluctuates because of diffusion of the molecules into and out of the confocal volume. Under these conditions, the correlation of fluorescence intensity is calculated in the function of time (the auto-correlation function) in fluorescence correlation spectroscopy (FCS). Quantitative information on the diffusion kinetics and local concentration of the molecule can be extracted from the auto-correlation function (1). Because the diffusion coefficient is dependent on the size of the molecules, binding of two molecules is estimated from the auto-correlation function when molecular size increases significantly upon binding (2).

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Fluorescence cross-correlation spectroscopy (FCCS) is a modification of FCS. Consider the case where two molecules are labeled with two different fluorophores such as green fluorescent protein (GFP) and red fluorescent protein (RFP). When their fluorescence is recorded separately in the green and red channels (G and R, respectively), the cross-correlation between the green and red fluorescence is calculated. The cross-correlation would be strong if two labeled molecules bind together, but absent if they move independently in a confocal volume. Note that when two molecules that have the same diffusion coefficients and concentrations move independently, their auto-correlations would be identical, having the same amplitudes and decaying together, but their cross-correlation would be absent. Thus, we can estimate the molecular interaction quantitatively by FCCS. Molecular interaction is often determined by another fluorescence technique, fluorescence resonance energy transfer (FRET), where the efficiency of energy transfer between two fluorophores is used as a measure of molecular interaction (3). However, application of FRET is sometimes limited because its efficiency is dependent on the relative orientation between the donor and acceptor fluorophores as well as the distance between them. On the other hand, FCS and FCCS are free from such complexity. Although the principle of FCS has been established for more than 30 years (4), it was not until the mid-1990s that FCS was successfully applied to biophysical research owing to improved laser technology, confocal optics, and computational ability (5). FCS is a versatile method in that it can be used for any macromolecules such as nucleic acids (6, 7), polypeptides (2, 8), and polysaccharides (1). It is also used in vitro (2, 6, 7) and in vivo and in different cellular compartments, such as cytoplasm (8), nuclei (9), plasma membrane (10, 11), and cell wall (1). For FCS, molecules of interest are either cross-linked to fluorophores (1, 2, 6, 7, 10) or genetically fused to fluorescent proteins (FP) (8, 9). We previously used FCCS to examine interactions between plant transcription regulators expressed in HeLa cytoplasm (8). Here, we describe FCCS of FP-fused proteins expressed in HeLa cells by the use of the LSM510-ConfoCor 2 system (Zeiss). In this system, fluorescence is detected by a single-photon counter, and correlation functions of the signal are calculated in real time by a built-in correlator. Because of singlephoton counting, the intensity of fluorescence is expressed in count rate (Hz). Since concentration of fluorophores can be measured from the auto-correlation, we can also estimate count rate per molecule (CPM) in FCS. ConfoCor 2 has been updated to ConfoCor 3.

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2. Materials 2.1. Cell Culture

1. Culture medium: Dulbecco’s Modified Eagle’s Medium (DMEM) (Sigma-Aldrich, St. Louis, MO) supplemented with 10% fetal bovine serum (Gibco/Invitrogen, Carlsbad, CA), 100 U/ml penicillin, and 0.1 mg/ml streptomycin (Penicillin-Streptomycin, 100 X; Sigma). Stored at 4°C. 2. 0.25% trypsin-EDTA solution (Sigma) stored at −20°C. 3. FuGENE 6 Transfection Reagent (Roche, Basel, Switzerland) stored at 4°C. 4. Opti-MEM I Reduced-Serum Medium (Gibco/Invitrogen) stored at 4°C. 5. CO2 incubators. 6. Phosphate-buffered saline, pH 7.2 (PBS) (Gibco/Invitrogen), stored at room temperature. 7. TE: 10 mM Tris-HCl, pH 7.5, containing 1 mM EDTA, stored at room temperature after sterilization. 8. Tissue culture dishes: CellStar Dish 664160, 100 × 20 mm (Greiner Bio-One, Frickenhausen, Germany). 9. Chambered cover slips: Lab-Teck #1.0 borosilicate cover glass system, 8 wells (Nalge Nunc, Rochester, NY). 10. 15-ml centrifuge tubes: #10-0152 (Biologix Research, KS).

2.2. Confocal Imaging

2.3. Plasmids

1. ~1 × 10−7 M Rhodamine 6G (Rho6G; Molecular Probe, OR) stored in the dark at room temperature. For FCCS analysis, each molecule of interest must be tagged by two different fluorophores. For measurement of protein–protein interactions, we recommend genetically encoded fluorophores, enhanced GFP (EGFP) for the green fluorescence and monomeric RFP (mRFP) tandem dimer (8, 12), mCherry (13), or mKeima (14) for the red fluorescence (see Note 1). The target proteins must be fused to each fluorophore. Plasmids that encode each non-fused FP are necessary for negative controls to estimate the minimum cross-correlation; a plasmid that encodes tandem dimer of two different FPs is also needed to measure the maximum cross-correlation. Thus, to measure the interaction between two proteins, at least five plasmids that encode the target proteins fused to FPs and three control plasmids are needed. For measurement in HeLa cells, constructs were made from pEGFP-C1 (Clontech, Mountain View, CA), in which target genes were constitutively expressed under the control of the cytomegalovirus (CMV) promoter in transfected HeLa cells.

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3. Methods 3.1. Cell Culture

1. Thaw stored HeLa cells in culture medium in a tissue culture dish. 2. Incubate at 37°C for 2 or 3 days in a CO2 incubator. 3. Discard culture medium using an aspirator. 4. Wash cells, by adding 5 ml of PBS, swirling, and discarding PBS with an aspirator. Repeat washing twice. 5. Add 0.5 ml of 0.25% trypsin and swirl it over the monolayer of cells. Incubate for a few minutes at 37°C. 6. Add 5 ml of culture medium to neutralize the trypsin digestion, transfer cells to a 15-ml tube, and spin them down by centrifugation at 180× g for 1 min. 7. Suspend the cell pellet with 5 ml of culture medium, and centrifuge again as above. 8. Resuspend the cell pellet with 5 ml of culture medium. 9. Transfer 300 µl of cells to a chamber of an 8-chambered cover slip.

3.2. Transfection

1. Add 1 µl of FuGENE6 to 10 µl of Opti-MEM. 2. Add 1–2 µl of water or TE containing 500 ng of plasmid DNA, mix well by gentle tapping, and leave for 15 min at room temperature. 3. Add all the solution to 300 µl of HeLa cells prepared as in Sect. 3.1. 4. Incubate at 37°C overnight in a CO2 incubator.

3.3. Confocal Setting

1. Place 15–20 µl of 10−6–10−7 M Rho6G on a chambered cover glass. 2. Adjust xy-axis at the Rho6G solution and z-axis at 200 µm above the surface of the cover glass. 3. Set the beam path to LP610 for R and BP505-530 for G; set the pinhole diameter to 78 nm for R and 40 nm for G (see Note 2); and set the intensity of 1 mW He-Ne laser (543 mm) and 25 mW Ar+ laser (488 nm) to 5 and 3% output, respectively. 4. Adjust pinhole position by auto-adjustment for each channel. 5. Measure fluorescence intensity of Rho6G for 15 s. 6. Perform curve fitting and data analysis as described in Sect. 3.5. except for fixing component at 1. 7. Check that the diffusion time is 10–30 µs and that fluorescence intensity in CPM is about 1.5–5 kHz for both G and R.

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1. Discard culture medium of the transfected HeLa cells in an 8-chambered cover slip using an aspirator. 2. Put 400 µl of Opti-MEM I in the chamber. 3. Select cells that express both green and red fluorescence with an appropriate intensity for the FCCS measurement (see Note 3 and Fig. 13.1). 4. Scan the cells using a water immersion objective (C-Apochromat, 40x, 1.2 NA; Zeiss). 5. Select a measuring point in a scanned image of the cell by setting xy-axis. 6. Set the intensity of the excitation lasers to 0.1% for both 488 and 543 nm. 7. Set bleach time to 0 s, measuring time to 15 s, and repeat count to 3. 8. Measure fluorescent intensity for each point (see Note 4 and Fig. 13.2).

Fig. 13.1. Fluorescent images of HeLa cells. EGFP and tandem mRFP dimer were coexpressed in (a) and (b), and EGFP fused to tandem mRFP dimer was expressed in (c) and (d). (a) and (c) and (b) and (d) show images in the green and red channel, respectively. Scale bar, 50 µm. Circles indicate cells suitable for FCCS analyses. The ratio of fluorescence intensity between the green and red channels varied depending on expression levels of EGFP and tandem mRFP dimer when they were co-expressed. The ratio observed when the fused protein was expressed corresponds to that obtained when the two proteins were expressed 1 to 1. Therefore, we should select cells that show a ratio of the fluorescent intensity similar to that observed when the fused proteins are expressed.

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Fig. 13.2. Examples of fluorescence correlation spectroscopy. In each panel, fluorescence intensity and its auto- and cross-correlation functions are shown in the upper and lower graphs, respectively. Grey lines show measurements in the green channel, and black lines show those in the red channel; dots represent a cross-correlation function between the two channels. In (a) to (c), EGFP and tandem mRFP dimer were co-expressed in HeLa cells; EGFP fused to tandem mRFP dimer was expressed in (d). Consequently, the minimum and maximum cross-correlation was observed in (a) and (d), respectively. Auto-correlation functions without convergence on 1 at longer time points were observed in (b) and (c). Fluorescence intensity varied with periods longer than 1 s in (b) because of movement of cells; photobleaching gradually decreased fluorescence intensity in (c). These correlation data without convergence on 1 should be discarded.

3.5. Curve Fitting

1. Fix structural parameter at 5 (see Note 5), component at 2, and triplet fraction at 0% for auto-correlation of G and R and cross-correlation between them. 2. Set fit limits between 10 to 100,000 µs (see Note 6).

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3. Analyze data with the ConfoCor 2 software (Zeiss). 4. Record the number of molecules for auto- and cross-correlation curve. 5. Calculate the relative cross amplitude, (Gc(0) − 1)/(Gr(0) − 1), which corresponds to a ratio of the associated molecules to the sum of the associated and the monomeric molecules (Nc/Ng) (see Note 7, Fig. 13.3, and Table 13.1). 3.6. Application of FCCS to Plant Cells

FCS is generally difficult in plant cells because of high levels of auto-fluorescence from pigments such as chlorophylls and carotenoids. So far, two FCS experiments have been done in plant cells, both using the tip region of growing root hair or moss protonemal cells, where chloroplasts are absent. Binding of fluorophore-labeled Nod factor (lipochito-oligosaccharide) to the cell wall was detected by FCS in root hair cells (1). It was also shown that a fraction of phytochrome at the cell periphery had a smaller diffusion coefficient, suggesting its binding to plasma membrane in moss protonemal tip cells (11). In preliminary FCCS experiments with moss protonemal cells, onion epidermal cells, and cultured tobacco BY2 cells, we found that it was important to use stronger laser excitation and to avoid cellular compartments with protoplasmic streaming. Figure 13.4 shows the minimum cross-correlation function between EGFP and mCherry co-expressed in nuclei of onion

Fig. 13.3. Effects of cross-talk on fluorescence cross-correlation spectroscopy. Only EGFP (a) or tandem mRFP dimer (b) was expressed in HeLa cells. For more details, see the legend to Fig. 13.2.

0.59 ± 0.07g

1.68 ± 0.19

0.24 ± 0.02

-

EGFP fused tR2

EGFP

tR2

1.16 ± 0.09

0.39 ± 0.03

Red Red

1.94 ± 0.09

1.89 ± 0.31

Red Green

1.95 ± 0.40

1.47 ± 0.12

Red Green

2.14 ± 0.49

Count per molecule (kHz)

Green

Channel

401 ± 74

411 ± 91

311 ± 120

366 ± 86

271 ± 6

221 ± 124

288 ± 13

28.9 ± 9.7

22.0 ± 5.5

18.1 ± 7.5

22.9 ± 6.5

18.2 ± 2.4

45.4 ± 20.0

17.8 ± 2.6

Diffusion time Diffusion coeffi(µs) cientc (µm2/s)

71.6 ± 25.5

74.2 ± 16.3

49.1 ± 0.2

84.0 ± 10.7

95.2 ± 4.3

64.1 ± 24.0

58.6 ± 33.1

Fractiond(%)

Presented values are mean and SD of three measurements in different cells. a Ratio of mean intensity in the red channel to that in the green channel. b Relative cross amplitude, which is the ratio of the associated molecules to the sum of the associated and the monomeric molecules. This corresponds to (Gc(0) − 1)/(Gr(0) − 1) = Nc/Ng (8). c Calculated from the published diffusion constant of Rh6G (DRh6G = 280 µm2/s) (15) and measured diffusion times of Rh6G (tRh6G) and fluorescent proteins (tfp), as follows (9): Dfp = DRh6G × tRh6G/tfp. d Fraction of the fast-diffusing component when auto-correlation function is analyzed with a two-component model (9). e Tandem dimer of mRFP. f Corresponds to the minimum cross-correlation.gCorresponds to the maximum cross-correlation.

-

1.22 ± 0.31

0.22 ± 0.04f

RCAb

Co-expression of EGFP and tR2e 1.52 ± 0.52

Sample

Signal ratioa (Red/Green)

Table 13.1 Examples for data analysis of FCCS

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Fig. 13.4. Fluorescence cross-correlation spectroscopy in nuclei of onion epidermal cells. EGFP and mCherry were co-expressed transiently in the epidermal cells by the use of particle bombardment. FCCS measurement was done at the same position of a nucleus with 0.1–10% intensities of both excitation lasers for 488 and 543 nm. Auto-correlation functions of the green and red channels are shown in (a) and (b), respectively, and cross-correlation functions are shown in (c); fluorescence intensity of the green (upper panel) and red (lower panel) channels are shown in (d). Auto- and crosscorrelation functions under 0.1 and 5% intensity of excitation lasers are shown in (e) and (f), respectively. In (a) to (d), lines are drawn in grey scale according to intensities of excitation lasers. For (e) and (f), see the legend to Fig. 13.2.

epidermal cells under the control of the 35S promoter. Noisier correlation functions were obtained in the standard measurement condition used for FCCS in HeLa cells (Fig. 13.4e). Under these conditions, fluorescence intensities of EGFP and mCherry were 0.5 and 0.4 kHz in CPM, respectively, which were much lower than those in HeLa cells (Table 13.1). Increasing the intensity of excitation lasers to 10% increased the signal to noise ratio (S/N) of auto-correlation of EGFP, but this did not change the amplitudes of each correlation function (Fig. 13.4a). Similar results were observed for auto-correlations of mCherry (Fig. 13.4b), except that the auto-correlation did not converge on 1 at 10% intensity of excitation laser because of photobleaching of mCherry (Fig. 13.4d, lower panel). Photobleaching of mCherry also prevented the cross-correlation function from converging on 1 (Fig. 13.4c). Thus, the optimal intensity of each excitation laser was 5% in this experimental system (Fig. 13.4f) with a relative cross amplitude of 0.33. In general, results with a sufficient S/N are observed

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Fig. 13.5. Free diffusion-independent fluctuation of fluorescence in the cytoplasm of plant cells. EGFP was transiently expressed in HeLa (upper panel), onion epidermal (middle panel), and tobacco BY2 (lower panel) cells, and its fluorescence was measured twice at the same position. Black and grey lines show the first and the second measurement, respectively.

when the fluorescence intensity of each fluorescence protein is more than 1.5 kHz in CPM and photobleaching does not occur. The requirement of a higher intensity of excitation lasers seems to be due to light-reflecting cuticle layers of the outer surface of plant cells. The fluorescence intensity changes drastically in the cytoplasm of onion epidermal and tobacco BY2 cells with periods longer than 1 s (Fig. 13.5). These cells seem to be in a state of either high or low fluorescence intensity. Some intracellular granules or extra-large molecules may slowly move across the confocal volume, so that they may block the light path of the excitation laser or the fluorescence from target molecules. Thus, the cell nucleus may be the best cellular compartment for FCS measurements. Treatments with inhibitors of protoplasmic streaming such as actinomycin D might also help FCS experiments.

4. Notes 1. Because FCCS is based on the counting of single photons at the single molecule level, the brightness of FP affects the success of measurement. Tandem dimer of mRFP compensates for weak fluorescence of mRFP (12). mCherry is one

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of the mFruits, the second-generation mRFPs that have improved brightness and photostability compared with the first-generation mRFP1 (13). On the other hand, mKeima facilitates single-laser FCCS (14), overcoming difficulties in aligning two laser lines and emission crossover between two FPs, described in Note 7. 2. The optimal pinhole diameter is primarily dependent on the numerical aperture (NA) number of an objective lens and the wavelength of each excitation laser used in FCCS. In our system using a water immersion objective (C-Apochromat, 40x, 1.2 NA; Zeiss), the theoretical pinhole size is 78 nm for R (wavelength of the excitation laser is 543 nm) and 70 nm for G (wavelength of the excitation laser is 488 nm), which correspond to one Airy unit. We measured the maximum cross-correlation with various pinhole sizes of G, using a tandem dimer of green (EGFP) and red (tandem mRFP dimer) FPs expressed from a positive control plasmid. We found the highest cross-correlation when the pinhole size of G was 40 nm, narrowing the detection volume for G. 3. HeLa cells appropriate for FCCS are shown by circles in Fig. 13.1. They should show a rather low expression level of FPs. In our system, the optimal intensity of fluorescence is 100–500 kHz in count rate. When HeLa cells are co-transfected (Fig. 13.1a and Fig. 13.1b), select cells in which the intensity of red fluorescence is 1.5–2 times higher than the intensity of green fluorescence, because cells expressing EGFP fused to a tandem mRFP dimer emit the two fluorescent lights with this ratio of intensity (Fig. 13.1c and Fig. 13.1d, Table 13.1). 4. Auto-correlation functions of G and R must have significant amplitudes at near 0 s and converge on 1 at a longer time point as a result of the Brownian motion of the target molecules (Fig. 13.2a and Fig. 13.2d). Fluctuation of fluorescence intensity with periods longer than 1 s (Fig. 13.2b, upper panel) does not depend on the free diffusion of the target molecules, but is brought about by the movement of HeLa cells or vibration of a microscopic equipment during measurement. This causes odd correlation data without convergence on 1 (Fig. 13.2b, lower panel). Gradual decrease in fluorescence intensity by photobleaching of FPs also causes odd results (Fig. 13.2c). Photobleaching often occurs when the intensity of red fluorescence is higher than 500 kHz. If odd data are obtained in more than two of the three repeated measurements, change the measuring points or select another cell. If two measurements are successful, omit the other odd data.

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5. The structural parameter (also called the axial ratio) is a ratio of the radius to half height of the detection volume, which, for simplification, is thought to be a cylinder. It can be determined for each G and R by measuring Rho6G (see Sect. 3.3) of which the diffusion coefficient is known in solution (15). Thus, the diffusion coefficient of a protein can be calculated from the diffusion time measured by FCCS using Rho6G as a reference. However, the detection volume of cross-correlation, which is overlapping two detection volumes of G and R, cannot be determined. Thus, the structural parameter must be fixed simply to 5 for both auto- and cross-correlation. 6. Fluorescence fluctuates also due to transition of a fluorophore from a singlet excited state to a triplet state, which affects the correlation function. This fluctuation occurs within a few µs, which is shorter than the diffusion time of measured molecules through the detection volume. Thus, curve fitting should be done in a time period longer than 10 µs. 7. In our measurements, for example, the minimum and maximum values of relative cross amplitude were 0.2–0.3 and 0.6–0.75, respectively (Table 13.1, 8). The minimum value of relative cross-amplitude above 0 is mainly due to cross-talk between two channels, as observed in Fig. 13.3a. Because EGFP has an emission spectrum longer than 610 nm, the fluorescence of EGFP is also detected in R with a ratio of the red fluorescence to the green fluorescence being 0.25, even when HeLa cells express only EGFP (Fig. 13.3a, upper panel, and Table 13.1). Because both signals come from EGFP, cross-correlation showed the maximum amplitude (Fig. 13.3a). On the other hand, when HeLa cells express only mRFP, no signals are detected in G, resulting in no amplitude of auto-correlation in G and cross-correlation (Fig. 13.3b). Therefore, to minimize the effects of crosstalk on the estimation of protein–protein interaction, the signal intensity of R should be kept higher than that of G.

Acknowledgements This work was supported in part by Grants-in-Aid for Scientific Research from the Japan Society for the Promotion of Science to H.M. (17770026) and M.K. (15370062) and from the Ministry of Education, Culture, Sports, Science, and Technology to M.K. (17050001) and K.T.Y. (14036201), 19060008.

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References 1. Goedhart, J., Hink, M. A., Visser, A. J. W. G., Bisseling, T. and Gadella, T. W. J., Jr. (2000) In vivo fluorescence correlation microscopy (FCM) reveals accumulation and immobilization of Nod factors in root hair cell walls. Plant J. 21, 109–119. 2. Konno, H., Murakami-Fuse, T., Fujii, F., Koyama, F., Ueoka-Nakanishi, H., Pack, G. -G., Kinjo, M. and Hisabori, T. (2006) The regulator of the F1 motor: inhibition of rotation of cyanobacterial F1-ATPase by the e subunit. EMBO J. 25, 4596–4604. 3. Piehler, J. (2005) New methodologies for measuring protein interactions in vivo and in vitro. Curr. Opin. Struc. Biol. 15, 4–14. 4. Magde, D., Elson, E. and Webb, W. (1972) Thermodynamic fluctuations in a reacting system -- Measurement by fluorescence correlation spectroscopy. Phys. Rev. Lett. 29, 705–715. 5. Rigler, R. (1995) Fluorescence correlations, single molecule detection and large number screening. Applications in biotechnology. J. Biotechnol. 41, 177–186. 6. Kinjo, M. and Rigler, R. (1995) Ultrasensitive hybridization analysis using fluorescence correlation spectroscopy. Nucl. Acids Res. 23, 1795–1799. 7. Takagi, T., Kii, H. and Kinjo, M. (2004) DNA measurements by using fluorescence correlation spectroscopy and two-color fluorescence cross-correlation spectroscopy. Curr. Pharm. Biotechnol. 5, 199–204. 8. Muto, H., Nagao, I., Demura, T., Fukuda, H., Kinjo, M. and Yamamoto, K. T. (2006) Fluorescence cross-correlation analyses of molecular interaction between Aux/IAA protein and protein-protein interaction domain of auxin response factors of Ara-

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15.

bidopsis expressed in HeLa cells. Plant Cell Physiol. 47, 1095–1101. Pack, C., Saito, K., Tamura, M. and Kinjo, M. (2006) Micro-environment and effect of energy depletion in the nucleus analyzed by mobility of multiple oligomeric EGFPs. Biophys. J. 91, 3921–3936. Bacia, K., Majoul, I. V. and Schwille, P. (2002) Probing the endocytic pathway in live cells using dual-color fluorescence cross-correlation analysis. Biophys. J. 83, 1184–1193. Böse, G., Schwille, P. and Lamparter, T. (2004) The mobility of phytochrome within protonemal tip cells of the moss Ceratodon purpureus, monitored by fluorescence correlation spectroscopy. Biophys. J. 87, 2013–2021. Saito, K., Wada, I., Tamura, M. and Kinjo, M. (2004) Direct detection of caspase-3 activation in single live cells by cross-correlation analysis. Biochem. Biophys. Res. Commun. 324, 849–854. Shu, X., Shaner, N. C., Yarbrough, C. A., Tsien, R. Y. and Remington, S. J. (2006) Novel chromophores and buried charges control color in mFruits. Biochemistry 45, 9639–9647. Kogure, T., Karasawa, S., Araki, T., Saito, K., Kinjo, M. and Miyawaki, A. (2006) A fluorescent variant of a protein from the stony coral Montipora facilitates dual-color single-laser fluorescence cross-correlation spectroscopy. Nature Biotechnol. 24, 577– 581. Rigler, R., Mets, Ü., Widengren, J. and Kask, P. (1993) Fluorescence correlation spectroscopy with high count rate and low background: analysis of translational diffusion. Eur. Biophys. J. 22, 169–175.

Chapter 14 The Determination of Protein–Protein Interactions by the mating-based Split-Ubiquitin System (mbSUS) Christopher Grefen, Petr Obrdlik, and Klaus Harter Abstract Dynamic and reversible protein–protein interactions have a pivotal function in all living cells. For instance, protein–protein interactions are involved in the assembly and regulation of multimeric enzymes and transcription factors, various signal response pathways, intracellular sorting and movement of proteins and membrane vesicles, cell-to-cell protein transport, and many others. Here we provide a detailed protocol for the mating-based split-ubiquitin system (mbSUS), which is a sensitive and user-friendly alternative to the classical yeast two-hybrid system in particular. mbSUS relies on the ubiquitin-degradation pathway as a sensor for protein–protein interactions. Thus, mbSUS is predominantly suitable for the determination of full-length proteins localized in the cytoplasm and in or at membrane compartments, without the need for their truncation and nuclear mislocation. In addition, we present a set of Gateway®-compatible mbSUS vectors that allow the rapid generation of constructs for fast and efficient interaction studies. An additional vector is introduced that allows the extension of mbSUS for the analysis of oligomeric protein complex formation and competition assays in vivo. In summary, mbSUS provides an additional versatile tool for protein–protein interaction studies, which is complementary to in planta assays such as BiFC and FRET. Key words: Protein–protein interaction, mbSUS, yeast transformation, bridge assay, mating, Gateway®.

1. Introduction To evaluate the function of a protein one can choose from various genetic, physiological, cellular or biochemical approaches. Throughout the different disciplines of molecular biology one method of investigation is vital for a functional examination – the study of protein–protein interactions. Numerous techniques

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have been developed for identifying putative interaction partners. Amongst them are methods such as co-immunoprecipitation (coIP, 1), yeast-two-hybrid (Y2H, 2), bimolecular fluorescence complementation (BiFC, 3–5) or fluorescence resonance energy transfer (FRET, 6). This protocol describes the mating-based split-ubiquitin system (mbSUS, 7) in yeast as an inexpensive, fast and reliable method for protein–protein interaction studies and recommends it as an alternative to the methods previously mentioned. Its advantages become obvious when the system’s features and possibilities are compared to those of other methods. Splitting the ubiquitin into two halves, and mutating the N-terminal half to avoid spontaneous reassembling (NubG, NubA), yields a sensitive detection system for protein–protein interaction (see Fig. 14.1; 8). The fusion of a bait protein to the C-terminal half of the ubiquitin (Cub) co-expressed with a prey fusion at the N-terminal half (Nub) enables the tracking of such an interaction.

Fig. 14.1. Schematic representation of the split-ubiquitin system. (a) N-terminal wild-type ubiquitin (NubWt) spontaneously re-assembles with C-terminal ubiquitin (Cub) and therefore the transcription reporter complex PLV (ProteinAlexA-VP16) is released through cleavage by ubiquitin-specific proteases (USPs). To apply this system as a tool to study protein–protein interactions, the capacity for spontaneous re-assembly of Cub and NubWt has to be reduced or enabled. (b) Mutating the isoleucine at position 13 of Nub to alanine (NubA) or glycine (NubG) decreases or even inhibits the interaction between Nub and Cub. (c) Fusion of two (X, Y) proteins, which interact with one another, to Nub and Cub respectively, enables re-assembling to a functional ubiquitin and hence switching on the reporter gene activity that can be monitored with several methods.

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If bait and prey interact Cub and Nub are brought into proximity and hence recognized as a functional ubiquitin leading to the release of a transcriptional activator (lexA), which is fused to the Cub and regulates the transcript activation of different reporter genes (ADE2, HIS3, lacZ). Due to the use of the ubiquitin-degradation pathway as an interaction sensor, the localization of the fusion proteins, which may be cytosolic or membrane-bound, facilitates the testing of proteins that are neither mislocalized nor truncated, as they would be in the Y2H system where nuclear localization is obligatory. Furthermore, the uncomplicated selection of positive interaction pairs and the fast subcloning of those identified via screening are more easily possible in yeast than in

Z

A

ZZ X

Y

X

Y USPs

NubG

B

Cub

PLV

NubG

Cub

Z

Y

PLV

Z X

Y

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USPs NubG

Cub

PLV

C

NubG

X

Cub

Y USPs

Z NubG

X

PLV

Cub

PLV

Y NubG

Cub

PLV

Z

Y Cub

PLV

Fig. 14.2. Schematic representation of the SUS “Bridge Assay”. Trimeric protein complexes can be examined when a third protein is constitutively expressed together with the bait and prey fusions of the split-ubiquitin system. Three modes of interaction are possible. (a) “Enhancing”: Bait (Y) and prey (X) fusion proteins already interact but coexpression of a third protein (Z) stabilizes the interaction. In such cases the interaction should be measured with an oNPG-assay. (b) “Facilitating”: The classical “Bridge Assay” can be performed if the bait and prey protein do not interact with one another, but expression of a third protein leads to the formation of a trimeric complex. (c) “Repressing (competing)”: A third possibility is that the bait and prey proteins interact with one another and a third protein is expressed that is able to interact with either the bait or the prey, reducing the amount of complex formation. This can be measured using quantitative assays.

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other eukaryotic systems (e.g., plant cells). The yeast strains THY. AP4 and THY.AP5 have another advantage that is expedient for screening large quantities of proteins: Being haploid and of different mating types, the Y-Cub fusion proteins are transformed in THY.AP4 and the X-Nub in THY.AP5. After transformation has been verified, these yeast clones can be mated rapidly and efficiently, enabling the researcher to screen a large number of putative interaction pairs. Besides the advantages of this system, we present here for the first time another feature that can be exploited: a Gateway®compatible yeast vector that enables the expression of a third unmodified protein besides the prey and bait fusion. This enables the investigation of putative oligomers through a so-called “Bridge-Assay” (see Fig. 14.2). Hereby the constitutively expressed protein can facilitate, enhance, repress or compete for the interaction of the bait and prey proteins.

2. Materials 2.1. Vectors and Strains

1. Vectors for transformation of Escherichia coli (creating Entryclones see Note 1):

Plasmid name

Origin Selection

Function

pDONR201

pUC

Kan/Cm

Cloning via BP-reaction/PCR-product with attB1/B2-sites

pDONR207

pUC

Gen/Cm

Cloning via BP-reaction/PCR-product with attB1/B2-sites

pENTR/D-TOPO

pUC

Kan

Cloning via TOPO-isomerase

2. Destination vectors for transformation of yeast (following LR-reaction; see Fig. 14.3):

Origin

Selection

Plasmid name

Promoter

E. coli

Yeast

E. coli

Yeast

Function

pMetYC-Dest

met25

pUC

ARS1/ CEN4

Amp

LEU2

Fusion protein with C-terminal Cub-PLV

TRP1

Fusion protein with N-terminal NubG

pNX32-Dest

ADH1

pUC

2µ

Cm Amp Cm

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Plasmid name

Promoter

E. coli

Yeast

E. coli

Yeast

Function

pXN22-Dest

ADH1

pUC

2µ

Amp Cm

TRP1

Fusion protein with C-terminal NubG

pXNubA22Dest

ADH1

pUC

2µ

Amp Cm

TRP1

Fusion protein with N-terminal NubA

pNubWt-Xgate ADH1

pUC

2µ

Amp

TRP1

NubWt protein not Gateway®-compatible

pVTU-Dest

pUC

2µ

Amp Cm

URA3

Protein expression in yeast

ADH1

XbaI (373)

A

attR1

C

EcoRI (833)

attR1

met25

C pU

pUC

Cm R

EcoRI (840)

R

B ccd

Am pR

Cm

Am pR

PvuII (740)

ADH

pVTU-Dest

Cub-PL V

pMetYC-Dest

ccd B

attR2

10186 bp

attR2

8587 bp

PvuII (2116) HindIII (2120)

U2 LE

2µ

EcoRV (3037)

EcoRV (6742)

EcoRI (3278)

UR A3

ARS1 / CEN4

EcoRV (5097) PvuI (4037)

Cub

B

EcoRV (8150)

ProteinA

lexA

VP16

XhoI(358)

PvuI (8012)

ADH

f1

pNX32-Dest

NubG attR1

CmR

ccdB

attR2 3HA

pXN22-Dest

attR1

CmR

ccdB

attR2 NubG 3HA

pXNubA22-Dest

attR1

CmR

ccdB

attR2 NubA 3HA

XbaI (6822)

TR P1

EcoRV (7023)

8496 bp

XbaI (5833)

2µ

C pU AmpR PvuI (4306)

Fig. 14.3. Maps of Gateway®-compatible mbSUS vectors. (a) pMetYC-Dest enables the expression of the bait-Cub-PLV fusion protein under the control of the met25 promoter. (b) Scheme of the different Nub-vectors that enable the expression of N- or C-terminal Nub prey fusion proteins. (c) pVTU-Dest allows the expression of a tagless protein under the control of the adh promoter.

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3. E. coli and yeast strains used (see Note 2): Name

Organism

Genotype

Function

ccdBsurvival E. coli

F−mcrA ∆(mrr-hsdRMS-mcrBC) φ80lacZ∆M15 ∆lacX74 recA1 ara∆139 ∆(araleu)7697 galU galK rpsL (StrR) endA1 nupG tonA::Ptrc–ccdA

Used for amplification of donor and destination vectors

TOP10

E. coli

F−mcrA ∆(mrr-hsdRMS-mcrBC) ϕ80lacZ∆M15 ∆lacX74 recA1 araD139 ∆(araleu) 7697 galU galK rpsL (StrR) endA1 nupG

Used for amplification of entry and destination clones

THY.AP4

S. cerevisae MATa; ade2−, his3−, leu2−, trp1−, ura3−; lexA::ADE2, lexA::HIS3, lexA::lacZ

Used for transformation of Cub-clones; can be used for the “Bridge-Assay”

THY.AP5

S. cerevisae MATα

used for transformation of Nub-clones

ade2−, his3−, leu2−, trp1−

2.2. Yeast Transformation and Mating

1. YPD media: 2% peptone, 2% glucose, 1% yeast extract, 2% agar (pH = 6.0) 2. Sterile de-ionized water. 3. 1 M lithium acetate [...] (LiAc): dissolve lithium acetate in de-ionized water. Adjust the pH to 7.5 with acidic acid, and sterilize by filtration. 4. 50% PEG 3350: dissolve PEG 3350 in de-ionized water to a final concentration of 50% (w/v), sterilize by filtration and avoid water loss during storage as this significantly decreases the transformation efficiency. 5. ssDNA: dissolve 10 mg/ml ssDNA in de-ionized water, sonicate and/or boil for 5 min following cooling on ice before use. 6. SC-ade−, his−, trp−, leu−, ura−, met−-dropout mix: grind 2.0 g of alanine, arginine, asparagine, aspartic acid, cysteine, glutamine, glutamic acid, glycine, inositol, isoleucine, lysine, phenyl alanine, proline, serine, threonine, tyrosine, valine and 0.2 g of p-aminobenzoic acid; store dry and dark at 4°C. 7. Chemicals for auxotrophy selection, each dissolved in 100 ml of water and sterilized by filtration; store in darkness at 4°C: ADE: 0.2 g of adenine sulphate (add 10 ml per litre media) URA: 0.2 g of uracile (add 10 ml per litre media) LEU: 1.0 g of L-leucine (add 10 ml per litre media) TRP: 1.0 g of L-tryptophane (add 2 ml per litre media)

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HIS: 1.0 g of L-histidine (add 2 ml per litre media) MET: 1.5 g of L-methionine (equals a 0.1 M stock solution, add appropriate amount to obtain 75, 150 and 300 µM final concentrations) 8. SC-minimal media (1 l): 1.7 g yeast nitrogen base (without amino acids), 5 g ammonium sulphate, 20 g glucose, 1.5 g of SC-dropout mix (see Sect. 2.2.6); adjust pH to 6.0– 6.3 with KOH; add 20 g agar if solid media is needed; add appropriate auxotrophy selection chemical (see Sect. 2.2.7) e.g., ADE, HIS and LEU for transformation of Nub-clones in THY.AP5. 2.3. Detection Assays

1. SC-minimal media plates and SC-plates containing adenine and histidine (add methionine if needed). 2. Z-buffer: 60 mM Na2HPO4 2H2O, 40 mM NaH2PO4X H2O, 10 mM KCl, 1 mM MgSO4 7H2O; adjust pH to 7.0 (NaOH) and autoclave. 3. 0.1% SDS. 4. Chloroform. 5. ortho-Nitrophenylgalactopyranoside (oNPG) solution: 1 mg oNPG per ml Z-buffer. 6. 1 M Na2CO3.

2.4. Western Blot Analysis

1. “Lyse & Load” (LL-) buffer: 50 mM Tris-HCl (pH 6.8), 4% SDS, 8 M urea, 30% glycerol, 0.1 M DTT, 0.005% bromophenol blue; store at −20°C. 2. Acid-washed glass beads (0.25–0.5 mm). 3. 20 ml SDS-PAGE-resolving-gel (10%): 7.9 ml H2O, 6.7 ml acrylamide mix (30%), 5.0 ml 1.5 M Tris (pH 8.8), 0.2 ml SDS (10%), 0.2 ml (NH4)2S2O8 (10%), 0.008 ml TEMED. 4. 5 ml SDS-PAGE-stacking-gel (5%): 3.4 ml H2O, 0.83 ml acrylamide mix (30%), 0.63 ml 1 M Tris (pH 6.8), 0.05 ml SDS (10%), 0.05 ml (NH4)2S2O8 (10%), 0.005 ml TEMED. 5. 10 X running buffer (1 l): 30 g Tris, 144 g glycine, 15 g SDS. 6. 100% methanol. 7. PVDF membrane (e.g., Immobilon-P, Millipore). 8. Transfer-buffer (1 l): 14.4 g glycine, 3.03 g Tris, 200 ml methanol. 9. 10 X PBS: 1.4 M NaCl, 27 mM KCl, 100 mM Na2HPO4, 18 mM KH2PO4; adjust pH to 7.3 (KOH). 10. Washing buffer: 1 X PBS-Tween: 100 ml 10 X PBS, 900 ml H2O, 0,1% Tween-20. 11. Blocking buffer: 1 X PBS-Tween, 5% milk powder.

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12. Antibodies: (dilute 1:1000 in 1 X PBS-Tween, add 0.1% NaN3). i.

Primary: VP16 (rabbit polyclonal; Abcam), HA (rat monoclonal; Roche)

ii.

Secondary: anti-rabbit IgG alkaline phosphatase, antirat IgG alkaline phosphatase (Sigma)

13. Staining buffer: 100 mM Tris-HCl (pH 9.5), 100 mM NaCl, 5 mM MgCl2. 14. NBT-solution: 50 mg/ml nitro blue tetrazolium chloride in 70% DMF; aliquot and store at −20°C. 15. BCIP solution: 50 mg/ml 5-bromo-4-chloro-3-indoylphosphate-p-toluidin in 100% DMF; aliquot and store at −20°C. 16. Staining solution: 66 µl NBT-solution, 33 µl BCIP-solution in 10 ml staining buffer; always prepare freshly.

3. Methods 3.1. Yeast Transformations 3.1.1. mbSUS Transformation

Before starting your protein–protein interaction study using this protocol, we highly recommend that you first consult and consider the Notes section. Notes 3, 4 and 7 are relevant for choosing interaction couples. 1. Inoculate THY.AP4 and AP5 in 5 ml YPD each and incubate, shaking overnight at 28-30 °C. 2. After approx. 14 h transfer the pre-culture in fresh 95 ml YPD and incubate shaking for 3–5 h until OD600 0.6–0.8. 3. Harvest cells by centrifugation (10 min at 2000 g), and discard supernatant. 4. Wash with 20 ml of sterile water and centrifuge again; discard supernatant. 5. Resuspend the cells with 1 ml of 0.1 M LiAc and transfer to a 2-ml Eppendorf tube, spin down (5 min at 1000 g) and discard supernatant. 6. Add appropriate amount of 0.1 M LiAc (multiply number of transformation with 20 µl) and incubate at room temperature for 30 min (hence called “competent yeast”). 7. Meanwhile prepare sterile PCR tubes, stripes or plates with 10 µl of ssDNA and 5 µl of plasmid DNA (approx. 200 ng/ µl) for each transformation. 8. Prepare mastermix: mix 70 µl of 50% PEG, 10.5 µl of 1 M LiAc, 1.5 µl of ssDNA and 18 µl of competent yeast for each transformation.

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9. Distribute the mastermix, 100 µl for each transformation, carefully mixing with the prepared DNA-mixture. 10. Incubate at 30°C for 30 min using a PCR-cycler. After 20 min mix each reaction using a multi-channel pipette but take great care in not cross-contaminating the tubes. 11. Heat shock cells at 43°C for 15 min. 12. Spin down cells at 1000 g for 5 min, and discard supernatant using a pipette. 13. Wash cells with 100 µl of sterile water, spin down and discard supernatant. 14. Resuspend cells in 100 µl of sterile water. 15. Plate 50 µl on appropriate SC-plate (SC-ADE+, HIS+, TRP+, URA+ for THY.AP4 and SC-ADE+, HIS+, LEU+ for THY. AP5). 16. Incubate for 48–72 h at 28-30 °C. 3.1.2. Bridge Assay Transformation

The transformation protocol is the same as that in Sect. 3.1.1 with the following exceptions: 1. Use haploid THY.AP4-cells exclusively. 2. Prepare DNA-mix with only 9 µl of ssDNA, which leaves 2 µl for each of the three constructs you would like to transform (use more highly concentrated plasmid DNA, if problems occur, e.g., too few colonies; see Note 8). 3. Plate cells on SC-ADE+, HIS+-media. Alternatively, if in all reactions one or even two clones remain the same (e.g., several different bridge proteins should be tested for a certain Cub-Nub-pair), the yeast containing these can be directly used for transformation by making it competent following the protocol above and transforming only the plasmid DNA encoding the bridge proteins. In general, it should be kept in mind that transforming three plasmids at once results in lower transformation efficiency and that it is not possible to use the mating system here, as the THY.AP5 strain contains a wild-type URA3 gene in its genome, which would suspend the necessity for an episomal URA3-gene.

3.2. Testing the Bait Fusion for Background Activity (Optional)

1. Pick approx. five representative single colonies per THY.AP4 transformations (Cub-PLV fusion) and grow overnight in 5 ml SC-ADE+, HIS+, TRP+, URA+ media at 28-30 °C. 2. Pick several colonies of the transformed “empty” pNX32DEST, expressing only NubG in THY.AP5 and pool them in 5 ml (or more depending on the number of bait fusion colonies to be tested) of SC-ADE+, HIS+, LEU+. Grow cells overnight by shaking at 28-30 °C.

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3. Harvest 2 ml of bait fusion and deep-freeze it for later Western Blot analysis. 4 . Harvest 2 ml of bait fusion and resuspend it in 200 µl YPD. 5. Harvest twice 2 ml of the NubG expressing yeast and resuspend it in enough YPD to mate with all bait samples (20 µl per each mating is needed). 6. Prepare a 96-well-plate with 20 µl NubG yeast cells in each well and add 20 µl of each bait fusion, and mix carefully. 7. Immediately drop 4 µl of each mating onto a 12 × 12 cm YPD plate (use only 64 spots per each square plate). 8. Incubate overnight (12–16 h) at 28-30 °C. 9. Use a replicator stamp with a sheet of sterile velvet to transfer cells onto SC-ADE+, HIS+-media (see Fig. 14.4). 10. Grow for 48–72 h at 28-30 °C, and then transfer the cells via replicator stamp onto SC-minimal media. 11. Check for growth after 3–5 days. Growing cells indicate that your bait protein is producing background activity, which will result in false positive results if used in your assay. The

Fig. 14.4. Replica plating of diploid yeast cells. (a) Material that is needed for the fast replica plating of up to 64 yeast colonies: sv = sterilized velvet sheets; rs = replicator stamp, a PVC-block with the measurements 11.5 × 11.5 × 6.0 cm to match the size of commercially available square plates (12 × 12 cm); ypd = YPD plate with the matings from the previous day; sc = SC-ADE+, HIS+-plate on which the matings will be transferred. (b) Stamp with velvet sheet prepared for sterile replica plating of the yeast colonies. (c–e) Execution of the replica plating: Put the YPD plate applying slight pressure on the stamp; check that the colonies are being equally absorbed by the velvet. Keep the orientation of your numbering in mind. Carefully put the pre-dried SC-ADE+, HIS+-plate on the velvet and use slight pressure to transfer the colonies onto the plate. It is useful to always produce two replicas of one velvet sheet (see Color Plates ).

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use of different methionine concentrations could solve the problem; see Note 9). 3.3. Mating

1. Use a bait colony that passed the test of background activity and inoculate 3 ml of SC-ADE+, HIS+, TRP+, URA+ media; grow overnight at 28-30 °C. 2. Pick one representative single colony for each prey fusion and inoculate 5 ml SC-ADE+, HIS+, LEU+. Grow overnight at 28-30 °C. 3. Harvest 2 ml of the THY.AP5 cells and deep-freeze it for later Western Blot analysis. 4. Harvest remaining cells and resuspend them in YPD (for each mating 20 µl of YPD should be calculated). 5. Mix 20 µl of each, bait and prey, in a 96-well-plate for any desired combination. 6. Immediately drop 4 µl of each mating onto a 12 × 12 cm YPD plate (use only 64 spots per each square plate). 7. Incubate overnight (12–16 h) at 28-30 °C. 8. Use a replicator stamp with a sheet of sterile velvet to transfer cells onto SC-ADE+, HIS+-media (see Fig. 14.4). 9. Grow for 48–72 h at 28-°C, then transfer the cells via replicator stamp onto SC-minimal media and SC-ADE+, HIS+ as growth control. 10. Check for growth after 3–7 days. Growing cells indicate a positive reporter gene activity that should result from an interaction of the corresponding bait and prey fusions. Using different methionine concentrations in the SC-minimal media can further increase the signal-to-noise ratio (see Note 9).

3.4. Detection Assays

3.4.1. oNPG-Assay (Quantitative)

To decide which detection method is the best for your screen you should refer to Notes 5 and 6. 1. Incubate for each clone three independent colonies in 5 ml selective media overnight (12–16 h) at 28-30 °C (see Note 6 and 10). 2. Try to collect all yeast cells at the same OD600 (0.8–1.0). 3. Wash cell pellets with 1 ml 4°C-cold Z-buffer. 4. Resuspend pellets in 650 µl Z-buffer. 5. Freeze in liquid nitrogen, thaw at 37°C for 60 s and repeat this freeze-and-thaw-cycle twice. 6. Add 50 µl 0.1% SDS, 50 µl of chloroform and vortex at the highest setting for 1 min. 7. Centrifuge at 10,000 g for 10 min at 4°C; transfer 600 µl of supernatant into a new tube, and keep on ice. 8. Estimate protein concentration via Bradford assay:

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i.

Use 15–30 µl of each yeast extract; fill up to 800 µl with H2O

ii.

Add 200 µl of Bradford solution

iii. Incubate 15 min at room temperature iv.

Measure extinction at 595 nm

v.

Compare with freshly prepared BSA standard

9. For each reaction prepare 800 µl oNPG-solution in an incubator at 37°C. 10. Add 100–200 µl of yeast extract (final volume 1 ml) incubating at 37°C. 11. Monitor the incubation time (2–60 min). 12. Stop the reaction with 0.5 ml 1 M Na2CO3 before the yellow colour is saturated. 13. Measure extinction at 420 nm; use an equally treated oNPGsolution (without extract) as blank. 14. Dilute samples if extinction exceeds OD420 = 0.8–1.0. 15. Calculate specific enzyme activity using the following formula: Specific enzyme activity [U/mg] = (E420 × V)/(ε × d × v × t × P) V = volume of reaction (1500 µl) e = extinction coefficient of o-nitrophenol (4.5 ml/µmol cm) d = thickness of the cuvette (1 cm) v = volume of extract (20–100 µl) t = incubation time [min] P = protein concentration [mg/ml] 3.4.2. Growth Assay (Qualitative)

1. Use the replicator stamp to transfer cells from the SC-ADE+, HIS+-plates onto SC-minimal media (with or without methionine at different concentrations) and a fresh SCADE+, HIS+-plate as growth control (see Fig. 14.4). 2. Incubate plates at 28-30 °C for up to 8 days. 3. Start monitoring the growth (size of yeast drop) on day 3.

3.5. Western Blot

1. Harvest 2 ml of overnight yeast culture (or use deep-frozen aliquots). 2. Add 50 µl acid-washed glass beads; add 150 µl of LLbuffer. 3. Vortex 1 min at the highest setting. 4. Incubate for 10 min at 65°C while shaking. 5. Spin down and transfer supernatant in a fresh tube. 6. Load 10–15 µl on a SDS-PAGE gel; run gel with appropriate conditions. 7. Perform Western blotting of the gel (e.g., 3 mA/cm2 for 90 min).

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8. Block the membrane 30–60 min in blocking buffer at room temperature (or overnight at 4°C). 9. Transfer the membrane into primary antibody solution; incubate at least 1 h at room temperature or overnight at 4°C. 10. Wash three times, 10 min each with 1 X PBS-Tween. 11. Transfer membrane into secondary antibody solution; incubate at least 1 h at room temperature or overnight at 4°C. 12. Wash three times, 10 minutes each with 1 X PBS-Tween. 13. Equilibrate membrane for a short time (15–30 s) in staining buffer. 14. Transfer membrane into staining solution; incubate until clear signals can be monitored.

4. Notes 1. There are two possibilities for obtaining an Entry clone. Either by BP-reaction using a PCR product that has attB1 and attB2-sites attached to the 5′ and 3′-ends respectively, or by TOPO cloning, adding just a CACC-quadruplet to the 5′-end. Depending on the nature of the cDNA (length, structure, possible toxicity, etc.), either way can be more favourable (see corresponding Gateway®-manuals). 2. If you encounter difficulties while cloning in E. coli or observe abnormal errors in the cDNA (point mutations, deletions, transposons), the use of a specialized strain (e.g., CopyCutterTM, Epicentre) is highly recommended. Eukaryotic genes, encoding for membrane proteins in particular, can be toxic for E. coli and cloning may be time-consuming if at all possible. (We observed toxic effects of genes that were amplified as Entry-clones, although termination signals were anchored within the vector sequence to avoid such problems; this leads to the assumption that inconvenient effects can occur on the DNA level). 3. To study the interaction of two proteins by way of mbSUS, verification that the bait protein (Cub-PLV fusion) is neither soluble nor nuclear-localized is recommended. The latter would lead to background activity (false positives), as the lexA transcriptional regulator that is fused to the Cub would activate reporter genes without being cleaved by the ubiquitin-specific proteases (USPs). Additional consideration should be given to putative signal sequences. As the N-terminal end of the bait protein stays unmasked, puta-

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tive signal peptides will be recognized and accordingly processed, thus being displaced from their interaction partners in the cytosol (false negatives). A third point is the topology of the proteins used. The C-terminal end of the bait bearing the Cub-PLV has to be in the cytosol (the same applies to the prey fusions as a matter of course) and should not be in the lumen of ER or other compartments, because the interaction between Cub and Nub can only be assayed in the cytosol in which the USPs can recognize the reconstituted ubiquitin. We therefore recommend using prediction programs like SignalP (http://www.cbs.dtu.dk/services/SignalP/) and TMHMM (http://www.cbs.dtu.dk/services/ TMHMM/) before deciding which bait should be used. (If necessary the bait can be modified, e.g. the signal peptide truncated, to avoid secretion. However, the physiological relevance of an interaction found with such a bait should be carefully considered regarding the in vivo localization of bait and prey. (see Fig. 14.5). 4. The topology of the prey fusion is just as important as that of the bait. However, we designed a set of vectors with either N-terminal or C-terminal NubG. In addition, an N-terminal NubA vector with an increased affinity to Cub compared with NubG (12) was created, as it appears that the C-terminally fused Nub has a decreased affinity towards Cub (see Fig. 14.5). If only a C-terminally fused Nub can be used for the prey fusion, an assay of the interaction with both Nub variants would be highly recommended. 5. The method to detect positive interaction in the mbSUS is by default growth on interaction-selective (minimal) media using the lexA-driven ADE2 and HIS3 reporter genes that complement the corresponding knockout alleles of the yeast strain. As an additional tool to monitor and quantify the interaction, the lexA-driven lacZ gene that encodes for the b-galactosidase was introduced into the yeast strain THY.AP4. The sensitivity of the lacZ reporter is, however, decreased compared with that of the growth assay and therefore only expedient if strong interaction partners are to be compared. An additional indication for interaction is the colour of the diploid cells that lack the phospho-ribosylaminoimidazole carboxylase, which is encoded by the ADE2 gene and catalyzes the de novo synthesis of adenine. If the ADE2 gene is not activated, cells accumulate a red pigment, whereas the interacting cells become white (see Fig. 14.5). However, this method should only be used as a preliminary tool to estimate the chance of a positive/negative interaction. 6. If you encounter difficulties with the oNPG protocol that is provided in this manuscript, we recommend scaling up

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Fig. 14.5. Model mbSUS analysis of the Arabidopsis ethylene receptor ETR1. (a) Schematic representation of the proposed ER membrane-localized native ETR1 homodimer (far left; 10, 11) and its mbSUS derivates (right). (b) Growth of mated yeast clones expressing the indicated fusion proteins. The homodimerization of the ethylene receptor is observed only when ETR1-Cub is co-expressed with the truncated ∆ TM-ETR1-NubG version. Probably due to steric hindrance, the formation of ETR1 homodimers was not detected when ETR1-Cub was co-expressed with ETR1-NubG, although both fusion proteins accumulate in yeast (see C). As positive control the expression of NubWt (7) was used; as negative control NubG. Note that the different phenotypes of the non-interacting yeast clones can be seen on the SC-ADE+, HIS+-media (red/white-selection). (c) Western blot analysis of the yeast clones shown in (b). The data reveal that the ethylene receptor proteins are unstable in yeast when NubG is fused to the N-terminus of the full-length (NubG-ETR1) or the truncated ethylene receptor (NubG-∆ TM-ETR1). The ETR1-Cub and NubG fusions were detected with an antibody against the VP16 domain within PLV or the HA-tag, respectively (NubWt and NubG-proteins do not contain a C-terminal HA-Tag). ∆ TM-ETR1: ETR1 without the transmembrane domains (aa 1 to 120); H: histidine kinase domain; D: receiver domain; black diamond = GAF-domain (data published in Ref. 9). (see Color Plates )

the amount of yeast (e.g., using an overnight culture of 50 ml). We found that certain interaction couples give such a weak signal that the detection of an interaction in the growth assay is possible but not in the oNPG-Assay. However, the use of a more concentrated yeast-protein extract facilitated the detection of galactosidase activity with differences comparable with those seen in the growth assay.

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7. The whole set of positive (NubWt) and negative controls should always be included in the screen. Not only an “empty” NubG as negative prey but also the “empty” pMetYC-Dest, which does not express a functional Cub should be mated with each Nub-prey fusion to exclude possible cross-contamination when working with large-scale approaches. 8. Our plasmids are purified via 7% PEG 8000 to get rid of contaminating nucleotides to measure the DNA concentration accurately by a spectrophotometer. We observed that low amounts of plasmid DNA (<3–500 ng) gave significantly less colonies in comparison with transformations with higher applications of DNA (>1 µg per plasmid). We recommend concentrating the plasmid DNA by either lyophilization or PEG precipitation. 9. The vector pMetYC-DEST contains a methionine-repressible promoter. Although the yeast maintains only one or two copies of the plasmid (due to the ARS/CEN origin), the expression of the fusion protein containing the PLV transcription-factor complex should be tightly regulated. This can be achieved by the application of exogenous methionine and should be done if you observe one of the two following cases: first, if your bait clone produces slight background activity with the negative control (e.g., the NubG control grows on interaction selective media; see Sect. 3.2) and second, if the signal-to-noise ratio should be further increased. 10. Pooling colonies of the same clone can increase the possibility of a positive interaction; however, we observed that weak interactions of problematic or toxic fusion proteins could not be detected if colonies were pooled. The yeast obviously had difficulty expressing the fusion protein and therefore only a minority of the colonies actually expressed the fusion proteins. While pooling resulted in no detectable growth on selective media after mating, choosing only those that expressed the construct (verified by Western Blot) yielded a clear signal.

Acknowledgements The authors wish to thank Felicity de Courcy, Achim Hahn and Patrick Williamson for critically discussing and reading the manuscript.

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References 1. Phizicky, E.M. and Fields, S. (1995) Protein-protein interactions: methods for detection and analysis. Microbiol. Rev. 59, 94–123. 2. Fields, S. and Song, O. (1989) A novel genetic system to detect protein-protein interactions. Nature 340, 245–246. 3. Hu, C.-D., Chinenov, Y., and Kerppola, T. (2002) Visualization of interactions among bZIP and Rel family proteins in living cells using bimolecular fluorescence complementation. Molecular Cell. 9, 789–798. 4. Bracha-Drori, K., Shichrur, K., Katz, A., Oliva, M., Angelovici, R., Yalovsky, S., and Ohad, N. (2004) Detection of protein–protein interactions in plants using bimolecular fluorescence complementation. Plant J. 40, 419–427. 5. Walter, M., Chaban, C., Schütze, K., Batistic, O., Weckermann, K., Näke, C., Blazevic, D., Grefen, C., Schumacher, K., Oecking, C., Harter, K., and Kudla, J. (2004) Visualization of protein interactions in living plant cells using bimolecular fluorescence complementation. Plant J. 40, 428–438. 6. Tsien, R.Y., Bacskai, B.J., and Adams, S.R. (1993) FRET for studying intracellular signalling. Trends Cell Biol. 3, 242–245. 7. Obrdlik, P., El-Bakkoury, M., Hamacher, T., Cappellaro, C., Vilarino, C., Fleischer, C., Ellerbrok, H., Kamuzinzi, R., Ledent,

8.

9.

10.

11.

12.

V., Blaudez, D., Sanders, D., Revuelta, J.L., Boles, E., Andre, B., and Frommer, W.B. (2004) K + channel interactions detected by a genetic system optimized for systematic studies of membrane protein interactions. Proc. Natl. Acad. Sci. U S A 101, 12242–12247. Johnsson, N. and Varshavsky, A. (1994) Split ubiquitin as a sensor of protein interactions in vivo. Proc. Natl. Acad. Sci. U S A 91, 10340–10344 Grefen, C., Städele, K., Rûžižka, K., Obrdlik, P., Harter, K., and Horak, J. (2008) Subcellular localization and in vivo interaction of the Arabidopsis thaliana ethylene receptor family. Molecular Plant, Mol Plant; 1:308–320 Schaller, G.E., Ladd, A.N., Lanahan, M.B., Spanbauer, J.M., and Bleecker, A.B. (1995) The ethylene response mediator ETR1 from Arabidopsis forms a disulfide-linked dimer. J. Biol. Chem. 270, 12526–12530. Chen, Y.F., Randlett, M.D., Findell, J.L., and Schaller, G.E. (2002) Localization of the ethylene receptor ETR1 to the endoplasmic reticulum of Arabidopsis. J. Biol. Chem. 277, 19861–19866. Raquet, X., Eckert, J.H., Muller, S., and Johnsson, N. (2001) Detection of altered protein conformations in living cells. J. Mol. Biol. 305, 927–938.

Chapter 15 Functional Complementation of Yeast Mutants to Study Plant Signalling Pathways Norbert Mehlmer, Elisabeth Scheikl-Pourkhalil, and Markus Teige Abstract The rapidly increasing amount of entirely sequenced genomes generates a need for fast and efficient methods to elucidate gene functions. Functional complementation of yeast mutants, displaying selectable phenotypes, has been used very successfully in the past years to isolate many plant genes involved in signalling, stress response or metabolic pathways. Using the well-characterized Hog1 pathway, a mitogen activated protein (MAP) kinase pathway required for adaptation to osmotic stress in budding yeast, as example, we describe here the isolation of plant protein kinases involved in abiotic stress adaptation in the model plant Arabidopsis thaliana. The osmo-sensitive phenotype of yeast mutants carrying a mutation in the Hog1 pathway allows an easy selection on high osmolarity media, containing i.e. 0.4 M NaCl. By using yeast mutants harbouring deletions in different components of the pathway, for example the MAP kinase kinase Pbs2 and the MAP kinase Hog1, it is furthermore possible to isolate consecutively acting components of a signalling pathway. Key words: Functional complementation, yeast mutants, plant stress response, plant signalling, regulation of transcription, metabolic adaptation.

1. Introduction With the rapidly increasing amount of entirely sequenced genomes the attribution of cellular functions of annotated genes becomes a major challenge in the post-genomic area. Accordingly, fast and simple methods for functional characterization of genes are more and more required in many fields of molecular biology. Since the pioneering work of Lee and Nurse in 1987 (1), which demonstrated cloning of a human cDNA by direct complementation of a fission yeast mutant, this technique has been very T. Pfannschmidt (ed.), Plant Signal Transduction, vol. 479 Humana Press, a part of Springer Science + Business Media, LLC 2009 DOI: 10.1007/978-1-59745-289-2_15

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successfully used many times for isolation of various genes from a great number of species ranging from worms (2, 3) or flies (3) to plants (4) and humans (1). Even though this technique sounds at a first glance quite “old-fashioned” today, many recent publications demonstrate its value in functional analysis and cloning of genes, particularly in the area of plant signalling or stress response. Some recent findings include genes involved in trehalose or proline metabolism (5) or revealed a new function of late embryogenesis abundant (LEA)-like proteins in oxidative stress tolerance (6,7). In the field of plant signalling, complementation of mutants in the Hog1 mitogen activated protein (MAP) kinase pathway from Saccharomyces cerevisiae, which is required for adaptation to hyperosmotic stress in yeast (8,9), was used to isolate components of a plant MAP kinase pathway required for plant cytokinesis (10) or for an MAP kinase pathway, which regulates adaptation to salt- and cold stress in Arabidopsis (11). As already discussed by Minet et al. (4) for the complementation of yeast auxotrophic mutants, this technique has some limitations, mainly due to the fact that yeast is only a unicellular eukaryote and has a less complex genetic system. However, it should also be noted here that even general principles in the regulation of gene expression such as chromatin remodelling are conserved between yeast and higher eukaryotes, thus enabling functional complementation also in this context of transcriptional regulation in a general stress response (12). The most important prerequisite for this approach is a selectable phenotype of the appropriate yeast mutant, which is suited for the approach. Here we will describe the method using mutants of the Hog1 MAP kinase pathway, which display an osmosensitive phenotype enabling an easy selection on media containing 0.3–0.4 M NaCl. The general strategy of the screening and subsequent further analysis of isolated genes are displayed in Fig. 15.1. The following protocol describes the detailed screening procedure for this screen using the Arabidopsis cDNA library in the yeast expression vector pFL61 from Minet et al. (4), which has been used very successfully in a number of different screens. The library uses the yeast URA3 gene as auxotrophic marker for selection in yeast and ampicillin resistance for bacterial selection. More general considerations in the growth or transformation of yeast cells and yeast two-hybrid methods extending the application of functional complementation for gene isolation, which is described here in all necessary details, were recently summarized by Gietz and Woods (13). Before starting the screen the conditions used for selection should be tested. In the case we discuss here this can easily be done by transforming the yeast mutants (i.e. pbs2D) and the parental wild type with an empty library plasmid (see Note 1) and plating serial dilutions (see Note 2) on plates with increasing

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Fig. 15.1. Overview of the general strategy for isolation of plant stress response genes. Here the isolation of an Arabidopsis MAP kinase kinase by functional complementation of the osmo-sensitive pbs2D mutant on selective plates containing 0.4 M NaCl is shown. The two critical steps of the entire procedure are the library screen using a high-efficiency yeast transformation protocol (1), and the subsequent plasmid rescue procedure (2), leading to the plant cDNA clone, which could be used for DNA sequencing and further functional studies (in vitro and in vivo) of the isolated gene. If a given plant gene should be tested only for a certain function, step (1) can be done using a simplified (fast) transformation protocol and the specific cDNA cloned into a suitable yeast expression vector.

salt concentrations and comparing growth. In an ideal case the mutant carrying the empty plasmid should not grow any more (top row in Fig. 15.1), and the wild-type strain should display normal growth (second row in Fig. 15.1). After determining the selection conditions, the cDNA library is transformed into the mutant strain at a large scale using a highly efficient method for yeast transformation. A total number of at least 300,000 transformants should be reached in the screen (see Note 3). Positive clones restoring growth under selective conditions (see clone 1 to 5 in Fig. 15.1) are used for plasmid rescue from yeast cells,

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enabling further studies of the isolated gene for further functional studies in vitro or in vivo.

2. Materials 2.1. Yeast Media

1. YPD medium: 1% yeast extract (BD/DIFCO), 1% peptone (BD/DIFCO) and 2% glucose (Sigma) (see Note 4) are used for routine growth of yeast strains if a selection for a transformed plasmid is not necessary. For plates 3% Bacto agar (BD/DIFCO) is added. 2. For selection of transformed plasmids a synthetic complete (SC) medium is used. The SC medium is prepared from Difco’s Yeast Nitrogen Base (BD/DIFCO) supplemented with glucose and an amino acid mixture, omitting the amino acids or bases used for selection. The amino acid mixture is prepared as 100 X stock solution (see Note 5) and contains the following: 6 g/l isoleucine, leucine and phenylalanine; 5 g/l threonine; 4 g/l tryptophane and lysine; 2 g/l arginine; 1 g/l histidine and methionine. All amino acids can be obtained from Sigma. The SC selection medium is composed of 6.7 g/l Yeast Nitrogen Base without amino acids, supplemented with 50 mg/l of adenine (Sigma) and tyrosine before autoclaving. After autoclaving and cooling to 60°C, 50 ml/l of an autoclaved 40% (wt/vol) glucose solution and 10 ml/l of sterile filtrated amino acid mixture are added.

2.2. Yeast Transformation

For high-efficiency yeast transformation Mix1 and 2 are prepared from sterile filtrated stock solutions according to Tables 15.1 and 15.2. 1. 60% Polyethyleneglycole 3.350 [PEG] (Sigma). 2. 1 M LiAc (Sigma).

Table 15.1 Trafo Mix1 (prepare 10 ml) Ingredient

Volume

Final conc.

Stock solution

LiAc

1 ml

100 mM

1M

Sorbitol

5 ml

1M

2M

TE

100 µl

10 mM

100 X

H2O

3.9 ml

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Table 15.2 Trafo Mix2 (prepare 15 ml) Ingredient

Volume

Final conc.

Stock solution

LiAc

1,5 ml

100 mM

1M

PEG 3350

10 ml

40%

60%

TE

150 µl

10 mM

100 X

H2O

3.35 ml

3. 2 M sorbitol (Sigma). 4. TE buffer (10 mM Tris-HCl, 1 mM EDTA, pH 8.0). 5. 1 M DTT solution (1.54 g DTT in 10 ml of 10 mM NaAc, pH 5.2). 6. Transformation Mix 1 contains the following: 100 mM LiAc (Sigma), 1 M sorbitol (Fluka), 10 mM Tris-HCl (Applichem), 1 mM EDTA (Applichem), pH 8.0. 7. Transformation Mix 2 contains: 100 mM LiAc, 40% Polyethyleneglycole 3.350 [PEG] (Sigma), 10 mM Tris-HCl, 1 mM EDTA, pH 8.0; bacterial RNA (10 mg/ml) to be used as single stranded carrier (see Note 6). 8. Bacterial RNA is prepared from bacteria using the alkaline lysis method, which is used for plasmid preparations, but omitting the RNase treatment. This procedure results in purification of large amounts of bacterial RNA, which is extracted once with phenol/chloroform, washed, dissolved in TE buffer, and diluted to 10 µg/ml. 2.3. Plasmid Rescue from Yeast Cells

1. Lyticase solution for cell wall digest: Dissolve 4 mg Zymolyase 200T (from A. luteus, Seikagu) in 2 ml Z-buffer (50 mM sodium phosphate, 10 mM KCl, 1 mM MgSO4). DNAse free RNase A (10 mg/ml) according to (14). 2. Buffer P1 (100 mM Tris-HCl, 10 mM EDTA, pH 8). 3. Buffer P2 (0.2 M NaOH, 1% SDS); buffer P3 (3 M KAc, pH 5.5). 4. A DNA purification kit, which is usually used for purification of PCR fragments; we usually use the PCR/Gel elution kit from Promega.

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3. Methods A number of different procedures for transformation of yeast cells have been described in the literature. In recent years the transformation protocol using lithium acetate, PEG, and single stranded carrier DNA, referred to as LiAc/SS Carrier DNA/PEG method, is the most commonly used method for transformation of yeast cells and has been adapted in many ways (13,15). We describe here the method in detail for functional yeast complementation using an osmosensitive yeast strain and the Arabidopsis cDNA library from Minet at al. (4), which is a modification of the LiAc/SS Carrier DNA/PEG method described by Gietz and Woods (13). 3.1. High-Efficiency Yeast Transformation for Library Screening (see Note 7)

1. Grow 30–50 ml yeast culture in appropriate medium (see Note 8) overnight. 2. On the next morning dilute cells to an OD600 of 0.2 and incubate further at 30°C with vigorous shaking until they reach an OD600 of 0.4–0.6 (takes usually 4–6 h). 3. Collect cells in four 50-ml plastic tubes (Falcon tubes) and harvest by centrifugation at 2000× g for 10 min at room temperature (see Note 9). 4. Re-suspend the pellets in 25 ml TE by vortexing, pool and again spin down for 5 min at 2000× g. 5. Re-suspend the cells in 1 ml Mix 1 by pipetting, transfer in a 1.5-ml microcentrifuge tube and spin down for 1 min at 2500× g. 6. Re-suspend cells by pipetting in the same volume of the pellet (about 150 µl, see Note 10) in Mix 1 and incubate for 10 min. 7. Transformation: Mix 250 µl carrier RNA with 40 µg cDNA library (1 µg/µl) and subsequently with 300 µl yeast cells (in Mix 1) by gentle pipetting (see Note 11). 8. Add this cell suspension to 1.75 ml Mix 2 in a 15-ml tube, vortex gently and incubate at 30°C for 30 min. 9. Add 220 µl DMSO and mix gently before applying the heatshock for 15 min at 42°C. 10. Add 10 ml of sterile water and spin down cells at 2000× g for 5 min at RT. 11. Re-suspend cells in 10 ml YPD and incubate for 60 min at 30°C with shaking. 12. Spin down cells at 2000× g for 5 min, re-suspend in an appropriate volume (see Note 12) of SC-ura and plate on selective media. Positive colonies will appear after 3–5 days of incubation at 30°C. Calculate transformation efficiency as described (Note 3).

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If the efficiency was not sufficient so far to generate a total number of 300,000 transformants, the procedure should be repeated. At this point the conditions for selection might also be adjusted less or more stringently; here, for example by simply decreasing or increasing the salt concentration in the selective plates. The next step of the procedure is to isolate (rescue) the library plasmid from the transformed yeast strain. If only one plasmid is present, the procedure can directly proceed with Sect. 3.2. A problem arises if another yeast plasmid has been transformed before carrying out the complementation screen, for instance if an interaction partner of a known gene should be isolated. In this case it has to be verified that only the unknown library plasmid is isolated and not the known input. A pre-selection is usually not possible (see Note 13), but yeast cells lose their plasmids quite rapidly without selective pressure. Thus, the selection is carried out only for the library plasmid (here SC-ura) for 2 days in liquid media before the following plasmid rescue is performed. The transformation of bacteria with isolated plasmid DNA from yeast cells is usually inefficient and a critical step in this procedure and in most yeast two-hybrid screens. Therefore, it is essential to further clean up the isolated plasmid DNA using a purification kit before transformation into E. coli. After this additional cleaning step has been done, the transformation works with chemically competent cells as well as electroporation. 3.2. Rescue of the Library Plasmid

1. Harvest 2 ml yeast cells in a 2-ml microcentrifuge tube by centrifugation at 5000× g for 5 min at room temperature, resuspend cells in 1 ml TE, transfer into a 1.5-ml microcentrifuge tube and again spin down at 2000× g for 5 min at room temperature. 2. Re-suspend the pellet in 100 µl of lyticase solution, supplemented with RNase, and incubate at 37°C for 30 min with shaking (check progress of digestion using a microscope). 3. Mix the cell suspension with 150 µl P2 and incubate for 30 min at RT. 4. Mix the suspension with 150 µl P3 and incubate for 15 min at 4°C. 5. Spin down the suspension at 15,000× g for 15 min at 4°C. 6. Transfer the supernatant into a new 1.5-ml microcentrifuge tube and mix with the same volume of DNA binding solution from the PCR/Gel elution kit. 7. Purify the plasmid DNA according to the protocol from the kit. 8. Elute plasmid DNA from the column with 40 µl sterile ddH2O and use for transformation into E. coli. Yeast cells are genetically very flexible and have the tendency to circumvent many metabolically challenging situations by a rapid

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accumulation of suppressors or other kinds of mutations. Thus, it is essential now to verify that the encoded protein from the isolated plasmid itself complements the phenotype and not any other effect. For this purpose, the plasmid is again transformed into the yeast mutant and the complementation is compared to the mutant, transformed only with the empty plasmid. At this stage it will also be indicated whether the correct plasmid DNA was rescued, because of the presence of the expected auxotrophic marker (Ura3 for the cDNA library in this case). It has also to be considered that plant genes could be active in different pathways in yeast cells and require interaction with endogenous components to be functional. Examples for this phenomenon are illustrated in Fig. 15.2. In Fig. 15.2a the

A

complementation of hog1 ∆ 0 M NaCl

0.4 M NaCl

0.6 M NaCl

wt hog1 ∆ + MPK6 + MPK6 / MKK2 + MPK4 + MPK4 / MKK2

B

complementation of mpk1∆ control

4 mM caffeine

wt mpk1 ∆ + MPK6 + MPK6 / MKK2 + MPK4 + MPK4 / MKK2

Fig. 15.2. Functional complementation of yeast MAP kinase mutants with Arabidopsis MAP kinases and MAP kinase kinases. (a) Functional complementation of hog1D mutants with Arabidopsis MPK4 and MPK6 in combination with Arabidopsis MAP kinase kinase MKK2 and selection on high-salt media. 4 µl of a logarithmically growing culture was spotted in serial dilutions (1, 1:10, 1:100) on selective plates, either without salt as control for equal loading and viability (left panel) or on plates supplemented with 0.4 and 0.6 M NaCl. (b) Complementation of mpk1D mutants with the same kinases as shown above. The experimental procedure was the same as described above, but in this case the selection for functional complementation was done on plates containing 4 mM caffeine corresponding to the phenotype of the mpk1D mutant.

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complementation of the hog1D mutant by different Arabidopsis MAP kinases is shown. The first quite unexpected result is that both Arabidopsis MAP kinases (MPK6 and MPK4) alone are not able to complement the osmo-sensitive phenotype of the hog1D mutant despite the presence of the endogenous upstream activating MAP kinase kinase PBS2. Only if the plant-specific activator MKK2 is co-transformed, the complementation works. In such a case a screen, for example the complementation of the hog1D mutant, would certainly fail. The second aspect is the question of specificity within a given pathway. As can be seen in Fig. 15.2a the combination of MPK6 and MKK2 can complement the osmo-sensitive phenotype of the hog1D mutant much better than the combination of MPK4 and MKK2. However, from yeast two-hybrid data (11), we knew that MKK2 interacts stronger with MPK4 than MPK6. Therefore, we tested if MKK2 and MPK4 might be more functional in another yeast pathway and tested the yeast mpk1D mutant, which is sensitive to caffeine in a similar complementation approach (Fig. 15.2b). It turned out that in this case the combination of MKK2 complements the caffeine sensitivity of the mpk1D mutant much better than MKK2 plus MPK6.

4. Notes 1. It is only important here that the plasmid contains the same marker for selection, so any comparable plasmid carrying the URA3 marker is suitable here. 2. About 4 µl of 3 serial dilutions (1, 1:10, 1:100) of a logarithmically growing culture adjusted to an OD600 of 0.1 were spotted onto appropriate plates. 3. The transformation efficiency can be determined by plating a small aliquot (30 µl) of three dilutions (1, 1:10 and 1:100) on selective media without restricting conditions (like NaCl). The number of transformants per 1 µg cDNA can be calculated. A small aliquot (5–10 µl) of the transformed cells on plates, which select only for the plasmid but not for functional complementation, i.e. SC-ura in the procedure, is described here. 4. YPD medium might be supplemented with adenine hemisulphate (40 mg/l) to prevent fast reversion of the ade2 to ADE2 reversion. 5. The amino acid mixture (100 X) requires a short heating, which might easily be done in a microwave oven, for some amino acids to solve. The mixture is sterilized by filtration and kept at room temperature in the dark. Amino acids represented with italics in the list might be used for plasmid

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selection and have to be omitted from the mixture in this case. For more flexibility, we prepare the 100 X amino acid mixture usually without adenine, uracil, leucine, tryptophane and histidine and prepare sterile filtrated 100 X stock solutions of histidine and leucine, and uracil, and a 50 X stock solution of adenine, which can be added to the appropriate SC medium when required. Due to their low solubility in stock solutions, we add adenine, tyrosine and uracil (if not needed for selection!) directly to the SC medium (50 mg/l) before we autoclave it. 6. The use of bacterial RNA instead of single-stranded carrier DNA results in at least similar transformation efficiencies and has the advantage that sonication and boiling before usage are not necessary any more. 7. The high-efficiency transformation protocol needs to be done only if a library screen is performed. Much faster methods for simple introduction of a plasmid into yeast are available if the given plasmids are only to be tested for complementation of a certain mutant (13). 8. Selective media are required only if selection for a plasmid is necessary. 9. All steps during the transformation procedure are carried out at room temperature if not explicitly stated otherwise. 10. It is essential here to re-suspend the yeast cells in a small volume (one- maximal two- fold of the size of the pellet) to obtain a high transformation efficiency. 11. 300 µl of yeast cells are re-suspended in the half volume in Mix 1. This will be sufficient for 10 large plates (140 mm diameter). 12. To calculate: 300 µl cell suspension will be plated on one large plate with selective media. 13. It is possible to select specifically for the yeast LEU2 marker, which can complement the leu-phenotype of the bacterial strain HB101 on minimal media. This is used in yeast twohybrid systems, but does not help in this case where the library carries the URA3 marker for selection in yeast and the ampicillin resistance for selection in bacteria, which is also present in all commonly used yeast plasmids.

Acknowledgments We thank F. Lacroute for providing the excellent Arabidopsis cDNA library in pFL61. Work in our laboratory is supported by grants from the Austrian Science Foundation to M.T. (P16963-B12 and P19825-B12).

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References 1. Lee, M. G., and Nurse, P. (1987) Complementation used to clone a human homologue of the fission yeast cell cycle control gene cdc2. Nature 327, 31–35. 2. Ewbank, J. J., Barnes, T. M., Lakowski, B., Lussier, M., Bussey, H., and Hekimi, S. (1997) Structural and functional conservation of the Caenorhabditis elegans timing gene clk-1. Science 275, 980–983. 3. Ohi, R., Feoktistova, A., McCann, S., Valentine, V., Look, A. T., Lipsick, J. S and., Gould, K. L. (1998) Myb-related Schizosaccharomyces pombe cdc5p is structurally and functionally conserved in eukaryotes. Mol. Cell Biol. 18, 4097–4108. 4. Minet, M., Dufour, M. E., and Lacroute, F. (1992) Complementation of Saccharomyces cerevisiae auxotrophic mutants by Arabidopsis thaliana cDNAs. Plant J. 2, 417–422. 5. Shima, S., Matsui, H., Tahara, S., and Imai, R. (2007) Biochemical characterization of rice trehalose-6-phosphate phosphatases supports distinctive functions of these plant enzymes. Febs J. 274, 1192–1201. 6. Deuschle, K., Funck, D., Hellmann, H., Daschner, K., Binder, S., and Frommer, W. B. (2001) A nuclear gene encoding mitochondrial Delta-pyrroline-5-carboxylate dehydrogenase and its potential role in protection from proline toxicity. Plant J. 27, 345–356. 7. Mowla, S. B., Cuypers, A., Driscoll, S. P., Kiddle, G., Thomson, J., Foyer, C. H and., Theodoulou, F. L. (2006) Yeast complementation reveals a role for an Arabidopsis thaliana late embryogenesis abundant (LEA)-like protein in oxidative stress tolerance. Plant J. 48, 743–756.

8. Hohmann, S. (2002) Osmotic stress signaling and osmoadaptation in yeasts. Microbiol. Mol. Biol. Rev. 66, 300–372. 9. O’Rourke, S. M., Herskowitz, I., and O’Shea, E. K. (2002) Yeast go the whole HOG for the hyperosmotic response. Trends Genet. 18, 405–412. 10. Soyano, T., Nishihama, R., Morikiyo, K., Ishikawa, M., and Machida, Y. (2003) NQK1/NtMEK1 is a MAPKK that acts in the NPK1 MAPKKK-mediated MAPK cascade and is required for plant cytokinesis. Genes Dev. 17, 1055–1067. 11. Teige, M., Scheikl, E., Eulgem, T., Doczi, R., Ichimura, K., Shinozaki, K., Dangl, J. L and., Hirt, H. (2004) The MKK2 pathway mediates cold and salt stress signaling in Arabidopsis. Mol. Cell 15, 141–152. 12. Stockinger, E. J., Mao, Y., Regier, M. K., Triezenberg, S. J., and Thomashow, M. F. (2001) Transcriptional adaptor and histone acetyltransferase proteins in Arabidopsis and their interactions with CBF1, a transcriptional activator involved in cold-regulated gene expression. Nucleic Acids Res. 29, 1524–1533. 13. Gietz, R. D., and Woods, R. A. (2006) Yeast transformation by the LiAc/SS Carrier DNA/PEG method. Methods Mol. Biol. 313, 107–120. 14. Sambrook, J., and Russell, D. W. (2001) Molecular Cloning, A Laboratory Manual. Cold Spring Harbour Laboratory Press, Cold Spring Harbour, New York. 15. Gietz, R. D., Schiestl, R. H., Willems, A. R., and Woods, R. A. (1995) Studies on the transformation of intact yeast cells by the LiAc/ SS-DNA/PEG procedure. Yeast 11, 355–360.

Chapter 16 Phosphatase Activities Analyzed by In Vivo Expressions Alois Schweighofer, Zahra Ayatollahi, and Irute Meskiene Abstract Protein phosphatases act to reverse phosphorylation-related modifications induced by protein kinases. Type 2C protein phosphatases (PP2C) are monomeric Ser/Thr phosphatases that require a metal for their activity and are abundant in prokaryotes and eukaryotes. In plants, such as Medicago and Arabidopsis PP2Cs control several essential processes, including ABA signaling, development, and wound-induced mitogen-activated protein kinase (MAPK) pathways. In vitro assays with recombinant proteins and yeast two-hybrid systems usually provide initial information about putative PP2C substrates; however, these observations have to be verified in vivo. Therefore, a method for transient expression in isolated Arabidopsis suspension cell protoplasts was developed to assay PP2C action in living cells. This system has proven to be very useful in producing active enzymes and their substrates and in performing enzymatic reactions in vivo. Transient gene expression in isolated cells enabled assembly of functional protein kinase cascades and the creation of phosphorylated targets for PP2Cs. The method is based on the co-transformation and transient co-expression of different PP2C proteins with MAPK. It shows that epitope-tagged PP2C and MAPK proteins exhibit high enzymatic activities and produce substantial protein amounts easily monitored by Western blot analysis. Additionally, PP2C phosphatase activities can be directly tested in protein extracts from protoplasts, suggesting a possibility for analysis of activities of new PP2C family members. Key words: PP2C, MAPK, MPK6, MKK4, MP2C, AP2C1, SIMK, SIMKK, NPK1, cell suspension culture, transient expression, protoplasts, Arabidopsis, plant phosphatase, dephosphorylation, in vivo expression.

1. Introduction Phosphorylation-related modifications of proteins are essential for conveying multiple environmental signals into functional cell responses. Protein kinases and protein phosphatases control phosphorylation of different target proteins. At the same time T. Pfannschmidt (ed.), Plant Signal Transduction, vol. 479 Humana Press, a part of Springer Science + Business Media, LLC 2009 DOI: 10.1007/978-1-59745-289-2_16

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transient activities of kinases often depend on their phosphorylation state emphasizing the role of phosphatases in the control of signal transmissions. Protein phosphatases of type 2C (PP2Cs) are monomeric serine/threonine (Ser/Thr) phosphatases that require Mn2+ or Mg2+ for their activity and are insensitive to known phosphatase inhibitors, including okadaic acid. Medicago sativa phosphatase 2C (MP2C) was originally isolated as a protein suppressing yeast pheromone arrest (1), suggesting that it may act on activated MAPK signaling cascades. Consequently, an alfalfa MAPK was found as interacting partner of MP2C in a yeast two-hybrid screen identifying SIMK–MP2C as an interacting protein pair (2). PP2Cs are implicated in MAPK inactivation in yeast (1, 3–6), Drosophila (7), and in mammalian cells, where they control stress-activated MAPKs (5, 6, 8) . In plants, stress-MAPKs are components of wound-induced signaling cascades and are activated by dual phosphorylation of the activation loop amino acids threonine and tyrosine, suggesting that dual-specificity, tyrosine, and serine/threonine phosphatases could reverse these phosphorylations. Accordingly, MP2C was capable of inactivating SIMK by dephosphorylating the phospho-Thr residue in the activation loop of the SIMK MAPK (2). To validate the observations from in vitro experiments it is essential to develop a system to study protein functions also in vivo. Transient gene expressions proved to be an ideal method for testing PP2C protein functions (2). They provided the basis for a model of inactivation of the wound-induced signaling pathway by dephosphorylation of the MAPK by phosphatase in plants (Fig. 16.1a). To study PP2C inactivation of MAPK in vivo, the wound signal transduction pathway including activated MAPK can be reconstituted using transient expression in protoplasts. For this purpose, epitope-tagged MAPK is transfected into protoplasts prepared from suspension cultured cells. After 10–16 h, MAPK protein is usually accumulated in transformed cells in an inactive form (Fig. 16.1b). Transiently expressed MAPK can be activated by co-transfection with an active upstream MAPKKK or MAPKK (Fig. 16.1b). After immunoprecipitation from protoplast extracts, kinase activities are tested by in vitro kinase assays using myelin basic protein (MBP) as a substrate. Due to activation with MAPKKK or MAPKK, MAPK activity highly increases (Fig. 16.1a); therefore, PP2C ability to inactivate MAPK can be tested. In previous experiments MAPK SIMK-HA was co-transfected into protoplasts with the MP2C-Myc construct (2). The activation of SIMK was achieved through SIMKK (9, 10); therefore, SIMK inactivation by MP2C was followed when SIMK was activated by co-expression of constitutively active SIMKK or delta-NPK1, an MAPKKK from tobacco (11). Co-transformation

Fig. 16.1. Activation of the MAPK pathway and its negative regulation by the 2C type protein phosphatase. (a) MP2C/AP2C1 control of activated MAPK. Plant wound-activated MAPK pathway contains a three-kinase cascade that comprises the MAPKKK, MAPKK, and MAPK. The MAPKK (SIMKK or MKK4) activates MAPK (SIMK or MPK6) by phosphorylation of essential threonine (Thr) and tyrosine (Tyr) in the kinase activation loop TEY. PP2C-type phosphatase MP2C or AP2C1 inactivates MAPK by targeting phospho-Thr and removing the phosphate, thereby inactivating the pathway. MP2C/AP2C1 expression is transcriptionally upregulated upon wounding at the time of the inactivation of MAPK in leaves, suggesting the existence of a negative feedback mechanism. (b) Detection of PP2C inactivation of MAPK by co-expressions in protoplasts. AP2C1 inactivates MPK6 in vivo. Arabidopsis protoplasts isolated from suspension cells were transfected with 5 µg of MPK6-HA plasmid, 5 µg of MKK4-MYC plasmid (MKK4 as activator for MPK6), and 1 µg each of AP2C1-GFP, ABI2-GFP, or HAB1-GFP plasmid. After 14 h MAPK was immunoprecipitated ex vivo with anti-HA specific antibody and kinase assays were performed using myelin basic protein (MBP) as the substrate. Expression levels of the proteins were detected by immunoblotting: for MPK6 protein HA antibody, for MKK5 c-myc antibody, and for the phosphatases GFP antibody were used. (c) Phosphatase activity measurement of transiently expressed PP2C phosphatases in vivo. Arabidopsis protoplasts were transformed with 5 µg of AP2C1, HAB1, or ABI2 plasmids. A sample transformed with plasmid expressing GFP was used as a control. Phosphatase assays with labeled casein were performed using total protein extracts and the release of free phosphate (Pi) was measured at indicated time points.

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of SIMK with SIMKK into protoplasts activated SIMK without the necessity of an additional stimulus; however, co-transformation with MP2C resulted in complete inactivation of the MAPK, indicating that MP2C inhibits MAPK activity in vivo. Inactivation of SIMKK by MP2C is excluded by observations of non inhibition and noninteraction between these proteins (2). To assess if MAPK inactivation can also be performed by other PP2Cs, protein phosphatases from Arabidopsis, such as AP2C1, which is the closest ortholog of alfalfa MP2C, and phosphatases from cluster A (12), HAB1 (13) and ABI2 (14, 15), were tested. Protoplasts were co-transfected with Arabidopsis MAPK MPK6-HA, MAPKK MKK4-Myc as activator and each of the PP2C constructs (Fig. 16.1b). In this experiment it was found that co-transfection with AP2C1 led to a dramatic decrease in activation of MPK6, whereas HAB1 and ABI2 expression did not alter MPK6 activation, indicating that AP2C1 acts as an MAPK phosphatase similar to its alfalfa ortholog MP2C. To test if protein phosphatases with different abilities to inactivate MAPK are enzymatically active, their action toward common substrate phosphorylated casein can be measured. These measurements are performed directly in the extracts from protoplasts transiently expressing PP2C proteins (Fig. 16.1c). In this way we demonstrate that all of these PP2Cs are active phosphatases; however, not all are able to inhibit MAPK. The basic premise in this setup of experiments is if PP2C is a negative regulator of the pathway, it will inactivate the MAPK. The ex vivo assay used here relies on the observation that inactivation of the MAPK in plant cells is performed by MP2C/AP2C1 but not by evolutionary more distant phosphatases. We demonstrate here the following: isolation of protoplasts from Arabidopsis suspension cell culture, protoplast transformation with plasmids containing genes from the MAPK cascade, transient expression and activation of the substrate MAPK by co-expression of activating kinases in these cells, and expression of the active PP2Cs and measurement of their activities. PP2C, MAPKK, and MAPK synthesis in each sample is monitored by Western blotting using the respective antibodies. The global genome bioinformatics analysis identified the gene family of PP2C phosphatases represented by 76 members (12) and 4 PP2C-like candidate genes (16) among at least 150 total protein phosphatases detected in Arabidopsis. Only a few of them have been characterized so far (13, 17– 23). This set of experiments provides the basis for studies and understanding of phosphatase functions in vivo, and together with in vitro, yeast two-hybrid and proteomic tools will serve to progress in our understanding of the functions of protein phosphatases in plants.

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2. Materials 2.1. Maintenance of Arabidopsis Suspension Cultures for Protoplast Isolation

1. Arabidopsis Medium: 4.4 g MS including B5 vitamins (Duchefa), 30 g saccharose (refined sugar from local supermarket), 1 mg 2.4-D per liter, pH 5.7 adjusted with KOH, autoclaved.

2.2. Protoplast Isolation from Suspension Culture Cells and Protoplast Transformation

1. B5-S: 0.28 M sucrose, 3.16 g B5 powder (Duchefa, G0210) per liter, pH 5.5 adjusted with KOH, autoclaved. 2. B5-GM: 0.34 M glucose, 0.34 M mannitol, 3.16 g B5 powder per liter, pH 5.5 adjusted with KOH, autoclaved. 3. Enzyme solution: 1% cellulase (Serva), 0.2% macerozyme (Yakult or Serva), in B5-GM, 0.45-µm filter sterilized (see Note 1). 4. 0.275 M Ca(NO3)2, pH 5.6 adjusted with KOH, autoclaved. 5. PEG solution: 25% PEG 6000, 0.45 M mannitol, 0.1 M Ca(NO3)2, pH 9.0 adjusted with KOH (see Note 2). 6. Falcon 50-ml tubes. 7. Round-bottom 12-ml tubes. 8. Rosenthal chamber (Hemacytometer). 9. Inverted light microscope. 10. Rotating platform.

2.3. Protein Extraction from Protoplasts

1. Lacus extraction buffer: 25 mM Tris-HCl, pH 7.8, 10 mM MgCl2, 15 mM EGTA, 75 mM NaCl, 1 mM dithiothreitol, 1 mM NaF, 0.5 mM NaVO3, 15 mM β-glycerophosphate, 0.1% [v/v] Tween-20, 15 mM p-nitrophenylphosphate, 0.5 mM phenylmethylsulfonylfluoride (PMSF), leupeptine (5 µg/ ml), aprotinin (5 µg/ml).

2.4. Ex Vivo Kinase Assay

1. Suc1 buffer: 50 mM Tris-HCl, pH 7.4, 250 mM NaCl, 5 mM EGTA, 5 mM EDTA, 0.1% Tween-20, 5 mM NaF, 0.1% Nonidet P-40, 0.5 mM PMSF. 2. Kinase buffer: 20 mM Hepes, Tris-HCl pH 7.5, 15 mM MgCl2, 5 mM EGTA, 1 mM dithiothreitol. 3. 4x SDS loading buffer: 200 mM Tris-HCl, pH 6.8, 400 mM DTT, 8% SDS, 40% glycerol, 0.1% bromophenol blue. 4. Coomassie Blue gel-staining solution: 0.25% [w/v] Coomassie brilliant blue R-250, 45% [v/v] methanol, 10% [v/v] acetic acid. 5. Gel destaining solution: 45% [v/v] methanol, 10% [v/v] acetic acid.

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2.5. Western Blotting

1. Polyvinylidene difluoride (PVDF) membranes (Millipore). 2. Buffer-tank–blotting apparatus. 3. Ponceau S: 0.5% [w/v] Ponceau S, 1% [v/v] glacial acetic acid. 4. TBS-T: 50 mM Tris base, 150 mM NaCl, 0.1% [v/v] Tween-20. 5. TBS-T buffer containing 5% [w/v] nonfat dried milk obtained from local supermarket. 6. Monoclonal anti-GFP antibody (Roche, Cat. No. 1814460), dilution 1:5000 in TBS-T containing 1% [w/v] nonfat dried milk. 7. Secondary anti mouse IgG developed in goat (Sigma, A-5153), dilution 1:5000 in TBS-T containing 1% [w/v] nonfat dried milk. 8. CDP-star detection reagent (Amersham Biosciences). 9. X-ray film: Hyperfilm-ECL film (Amersham Biosciences).

2.6. Ex Vivo Phosphatase Assay 2.6.1. Substrate (Casein) Labeling

1. Kinase buffer: 50 mM Tris-HCl pH 7.0, 5 mM MgCl2. 2. Protein kinase A catalytic subunit from bovine heart (PKA; Sigma P-2645). 3. Casein solution from bovine milk, 5% (Sigma C-4765). 4. γ-32P-ATP from Amersham Biosciences (AA0018, 500 µCi).

2.6.2. Casein Dephosphorylation

1. 20% TCA in 20 mM NaH2PO4. 2. 10x buffer for dephosphorylation reaction: 600 mM TrisHCl pH 7.0, 1 mM EGTA, 100 mM β-mercaptoethanol, 100 mM MgCl2, 5 µM okadaic acid. 3. Norit A solution: 900 mM HCl, 90 mM Na-pyrophosphate, 2 mM NaH2PO4, 4% [v/v] Norit A (activated charcoal).

2.7. Plasmids

Expression plasmids: full-length PP2Cs and MAPKs were cloned under control of 35S promoter in pRT100 plant expression vector from Ref. (24), which was modified by inserting the translational enhancer (5′UTR) from the Tobacco etch virus (TEV) (25) or the omega element from tobacco mosaic virus (TMV) (26). Kinases were tagged at their C-termini with triple HA (influenza hemagglutin) or c-Myc epitope; PP2Cs were tagged at their C-termini with green fluorescent protein GFP (27). Plasmid DNAs used for transformation were isolated with Genomed or Qiagen DNA isolation kits according to the manufacturer’s recommendations and diluted to a concentration of 1 µg/µl.

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3. Methods The strategy described below combines several methods outlined as follows: 1. The maintenance of Arabidopsis suspension cultures for protoplast isolation 2. The isolation of protoplasts from the cultivated cells 3. The transformation of the protoplasts 4. The extraction of the proteins from protoplasts 5. The immunoprecipitation and ex vivo kinase assay 6. The Western blotting 7. The ex vivo phosphatase assay 3.1. Maintenance of Arabidopsis Suspension Cultures for Protoplast Isolation

1. Propagate Arabidopsis suspension cells each week by diluting 1:5 in Arabidopsis Medium (5 ml of 7-days-old cells with 20 ml fresh medium) in 250-ml Erlenmeyer flasks under sterile conditions (see Note 3). Cultivate cells on a shaker at 150 rpm in darkness at 22°C.

3.2. Protoplast Isolation from the Cells

1. Collect 4- to –5-days-old Arabidopsis cell cultures. 2. Spin 25 ml of culture in a 50-ml Falcon tube for 5 min at 250 × g. 3. Discard the supernatant by decanting and keep the cells (see Note 4). 4. Add 25 ml of enzyme solution to the cells and fill up to 50 ml with B5-GM. 5. Split the content and place in two large Petri plates. 6. Add 25 ml of B5-GM to each plate. 7. Place plates on the rotating platform for ~3 h (see Note 5). 8. Transfer the content from each plate to 50-ml Falcon tubes. 9. Spin the tubes for 5 min at 150 × g (see Note 6). 10. Discard the supernatant by decanting. Resuspend the pellet in 15 ml of B5-S by carefully ticking the tube and then fill up to 35–40 ml with B5-S. 11. Spin the tubes for 8 min at 70 × g, transfer the floating cells with a plastic wide-mouth Pasteur pipette (~2–3 ml) to the new 15-ml Falcon tube, and fill up to ~10 ml with B5-S. 12. Spin the tubes again for 5–7 min at 70 × g. 13. Transfer the floating cells to the new 15-ml falcon tube. It should give ~2–3 ml of protoplasts.

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14. Check protoplast quality under the light microscope (see Note 7) and count their concentration in a Rosenthal chamber. The final concentration of protoplasts should be adjusted to 4 × 106/ml with B5-S (see Note 8). 3.3. Protoplast Transformation

1. Use 1–5 µg of each plasmid per tube for transformation. Set the final volume to 15 µl in the microcentrifuge tubes (see Note 9). Usually protoplast transformations are performed in triplicates. 2. Add 1–2 × 105 of protoplasts in 50 µl per tube, and mix with the plasmids by carefully ticking against the tube. 3. Add 150 µl PEG solution immediately and mix well by ticking against the tube several times. 4. Incubate the suspension 5–15 min at room temperature. 5. Dilute the PEG by adding stepwise 0.275 M Ca(NO3)2 (twice 0.5 ml) and mix well by inverting the tubes several times. 6. Spin at 70 × g for 7 min. 7. Remove the supernatant (see Note 10). 8. Add 0.5 ml of B5-GM to cultivate protoplasts for 3–14 h (see Note 11).

3.4. Protein Extraction from Protoplasts

1. Combine protoplasts transformed in triplicates using a pipettes with wide-mouth tip. Harvest the protoplasts by centrifugation at 100 × g for 1 min and remove the supernatant by aspiration. 2. Add 100 µl of Lacus extraction buffer, resuspend the pellet quickly by flicking the tube, and freeze in liquid nitrogen. Store the samples at −80°C. 3. Proteins are extracted from the protoplasts by short vortexing (see Note 12). 4. Spin down the cell debris for 20 min at 30,000 × g at 4°C. 5. Transfer the supernatant into new microcentrifuge tubes whereby 80–90 µl of protein extract is used for immunoprecipitation of kinases and 10 µl of protein extract should be combined with 3.3 µl 4×SDS PAGE sample buffer for Western blot analysis (see Note 13).

3.5. Ex Vivo Kinase Assay 3.5.1. Immunoprecipitation of Kinase

1. Prepare protein extracts as described in Sect. 3.4. 2. Take About 90 µl of the protein extract from a triple protoplast sample (3 combined tubes) for immunoprecipitation. 3. Pre-clean the extract with 20 µl 50% slurry of sepharose beads (composed of a 1:1 mixture of protein A and protein G sepharose beads (Amersham, Sweden)) in an equal volume of Suc1 buffer, rolling the tube horizontally on a tube turner wheel with rotary agitation for 1 h at 4°C. 4. Centrifuge at 250 × g for 2 min at 4°C in a swing out rotor.

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5. Transfer the supernatant into a new microcentrifuge tube containing 20 µl protein G sepharose beads and 3 µL of antibody, and roll the tubes like in Step 3 horizontally 3–4 h or overnight at 4°C. 6. Spin down the beads at 250× g at 4°C for 1 min, aspirate the supernatant by a pipette or just a tip connected to a vacuum pump (takes few seconds), and wash the beads three times with 500 µl ice-cold Suc1 buffer. 7. Wash the beads once with kinase buffer and remove the leftover liquid from the beads with a syringe. 3.5.2. In Vitro Phosphorylation Reaction of MBP

1. Add 1.5 µg MBP, 15 µl kinase buffer, 0.015 µl 1 M DTT, 3 µCi of γ-32P-ATP [0.2 µl of 500 µCi], and 0.15 µl 10 mM ATP into tubes containing beads prepared from Sect. 3.5.1, and incubate the kinase reaction for 30 min at room temperature. 2. Add 5µl 4× SDS loading buffer. 3. Heat the sample at 95°C for 3 min and chill it on ice. 4. Spin the sample at 16,000× g for 1 min.

3.5.3. SDS-PAGE of MBP

1. Load 6 µl of the sample on a 12.5% SDS-PA gel. 2. Run electrophoresis with 15 mA until the dye runs out. 3. Stain the gel with gel staining solution for 20 min. 4. Destain with destainer solution for 2 h, changing the solution every 20 min. 5. Place the gels on Whatman 3MM paper and vacuum dry at 80°C for 45 min. 6. The phosphorylation of MBP is analyzed by autoradiography exposing the gel to an x-ray film for 2–4 h overnight.

3.6. Western Blotting

1. Following electrophoresis, transfer proteins from the polyacrylamide gel to polyvinylidene difluoride membranes (Millipore) in a buffer-tank–blotting apparatus at 75 V for 2.5 h. 2. Check transfer efficiency by staining proteins on the membrane in Ponceau S for 30 s, and destain in water for 30 s. 3. Incubate membranes in blocking solution TBS-T with 5% nonfat dried milk for 2 h at room temperature with gentle agitation. 4. Probe membranes with monoclonal antibodies diluted 1:5000 in TBS-T buffer containing 5% [w/v] nonfat dried milk for 12–16 h at 4°C with gentle agitation. 5. Wash the membranes in TBS-T buffer containing 1% [w/v] nonfat dried milk by changing the washing solution four times every 15 min. 6. Add secondary anti mouse IgG diluted at 1:5000 in TBS-T containing 1% [w/v] nonfat dried milk and incubate for 1 h at room temperature.

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7. Wash the membranes in TBS-T buffer by changing the buffer eight times every 20 min. 8. Perform chemiluminescent detection reaction using CDPstar detection reagent (Amersham Biosciences) according to the manufacturer’s recommendations. 9. Expose membranes to Hyperfilm ECL (Amersham Biosciences) for 1–10 min. 3.7. Ex Vivo Phosphatase Assay 3.7.1. Substrate (Casein) Labeling

1. Reconstitute PKA catalytic subunit by adding 50 mM DTT to make 5 U/µl solution (see Note 14). 2. Add ~1 mg of casein to the reaction mixture containing ~100 µCi γ-32P-ATP and ~25 units of catalytic subunit of PKA in a buffer containing 50 mM Tris-HCl pH 7.0 and 5 mM MgCl2 in a total volume of 100 µl. 3. Incubate the reaction for 30 min at 30°C. 4. Stop the reaction by adding 500 µl 20% TCA/20 mM NaH2PO4 and chill 20 min on ice. 5. Spin 10 min at 10,000 × g at room temperature. 6. Remove supernatant. 7. Wash in several consecutive steps by adding 700 µl 20% TCA/20 mM NaH2PO4 and spinning for 1 min at 8000 × g until the supernatant reaches a radioactivity below ~300 counts per minute (cpm) (see Note 15). 8. Remove the supernatant completely and dissolve the casein pellet in 400 µl of 0.2 M Tris-HCl pH 8.0 and 500 nM okadaic acid. 9. Measure an aliquot of labeled casein in a microcentrifuge tube by mixing 5 µl of labeled casein (~2.5 mg/ml) with 750 µl Norit A mixture in a microcentrifuge tube and incubate for 10 min at room temperature (see Note 16). 10. Centrifuge for 5 min at 10,000 × g. 11. Use 500 µl of the supernatant for measurement in a scintillation counter and compare with counts from the pellet (see Note 17).

3.7.2. Casein Dephosphorylation

1. Prepare the reaction in a total volume of 50 µl by mixing on ice: ~5 µl (~12.5 µg) of radiolabeled casein (>200,000 cpm) and 5 µl protein extract (1.2–1.5 µg) in 1x dephosphorylation buffer (5 µl 10× buffer and H2O to reach the final reaction volume of 50 µl). 2. Incubate the reaction for 30 min to 2 h at 28°C (dependent on the activity of the phosphatase) (see Note 18). 3. Stop the reaction by adding 750 µl Norit A mixture; incubate it for 10 min at room temperature.

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4. Spin the sample for 5 min at 10,000× g and immediately transfer 500 µl of the supernatant to a new tube, avoiding any charcoal particles that may remain from the Norit A mixture. 5. Measure the supernatant activity by placing the microcentrifuge tube directly into a scintillation vial (see Note 17).

4. Notes Originally developed for parsley suspension cell culture protoplasts (Petroselinum crispum) (28), this method has been modified and established for Arabidopsis suspension. Other procedures involving PP2C expressions were also reported for mesophyll protoplasts isolated from maize (29). 1. Enzyme solution has to be prepared fresh, stirred slowly for 1 h, filtered through Whatman paper, and then sterilized through a 0.45-µm filter. 2. After stirring 4 h or overnight (the pH has to be remeasured), the solution is sterilized by filtration. 3. Dilution depends on the specificity of the suspension culture; for slower growing cells it may be necessary to dilute cells 1:3 only. 4. Cells should be in approx. 10- to 5-ml pellets. 5. Rotate plates slowly at 30–40 rpm/min and check protoplasts under the inverted light microscope every 30 min to decide on the time for harvesting. If 80% of the cells start to round up (meaning that they are fully protoplasted), begin with the protoplast isolation. 6. All centrifugation steps for protoplasts should decelerate from any speed to zero without a break. Cells should float. 7. 90 % the cells should be fully rounded. The cell number in 16 small squares × 5 × 103 in a chamber (depth 0.2 mm, surface area of smallest square 0.0625 mm2) gives the number of protoplasts per microliter. 8. Protoplasts are usually kept at 4°C and can be transformed on the same day or stored overnight. The transformation efficiency decreases with storage time, but they can be stored for 2–3 days and used, depending on the experimental goals. 9. Several constructs can be combined for co-transformation. Optimal plasmid size is 4–6 kB. Plasmid(s) are isolated by Qiagen or Genomed DNA isolation kits and dissolved after ethanol precipitation in sterile H2O. Set the volume to 15 µl in the microcentrifuge tubes. 10. Remove the supernatant by aspiration with a vacuum pump or with a pipette.

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11. Incubate cells for 10–14 h at 23°C in darkness. Transient expression can be monitored by fluorescence microscopy after ~4–6 h. Transformation efficiency is always tested by transforming a control sample with a 35S::GFP plasmid. Good transformation of protoplasts gives 30–80% transformation efficiency (counting transformed versus untransformed protoplasts by visual detection with the microscope). 12. All the extraction steps are carried out at 0 or 4°C. 13. Samples for immunoprecipitation have to be used right away and samples for Western blot can be stored at −20°C. 14. PKA is purchased as lyophilized powder. Dissolving in 50 mM DTT reconstitutes the active protein. Use reconstituted PKA right away or store up to 3 days on ice. 15. It takes ~10 washing steps until the supernatant activity is below 300 cpm. The labeled casein can be stored at −20°C for 1–2 weeks in TCA/NaH2PO4. 16. Use a magnetic stirrer to stir Norit A solution to have equal distribution of carbon particles. Determination of the efficiency of 32P incorporation into casein: good labeling should give >200,000 cpm/5 µl. 17. Take care not to aspirate the carbon particles, as this will increase the background. The β-emissions of the pellets (labeled casein) and the supernatant (unincorporated γ-32P-ATP, i.e., background) are quantified as counts per minute (cpm) in a Tri-carb 1600 TR Liquid scintillation counter (Packard). The background usually is ~0.1%, i.e., the labeled casein gives ~200,000 cpm, whereas the background ~200–300 cpm. 18. Phosphate release can be detected after 30 min. However, if the analyzed enzyme is not highly active, longer incubation time is advisable. Dephosphorylation may continue up to 2 h, which might be favorable when comparing different samples.

Acknowledgements We thank Austrian FWF and Lithuanian State Science and Studies Fund for support to I.M., FWF Erwin-Schrödinger-Fellowship to A.S. and Vienna University Fellowship to Z.A.

References 1. Meskiene, I., Bogre, L., Glaser, W., Balog, J., Brandstotter, M., Zwerger, K., Ammerer, G., and Hirt, H. (1998) MP2C, a plant protein phosphatase 2C, functions as a negative regulator of mitogen-activated protein

kinase pathways in yeast and plants. Proc. Natl. Acad. Sci. U S A 95, 1938–1943. 2. Meskiene, I., Baudouin, E., Schweighofer, A., Liwosz, A., Jonak, C., Rodriguez, P. L., Jelinek, H., and Hirt, H. (2003) The Stressinduced protein phosphatase 2C is a negative

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regulator of a mitogen-activated protein kinase. J. Biol. Chem. 278, 18945–18945. Warmka, J., Hanneman, J., Lee, J., Amin, D., and Ota, I. (2001) Ptc1, a type 2C Ser/ Thr phosphatase, inactivates the HOG pathway by dephosphorylating the mitogenactivated protein kinase Hog1. Mol. Cell Biol. 21, 51–60. Nguyen, A. N. and Shiozaki, K. (1999) Heat-shock-induced activation of stress MAP kinase is regulated by threonine- and tyrosine-specific phosphatases. Genes Dev. 13, 1653–1663. Takekawa, M., Maeda, T., and Saito, H. (1998) Protein phosphatase 2Calpha inhibits the human stress-responsive p38 and JNK MAPK pathways. Embo J. 17, 4744–4752. Takekawa, M., Adachi, M., Nakahata, A., Nakayama, I., Itoh, F., Tsukuda, H., Taya, Y., and Imai, K. (2000) p53-inducible wip1 phosphatase mediates a negative feedback regulation of p38 MAPK-p53 signaling in response to UV radiation. Embo J. 19, 6517–6526. Baril, C. and Therrien, M. (2006) Alphabet, a Ser/Thr phosphatase of the protein phosphatase 2C family, negatively regulates RAS/MAPK signaling in Drosophila. Dev. Biol. 294, 232–245. Bulavin, D. V. and Fornace, A. J., Jr. (2004) p38 MAP kinase’s emerging role as a tumor suppressor. Adv. Cancer Res. 92, 95–118. Kiegerl, S., Cardinale, F., Siligan, C., Gross, A., Baudouin, E., Liwosz, A., Eklof, S., Till, S., Bogre, L., Hirt, H., and Meskiene, I. (2000) SIMKK, a mitogen-activated protein kinase (MAPK) kinase, is a specific activator of the salt stress-induced MAPK, SIMK. Plant Cell 12, 2247–2258. Cardinale, F., Meskiene, I., Ouaked, F., and Hirt, H. (2002) Convergence and divergence of stress-induced mitogen-activated protein kinase signaling pathways at the level of two distinct mitogen-activated protein kinase kinases. Plant Cell 14, 1–10. Kovtun, Y., Chiu, W. L., Zeng, W., and Sheen, J. (1998) Suppression of auxin signal transduction by a MAPK cascade in higher plants. Nature 395, 716–720. Schweighofer, A., Hirt, H., and Meskiene, I. (2004) Plant PP2C phosphatases: emerging functions in stress signaling. Trends Plant Sci. 9, 236–43. Saez, A., Apostolova, N., Gonzalez-Guzman, M., Gonzalez-Garcia, M.P., Nicolas, C., Lorenzo, O., and Rodriguez, P.L. (2004) Gain-of-function and loss-of-function phenotypes of the protein phosphatase 2C

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HAB1 reveal its role as a negative regulator of abscisic acid signaling. Plant J. 37, 354–369. Leung, J., Merlot, S., and Giraudat, J. (1997) The Arabidopsis ABSCISIC ACIDINSENSITIVE2 (ABI2) and ABI1 genes encode homologous protein phosphatases 2C involved in abscisic acid signal transduction. Plant Cell 9, 759–771. Rodriguez, P. L., Benning, G., and Grill, E. (1998) ABI2, a second protein phosphatase 2C involved in abscisic acid signal transduction in Arabidopsis. FEBS Lett. 421, 185–190. Kerk, D. (2006) Genome-scale discovery and characterization of class-specific protein sequences: an example using the protein phosphatases of Arabidopsis thaliana. Methods Mol. Biol. 365, 347–370. Merlot, S., Gosti, F., Guerrier, D., Vavasseur, A., and Giraudat, J. (2001) The ABI1 and ABI2 protein phosphatases 2C act in a negative feedback regulatory loop of the abscisic acid signaling pathway. Plant J. 25, 295–303. Mishra, G., Zhang, W., Deng, F., Zhao, J., and Wang, X. (2006) A bifurcating pathway directs abscisic acid effects on stomatal closure and opening in Arabidopsis. Science 312, 264–266. Saez, A., Robert, N., Maktabi, M. H., Schroeder, J. I., Serrano, R., and Rodriguez, P. L. (2006) Enhancement of abscisic acid sensitivity and reduction of water consumption in Arabidopsis by combined inactivation of the protein phosphatases type 2C ABI1 and HAB1. Plant Physiol. 141, 1389–1399. Yoshida, T., Nishimura, N., Kitahata, N., Kuromori, T., Ito, T., Asami, T., Shinozaki, K., and Hirayama, T. (2006) ABA-hypersensitive germination3 encodes a protein phosphatase 2C (AtPP2CA) that strongly regulates abscisic acid signaling during germination among Arabidopsis protein phosphatase 2Cs. Plant Physiol. 140, 115–126. Kuhn, J. M., Boisson-Dernier, A., Dizon, M. B., Maktabi, M. H., and Schroeder, J. I. (2006) The protein phosphatase AtPP2CA negatively regulates abscisic acid signal transduction in Arabidopsis, and effects of abh1 on AtPP2CA mRNA. Plant Physiol. 140, 127–139. Stone, J. M., Trotochaud, A. E., Walker, J. C., and Clark, S. E. (1998) Control of meristem development by CLAVATA1 receptor kinase and kinase-associated protein phosphatase interactions. Plant Physiol. 117, 1217–1225.

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23. Ding, Z., Wang, H., Liang, X., Morris, E.R., Gallazzi, F., Pandit, S., Skolnick, J., Walker, J.C.., and Van Doren, S.R. (2007) Phosphoprotein and Phosphopeptide Interactions with the FHA Domain from Arabidopsis Kinase-Associated Protein Phosphatase. Biochemistry 46, 2684–2696. 24. Topfer, R., Matzeit, V., Gronenborn, B., Schell, J., and Steinbiss, H.H. (1987) A set of plant expression vectors for transcriptional and translational fusions. Nucleic Acids Res. 15, 5890. 25. Restrepo, M. A., Freed, D. D., and Carrington, J. C. (1990) Nuclear transport of plant potyviral proteins. Plant Cell 2, 987–998. 26. Holtorf, S., Apel, K., and Bohlmann, H. (1995) Comparison of different constitutive

and inducible promoters for the overexpression of transgenes in Arabidopsis thaliana. Plant Mol. Biol. 29, 637–646. 27. Chiu, W., Niwa, Y., Zeng, W., Hirano, T., Kobayashi, H., and Sheen, J. (1996) Engineered GFP as a vital reporter in plants. Curr. Biol. 6, 325–330. 28. Dangl, J. L., Hauffe, K. D., Lipphardt, S., Hahlbrock, K., and Scheel, D. (1987) Parsley protoplasts retain differential responsiveness to u.v. light and fungal elicitor. Embo J. 6, 2551–2556. 29. Sheen, J. (1998) Mutational analysis of protein phosphatase 2C involved in abscisic acid signal transduction in higher plants. Proc. Natl. Acad. Sci. U S A 95, 975–980.

Chapter 17 Chromatin Immunoprecipitation Experiments to Investigate In Vivo Binding of Arabidopsis Transcription Factors to Target Sequences Benjamin Fode and Christiane Gatz Abstract For understanding the mechanism of transcriptional regulation, it is essential to know which transcription factor is bound in vivo to the promoter to be analysed. If transcription from a given promoter is regulated by developmental or environmental stimuli, the question of inducible versus constitutive binding has to be answered, particularly if the transcriptional regulator is expressed both under uninduced and induced conditions. Chromatin immunoprecipitation (ChIP) assays constitute the most adequate approach to address these issues as proteins are cross-linked to the DNA before disruption of the tissue. Thus, the DNA–protein interaction is stabilized during purification of the chromatin. The specific DNA– protein complex is immuno-enriched employing specific antibodies against the transcription factor to be analysed. After reversal of cross-links, the recovered DNA is amplified by PCR using specific primers that match sequences flanking the suspected binding site. The amount of PCR product is indicative of the relative abundance of the DNA–protein complex in vivo. A protocol for ChIP assays for Arabidopsis thaliana leaves is described. Key words: Chromatin immunoprecipitation, protein–DNA complex, transcription factor.

1. Introduction An important method to study the regulation of transcription in living cells is the chromatin immunoprecipitation (ChIP) assay (1). ChIP allows the analysis of the in vivo binding status of transcription factors or other DNA-associated proteins to certain DNA sequences (X-ChIP). Intact cells are treated with formaldehyde to cross-link promoter-associated proteins to the DNA. After isolation and shearing of the chromatin, protein– T. Pfannschmidt (ed.), Plant Signal Transduction, vol. 479 Humana Press, a part of Springer Science + Business Media, LLC 2009 DOI: 10.1007/978-1-59745-289-2_17

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DNA complexes are immunoprecipitated with specific antibodies against the protein of interest. The precipitated DNA fragments are subsequently purified and analysed by polymerase chain reactions (PCR) using primers flanking the (putative) binding site of the protein. The amount of PCR product obtained is indicative of the relative amount of protein bound to the DNA when the tissue was harvested. The procedure allows detection of quantitative differences in the relative amount of protein–DNA complexes, so that stimulus-induced binding can be detected. If the binding site (cis element) of the transcription factor is not known, the promoters of putative target genes can be identified by ChIP followed by microarray analyses (ChIP-Chip, (2)). In these experiments, new direct target genes can be identified, especially when whole genome tiling arrays are available (like for Arabidopsis thaliana). Alternatively, a library can be generated by cloning precipitated fragments after amplification by ligation mediated PCR (3). In addition to the analysis of transcription factor binding, multi-protein complexes associated with the DNA can be studied using ChIP. As formaldehyde also cross-links interacting proteins, multi-protein complexes that assemble on the promoter region can be mapped in vivo (4). In this context, the so-called SeqChIP can be used to address whether two proteins are simultaneously bound to a stretch of DNA (5). As ChIP is also used to detect modifications at histones (for review see Ref. (6)), a comprehensive snapshot of the events taking place during transcriptional activation can be obtained. The following protocol describes a classical X-ChIP approach performed with leaves from Arabidopsis thaliana.

2. Materials 2.1. Preparation of Samples 2.2. Cross-Linking of Proteins to DNA

Approximately 5 g of Arabidopsis thaliana leaf material is needed. 1. Cross-link buffer 1 (CLB1): 50 mM KH2PO4/K2HPO4, pH 5.8, 1% formaldehyde (see Note 1) 2. Cross-link buffer 2 (CLB2): 50 mM KH2PO4/K2HPO4, pH 5.8, 0.3 M glycine

2.3. Isolation of Nuclei

1. Twenty-five percent Triton X-100, stored at 4°C. 2. Nuclei extraction buffer (NEB): 1 M hexylene glycol (2-methyl-2,4-pentanediol), 50 mM PIPES-KOH, pH 7.2, 10 mM MgCl2. Before use, β-mercaptoethanol has to be added to a final concentration of 5 mM (see Note 2).

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3. Gradient buffer (GB): 0.5 M hexylene glycol (β-methyl-2,4pentanediol), 50 mM PIPES-KOH, pH 7.2, 10 mM MgCl2, 1% (w/v) Triton X-100. Before use, β-mercaptoethanol has to be added to a final concentration of 5 mM. 4. Percoll cushions: 75 and 35% (w/v) Percoll in GB. 5. Miracloth (Calbiochem, Merck Chemicals Ltd., Nottingham, UK). 6. Nuclear stain: 300 nM DAPI (4,6-diamidino-2-phenylindole) in water, to be stored at −20°C. 7. Tissue homogenize (e.g., MiccraRT from Art Labortechnik, Mülheim, Germany). 2.4. Extraction of Chromatin

1. Sonic buffer (SB): 10 mM Tris-HCl, pH 7.4, 1 mM EDTA (ethylenediamine tetraacetic acid) containing either 0, 0.25 or 0.5% (w/v) sodium dodecyl sulphate (SDS). The buffer is stored at room temperature. 2. Protease inhibitor mix for plants (P-9599, Sigma-Aldrich, St. Louis, MO). 3. MSE Soniprep 150 ultrasonic disintegrator (Sany-GallenKamp, Loughboro, Leicestershire, UK)

2.5. Purification of DNA

1. Proteinase K (20 mg/ml stock). 2. PCI-Mix: phenol (equilibrated, pH 7.6–8.0, AppliChem)/ chloroform/isoamyl alcohol 25:24:1 (v/v/v). The mixture is stored at 4°C. All work involving phenol and chloroform should be done under a hood. 3. CI-Mix: chloroform/isoamyl alcohol 24:1 (v/v). 4. 3 M Na-acetate. 5. Glycogen 10 mg/ml (G-8751, Sigma-Aldrich), store at 2–8°C.

2.6. Chromatin Immunoprecipitation

1. RIPA-F: 50 mM HEPES-NaOH, pH 7.4 (see Note 3), 140 mM NaCl, 1 mM EDTA, 1% Triton X-100, 0.1% sodium deoxycholate (DOC, minimum 97%, Sigma-Aldrich), 0.1% SDS; RIPA-F buffer without SDS is required for sample dilution before immunoprecipitation (see below). RIPA-F buffer may be used for up to 4 weeks when stored at 4°C. 2. Protein A Sepharose (from Staphylococcus aureus, SigmaAldrich); store at 2–8°C. 3. Elution buffer (EB): 0.1 M glycine (adjust with HCl to pH 2.5), 0.5 M NaCl, 0.05% Tween20. This buffer can be stored for up to 4 weeks at 4°C. 4. 10 mM Tris-HCl, pH 7.4 5. Proteinase K (20 mg/ml). Aliquots (100 µl) should be stored at −20°C.

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6. Rotary platform like e.g., an Intelli-Mixer, LTF Labortechnik, Wasserburg, Germany

3. Methods The protocol starts with infiltrating the material with a buffer containing formaldehyde. Formaldehyde is a very reactive bipolar compound that reacts with amino and imino groups of amino acids and DNA thus causing reversible cross-links between proteins and DNA or between proteins (1). Cross-linking is terminated by adding 0.3 M glycine. After the cross-linking, nuclei are prepared by two centrifugation steps of filtered whole cell extracts on Percoll cushions. Afterwards, the nuclear envelope is solubilized with detergents. The chromatin is sheared into 500- to 1000-bp fragments by sonication. After spinning down the insoluble debris, the supernatant is directly used for the immunoprecipitation. Immunoprecipitations can be either performed with affinity-purified antibodies or with the complete antiserum. When using unprocessed antisera, control precipitations with the respective pre-immune serum have to be done. Mutant plants lacking the transcription factor and/or amplification of a fragment lacking the putative binding site are also valuable tools to prove the specificity of the precipitation. After the immunoprecipitation, the DNA has to be purified for PCR analysis. A combination of de-cross-linking by heat treatment, protease A digestion, and phenol extractions serves to remove the protein moiety of the complex. Chromatin that is not subjected to the immunoprecipitation is purified in the same way and serves to demonstrate that the same amount of chromatin is used for each sample (input control). Quantitative analysis of the so-called IP-DNA by real-time PCR provides information about the extent to which a given transcription factor occupies its target sequence within a promoter. For the typical outcome of a ChIP analysis see Fig. 17.1. 3.1. Plant Growth

1. Grow Arabidopsis plants in a climate chamber under shortday conditions (see Note 4) for about 4–5 weeks until they have developed enough biomass (about 5 g per sample). 2. Depending on the biological question to be asked, subject your plants to the appropriate treatment (e.g., pathogens, hormones, etc.). 3. Harvest 5 g of leaf material.

3.2. In Vivo Cross-linking of Proteins to DNA

All steps for the cross-linking procedure are carried out at room temperature if not noted otherwise. 1. Put the leaves in a suitable device for subsequent vacuum-infiltration (see Note 5). This device should prevent floating of

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threshold cycle C(t)

18 19

promoter 1

20

promoter 2

21 22 23 24 25 26 27 28

wild-type

tga2tga5tga6

IP DNA

wild-type

tga2tga5tga6

Input DNA

Fig. 17.1. ChIP analysis to demonstrate specific binding of bZIP factor TGA2 to one of its target promoters (promoter 1). A primer pair directed against a control promoter that does not contain a TGA-binding site was used to demonstrate the specificity of the immunoprecipitation. In addition, wild-type plants were compared to mutant plants lacking TGA2 and related factors TGA5 and TGA6 (7). Amplification of the input DNA indicates that the same amount of chromatin of wild-type and mutant plants was used for the immunoprecipitation. The threshold cycles are shown on an inverted scale. Threshold cycles above 28 are omitted from the scale, as they reflect unspecific amplifications as determined by control IP reactions with pre-immuneserum.

the leaves to the surface in order to make sure that the buffer, and not the air, is sucked in when the vacuum is released. 2. Put the device containing the leaf material into a beaker filled with CB1 so that the leaves are submerged. 3. Put the beaker in a desiccator and apply vacuum for 5 min using an oil pump. Release the vacuum, re-apply vacuum for an additional 5 min, release the vacuum and allow the crosslinking to proceed for 20 min (see Note 6). 4. Discard CB1 and wash the samples with CB2. Infiltrate CB2 into the leaves by applying vacuum for 5 min, release the vacuum and incubate the samples for another 5 min. 5. Discard CB2 and wash the samples twice with distilled water. After removing as much water as possible, the material can be frozen and stored in liquid nitrogen. 3.3. Isolation of Nuclei

1. All steps are carried out at 4°C. Working in a cooling chamber is recommended. 2. Grind the plant material in liquid nitrogen. 3. Transfer the leaf powder into 50-ml tubes filled with 20 ml of NEB and mix with a glass pipette until the powder is completely submersed in the buffer.

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4. Homogenize the sample with a tissue homogenizer at low power at 14,000 rev/min for 5 min. High-power mixers are not recommended as they will heat up the samples. 5. Filter the suspension through two sheets of Miracloth and collect the filtrate in 100-ml beakers (a small spoon can be used for careful mixing to accelerate the flow-through process). Add NEB to a total volume of 24 ml. 6. Add 1 ml of Triton X-100 (25%) dropwise to the filtrate on a magnetic stirrer. This step serves to lyse the organelles while leaving the nuclei intact. Add the Triton slowly to avoid high local concentrations of the detergent, which might lead to disruption of the nuclear envelope. 7. Continue stirring for at least 30 min (this step can be extended up to 2 h). 8. In the meantime, prepare the Percoll cushions in fresh 50-ml tubes. Overlay 6 ml of Percoll-cushion (75%) carefully with 6 mL of Percoll-cushion (35%). 9. Carefully place the lysate (Step 5) on top of the Percoll cushions. 10. Centrifuge at 2,100× g and 4°C for 30 min in a swing-out bucket rotor (soft start and stop). The nuclei should appear as a greyish layer at the interface between the cushions. 11. Recover the nuclei with a blue tip (1 ml), transfer them into a fresh 50-ml tube and add GB to a total volume of 20 ml. This step might work better when cut tips are used to recover the nuclei. 12. Add 6 ml Percoll-cushion (35%) to fresh tubes. 13. Overlay the Percoll-cushion (35%) with the resuspended nuclei. 14. Centrifuge at 2,100× g and 4°C for 10 min in a swing-out bucket rotor (soft start and stop). 15. Discard the supernatant, resuspend the nuclei in 1 ml GB, and transfer them into 10-ml tubes. Take an aliquot to stain the nuclei with DAPI (see Note 7). The isolated nuclei appear under the fluorescent microscope as crescent shaped structures. 16. Centrifuge at 2,100× g and 4°C for 10 min (soft start and stop) and discard the supernatant. Nuclei can be stored for 1 day at −80°C. 3.4. Preparation of Chromatin

All preparation steps should be carried out at 4°C. 1. Resuspend the nuclei in 1 ml of SB, 0.5% SDS and add the protease inhibitor mix (1:100). Incubate under gentle agitation for 20 min at 4°C. Dilute (1:1) with 1 ml SB lacking SDS to get a final concentration of 0.25% SDS.

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2. Sonicate four times for 20 s with a power of 100 W using the MSE Soniprep 150 ultrasonic disintegrator (see Note 8). Cool the samples in ice/EtOH during and between the sonication pulses. If foaming is a problem, reduce the percentage of SDS in the sample. 3. Centrifuge at 11,200× g and 4°C for 15 min to separate the soluble chromatin from the debris of the nuclei. 4. Freeze aliquots of 200 µl in liquid nitrogen and store at −80°C. Take one aliquot of 50 µl for the quantification of the DNA content. 3.5. Quantification of DNA in Chromatin Samples

1. Add 200 µl SB, 0.25% SDS and 5 µl of Proteinase K (from a 20 mg/ml stock solution) to 50 µl of chromatin. 2. Incubate samples for 1 h at 37°C and 16 h at 42°C in a heating block to reverse the cross-links. Proteinase K treatment can also be done after the reversion of the cross-links. 3. Extract with 250 µl PCI, vortex rigorously for 30 s and separate the phases by centrifugation in a microcentrifuge (5 min at 16,000× g and room temperature (RT)). Transfer the supernatant to a fresh microcentrifuge tube. 4. Extract with 250 µl CI, vortex rigorously for 30 s and separate phases by centrifugation in a microcentrifuge (5 min at 16,000× g, RT). Transfer the supernatant to a fresh microcentrifuge tube. 5. Optional: Add 1 µl RNaseA (10 mg/ml) to the supernatant and incubate at RT for 15 min. 6. Add 1/10 volumes of 3 M Na-acetate, 1 µl glycogen (10 mg/ml) and 2 volumes of absolute ethanol, mix by inverting the tubes for –six to eight times and precipitate for 2–4 h at −80°C. 7. Pellet the DNA by centrifugation in a microcentrifuge (35 min at 16,000× g, 4°C) and wash with 800 µl 70% ethanol. 8. Pellet the DNA by centrifugation in a microcentrifuge (20 min at 16,000× g, RT) and discard the supernatant. Place the open microcentrifuge tube for 10 min at 37°C and resuspend the dried pellet in 50 µl water (ultrapure). 9. Measure the OD260 and analyse the size of the DNA fragments on a 1% agarose gel (see Fig. 17.2 and Note 8).

3.6. Immunoprecipitation of Protein DNA Complexes

1. Thaw chromatin samples (from Sect. 3.4) on ice (this will take some time). 2. Bring equal amounts of chromatin as measured by DNA content (15 µg) to a total volume of 200 µl with SB, 0.25% SDS and add 300 µl RIPA-F lacking SDS. 3. Incubate for 1 h with 5 µl pre-immune serum (PPI) with slow rotation at 4°C on a rotation platform.

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3000 2000 1000 500 300

Fig. 17.2. Four independent samples of purified DNA obtained from chromatin. 5 µl of DNA (see Sect. 2.5) was analysed by gel electrophoresis in a 1% agarose gel and subsequent staining with ethidium bromide. As a size marker, O’GenerulerDNA Ladder Mix from Fermentas, Burlongton, Canada, was loaded.

4. In the meantime, add 1 ml RIPA-F to 50 µl of 50% protein A-Sepharose beads. You will need 2 × 50 µl protein A-Sepharose beads per sample: first, the complexes associated with the PPI are captured, and afterwards the specific complexes are enriched with the immuneserum. To equilibrate the beads, let them sit for 15 min with slow rotation at 4°C. Centrifuge in a microcentrifuge at 2,000× g for 3 min, discard RIPA-F (works nicely with an insulin needle) and wash the beads once again with 1 ml RIPA-F (slowly rotate for 5 min at 4°C). Discard the supernatant like before and dissolve the beads with RIPA-F to get a 50% beads solution. 5. Add 50 µl of the equilibrated 50% protein A-Sepharose beads to the samples and incubate them for 1 h at 4°C with slow rotation. This step serves to remove complexes from the chromatin that interact with the PPI. 6. Centrifuge the samples in a microcentrifuge for 2 min at 16,000× g. Take 50 µl of the supernatant for later use as an input control (see Note 7) and use the remainder of the supernatant for the immunoprecipitation (see Note 8). 7. Add 400 µl of SB, 0.25% SDS to the input control samples. 8. Transfer the remainder of the supernatant into a fresh microcentrifuge tube, add the antibody (1–5 µl) and incubate for 2 h at 4°C with slow rotation (see Note 10). 9. Add 50 µl of the 50% protein A-Sepharose beads and let the samples rotate for additional 2 h at 4°C. 10. Centrifuge for 3 min at 2,000× g in a microcentrifuge and remove the supernatant with an insulin needle until the Sepharose is half-dry. 11. Wash the beads by adding 1 ml RIPA-F and let them rotate for 5 min at 4°C, centrifuge for 3 min at 2,000× g in a

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microcentrifuge and discard the supernatant with an insulin needle. Repeat this washing step for two more times. 12. Add 800 µl RIPA-F and transfer the samples into a fresh microcentrifuge tube. Incubate for 5 min at 4°C with slow rotation, centrifuge for 3 min at 2,000× g and discard the supernatant like before. 13. Add 150 µl EB to the beads, vortex rigorously for 30 s, centrifuge for 1 min at 16,000× g in a microcentrifuge, transfer the supernatant with a yellow tip and into a microcentrifuge tube filled with 150 µl 10 mM Tris-HCl, pH 7.4, repeat the elution and combine the samples. 14. Add 5 µl proteinase K (20 mg/ml; add also to input controls set aside in Step 7) and incubate samples for 1 h at 42°C. Reverse the cross-links for about 4 h at 65°C. 3.7. Purification of DNA for PCR

1. Extract with 450 µl PCI, vortex rigorously for 30 s and separate the phases by centrifugation in a microcentrifuge (5 min at 16,000× g, RT). Remove the supernatant with a blue tip while holding the microcentrifuge tube in an angle of about 45° and transfer the supernatant to a fresh microcentrifuge tube. 2. Add 450 µl CI, vortex rigorously for 30 s and separate phases by centrifugation in a microcentrifuge (5 min at 16,000× g, RT). Transfer the supernatant into a fresh microcentrifuge tube. 3. Optional: Add 1 µl RNase A (10 mg/ml) and incubate at RT for 15 min. 4. Add 1/10 volumes of 3 M Na-acetate, 1 µl glycogen (10 mg/ml) and two volumes of absolute ethanol, mix by inverting the tubes –six to eight times and precipitate for 2–4 h at −80°C. 5. Pellet the DNA by centrifugation in a microcentrifuge (35 min at 16,000× g, 4°C) and wash with 800 µl 70% ethanol. 6. Pellet the DNA by centrifugation in a microcentrifuge (20 min at 13,000× g, RT) and discard the supernatant completely. Dry the pellet for 10 min at 37°C in an open microcentrifuge tube. Dissolve IP DNA in 35 µl and input DNA in 175 µl of water (ultrapure). 7. Resuspend the DNA at 65°C for 15 min while shaking at low speed. The purified DNA should be stored at −20°C.

3.8. Analysis by PCR or Real-Time PCR

The analysis of ChIP experiments by real-time PCR (qPCR) is recommended. If the PCR products are analysed by agarose gel electrophoresis, different numbers of cycles have to be run for each sample to ensure that the reaction is in the linear range. The conditions for the PCR depend on the primers that are used

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for amplification of the promoter DNA sequence. It is recommended to use primers that amplify a fragment of about 250 bp (see Note 11). 1. For qPCR, use the following protocol as a start; some optimizations might be necessary depending on the sequence of the primers: Use 2.5 µl each of the purified IP-DNA and the input DNA. For a typical pipetting scheme see Table 17.1. 2. A typical qPCR program is 6 min at 95°C (+2 min when no fluoresceine measurement is performed) 40 cycles of 20 s at 95°C, 20 s at 60°C, 28 s at 72°C 4 min at 72°C 1 min at 95°C 1 min at 55°C This program should be followed by a melting curve measurement to make sure that the primers have amplified a specific product. The initial denaturation time (in this case 6 min) depends on the polymerase that is used. The annealing temperature has to be adapted to the primers used. Threshold cycles, which define the beginning of the exponential phase of the PCR, are used as units to indicate the relative amount of DNA in the samples (see Fig. 17.1).

Table 17.1 Pipetting scheme for a real-time PCR Stock

Final conc.

µL

Water

—

—

Xa

Buffer

10 x

1x

2.5

dNTPs

100 µM

10 µM

0.25

1. Primer

10 mM

0.25 µM

0.625

2. Primer

10 µM

0.25 µM

0.625

SybrGreen 1:1000

10 x

0.1 x

0.25

MgCl2

1M

Xa

Xa

Taq polymerase

5 U/µl

1.25 U

0.25

Fluoresceine

1x

1x

0.25

DNA Final volume

2.5 25 µl

a The MgCl2 concentration to be used depends on the type of Taq polymerase. We usually use Immolase polymerase mix from Bioline (Bioline, Luckenwalde, Germany). For Immolase, a concentration of 4 mM provides good results. The reaction buffer is usually supplied with the enzyme. SybrGreen is from Cambrex Bio Science Rockland Inc., Rockland, ME, USA.

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4. Notes 1. Take special care when working with formaldehyde. It is highly toxic and has to be disposed separately. All work should be done under a hood. Wear protection clothes all the time. 2. It is recommended to prepare NEB and GB freshly for each preparation. Add the β-mercaptoethanol just before you start the experiment. Work carefully with β-mercaptoethanol as it is harmful and causes damage to the environment. Waste should be disposed separately. 3. The use of NaOH instead of KOH to adjust the pH of the RIPA-F buffer is recommended as potassium forms precipitates with SDS in the buffer. RIPA-F contains a combination of denaturing and non-denaturing detergents (Triton, DOC, SDS) to solubilize the chromatin. 4. Short-day conditions are 8/16 h light/dark period, 22°C. Grow the plants on soil with ~20 plants per pot. It is also possible to grow the plants under long-day conditions, but the fraction containing the nuclei might be less clean as the cells accumulate starch under long-day conditions. 5. One possibility is to use nylon stockings. 6. The incubation time with formaldehyde depends on the proteins that should be cross-linked. Shorter periods (of about 10 min) are sufficient for nucleosomal proteins; for cross-linking other proteins, the tissue should be treated for 20 min to 1 h. The cross-linking should not be extended to longer periods as proteins get denatured or masked by formaldehyde. If problems (low yield of chromatin, or low yield of IP-DNA) occur, a time-course experiment should be performed. 7. As DAPI (4,6-diamidino-2-phenylindole) moves slowly through membranes, an incubation time of 1.5 h for the samples (at 4°C, in the dark) is recommended. The excitation wavelength is 358 nm; emitted light has a wavelength of 461 nm. Emission at 400 nm is due to binding of DAPI to RNA and should be subtracted by an appropriate filter. 8. The shearing of the chromatin by sonication should be controlled by gel electrophoresis on a 1% agarose gel after the precipitation of the DNA. The fragments should appear as a smear from 2 kb down to 300 bp (see Fig. 17.2). Depending on the sonication device, the power and the length and number of pulses need to be optimized. 9. This so called “cleaning” step with the pre-immune serum is strongly recommended when an antiserum is used instead of a purified antibody.

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10. The performance of the ChIP largely depends on the antibody that is used. Some optimization is necessary to determine the best concentration for each antibody or antiserum. 11. We perform real-time PCR quantification using the SYBR Green technology in a MyiQ Single-Color Real-Time PCR Detection System (Bio-Rad, Munich, Germany). The SYBR Green stock is from Cambrex Bio Science, Rockland, USA, and the Immolase DNA polymerase mix and the reaction buffer are from Bioline, Randolph, USA. Fluoresceine is added to calibrate for equal efficiency of fluorescence in each well.

References 1. Orlando, V. (2000) Mapping chromosomal proteins in vivo by formaldehyde-crosslinkedchromatin immunoprecipitation. Trends Biochem. Sci. 25, 99–104. 2. Thibaud-Nissen, F., Wu, H., Richmond, T., Redman, J.C., Johnson, C., Green, R., Arias, J. and Town, C.D. (2006) Development of Arabidopsis whole-genome microarrays and their application to the discovery of binding sites for the TGA2 transcription factor in salicylic acid-treated plants. Plant J. 47, 152–162. 3. Wang, H., Tang, W., Zhu, C. and Perry, S.E. (2002) A chromatin immunoprecipitation (ChIP) approach to isolate genes regulated by AGL15, a MADS domain protein that preferentially accumulates in embryos. Plant J. 32, 831–843. 4. Rochon, A., Boyle, P., Wignes, T., Fobert, P.R. and Despres, C. (2006) The Coactiva-

tor Function of Arabidopsis NPR1 Requires the Core of Its BTB/POZ Domain and the Oxidation of C-Terminal Cysteines. Plant Cell 18, 3670–1385. 5. Geisberg, J.V. and Struhl, K. (2004) Quantitative sequential chromatin immunoprecipitation, a method for analyzing cooccupancy of proteins at genomic regions in vivo. Nucleic Acids Res. 32, e151. 6. Kuo, M.H. and Allis, C.D. (1999) In vivo cross-linking and immunoprecipitation for studying dynamic Protein:DNA associations in a chromatin environment. Methods (San Diego, Calif.) 19, 425–433. 7. Zhang, Y., Tessaro, M.J., Lassner, M. and Li, X. (2003) Knockout analysis of Arabidopsis transcription factors TGA2, TGA5, and TGA6 reveals their redundant and essential roles in systemic acquired resistance. Plant Cell 15, 2647–2653.

Chapter 18 Fluorescence-Based Electrophoretic Mobility Shift Assay in the Analysis of DNA-Binding Proteins Sebastian Steiner and Thomas Pfannschmidt Abstract Changes in gene expression mediated by DNA-binding protein factors are a crucial part of many signal transduction pathways. Generally, these regulatory proteins are low abundant and thus their purification and characterisation is labour- and time-intensive. Here we describe a workflow for purification, characterisation and identification of DNA-binding proteins. We show the use of a fluorescence-based electrophoretic mobility shift assay (fEMSA) and describe its advantages for a rapid and convenient screening for regulatory cis–elements. This involves a crude enrichment of nucleic acid binding proteins by heparin–Sepharose chromatography and the characterisation of fractions using overlapping fluorescence-labelled DNA probes spanning the promoter region of interest. The determined protein-binding sites can then be used for sequence-specific DNA-affinity chromatography to purify specifically interacting proteins. Finally, the DNA-binding complexes can be characterised and identified using two-dimensional EMSA, UV-cross-linking and mass spectrometry. Key words: DNA-binding proteins, transcription factors, fluorescence-EMSA, 2D-EMSA, gel shift, DNA-affinity chromatography, UV-cross-linking.

1. Introduction Cellular responses of plants to many biotic and abiotic factors often involve changes in transcription. Key elements in their regulation are DNA-binding protein factors, which mediate enhancement or inhibition of basal transcription. They operate in response to up-stream signalling pathways by binding to distinct cis-elements close to or within promoters. Furthermore, such

T. Pfannschmidt (ed.), Plant Signal Transduction, vol. 479 Humana Press, a part of Springer Science + Business Media, LLC 2009 DOI: 10.1007/978-1-59745-289-2_18

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factors can interact with the transcription apparatus or induce changes in the topology of DNA. A common method to study nucleic-acid–protein interactions is the electrophoretic mobility shift assay (EMSA), first described for the lac-repressor by Fried and Crothers in 1981 (1). Later on, it was also applied to study RNA–protein and protein–protein interactions (2,3). The method is based on the retarded mobility of protein-bound DNA fragments when compared with the same but free DNA after separation on a non-denaturing gel. Thus, the interaction of a protein with a DNA fragment can be observed as an additional band in the gel (see Fig. 18.1a). This principle allows in vitro characterisation of DNA-binding properties of transcription factors with respect to specificity, binding region, ionic milieu, influences of post-translational modifications and kinetics of binding constants. Due to its high sensitivity it can be also used to resolve DNA–protein complexes of different stoichiometries or conformations (4).

Fig. 18.1. (a) Principle of the electrophoretic mobility shift assay (EMSA). Depicted is a schematic EMSA gel loaded as follows: Lane 1: free DNA probe (F), Lane 2: protein extract (P) + DNA probe; By binding of proteins the mobility of the DNA probe is retarded; Lane 3: protein extract (P) + DNA probe + unlabelled competitor-DNA (C); Unspecific signals disappear whereas remaining signals are sequence-specific; Lane 4: as Lane 3 + antibody against binding protein (a); the DNA–protein–antibody complex exhibits a more reduced mobility in the gel than the DNA–protein complex alone (“supershift”). This indicates the involvement of a known binding factor within a DNA–protein complex. (B) Schematic example for “probe walking”. By choosing overlapping DNA probes spanning cis-elements of interest, fEMSA can be used to find binding sites and gives information for further enrichment of trans-acting factors. (C) Workflow scheme for the purification of sequence-specific DNA-binding proteins within e.g. chloroplasts.

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The EMSA is suitable to study known binding factors and novel factors within crude protein extracts. Thus, it provides a tool to monitor the purification process of a protein of interest. By combining EMSA with interference or mutational assays one can determine the exact binding site and its essential nucleotides. The additional use of antibodies helps to identify involved protein factors, by causing a so called “supershift” (an additional retardation of the DNA–protein complex). This supershift is caused by the increase in size of the protein complex on specific protein recognition and binding of the antibody (see Fig. 18.1a). However, the approach can also lead to an abolishment of the complex depending on the antibody binding site, its affinity and protein concentration. A number of different applications of this method have been published (5). Usually, the DNA probe is radioactively labelled with 32P using T4 polynucleotide-kinase or Klenow fragment. In addition, there also exist other labelling or detection systems like staining with EtBr, Biotin/Hapten/Dig-labelling and others. These procedures exhibit a number of disadvantages, like handling of radionucleotides, long exposition times, short half-lives of isotopes or time-consuming blotting of DNA–protein complexes followed by detection through chemiluminescent techniques or a low sensitivity. Here we describe a rapid and easy modification of the EMSA principle using fluorescent DNA probes that allow a convenient and fast analysis of putative regulatory DNA sequences and their binding proteins, which is applicable also to high throughput screening approaches. In contrast to radiolabelled probes fluorescence-labelled probes are non-hazardous and stable for a long time. Their detection is rapid and can be done directly in the gel within the glass plates. No time-consuming gel drying and exposition to autoradiography films are necessary. In addition, one can scan gels and run it further on to increase resolution if necessary. The sensitivity is very high (down to pg) whereby the signal is linear in a broad range, which is ideal for quantification (6). By using dyes with different fluorescence one can add loading controls in the same gel lane or one can perform competition, interference or mutational assays in a single sample. Another advantage is that the fluorophores are not susceptible for cleavage by phosphatases like probes end-labelled with 32P. Most fluorophores are also suitable for UV-cross-linking, allowing direct determination of the molecular weight of a DNA-binding protein by SDS-PAGE. In this case one can also scan the gel directly in the glass plates before staining the proteins. Due to the stability of most fluorophores, dried gels can be usually also scanned at later time points. These properties make fluorescence-based EMSA a real advancement in the investigation of nucleic acid binding proteins. Furthermore, it is possible to scan easily putative regulatory sequence elements

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(cis-elements) like promoter regions for trans-acting factors by “probe walking” (see Fig. 18.1b). Such regulatory sequences can be found by primer extension analyses in the case of promoters or by deletion analyses with promoter::reporter gene constructs. There are also prediction tools available for prediction of transcription factor binding sites (7–9). EMSAs are also a useful tool to develop purification strategies for identification of novel nucleic acid binding proteins (see Fig. 18.1c). Here we describe the workflow for identification of DNAbinding proteins in chloroplasts as an example; however, the procedure is applicable to protein extracts from other compartments of plant cells as well. This includes crude enrichment of nucleic acid binding proteins by heparin–Sepharose chromatography followed by characterisation of such fractions by fluorescence-based EMSA (fEMSA). We use the LI-COR Odyssey Infrared Laser Imaging System for detection of the IRDye 800 and DY-781 end-labelled DNA probes, but the protocol in general is also applicable to other dyes and fluorescence detection systems.

2. Materials 2.1. Heparin–Sepharose Chromatography

1. XK 16 column (GE Healthcare). 2. Heparin–Sepharose (GE Healthcare). 3. Peristaltic pump; e.g., P1 (GE Healthcare). 4. Column buffer A; 50 mM Tris-HCl pH 7.6, 0.1 mM EDTA, 1 mM 2-mercaptoethanol or DTT, 0.1% (v/v) Triton X-100, 80 mM (NH4)2SO4, 10 mM MgCl2, 10% (v/v) glycerol. Store at 4°C. 5. Elution buffer, column buffer A but 1.2 M (NH4)2SO4. Store at 4°C. 6. ZelluTrans T1 Dialysis membrane MWCO 4000–6000 (Roth). Wash three times with ddH2O before use. 7. Dialysis buffer 50; 50 mM Tris-HCl pH 7.6, 0.1 mM EDTA, 1 mM 2-mercaptoethanol or DTT, 0.1% (v/v) Triton X-100, 50% (v/v) glycerol. Pre-cool to 4°C. 8. Lowry Protein Assay Kit (Bio-Rad).

2.2. fEMSA (fluorescence Electrophoretic Mobility Shift Assay)

1. Electrophoresis unit; e.g. Hoefer SE 600 (GE Healthcare). 2. Infrared Laser-Imaging System; e.g., Li-Cor Odyssey Infrared Imaging System. 3. End-labelled fluorescent oligonucleotides with the desired binding site; IRDye 700, IRDye 800, DY-681, DY-781 or other compatible dyes (MWG-Biotech; biomers.net).

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4. TE buffer; 10 mM Tris-HCl pH 7.6, 1 mM EDTA. 5. Forty percent polyacrylamide (29:1) (Roth). Store at 4°C. Acrylamide is toxic, so handle with care. 6. 3 X EMSA gel buffer; 1.5 M Tris-HCl pH 8.85. 7. Ten percent ammonium persulfate; prepare 10% solution in ddH2O, and store in aliquots at −20°C. 8. N,N,N′,N′-tetramethylethylenediamine (TEMED); store at 4°C. 9. 10 X native running buffer; 250 mM Tris, 1.92 M glycine. 10. 10 X binding buffer; 300 mM Tris-HCl pH 7, 50 mM 2-mercaptoethanol, 5 mM EDTA; store at 4°C. 11. Poly (dIdC) . poly (dIdC) (GE Healthcare). Dissolve in ddH2O and adjust concentration to 1 µg/µl; store at −20°C. 12. 500 mM MgCl2 13. Colour marker; 600 mM Tris-HCl pH 6.8, 0.4% bromophenol blue, 50% glycerol. 2.3. DNA Affinity Chromatography

1. K9 column (GE Healthcare). 2. CNBr-activated Sepharose 4B (GE Healthcare). Cyanbromide is toxic, so handle in a fumehood. 3. (5′-NH2-) oligonucleotides or PCR fragment with desired binding site (min. 50 nmol/g Sepharose). 4. Coupling buffer; 100 mM NaHCO3, 500 mM NaCl. 5. Blocking buffer; 100 mM Tris-HCl pH 8 or 1 M ethanolamine. 6. Washing buffer I; 1 mM HCl pH 3. 7. Washing buffer II; 100 mM natriumacetate/acetate pH 4, 500 mM NaCl. 8. Washing buffer III; 100 mM Tris-HCl pH 8, 500 mM NaCl. 9. Column buffer A (see Sect. 2.1). 10. Elution buffer (see Sect. 2.1). 11. Dilution buffer; 50 mM Tris-HCl pH 7.6, 0.1 mM EDTA, 1 mM 2-mercaptoethanol or DTT, 0.1% (v/v) Triton X-100. Pre-cool to 4°C. 12. Centricon YM-10 (Millipore).

2.4. 2D-EMSA

1. Electrophoresis unit; e.g. Hoefer SE 660 (GE Healthcare) 2. Forty percent polyacrylamide (29:1) (Roth) 3. 3 X stacking buffer; 0.5 M Tris-HCl pH 6.8, 0.4% SDS 4. 3 X separation buffer; 1.5 M Tris-HCl pH 8.8, 0.4% SDS

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5. 10 X SDS running buffer; 250 mM Tris, 1.92 M glycine, 1.5% SDS 6. Denaturation buffer; 75 mM Tris-HCl pH 6.8, 2.5% SDS, 3% β-mercaptoethanol, 2% glycerol 7. 5 X SDS sample buffer; 250 mM Tris-HCl pH 6.8, 20% glycerol, 10% SDS, 5% 2-mercaptoethanol, 0.1% bromophenol blue 8. Protein molecular weight marker; e.g., Sigma Wide Range Marker (Sigma-Aldrich) 9. 1% agarose in 1 X SDS running buffer 2.5. UV-Cross-Linking

1. 3 UV-lamps; e.g. HL-6-KM (Bachofer Laborgeräte) 2. Saran wrap (Roth)

3. Methods 3.1. Heparin– Sepharose Chromatography (HSC)

Heparin–Sepharose chromatography is a widely used technique for crude enrichment of nucleic acid binding proteins, and also for other proteins. Heparin works as an anion exchanger, but it also resembles the phosphodiester-backbone of DNA. As an example, here we use chloroplast lysates, prepared according to (10). However, the protocol should be easy to alter for other preparations like nuclei or mitochondria. 1. Wash column material (solid: 3.5 g, slurry: ~20 ml) and pack column according to manufacturer’s recommendations to ~12 ml bed volume. Pre-cool to 4°C (see Note 1). 2. Equilibrate column with column buffer. 3. Adjust lysate to 80 mM (NH4)2SO4 (and 10 mM MgCl2). 4. Subject lysate (approx. 60 mg protein) to the column and adjust flow rate to 500 µl/min (see Note 2). 5. After the lysate is passed, wash extensively with at least 5–10 bed volumes of column buffer (see Note 3). 6. Elute the proteins with elution buffer and take samples. Sanitise and store column material according to manufacturer’s recommendations. 7. Dialyse the fractions overnight at 4°C against dialysis buffer 50. 8. Determine protein concentration with a Lowry Assay Kit. Pool the peak fractions and store aliquots at −20°C.

3.2. fEMSA (fluorescence-EMSA)

Here we describe the workflow for a fluorescence-based gel shift assay. This technique can be used to rapidly scan DNA regions for

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binding of proteins and their binding characteristics (see Fig. 18.2). By using overlapping DNA probes spanning the region of interest one can roughly characterise binding sites, which can be subsequently used for sequence-specific DNA-affinity chromatography. 3.2.1. Preparation of a Fluorescent DNA Probe

Order commercially available fluorescence-labelled oligonucleotides (e.g., with IRDye700 or 800, DY-681, DY-781, Cy5.5 or Alexa Fluor 750 or other dyes compatible to your imaging system) spanning the region of interest and the unlabeled antisense oligonucleotide. 1. Mix equimolar amounts of the complementary oligonucleotides solved in ddH2O or TE buffer. 2. Heat up to 95°C for 1 min. 3. Allow the mixture to cool down slowly to RT. 4. Adjust DNA concentration to approx. 10 ng/µl using a photometer.

Fig. 18.2. Grey-scaled infrared image of a fluorescence-EMSA with mustard chloroplast heparin–Sepharose fractions (18 µg) and five different DY-781-labelled DNA probes (Lane 1–5, 10 ng) spanning the psbA-promoter region (“probe walking”). The samples were mixed and incubated according to the protocol described in Sect. 3.2.3 and loaded onto a 6% non-denaturing polyacrylamide gel. After the electrophoresis, the gel was scanned between the glass plates using a Li-Cor Odyssey Laser scanner. The scanning parameters were as follows: channel: 800 nm, focus height 3.5 mm, intensity 5. Free DNA is recognised at the bottom of the gel; several DNA protein complexes appear in the middle of the gel and a high molecular protein complex binding the probe at the top of the gel represents the plastid encoded RNA polymerase complex (PEP) (14). Beside the prominent signals several weaker signals appear. Areas with saturated signals (e.g. free DNA) appear white.

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3.2.2. Preparation of a Native Polyacrylamide Gel

Here we describe the casting of a native midi-gel for a Hoefer SE 600 electrophoresis unit. For other electrophoresis units adjust volumes accordingly. Gel concentration must be chosen with respect to the length of the probe or the size of the DNA–protein complex. Five percent gels are a good starting point for typical oligonucleotides of 30–50 bp in size. For larger probes or large protein complexes, use lower percentage gels or reduce cross-linking of polyacrylamide by decreasing the bisacrylamide concentration. 1. Prepare a 5% gel (1.5 mm thick) by mixing 3.12 ml 40% acrylamide/bisacrylamide (29:1), 8.33 ml of 3 X gel buffer and 13.55 ml of ddH2O. Add 150 µl 10% ammoniumpersulfate and 150 µl TEMED. Mix and pour between the glass plates. Add a comb. The gel should polymerise within 5 min. 2. Rinse the wells with 1 X native running buffer after removing the comb. 3. Equilibrate gel, running buffer and electrophoresis unit to the temperature at which you want to run the gel. Pre-run the gel with 30 mA for 15–30 min to remove remaining APS and TEMED.

3.2.3. Binding Reaction and Gel Run

DNA–protein interactions are affected by many biophysical and physiological parameters. Therefore, it is mandatory to titrate the best binding conditions for every protein. 1. Mix the binding reaction according to Table 18.1. 2. Incubate at RT for 15–30 min. 3. Load the sample on the prepared native gel (see Note 8). 4. Add 5 µl of colour marker in a free lane to make the run observable (see Note 9).

Table 18.1 Setup of a typical EMSA sample (see Note 7) Substance

Volume

Final concentration

10 X binding buffer

5 µl

1X

500 mM MgCl2 (see Note 4)

1 µl

10 mM

1µg/µl poly(dIdC) • poly(dIdC) (optional) (see Note 5)

1 µl

20 ng/µl

DNA probe (20 ng/µl)

1 µl

0.4 ng/µl

Protein sample (see Note 6)

15–30 µl

—

ddH2O

fill up to 50 µl

—

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5. Run the gel at 4°C with 30 mA or overnight at 7 mA until the blue colour reaches the bottom of the gel (see Note 10). 3.2.4. Scanning the Gel

1. When the gel run is finished dry and clean the outside of the glass plates. 2. Place the gel within the glass plates on the scanning screen of the fluorescence imager. 3. Adjust the scanning focus to the height of the gel between the glass plates. Adjust scanning channels, intensities and scanning time. 4. Start scanning.

3.3. Preparation of DNA Affinity Column Media

After screening possible regulatory DNA sequences by fEMSA, protein-binding sites might be identified. By using tailor made sequence-specific DNA-affinity chromatography it is possible to further enrich such interacting factors.

3.3.1. Hybridisation of the DNA Ligand

For easy and rapid purification of sequence-specific DNA-binding factors, use synthesised oligonucleotides containing the desired binding site. To maximise the probability of terminal coupling of the oligonucleotides one can use commercially available 5′-NH2oligonucleotides. One can also couple purified PCR products or cloned fragments. To increase the amount of coupled binding sites use concatemeric fragments. 1. When using synthetic oligonucleotides mix equimolar amounts of both complementary DNA strands in ddH2O or coupling buffer. When using PCR products precipitate and solve them in coupling buffer. Fill up with coupling buffer to 5 ml/g Sepharose used. Do not use Tris-buffered solution or anything else containing primary amines (see Note 11). 2. Incubate the mixture at 95°C for a few minutes in a thermomixer or water bath. 3. Allow the mixture to cool down slowly to RT.

3.3.2. Preparation of the DNA Affinity Column Media

This step describes the coupling of the DNA ligand to CNBractivated Sepharose. 1. Incubate 1 g of CNBr-activated Sepharose with 50 ml of 1 mM HCl (pH 3). The material swells to a volume of 3–4 ml/g. 2. Remove the liquid using a filter and a vacuum pump. Repeat Step 1 and 2 two times to wash away additives/contaminants and activate the coupling group. 3. Dissolve the hybridised DNA in 5 ml coupling buffer per gram of Sepharose (see Sect. 3.3.1). 4. Incubate the washed and activated Sepharose with the coupling solution (ligand in 5 ml coupling buffer per gram of

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Sepharose) for 1–2 h at RT or overnight at 4°C. Rotate the mixture or apply any other gentle stirring. 5. Remove the liquid and wash two times with five volumes coupling buffer to remove unbound DNA. Determine the DNA concentration in the collected wash samples and calculate coupling efficiency. 6. Block all remaining active groups by incubation with blocking buffer or 1 M ethanolamine for at least 2 h at RT. 7. Wash the resin with alternating pH. Add wash buffer II (pH 4). Remove the solution and wash with wash buffer III (pH 8). Repeat this three times. 8. For use, equilibrate the resin with column buffer A or store it in column buffer A or 20% ethanol at 4°C. 3.3.3. Column Packing

1. Column media should have the temperature at which the chromatography will be performed (usually 4°C). 2. Equilibrate with one bed volume of column buffer A. Degas the media. 3. Flush all column parts (e.g., K9 Column (GE Healthcare)) with buffer to remove remaining air bubbles. 4. Fill the column with the degassed media in one step and adjust the desired flow rate. 5. Refill column buffer A until all material is settled down.

3.4. DNA-Affinity Chromatography (DAC)

Here we describe the enrichment of sequence-specific DNAbinding proteins with DNA-affinity chromatography using immobilised oligonucleotides containing the desired binding site found with fEMSA (see Note 12). We use heparin–Sepharose chromatography fractions (see Sect. 3.2) for further purification. The dialysed fractions need to be diluted to decrease viscosity by adding four volumes of dilution buffer. This adjusts the properties such as in column buffer A. For subsequent DNA-affinity chromatography without storage of heparin–Sepharose protein fractions, you can dialyse against dialysis buffer containing only 10% glycerol. 1. Adjust the protein solution to 80 mM (NH4)2SO4 and 10 mM MgCl2. Add an unspecific competitor repressing unspecific binding to a final concentration of 50 µg/ml. 2. Load the sample on the column and adjust a flow rate of 500 µl/min. 3. After passing of the sample wash the column with 5–10 total volumes of column buffer A. 4. Elute the binding fraction by applying 4–5 bed volumes of elution buffer. Discard a half bed volume and then take

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1–2 ml fractions until elution buffer runs out. Sanitise and store column material according to manufacturer’s recommendations. 5. Concentrate the collected sample and remove excessive salt by dialysis or centrifugation in Centricon cells and adjusting to dialysis buffer conditions, respectively. Store protein fractions at −20°C. 3.5. Two-Dimensional EMSA (2D EMSA)

After enrichment of specific DNA-binding proteins, one can visualise them by common SDS-PAGE and silver staining. To characterise different binding complexes occurring on the cis-element perform two-dimensional EMSA resolving the composition of different binding complexes in a second denaturing gel (11). In this technique in a first dimension fEMSA is done as explained above. The obtained signals or even the whole gel lane can then be excised and loaded onto a SDS-PAGE (see Fig. 18.3) as a second dimension (see Note 13). 1. Prepare a standard 1.5 mm 10% Laemmli SDS-polyacrylamide gel (Hoefer SE 660, 18 × 24 cm glass plates) ensuring that space remains above the stacking gel for gel pieces or lanes from the first dimension gel (see Note 14). 2. After scanning the fEMSA gel adjust contrast, intensity and gamma of the image and print it on the same scale as the original gel. 3. Place the printed image under the glass plates and remove the upper plate. 4. Excise the gel area of interest or the whole lane with a clean, sharp scalpel. 5. Denature and equilibrate the proteins in the cut out gel by incubating it in denaturation buffer for 5 min at RT. 6. Flush the gel piece with 1 X running buffer to remove excess 2-mercaptoethanol. 7. Load the gel piece onto the stacking gel of the second dimension gel. Remove all air bubbles under the gel piece (see Note 15). 8. Pipette a protein molecular weight marker onto a small piece of filter paper and place it beside the gel piece on the stacking gel. 9. Fix the gel piece and marker with 1% agarose in 1 X SDSrunning buffer. 10. Run the gel with 30 mA or overnight with 7 mA. 11. After the gel run scan the gel as described in Sect. 3.2.4 to visualise the DNA probe released from the protein–DNA complexes (see Fig. 18.3, bottom panel).

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12. Stain the gel with silver nitrate or Coomassie R 250 (see Note 16). 13. Compare the obtained protein spot pattern with corresponding protein–DNA complexes obtained by the first dimension fEMSA and the released DNA probe. Subtract background by comparison with the negative control without the DNA probe. Shifted protein spots or proteins above the corresponding signals of released DNA are promising candidates for interacting proteins (see Fig. 18.3).

Fig. 18.3. Principle of two-dimensional EMSA (2D-EMSA) and its advantage to separate DNA–protein–complexes of different protein composition and stoichiometries. An example with commercially available E. coli RNA polymerase core enzyme (Sigma) and a fluorescence-labelled psbA-core promoter fragment (15) is shown. After incubation of 1.5 µg E. coli RNA polymerase core enzyme with 20 ng of the labelled psbA-promoter fragment (see Sect. 3.2.3), the DNA–protein complexes were separated by mass and shape in first dimension (upper gel) by a native polyacrylamide gel (4.5%) and detected by imaging of the fluorescence-labelled DNA probe. The free DNA and two different complexes (1 and 2) are visible (top panel). The whole gel lane was cut out, denatured and applied onto a second denaturing SDS gel (10%) (see Sect. 3.5). The DNA–protein complexes were degraded and its components were separated by mass in a second dimension. The former bound DNA probe was released and can be detected at the bottom of the gel through fluorescence imaging (in-gel scan, bottom panel). Silver staining of the SDS gel visualised the proteins (middle panel) and thus the composition of the DNA–-protein complexes can be identified. In the case of the known E. coli RNA polymerase complex 1 corresponds to the α-, β-, β¢- and σ-structure (holo-enzyme) whereas complex 2 lacks the σ-subunit (core-enzyme) and thus has a higher mobility in the first dimension (16).

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14. Excise protein spots and perform in-gel digestion and MS analysis (12) or other protein identification procedures (N-terminal sequencing). UV Cross-Linking

UV-cross-linking is a well-established method to estimate the molecular weight of a DNA interacting factor and thus is very helpful for identification of such proteins even within crude extracts (13). In the first step the DNA–protein complexes are irradiated with UV light, leading to covalent bonds between the DNA and binding proteins that are in close vicinity. This also works with fluorescence-labelled DNA probes (see Fig. 18.4), but needs no time-consuming autoradiography. After the UV-crosslink the DNA-binding proteins can be separated on a denaturing SDS gel. Scanning the gel with an infrared imager indicates the location of the interacting protein by fluorescence of the crosslinked DNA probe (see Note 17). The binding protein can then be associated to a band after staining of the gel. 1. Set up a fEMSA sample as described in Table 18.1. Include negative controls, containing just the DNA probe or protein sample without the respective other. Also, show specificity with Proteinase K or DNase treatment. 2. After 30 min of incubation, cover the open tube with Saran wrap to prevent evaporation. 3. Place the tubes in a rack that is open on both sides. 4. Place 3 UV lamps (e.g. 3 bulbs providing 80 mW/cm2 254 nm UV light) above the tubes and one at each side of the rack. The distance between the lamp and sample should not exceed 5 cm. 5. Irradiate the samples with 254 nm UV light for 5 min to 3 h. The irradiation increases the temperature within the sample. Therefore, cool the samples with ice or irradiate in a cool room. Set up a time course to evaluate the best cross-link conditions for the protein of interest (see Note 18). 6. After irradiation denature the proteins by adding SDS-sample buffer and heating at 95°C for 5 min. 7. Load the samples on a standard 10% SDS-polyacrylamide gel and run the gel. 8. After the gel run, clean and dry the outside of the glass plates and place the gel within glass plates on the scanning screen of the fluorescence imager. Adjust the scanning focus to the height of the gel between the glass plates. Adjust scanning channels, intensities and scanning time. Start scanning. 9. After scanning stain the gel with silver or Coomassie R250. 10. Compare the obtained fluorescence signals with corresponding protein bands on the stained gel to identify DNAbinding proteins.

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Fig. 18.4. UV-cross-linking of fluorescence-labelled DNA probes with binding proteins. A grey-scaled infrared image (800-nm channel) of an SDS-PAGE gel (10%), loaded with different samples of UV-cross-linked E. coli RNA-polymerase core enzyme (Sigma) with DY-781 fluorescence-labelled psbA-promoter fragment (see Sect. 3.6), is shown. All samples were denatured with SDS-sample buffer for 10 min at 95°C before loading the gel. Lane 1: psbA-promoter probe (20 ng) incubated without protein after 60 min of UV irradiation. Lane 2: 3 µg E. coli RNA-polymerase core enzyme (Sigma) after 60 min of UV irradiation. No fluorescence is visible. Lane 3: mock control. About 20 ng of the probe was incubated with 3 µg of the RNA polymerase and loaded onto the gel without UV treatment. Lanes 4–7: Samples with the same protein content as in lane 3 but were irradiated with UV light (254 nm) for 10, 20, 30 and 60 min, respectively, before denaturing and loading onto the gel. Bands represent proteins coupled to the fluorescent DNA probe by UV light (compare (17)). Longer irradiation causes cross-linking of more binding subunits leading to signals representing high molecular weight protein–DNA complex (lane 5–7). Areas with saturated signals (e.g. free DNA) appear white.

4. Notes 1. It is highly important to avoid air bubbles in the column. Flush all parts with water before packing. Pre-cool all solutions and column material before packing. Degas the column material. Pack columns according to the manufacturer’s manual. 2. To prevent clogging of the column and aggregation of proteins or lipids use detergents. Centrifuge the lysate to remove insoluble ingredients. When the column flow rate decreases, use a peristaltic pump. If the column becomes clogged, be careful with the pump to avoid drying out the column by

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vacuum. In case of clogging, open the column, gently resuspend the column material and let it settle down again. 3. For chloroplast lysate the green colour is a good marker for washing. Rinse until column material is yellowish to white again. Alternatively, you can determine protein content in washing fractions using a UV monitor. Elute bound proteins when protein concentration reaches zero or a steady state. To remove more unspecific binding proteins raise ammoniumsulphate concentration up to 200 mM in the washing buffer. 4. In some cases it is useful to add a carrier protein like BSA. Some proteins require additional cofactors (Zn, second messenger etc.), other pH or a different ionic milieu. Titrate to find the optimal conditions. 5. To lower unspecific binding add unspecific competitors like poly (dIdC) ˙ poly (dIdC). Try different concentrations. To test sequence specificity of the obtained DNA–protein complexes, add an excess of the unlabelled DNA probe (10- to 100-fold). Sequence-specific DNA-binding proteins form complexes also in the presence of competitors. 6. The protein concentration should be 10–20 µg in crude protein extracts. However, this value depends on several aspects of the desired proteins like total concentration in protein extract, binding strength and specificity. 7. Always perform negative controls without a protein sample or DNA probe to check the background of the DNA probe or protein sample. Chlorophyll, often a contaminant in plant protein samples even after chromatography, exhibits fluorescence in the 700-nm channel of the Li-Cor Odyssey Infrared Imaging system. In this case use probes labelled with dyes giving fluorescence in the 800-nm channel like IRDye 800. 8. Because there is no stacking gel, the filling level determines more or less the signal broadness. To get sharp bands use concentrated protein samples and minimise the volume of the binding mixture or use thicker gels when scaling up. 9. Infrared Imaging Systems like Li-Cor Odyssey are able to detect bromophenol blue in the 700-nm channel. Choose Orange G loading dye when using the 700-nm channel. 10. When running the gel overnight use 1 ml 20% gel at the bottom as a stop gel to prevent leaking. 11. CNBr binds amines, preferentially primary amines. Therefore, it is necessary to prevent contact with amine-containing substances such as Tris-buffered solutions before blocking. 12. In some cases the purification result might be better when performing an unspecific DAC before a specific DAC. In this

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case use coupled calf thymus or salmon testis DNA instead of oligonucleotides. 13. For 2D EMSA always perform negative controls with all ingredients except the DNA probe in the mixture to have a reference for subtraction of the background. 14. Use a 7–17% gradient gel to increase resolution and to visualise big as well as small proteins in one gel. 15. The placing of the gel pieces onto the second dimension gel works better when you fill the space above stacking gel with 1 X running buffer. 16. Due to the native separation of proteins and the contamination with salts and DNA in the first dimension, the stained protein spots appear smeary and diffuse (see Fig 18.3). 17. For this purpose, do not use DNA probes longer than 50 bp. Otherwise the protein will show an altered mobility in the gel due to the mass increase by the bound DNA probe. Multisubunit cross-links are also problematic in interpretation. 18. Halogenated thymidin analogs such as bromodesoxyuridine (BrdU) have a higher photoreactivity and thus can increase cross-link efficiency. In some cases it might be useful to substitute thymidines with BrdU in the DNA probe. In this case, one can also irradiate with a longer wavelength (305 nm) preserving degradation of the proteins.

Acknowledgement This work was supported by grants of the Deutsche Forschungsgemeinschaft and the NWP of Thuringia.

References 1. Fried, M. and Crothers, D. M. (1981) Equilibria and kinetics of lac repressoroperator interactions by polyacrylamide gel electrophoresis. Nucleic Acids Res. 9, 6505–6525. 2. Setzer, D. R. (1999) Measuring equilibrium and kinetic constants using gel retardation assays. Methods Mol. Biol. 118, 115–128. 3. Erickson-Viitanen, S. and DeGrado, W. F. (1987) Recognition and characterization of calmodulin-binding sequences in peptides and proteins. Methods Enzymol. 139, 455–478.

4. Fried, M. G. (1989) Measurement of protein-DNA interaction parameters by electrophoresis mobility shift assay. Electrophoresis 10, 366–376. 5. Sambrook, J., Fritsch, E. F., and Maniatis, T. (1989) Molecular Cloning. A Laboratory Manual, Cold Spring Harbor Laboratory Press, Cold Spring Harbor, New York. 6. www.licor.com. 7. Kel, A. E., Gossling, E., Reuter, I., Cheremushkin, E., Kel-Margoulis, O.V., and Wingender, E. (2003) MATCH: A tool for searching transcription factor binding sites

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9.

10.

11.

in DNA sequences. Nucleic Acids Res. 31, 3576–3579. Matys, V., Fricke, E., Geffers, R., Gossling, E., Haubrock, M., Hehl, R., Hornischer, K., Karas, D., Kel, A. E., Kel-Margoulis, O. V., Kloos, D. U., Land, S., LewickiPotapov, B., Michael, H., Munch, R., Reuter, I., Rotert, S., Saxel, H., Scheer, M., Thiele, S., and Wingender, E. (2003) TRANSFAC: transcriptional regulation, from patterns to profiles. Nucleic Acids Res. 31, 374–378. Steffens, N. O., Galuschka, C., Schindler, M., Bulow, L., and Hehl, R. (2004) AthaMap: an online resource for in silico transcription factor binding sites in the Arabidopsis thaliana genome. Nucleic Acids Res. 32, D368–372. Tiller, K., Eisermann, A., and Link, G. (1991) The chloroplast transcription apparatus from mustard (Sinapis alba L.). Evidence for three different transcription factors which resemble bacterial sigma factors. Eur. J. Biochem. 198, 93–99. Pfannschmidt, T., Ogrzewalla, K., Baginsky, S., Sickmann, A., Meyer, H. E., and Link, G. (2000) The multisubunit chloroplast RNA polymerase A from mustard (Sinapis alba L.) - Integration of a prokaryotic core into a larger complex with organelle-specific functions. Eur. J. Biochem. 267, 253–261.

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12. Stauber, E. J., Fink, A., Markert, C., Kruse, O., Johanningmeier, U., and Hippler, M. (2003) Proteomics of Chlamydomonas reinhardtii light-harvesting proteins. Eukaryot. Cell 2, 978–994. 13. Chodosh, L. A., Buratowski, S., and Sharp, P. A. (1989) A yeast protein possesses the DNA-binding properties of the adenovirus major late transcription factor. Mol. Cell Biol. 9, 820–822. 14. Pfannschmidt, T., and Link, G. (1997) The A and B forms of plastid DNA-dependent RNA polymerase from mustard (Sinapis alba L.) transcribe the same genes in a different developmental context. Mol. Gen. Genet. 257, 35–44. 15. Homann, A., and Link, G. (2003) DNAbinding and transcription characteristics of three cloned sigma factors from mustard (Sinapis alba L.) suggest overlapping and distinct roles in plastid gene expression. Eur. J. Biochem. 270, 1288–1300. 16. Severinova, E., Severinov, K., Fenyo, D., Marr, M., Brody, E. N., Roberts, J. W., Chait, B. T., and Darst, S. A. (1996) Domain organization of the Escherichia coli RNA polymerase sigma 70 subunit. J. Mol. Biol. 263, 637–647. 17. Buckle, M., Geiselmann, J., Kolb, A., and Buc, H. (1991) Protein-DNA cross-linking at the lac promoter. Nucleic Acids Res. 19, 833–840.

Chapter 19 Analysis of Plant Regulatory DNA sequences by the Yeast-One-Hybrid Assay Dierk Wanke and Klaus Harter Abstract Regulatory DNA sequences harbor the essential information to control specific gene expression changes and integrate information derived from upstream signaling cascades. This regulatory potential is mediated by direct binding of proteins, e.g., transcription factors, to defined stretches of DNA motifs in regulatory regions. The analysis of these DNA regions, at which several signaling pathways could merge to orchestrate gene expression, is still a challenging task. To date, the combination of functional approaches in the laboratory and computer aided sequence evaluation is frequently used for regulatory sequence analysis. The yeast-one-hybrid method is a possible approach to test for direct binding of plant proteins to DNA in a heterologous system. Moreover, it is the most frequently used method for the identification of DNA-binding proteins targeting a given DNA sequence by screening a cDNA library. Key words: Promoter and regulatory sequence analysis, protein–DNA interaction, yeast-one-hybrid, transcriptional activationSaccharomyces cerevisiae.

1. Introduction Among the many model species, the yeast Saccharomyces cerevisiae remains one of the best known and understood organism. Its use as a eukaryotic genetic tool is of great importance. Many molecular methods profit from the ease of genetic manipulation based on homologous recombination or transformation with circular plasmid DNA (1, 2). In the area of functional analysis of transport proteins or protein interaction studies, yeast is an indispensable research tool. The powerful technique of using hybrid protein fusions was first established for yeast and is now commonly

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used for the determination of intracellular protein movement or the investigation of protein–protein interaction (3–9). Systematic screening for protein–protein interaction has been carried out in yeast using the yeast-two-hybrid method (7, 10–15). A related method for identifying protein–DNA interactions is the yeast-one-hybrid system, which is a variant of the yeast-two-hybrid system (7, 16–19). The yeast-one-hybrid system is based on a hybrid protein assay where the transcription activation domain (AD) of a eukaryotic transcription factor is fused translationally to the protein of interest (Fig. 19.1). A regulatory sequence, either from a native context or as a multimerized short stretch of DNA containing a cis-regulatory element (CRE), is cloned upstream of a reporter gene. If the fusion protein is able to bind to the DNA sequence present in this regulatory sequence in vivo, the reporter gene confers the growth of yeast on selective media. This basic principle can be used for the identification of DNA-binding proteins in a large-scale screening procedure using a heterologous expression library (20, 21). The screening for DNA-binding proteins is based on growth assays and the complementation of yeast auxotrophy markers by reporter gene activity (1, 2, 18). The use of the yeast-one-hybrid method for the analysis of any non-coding regulatory sequence and the identification of DNA-binding proteins are universal and not restricted to plant organisms. Although endogenous

Fig. 19.1. Schematic outline of the yeast-one-hybrid system. For testing protein–DNA interaction the hybrid protein of a transcription factor fused to the activation domain is constitutively expressed. In case of a corresponding DNA-binding motif present in the DNA-bait sequence of the reporter-plasmid, the reporter gene is transcribed and mediates the growth of the yeast on selective media. The specificity of the interaction can be tested by using a mutated regulatory sequence as bait.

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yeast proteins might recognize the introduced regulatory sequence, and hence producing false positives, the use of this heterologous system is considered beneficial (18, 21–24). Thus, commercial yeast-one-hybrid kits, which provide the scientist with expression and reporter-plasmid combinations, are available. The following protocols describe yeast-one-hybrid techniques in a general way without focusing on one or the other plasmid combination. Some of the protocols given here can also be used for the transient protoplast expression system described in chapter 20 of in this volume. To gain a detailed insight into the function of regulatory sequences, a combination of several techniques is recommended, which will minimize the risk of misleading results.

2. Materials 2.1. Yeast Strains

1. Saccharomyces cerevisiae strain Y190: MATa ura3-52 his3-200 ade2-101 lys2-801 trp1-901 leu2-3, 112 gal4∆ metgal80∆ cyhr2 LYS2::GAL1UAS-HIS3TATA-HIS3 URA3:: GAL1UAS-GAL1TATA-lacZ 2. Saccharomyces cerevisiae strain PJ69-4A: MATa ura3-52 his3200 trp1-901 leu2-3, 112 gal4∆ gal80∆ LYS2::GAL1UAS-GAL1TATA-HIS3 GAL2-ADE2 met2::GAL7-lacZ 3. Saccharomyces cerevisiae strain AH109: MATa ura3-52 his3-200 trp1-901 leu2-3, 112 gal4∆ gal80∆ LYS2::GAL1UAS-HIS3TATAHIS3 GAL2-ADE2 URA3:: MEL1UAS- MELTATA-lacZ MEL1 (see Note 1) 4. Saccharomyces cerevisiae strain Y187: MATα trp1-901 ura352 his3-200 ade2-101 leu2-3, 112 gal4∆ met-gal80∆ URA3:: GAL1UAS- GAL1TATA-lacZ MEL1 (see Note 1)

2.2. Media

1. YPD liquid media: 1% yeast extract, 2% peptone, 2% glucose (see Note 2), autoclave. 2. YPD plates: YPD liquid media containing 1.5% agar, autoclave. 3. SD drop-out media (see Note 3): synthetic growth media for yeast containing 0.67% yeast nitrogen base and specifically lacking essential amino acids, uracil, or adenine for selection of transformants.

2.3. Solutions and Reagents

1. Restriction endonucleases and 10 X reaction buffer: SpeI and XbaI or XhoI and SalI; 10 U/µl. Store at −20°C. 2. T4-DNA ligase and 10 X ligation buffer: 2 U/µl. Store at −20°C.

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3. T4-polynucleotide kinase and 10 X T4-kinase buffer: 10 U/ µl. Store at −20°C. 4. 0.5 M EDTA, pH 8.0, autoclave. 5. 1 M Tris-HCl, pH 8.0, autoclave. 6. TE: 1 mM EDTA, 10 mM Tris-HCl, pH 8.0, autoclave. 7. 1 M NaCl, autoclave. 8. 5 mM ATP. Prepare small aliquots with sterile water. Store at −20°C. 9. 10 X DNA loading buffer: 0.25% bromophenolblue, 50% glycerol. Prepare small aliquots in water, heat buffer to 70°C for 5 min. Store at 4°C. 10. 10 X TBE: 108 g/l Tris-Base, 55 g/l borate, 40 mM EDTA. 11. Polyacrylamide stock solution: 30% acrylamide in water. Add 5–15% of bis-acrylamide according to the size of the multimerized oligonucleotides (see Sect. 3.1). Place in excicator and degas in vacuum for 20 min. Store at 4°C. 12. 10% APS (ammonium persulfate). Prepare small aliquots in water. Store at 4°C. 13. 1% ethidium bromide. Prepare aliquots in water. Store in darkness. 14. Glycogen solution: Prepare aliquots of 2% glycogen in water. Store at −20°C. 15. 3 M Na-acetate, pH 5.2, autoclave. 16. PEG 4000 solution: 50% (w/v) PEG 4000, autoclave. 17. 10 X Li-acetate: 1 M Li-acetate, pH 7.5, sterilize by filtering through syringe filter FP 30 CA (Schleicher & Schuell), 0.2 µm pore. 18. Carrier DNA solution: dissolve 10 mg/ml of sheared salmon or herring sperm DNA, boil for 10 min, and prepare aliquots (see Note 4). Store at −20°C. 19. PEG transformation mix: 666 ml PEG 4000 solution, 99 ml 10 X Li-acetate and 194 ml TE. Sterilize through filter (25). 20. Z-buffer: 16.1 g/l Na2HPO4, 5.5 g/l NaH2PO4, 0.25 g/l MgSO4, 0.75 g/l KCl, pH 7.0, autoclave. 21. X-gal stock solution: 20 mg/ml X-GaL (5-bromo-4-chloro3-indol-β-d-galactopyranoside) in N,N-dimethylformamide (DMF). Store in darkness at −20°C. 22. Z-buffer/X-gal solution: 100 ml Z-buffer, 0.27 mL β-mercaptoethanol, 1.67 ml X-gal stock solution. This solution has to be prepared freshly. 23. β-mercaptoethanol. Store in darkness at 4°C.

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24. ONPG-stock solution: 4 mg/ml ONPG (ortho-nitro-phenylgalactoside). Prepare aliquots in water. Store at −20°C. 25. 1 M Na2CO3, prepare aliquots in water. 2.4. Plasmids

Several different plasmid combinations are used and some of them are given in Table 19.1. There are more yeast-one-hybrid plasmids available than those listed, but they use similar principles. The plasmids differ in their auxotrophy markers for selection of positive protein–DNA interaction, the expression level of the hybrid proteins, or in their reporter enzyme (1, 2, 18, 26). Different plasmids are available for the Matchmaker system (Clontech) that make use of the highly sensitive α-galactosidase activity as a reporter instead of the β-galactosidase reporter (21, 27, 28). Recently, a combination of Gateway technique compatible yeast-one-hybrid plasmids (Invitrogen) has been published (24). Another vector system described by Sieweke (26) is capable of reducing the possibility of identifying false positive interactions by the galactose inducible expression of the target transcription factor (18, 29). The protocols given here describe the transformation of the hybrid-protein expression vector into a yeast reporter strain. Alternatively, different vector systems also open up the possibility to combine the expression plasmids with the reporter-plasmids by mating of the yeast (1, 21, 28, 30).

Table 19.1 List of frequently used driver- and reporter-plasmid combinations for the yeast-one-hybrid system Reporterplasmid

Auxotrophy marker Expression plasmid

Auxotrophy marker

Interaction marker

pHIS

Trp

pGADT7-Rec2, pGADT7, Leu pGAD424, pACT, pACT2, pGAD GH, pGAD GL, and pGAD10

Trp, Leu, His, α-Gal

(21, 28)

pINTHIS3NB

Trp

pGADT7-Rec2, pGADT7, Leu pGAD424, pACT, pACT2, pGAD GH, pGAD GL, and pGAD10

Trp, Leu, His, α-Gal

(18, 21)

pMSv3

His

pMSe4

Ura, galactose inducible expression

Trp, His, Ura, β-Gal

(26)

pDest6

Ura

pDest22

Leu

Leu, His, Ura, β-Gal

(24)

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3. Methods 3.1. Multimerization of Target DNA

Multimerization of a DNA pattern has been shown to be useful in cases where the fragments under investigation are shorter than about 200 bps in length (31). As some elements appear to possess a position-dependent functionality (32), multimerization increases the chance of matching a semi-natural positioning. It has been shown that a single element confers functionality, but its regulatory capacity was in direct relation to the number of multimers used (33). We therefore recommend the use of dimers for larger fragments and of tetra- to hexamers for short DNAfragments of less than 30 bps. Larger segments of endogenous regulatory sequences can directly be used without multimerization, but reporter activity might be significantly lower. The following protocol describes how to multimerize DNA for the in vivo assays used here: 1. Design oligonucleotides in sense and antisense strands for short DNA-fragments containing the regulatory sequence under investigation. Include compatible restriction sites, e.g., for SpeI and XbaI or XhoI and SalI as shown in Fig. 19.2. As a control, design equivalent oligonucleotide pairs, but with

Fig. 19.2. Principle of the multimerization of the target DNA. Compatible restriction sites are indicated as site I and site II. Possible combinations of restriction enzymes are SpeI/XbaI or XhoI/SalI. (A) Example of two hybridized oligonucleotides for multimerization as used by Rushton et al. (33) for the analysis of the W1-box sequence. (B) Examples of resulting multimers.

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nucleotide substitutions inside the regulatory sequence that abolish transcription factor binding. Follow the protocol for endogenous and mutated DNA-fragments simultaneously. 2. Adjust oligonucleotide concentration to 5 µg/µl. Mix 20 µl of sense and antisense oligonucleotides in 1.5 ml reaction tube with 2.5 µl 1 M NaCl and 7.5 µl TE (total volume 50 µl) (see Note 5). 3. Heat reaction mix to 80°C for 5 min and allow slow cooling to room temperature for 30 min. The reaction mix now contains a high proportion of double-stranded oligonucleotides. 4. Transfer 10 µl of the reaction mix to a new tube and add 2 µl 10 X T4-kinase buffer, 4 µl 5 mM ATP, 2 µl water, and 2 µl T4-polynucleotidekinase (see Note 6). Incubate for 1 h at 37°C. Put reaction tube on ice. 5. Add 10 µl 10 X ligation buffer (containing ATP), 2 µl of restriction endonuclease for site I (according to the example in Fig. 19.2: SpeI or XhoI), 2 µl of restriction endonuclease for site II (according to the example in Fig. 19.2: XbaI or Sal, respectively), 4 µl of T4-DNA ligase, and 62 µl water. Incubate for 2 h at 37°C (see Note 7). 6. Add 10 µl 10 X DNA loading buffer to the sample and mix well. Split the samples equally and load them onto a nondenaturing TBE polyacrylamide gel. The polyacrylamide concentration required depends on the size of the DNA fragments you use (see Note 8). Pour a 15% polyacrylamide gel for oligonucleotides smaller than 30 nucleotides. Run gel at 1–8 V/cm to separate DNA fragments well. 7. Stain gel in an ethidium bromide containing TBE bath. Gently agitate for 15–30 min. 8. Transfer gel to a UV-light chamber. Cut out bands corresponding to 2, 4, and 6 copies of multimerized oligonucleotides or DNA fragments. 9. Transfer each slice independently into a fresh 1.5-ml reaction tube and add 700 µl TE. Agitate overnight at 50°C. 10. Transfer DNA containing TE buffer into a fresh 1.5-ml reaction tube. Add 0.5 µl of glycogen solution, 35 µl 3 M Naacetate solution, and 700 µl isopropanol. Invert tube several times to mix well. Incubate at −20°C for 20 min to precipitate DNA. Spin for 15 min at 4°C in a tabletop centrifuge, discard supernatant, wash with 70% ethanol, and dry the pellet. Resuspend it in 10 µl TE (see Note 9). 3.2. Generation of Yeast Reporter Strain

1. Streak out the desired yeast strain on YPD plates, so that single colonies are separated (see Note 2). Incubate at 30°C for 1–2 days.

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2. Pick one colony and grow in 3 ml YPD liquid media at 30°C. The density should be at 0.6–0.8 OD600. 3. Sediment the cells by centrifugation for 2 min at 3,000× g. Resuspend yeast cells in 500 µl of water. 4. Adjust reporter-plasmid concentration to 1 µg/µl (see Note 10). 5. Use a 1.5-ml reaction tube. Filter-sterilize 240 µl PEG 4000 solution, add 36 µl 10 X Li-acetate, 20 µl TE, 50 µl of carrier DNA solution, and 2 µg reporter-plasmid (see Note 4 and 11). Add 12 µl of yeast cell suspension (see Note 12). Mix well. Shake at 30°C for 1 h. 6. Perform heat shock in a water bath for 20 min at 42°C. Place on ice for 2 min. 7. Centrifuge with short spin (less than 1 min) in a tabletop centrifuge to sediment yeast cells. Remove supernatant with a pipette and discard it. Resuspend pellet in 1 ml of YPD liquid media. Incubate at 30°C for 1 h. 8. Centrifuge 1 min at 3,000× g to sediment yeast cells. Resuspend cells in 1 ml of sterile water for washing. Centrifuge again. Remove 900 µl supernatant with a pipette and discard. 9. Resuspend yeast cells in the remaining volume and plate out on appropriate SD drop-out media plates lacking the essential amino acid for selection of the reporter-plasmid transformants. Incubate at 30°C until colonies can be seen. 10. Use colonies of positively generated yeast reporter strain for quantitative colorimetric ONPG-assays (see Sect. 3.7) to estimate background reporter activity. 3.3. Transformation of Driver-Plasmid

1. For transformation of a driver-plasmid containing the cDNA of an effector protein under the control of an inducible or constitutive promoter, follow the yeast transformation protocol in Sect. 3.2 and start this time with a colony of the previously generated yeast reporter strain. 2. If you have generated yeast reporter strains for a wild-type and a mutant regulatory sequence, carry on driver-plasmid transformation with both strains. Include an empty vector control strain. 3. Note that yeast reporter strains have to be grown on selective SD media lacking one amino acid. Transformed strains, positive for reporter-plasmid and driver-plasmid, have to be selected on media lacking supplements for both auxotrophy markers.

3.4. Transformation of Expression Library

1. To identify DNA-binding proteins that bind to the regulatory sequence under investigation, a population of driver-plasmids containing a cDNA library is transformed into the yeast

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reporter strain (see Note 13). Adjust plasmid DNA concentration to 1 µg/µl. 2. Prepare a total of 60–100 µg of the cDNA library containing driver-plasmids for transformation. Estimate the transformation efficiency following the protocol in Sect. 3.2 with a dilution series of the library plasmid. Start with the previously generated yeast reporter strain and transform 0.1 µg, 10 ng, and 1 ng of library plasmid DNA (see Note 14). 3. Note that the yeast reporter strain has to be grown on selective SD media lacking one amino acid. Identify colonies positive for reporter-plasmid and library plasmid on media lacking supplements for both auxotrophy markers. 4. Grow a colony of the previously generated yeast reporter strain in 5 ml liquid selective SD media lacking one amino acid (auxotrophy marker) overnight at 30°C (starter culture). 5. Transfer the starter culture into 4 l of selective SD media and grow cells to a density of 0.8 OD600. 6. Sediment the cells at 3,000× g for 5 min and resuspend the pellet in 4 l of freshly made YPD liquid media (see Note 15). Let cells grow at 30°C to an optical density of 1 OD600. 7. Sediment the yeast cells by centrifugation at 3,000× g for 5 min and resuspend pellet in 800 ml of sterile water for washing. Use 50-ml reaction tubes and split the cell suspension equally into 16 aliquots. Again, sediment the cells by centrifugation at 3,000× g for 5 min. 8. Resuspend each pellet in 45 ml PEG transformation mix. Shake at 30°C for 30 min. 9. Add 500 µl of carrier DNA and 6 µg library plasmid to each reaction tube. Mix well. Shake at 30°C for 1 h. 10. Perform heat shock in a water bath for 20 min at 42°C. Place on ice for 2 min. 11. Sediment the yeast cells by centrifugation at 3,000× g for 5 min and resuspend each pellet in 50 ml sterile water for washing. Again, sediment the cells by centrifugation at 3,000× g for 5 min. 12. Resuspend each pellet in 20 ml of YPD liquid media (see Note 16). Incubate at 30°C for 30 min. 13. Sediment the yeast cells by centrifugation at 3,000× g for 5 min and resuspend each pellet in 50 ml of sterile water for washing. Again, sediment the cells by centrifugation at 3,000× g for 5 min. Resuspend pellets in 5 ml of sterile water. 14. Merge yeast cells from all 16 reaction tubes. 15. Place a layer of nitrocellulose filter membranes onto 20 × 20 cm SD triple drop out media, lacking both auxotrophy markers, of

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the reporter-plasmid and the library containing driver-plasmid, as well as the marker for reporter gene activity (see Note 17). 16. Use 20 plates containing SD triple drop out media. Plate 4 ml of yeast cell suspension onto the nitrocellulose filter membranes. Spread cells equally with a glass spreader, which will later facilitate the identification of single positive colonies. Up to 50,000 colonies can be separated on plates like these, so that a total of 106 transformants could be monitored. 17. To estimate transformation efficiency, plate 100, 10, and 1 µl of yeast suspension on Petri dishes containing SD double drop out media lacking auxotrophy markers of the reporter-plasmid and the library containing driver-plasmid. A sufficient transformation should yield about 1000 colony forming units from a 100 µl cell suspension. 18. Incubate the plates covered with nitrocellulose filter membranes at 30°C for about 36–48 h until the first small colonies become visible. 3.5. Identification of Positive Clones

1. Transfer nitrocellulose filter membranes to selective SD media plates appropriate for your plasmids and the selective conditions for all three auxotrophy markers (see Note 18 and 19). Avoid air bubbles during transfer. Yeast colonies that do not support growth for all three selective auxotrophy markers will arrest in growth. 2. Incubate the plates at 30°C for an additional 36 h. Then pick well-growing colonies and streak them about 1 cm over the surface of an additional selective SD media plate. Include control colonies that have been transformed with the reporter-plasmid only. 3. Incubate plates at 30°C for about 36–48 h. Yeast cells that contain hybrid proteins positively interacting with the DNAbait of the reporter-plasmid should yield in well-growing colonies. The control colonies should not grow. 4. Pick colonies into 5 ml liquid SD media selective for all auxotrophy markers of your plasmids. As controls pick a yeast colony transformed with the reporter-plasmid and a positive control, if you have one, in appropriate SD media. Follow the protocol with all colonies simultaneously. Grow overnight at 30°C with vigorous shaking to a density of about 0.8 OD600. 5. Sediment the yeast cells by centrifugation at 3,000× g for 5 min. Discard the supernatant and resuspend each pellet in 1 ml of sterile water. Use vortex to properly disperse clumping cells. 6. Adjust 0.5 ml of each of the yeast cell suspensions to a density of 0.5 OD600. 7. Make serial dilutions for all cell suspensions of 1:10, 1:100 and 1:1000 using sterile water.

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8. Prepare two kinds of SD media plates: One being selective for the auxotrophy marker of the reporter-plasmid only and a second kind selective for all auxotrophy markers. The surface of the media should be dry. 9. Place one drop of 7.5 µl for each of the cell suspensions and for each of the dilutions on both kind of SD media plates. Try to drop the dilution series in one line with equal distance between each spot (see Fig. 19.3). As a guideline, stick a print-out of a grid to the bottom of the plates.

Fig. 19.3. Example of a yeast-one-hybrid experiment. (A) Display of a yeast growth assay on selective and control media plates. Yeast of the negative control show some background growth in the lowest dilution on selective media. Two interactors have the capacity to mediate strong growth of the yeast cells. (B) Relative β-galactosidase activity measured in the yeast cells from the above growth assay using ONPG as a substrate. Each bar represents the average of four measurements ± standard error of the mean.

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10. Incubate the plates at 30°C for additional 36–48 h. Stop incubation, when clear colonies grow on the SD media plate with only the auxotrophy marker for the reporter-plasmid. Compare growth on both kind of plates, between positive and negative controls and the colonies displaying a positive protein–DNA interaction. 11. Use colonies on the second kind of SD media plate selective for all auxotrophy markers for filter-lift assay (Sect. 3.6). 3.6. Filter-Lift Assay with X-Gal

1. A positive interaction in yeast hybrid-protein assays can be estimated by the rapid detection of β-galactosidase activity. For a filter-lift assay use round nitrocellulose filter-membranes that fit inside your Petri dishes. Grow yeast freshly on SD media appropriate for your plasmids and the selective conditions for all auxotrophy markers. 2. Label your nitrocellulose filters with a pencil or ball-pen and properly label your Petri dishes accordingly. Use a clear labeling system, especially if you have several plates to analyze, and avoid long names. 3. Use forceps to carefully place a filter onto the yeast colonies. Allow it to wet completely (see Note 17). Avoid whatever will lead to a smearing of the colonies. Poke holes asymmetrically through the filter and into the agar, which will help to orientate the membrane after the staining procedure. 4. Lift the filter off the plate. At best, lift it from one side with a constant movement. 5. Place the filter in liquid N2 for ~5 s to break up and permeabilize the yeast cells (see Note 20). 6. Cover the bottom of a Petri dish with 3MM chromatography paper and allow to soak with 3 ml Z-buffer/X-gal solution. Place nitrocellulose filters with the cell-side up onto the 3MM paper. 7. Incubate at 30°C and monitor the development of the blue color. Depending on the strength of interaction, this step might take several minutes or up to several hours (see Note 21). 8. Colonies of β-galactosidase expressing yeast cells can be identified by aligning the filter with the agar plate using the orientation marks.

3.7. Quantitative Colorimetric ONPG Assay for Reporter Activity

1. Pick fresh colonies into 5 ml liquid SD media appropriate for your plasmids and the selective conditions for all auxotrophy markers (see Note 22). Grow overnight at 30°C with vigorous shaking. 2. Transfer 1 ml overnight culture into a fresh tube with 4 ml of liquid SD media and continue shaking at 30°C for 3–5 h. Grow cells to a density of 0.7 OD600.

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3. Split the samples equally (1 ml each) in three 1.5-ml reaction tubes (see Note 23). Follow the protocol with all samples simultaneously. 4. Sediment the yeast cells by centrifugation at 3,000× g for 5 min, decant the supernatant and resuspend each pellet in 1.5 ml sterile Z-buffer. Again, sediment the cells by centrifugation at 10,000× g for 30 s. 5. Discard supernatant and resuspend each pellet in 1 ml Z-buffer. Vortex rigorously to properly disperse clumping cells. 6. Adjust 1 ml of the yeast cells to a density of 0.5 OD600. 7. Sediment the yeast cells by centrifugation at 3,000× g for 5 min, remove the supernatant and resuspend each pellet in 300 µl Z-buffer. 8. Freeze cells in liquid N2 and thaw again in a 37°C water bath for 1 min. Repeat this twice to ensure quantitative breaking of the cells; then place tubes on ice. 9. Prepare 1.5-ml reaction tubes with 500 µl Z-buffer and 10 µl β-mercaptoethanol. Add 100 µl of ONPG-stock solution and mix well. Prepare one additional tube as blank tube. 10. Start a timer immediately before the addition of each cell suspension to the ONPG pre-mixes. Briefly vortex each cell suspension and transfer 100 µl into each of the prepared reaction tubes. Add 100 µl of Z-buffer to the blank control tube. Mix the reaction tubes carefully and incubate at room temperature. 11. Take the exact time needed for each of the tubes to develop the yellow color. 12. Add 300 µl 1 M Na2CO3 to the reaction and the blank tubes to stop the reaction. Record the elapsed time in minutes. 13. Sediment cellular debris by centrifugation at 10,000× g for 15 min. 14. Transfer supernatant into a fresh 1.5-ml reaction tube and measure the OD420 (see Note 24). Use the control tube that did not contain any yeast lysate as a blank for the measurement. 15. Calculate β-galactosidase units according to the formula 1000 × OD420 , where t is the elapsed time in minutes, U = t × OD600 × V V is the volume 0.1 ml × dilution factor and OD600 is adjusted to 0.5 OD600. The dilution factor is 3 for this protocol related to 1 ml of cell suspension (see Note 25). 3.8. Characterization of Positively Interacting Hybrid-Protein

The identity of a positively interacting protein isolated from an expression library can be clarified by sequencing the cDNA-insert of the driver-plasmid (1, 34). There are several commercial kits

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available for the extraction of plasmid DNA from yeast. Design driver-plasmid-specific primer pairs flanking the cloned cDNA sequence and amplify the cDNA-insert from the isolated plasmid DNA by PCR. In most cases the purified PCR product can be used as a template for direct sequencing by using one of the specific primers flanking the insert.

4. Notes 1. Several different yeast strains exist that are frequently used for the yeast-one-hybrid technique. The Matchmaker System (28) makes use of yeast strains that contain the MEL1 reporter gene for highly sensitive α-galactosidase assay. This allows direct quantitative screening for interaction by colorimetric measurements of α-galactosidase activity (27). We do not integrate α-galactosidase protocols, as most of the literature on yeast-one-hybrid makes use of the less sensitive β-galactosidase activity assay instead. 2. If the yeast strain used carries an ade2 mutation, the amount of adenine in the media might be limiting. In this case a red colored intermediate might accumulate rapidly to toxic concentrations. If cells are not white and turn yellow-red, pink, or brown, it has been recommended to add extra adenine (20 mg/l) to the media (1, 26). 3. The synthetic SD medium is supplied with all essential amino acids as nitrogen source, trace elements, salts, and vitamins. It can be purchased from several companies as full or dropout media for growth selection, e.g., Sigma-Aldrich, DifcoLabo-ratories. SD drop-out media compositions that lack all specific amino acids, uracil, or adenine are available. This allows choosing the specific medium composition appropriate for your plasmids and the selective conditions for the auxotrophy markers present on the driver-plasmid, reporter-plasmid, and for the reporter gene. Amino acids lacking in the SD drop-out medium can directly be added, which reduces the risk of contamination. Traces of selective nutrients in the drop-out medium are a frequent cause for problems with selection of transformants. We therefore recommend keeping yeast media components, like SD drop-out media, amino acids, uracil and adenine, separate from other regularly used chemicals. It is advisable to use chemicals for yeast media exclusively and not to mix with others, e.g., plant growth media. 4. The purity and size of carrier and expression vector DNA are important factors for transformation efficiency. The

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recommended size of sheared carrier DNA should average at about 1 kb in length. This can be achieved by sonication or mechanical shearing. We recommend sonicating for different time periods and checking the size of carrier DNA on an agarose gel. For preparation of plasmid DNA use silica matrix-based spin columns. 5. Be sure to mix the reaction well. The use of higher concentrated NaCl solution has been recommended in other protocols. We cannot support this, as the salt solution has a higher density, it will likely “flow” to the bottom of the tube, and will not mix during heating. Hence, under low salt conditions the primer hybridization process will not take place at a sufficient rate. If oligonucleotide annealing fails, improper mixing is a frequent cause. 6. This step attaches phosphate residues to the double-stranded oligonucleotides to allow subsequent ligation. It can be omitted if the oligonucleotides have been ordered phosphorylated. 7. The combined restriction and ligation mix allow only directional multimerization. Compatible site I - site I or site II - site II ligation will subsequently be cut by the appropriate site I or site II specific endonucleases present in the reaction mix (Fig. 19.2). Hence, only compatible site I - site II ligations are permissive. 8. Pour a 12% polyacrylamide gel for oligonucleotides smaller than 50 nucleotides, a 8% gel for DNA fragments smaller than 100 bps, or a 5% polyacrylamide gel for DNA fragments larger than that. 9. The multimerized DNA is now ready for cloning to appropriate vectors. If restriction sites are not suitable for direct cloning, fill-in reaction with DNA-polymerases and subcloning might be required. For GATEWAY compatible cloning, we recommend fill-in reaction and cloning to a pENTR-Topo vector (Invitrogen) containing appropriate att-recombination sites. 10. If an integrative vector is used that needs to be integrated at a homologous site into the yeast genome, the plasmid usually needs to be linearized with an endonuclease and gel purified before transformation. 11. Keep in mind that transformation has to be carried out with a wild-type and a mutated reporter-plasmid, where applicable. The generation of two reporter strains with wild-type and mutant regulatory sequence will greatly enhance the validation process. Additionally, transform an empty reporter-plasmid lacking a regulatory sequence as control. 12. Instead of adding 12 µl yeast suspension, one can directly pick a freshly grown colony from the plate, transfer it into the transformation mix, and resuspend the cells carefully. Accordingly increase volume of TE to 32 µl.

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13. For plasmid DNA isolation of the driver-plasmid population containing the cDNA library from bacteria, use a silica matrix-based MIDI preparation kit. Adjust plasmid concentration to 1 µg/µl and test the integrity by re-transforming 0.1 µg into bacteria. The transformation should result in a minimum of 5 × 104 bacterial colonies. 14. Calculate the amount of total cDNA library containing driver-plasmid needed for transformation to yield a total of 106 colonies. Based on experience, 60–100 µg plasmid DNA will be needed. 15. This centrifugation step removes dead cells and cellular debris from the culture. The culture is now enriched for living and thriving yeast cells. 16. This step might well be skipped. To our experience, it increases the chance of detecting interaction partners that have growth inhibitory effects in yeast or that are toxic. 17. Placing the nitrocellulose filters on top of the plates can be difficult and we recommend preparing the plates the day before. Air bubbles should be avoided and plates should not be too wet. Best results could be achieved when the sides of the filter are bent inward with the help of two tweezers and then allowing the filter to touch the middle line of the plates first. Carefully lower the nitrocellulose filters, while they soak the liquid from the plate’s surface. 18. The surface of the media should be relatively dry. Excess fluid should be removed either by placing the plates in a sterile work bench until the surface gets dry or by placing plates upside down in a fridge and wait until fluid accumulates in the lid. Then dispose the wet lid and use a fresh one from another Petri dish. 19. If your yeast has overgrown and forms big colonies on the nitrocellulose filter, it is not recommended to transfer the filter to selective SD media plates with triple drop out selection, but to blot the colonies onto the drop-out plate. Take the filter with two forceps and carefully place it cellside-down on the appropriate triple SD drop out. Turn forceps and gently move over the filter to transfer yeast cells from the filter onto the media’s surface. 20. Be very careful because the frozen filter is brittle. When holding it, use forceps with rubber or flat tips. 21. For impatient researchers, the development of blue color can be speeded up by incubating at 37°C. However, be careful: For strong interactions, this might lead to exaggerative staining. The intensity of staining does not only depend on the strength of protein–DNA interaction, but also on the kind of sequence you are investigating: When using a short

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multimerized sequence, you can expect a blue staining to appear within 5–20 min. Instead, the development of a blue color that is indicative of β-galactosidase activity can take several hours when a native promoter of 1 kb or longer is used in the experiment. Incubation times exceeding 8 h tend to give false positive signals. Therefore, including a negative control colony on each of the filters is essential, if a prolonged incubation time is necessary. Monitor the development of blue color. The negative control will accumulate blue color as well, but staining in case of a positive interaction should be more intense. 22. Besides the yeast colonies under investigation, include a negative and, where possible, a positive control (Fig. 19.3). 23. Use triplicates for each of the yeast colonies. This enables you to calculate average and standard error for each of the positive protein–DNA interactions. 24. The yellow color will become more intense with time. To preserve linearity, values should not exceed 0.8 OD420. If yellow color accumulates too quickly, yeast cell suspension has to be diluted: Instead of 100 µl use only 10 µl of cell suspension and repeat the assay. 25. One unit of β-galactosidase is defined as the amount that hydrolyzes 1 µmol ONPG to O-nitrophenol and d-galactose per minute per cell (35).

Acknowledgements The authors would like to thank Kenneth W. Berendzen, Sabine Hummel and Friederike Ladwig for their advice and helpful discussions. References 1. Guthrie, C. and Fink, G. R. (eds.), (1991) Guide to Yeast Genetics and Molecular Biology, Methods in Enzymology, Vol. 169, Academic Press, San Diego. 2. Adams, A., Gottschling, D. A., Kaiser, C. A., and Stearns, T. (1998) Methods in Yeast Genetics, A Cold Spring Harbor Laboratory Course Manual, 1997 Edition, Cold Spring Harbor Laboratory Press. 3. Fields, S. and Sternglanz, R. (1994) The two-hybrid system: An assay for proteinprotein interactions. Trends Genet. 10, 286–292.

4. Johnson, N. and Varshavsky, A. (1994) Split ubiquitin as a sensor of protein interactions in vivo. Proc. Natl. Acad. Sci. USA 91, 1034–1044. 5. Zhang, J. and Lautar, S. (1996) A yeast three-hybrid method to clone ternary protein complex components. Anal. Biochemistry 242, 68–72. 6. SenGupta, D. J., Zhang, V., Kraemer, V., Pochart, P., Fields, S. and Wickens, M. (1996) A three-hybrid system to detect RNA-protein interactions in vivo. Proc. Natl. Acad. Sci. USA 93, 8496–8501.

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7. Zhu, L. and Hannon, G. J. (eds.) (2000) Yeast Hybrid Technologies. Eaton Publishing, BioTechniques Books Division, Natick, MA. 8. Lehming, N. (2002) Analysis of proteinprotein proximities using the split-ubiquitin system. Brief Funct Genomic Proteomic. 1, 230–238. 9. Walter, M., Chaban, C., Schütze, K., Batistic, O., Weckermann, K., Näke, C., Blazevic, D., Grefen, C., Schumacher, K., Oecking, C., Harter, K. and Kudla, J. (2004) Visualization of protein interactions in living plant cells using bimolecular fluorescence complementation. Plant J. 40, 428–438. 10. Ma, J. and Ptashne, M. (1987) Deletion analysis of GAL4 defines two transcriptional activating segments. Cell 48, 847–853. 11. Fields, S. and Song, O. (1989) A novel genetic system to detect protein-protein interactions. Nature 340, 245–246. 12. Ruden, D. M., Ma, J., Li, Y., Wood, K. and Ptashne, M. (1991) Generating yeast transcriptional activators containing no yeast protein sequences. Nature 350, 250–252. 13. Li, J. J. and Herskowitz, I. (1993) Isolation of ORC6, a component of the yeast origin recognition complex by a one-hybrid system. Science 262, 1870–1874. 14. Lehming, N., Thanos, D., Brickman, J. M., Ma, J., Maniatis, T. and Ptashne, M. (1994) An HMG-like protein that can switch a transcriptional activator to a repressor. Nature 371, 175–179. 15. Bartel, P. and Fields, S. (1995) Analyzing protein-protein interactions using twohybrid system. METHODS: A Companion of Methods in Enzymology 254, 241–263. 16. Wilson, T., Padgett, K., Johnston, M. and Milbrandt, J. (1993) A genetic method for defining DNA-binding domains: application to the nuclear receptor NGFI-B. Proc. Natl. Acad. Sci. USA 90, 9186–9190. 17. Ausubel, F. M., Brent, R., Kingston, R. E., Moore, D. D., Seidman, J. G., Smith, J. A. and Struhl, K. (1995) Current Protocols in Molecular Biology, John Wiley & Sons. 18. Meijer, A. H., Ouwerkerk, P. B. and Hoge, J. H. (1998) Vectors for transcription factor cloning and target site identification by means of genetic selection in yeast. Yeast 14, 1407–1415. 19. Wei, Z., Angerer, R. C. and Angerer, L. M. (1999) Identification of a new sea urchin ets protein, SpEts4, by yeast one-hybrid screening with the hatching enzyme promoter. Mol. Cell. Biol. 19, 1271–1278.

20. Jolly, E. R., Chin, C. S., Herskowitz, I. and Li, H. (2005) Genome-wide identification of the regulatory targets of a transcription factor using biochemical characterization and computational genomic analysis. BMC Bioinformatics 6, 275. 21. Lopato, S., Bazanova, N., Morran, S., Milligan, A. S., Shirley, N. and Langridge, P. (2006) Isolation of plant transcription factors using a modified yeast one-hybrid system. Plant Methods 2, 3. 22. Fields, S. (1993) The two-hybrid system to detect protein-protein interactions. METHODS: A Companion to Methods in Enzymology 5, 116–124. 23. Vidal, M. and Legrain, P. (1999) Yeast forward and reverse “n”-hybrid system. Nucl. Acids Res. 27, 919–929. 24. Deplancke, B., Dupuy, D., Vidal, M. and Walhout, A. J. M. (2004) A Gateway-Compatible Yeast One-Hybrid System. Genome Res. 14, 2093–2101. 25. Gietz, D., St Jean, A., Woods, R. A. and Schiestl, R. H. (1992) Improved method for high efficiency transformation of intact yeast cells. Nucl. Acids Res. 20, 1425. 26. Sieweke, M. (2000) Detection of transcription factor partners with a yeast one hybrid screen. Methods Mol Biol. 130, 59–77. 27. Aho, S., Arffman, A., Pummi, T. and Uitto, J. (1997) A novel reporter gene MEL1 for the yeast two-hybrid system. Anal Biochem. 253, 270–272. 28. Matchmaker Library Construction and Screen Kit (2000) Clontechniques XV(4), 5–7. 29. Bartel, P., Chein, C., Sternglanz, R. and Fields, S. (1993) Elimination of false positives that arise in using the two hybrid system. Biotechniques 14, 920–924. 30. Bendixen, C., Gangloff, S. and Rothsein, R. (1994) A yeast mating - selection scheme for detection of protein-protein interactions. Nucl. Acids Res. 22, 1778–1779. 31. Liaw, G.J. (1994) Improved protocol for directional multimerization of a DNA fragment. Biotechniques 17, 668–670. 32. Berendzen, K. W., Stueber, K., Harter, K. and Wanke, D. (2006) Cis-motifs upstream of the transcription and translation initiation sites are effectively revealed by their positional disequilibrium in eukaryote genomes using frequency distribution curves. BMC Bioinformatics 7, 522. 33. Rushton, P. J., Reinstadler, A., Lipka, V., Lippok, B. and Somssich, I. E. (2002) Synthetic plant promoters containing defined

Analysis of Plant Promoters regulatory elements provide novel insights into pathogen- and wound-induced signaling. Plant Cell 14, 749–762. 34. Sathe, G., O’Brian, S., McLaughlin, M., Watson, F. and Livi, G. (1991) Use of polymerase chain reaction for rapid detection

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of gene insertions in whole yeast cells. Nucl. Acids Res. 19, 4775. 35. Miller, J. H. (1972) Experiments in Molecular Genetics. A Cold Spring Harbor Laboratory Manual, Cold Spring Harbor Laboratory Press.

Chapter 20 Analysis of Plant Regulatory DNA sequences by Transient Protoplast Assays and Computer Aided Sequence Evaluation Kenneth W. Berendzen, Klaus Harter, and Dierk Wanke Abstract The orchestrated regulation of hundreds of genes responding in a temporal, spatial, and conditional expression is in part mediated by the transient binding of transcription factors to their specific DNA motifs. The analysis of these cis-regulatory DNA sequences is still a challenging task. Therefore, the combination of the transient protoplast expression assay with computer aided sequence analysis is a preferred method for regulatory sequence analysis. The protocols given here describe the use of a reporter gene plasmid to investigate the effects of a transiently co-expressed transcription factor gene in planta. As a number of bioinformatic analysis tools for cis-elements are publicly available, we suggest a workflow for the electronic analysis of promoter sequences. This is in particular difficult, as most analysis programs have not been developed for the investigation of single sequences. We provide insight into bioinformatics tools that investigate cis-element presence, their distribution, and implications for further functional analyses. Key words: Transient protoplast expression assay, Promoter and regulatory sequence analysis, protein-DNA interaction, transcriptional activation, cis-regulatory elements.

1. Introduction The analysis of protein-DNA interaction is a crucial step for our understanding of how gene expression is regulated in a temporal and spatial manner. Transcription factors bind specifically to their cognate DNA motif to influence the transcriptional machinery and thereby orchestrate gene expression changes. The analysis of these cis-regulatory DNA sequences is still a challenging task and several methods have been described that provide a closer insight

T. Pfannschmidt (ed.), Plant Signal Transduction, vol. 479 Humana Press, a part of Springer Science + Business Media, LLC 2009 DOI: 10.1007/978-1-59745-289-2_20

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into the effects of transcription factor–cis-regulatory element (CRE) interaction. The identification of directly targeted DNA motifs that are specifically bound by a transcription factor in vivo, the X-ChIP method, can be used to immuno-precipitate short stretches of chromatin. To subsequently investigate whether the transcription factor acts as an activator or a repressor on transcription, transient gene expression in protoplasts provides a powerful and fast tool for identifying regulatory sequences (1–4). The technique of PEG-mediated DNA transfer for the transient expression analysis of transcription factor genes is a convenient alternative to stable transformation procedures (5). The protocol given here uses the uidA-Gene (GUS) as an in planta reporter enzyme to carry out qualitative and quantitative assays (6, 7). The positive or negative effects on gene expression reflected by enzymatic GUS activity are directly related to the amount of transcription factor binding to the regulatory region (Fig. 20.1) (2, 3, 8). We recommend integrating the functional analysis of plant regulatory DNA sequences into a bioinformatics analysis of cis-regulatory elements. The frequency, position, or orientation of cis-regulatory elements might also provide essential hints for detailed functional characterization of the regulatory sequence in the transient protoplast expression system (9). While many more DNA-motif analysis and prediction tools exist for the investigation of correlative promoter sequences, there are few that support the scientist with some information on the basis of only one single sequence to study. Here, we provide a workflow based on a single DNA sequence that will result in the identification of putative functional cis-regulatory elements and of DNA motifs that might constitute yet unknown regulatory sequences.

2. Materials 2.1. Plant Materials

The protocols given here start with Arabidopsis thaliana protoplasts from liquid cell suspension cultures. A suitable protocol for the isolation of such protoplasts has been described in another chapter of this volume by Schütze et al.

2.1.1. Media and Solutions for Transient GUS-Expression Assay in Protoplasts

1. W5: 154 mM NaCl, 125 mM CaCl2, 5 mM KCl, 5 mM glucose, pH 5.8–6.0 adjusted with KOH, autoclave. 2. MMM: 15 mM MgCl2, 0.1% MES, 0.5 M mannitol, pH 5.8 adjusted with KOH, autoclave. 3. PEG-solution: 40% PEG 4000, 0.4 M mannitol, 0.1 M Ca(NO3)2, pH 8–9 adjusted with KOH (the pH needs 1–2 h to stabilize), autoclave, store in aliquots at −20°C.

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Fig. 20.1. Principle of the transient protoplast expression assay. (A) Scheme of the co-transformation of driver- and reporter-plasmids into a plant protoplast. The driver-plasmid expresses the cDNA of a transcription factor that changes the reporter gene expression by direct interaction with the regulatory sequence present in the reporter-plasmid. (B) Example for the identification of an important regulatory region measuring relative GUS activity with 4-MUG as a substrate. Transient assays were done either with the reporter-plasmids alone or combined with a driver-plasmid constitutively expressing a transcription factor. Grey bars represent the GUS activity of the reporter-plasmid in the presence of the driver-plasmid. White bars represent GUS activity of the reporter-plasmid alone. Each bar represents the median of three independent transformations; the error bars are ± standard error of the mean. A region of regulatory importance is identified by a proportionally high difference in GUS activity, which is transcription factor dependent.

4. K3-medium (for 100 ml): 10 ml macro elements, 0.1 ml micro elements, 0.1 ml B5-Vitamins stock, 0.5 ml EDTA stock, 1 ml Ca-phosphate solution, 10 mg myo-inositol, 25 mg D(+)-xylose, 13.7 g sucrose, pH 5.6 adjusted with KOH and sterilized by filtering through bottle-top membrane filter unit (0.2 µm pore, Nalgene); store in 10 ml aliquots at −20°C. 5. Ca-phosphate solution (for 200 ml): for 200 ml Ca-phosphate stock: 1.26 g CaHPO4 • 2H2O dissolved in H2O, pH 3 adjusted with 25% HCl, autoclave and keep in the dark. 6. EDTA stock: for 1 l EDTA stock, dissolve 7.46 g EDTA in 300 ml H2O and heat to 60°C, 5.56 g Fe(II)SO4 • 7H2O dissolve

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in 300 ml of H2O under heat, mix, and add H2O up to 1 l. Autoclave and keep in the dark. 7. B5-Vitamins stock (100 ml): 100 mg nicotinic acid, 100 mg pyridoxin · HCl, 1 g thiamin · HCl, add H2O up to 100 ml, filter sterilize, and freeze at −20°C. 8. Micro-elements (100 ml): 75 mg KI, 300 mg H3BO3, 1 g MnSO4 • 7H2O (0.6 g MnSO4 • H2O), 200 mg ZnSO4 · 7H2O, 25 mg Na2MoO4 • 2H2O, 2.5 mg CuSO4 • 5H2O, 2.5 mg CoCl2 · 6H2O, add H2O up to 100 ml, filter sterilize and freeze at −20°C. 9. Macro-elements (1 l): 1.5 g NaH2PO4 • H2O, 9.0 g CaCl2 • 2H2O, 25 g KNO3, 2.5 g NH4NO3, 1.34 g (NH4)2SO4, 2.5 g MgSO4 • 7H2O, add H2O up to 1 l, and autoclave. 10. Extraction buffer: 200 mM Na-phosphate pH 7.0, 10 mM EDTA, 0.1% Triton X-100, 0.1% laurylsarcosine, 10 mM β-mercaptoethanol. 11. Reaction solution: 2 mM 4-MUG, 50 mM Na-phosphate pH 7.0, 1 mM EDTA, 0.1% Triton X-100, 10 mM β-mercaptoethanol. 12. Stop solution: 200 mM Na2(CO3)

3. Methods The following methods are divided into two parts, which describe the analysis of plant regulatory sequences. First, the transient protoplast expression assay provides one possible way for the functional analysis of regulatory sequences in planta (Sect. 3.1). Second, regulatory sequences can be analyzed by computational means, which might give hints that form a basis for further functional analysis (Sect. 3.2). 3.1. Transient GUS-Expression Assay in Protoplasts 3.1.1. Isolation and PEGMediated Transformation of Protoplasts

For the generation of protoplasts from liquid cell cultures we use the protocol described by Schütze et al. in this volume; therefore, only the method for assaying promoter activation is described here using the uidA reporter gene (GUS; E.C. 3.2.1.31). The GUS enzyme (and other glycosidase enzymes) hydrolyzes preferred sugars (10), which allow the development of fluorescent and chromogenic substrates. These enable direct measurements of the protein amount by measuring the substrate turnover (Fig. 20.1). The method described here uses a driver- (typically carrying a gene encoding a transcription factor) and a reporter-plasmid (typically a promoter of interest fused upstream to a reporter gene sequence). This workflow can be applied readily

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for the analysis of regulatory elements in the absence of a driverplasmid yet responding to applied stimuli that can be mapped to specific cis-elements using deletion and mutagenesis strategies. A protocol for the isolation and PEG-mediated transformation of protoplasts from liquid cell suspension cultures is described in this volume by Schütze et al. and the steps given below are to complement these standard methods. If one wishes to use a cell wall digestion devoid of any sugar, the reader is referred to the method of Dangl et al. (1987) (11). 3.1.2. Co-transfection of Driver- and GUS Reporter-Plasmids

1. For each of the driver- and reporter-plasmid combinations and control-plasmids, 250 µl of protoplast suspension adjusted to a density of 2 × 106 cells/ml is used per transformation experiment (see Note 1). This equals a total number of about 500,000 cells. 2. Slowly add 30 µg of total plasmid DNA dissolved in not more than 30 µl water (see Note 2). 3. Perform transfection of two plasmids (or more) with the same amount of each driver- and reporter-plasmids (here 10 µg each) and a standardization plasmid (e.g. 5 µg) (see Note 3). 4. Wait 5 min and then add very slowly 250 µl PEG-solution, mix gently, and incubate for an additional 15–20 min (see Note 4). 5. Gradually add a total volume of 10 ml of W5 solution; do not exceed 1 ml/min (see Note 5). 6. Centrifuge at 60–100 g for 5 min to collect protoplasts. 7. Resuspend the protoplasts in 2 ml of K3 medium. 8. Incubate at 26°C in darkness (see Note 6).

3.1.3. GUS-Enzyme Activity Assay

All standard protocols concerning measuring the amount of GUS activity are based on the original protocol by Jefferson et al. (1987) (6). The method is recapitulated here and the measurement of GUS activity is based on the fluorescent substrate 4-methylumbelliferyl (MU) released by the hydrolysis of 4-methlyumbelliferylβ-D-glucuronide (4-MUG).

3.1.4. Method for Measuring GUS-Enzyme Activity

1. Slowly add 10 ml of MMM solution to the 2 ml of protoplast suspension (~500,000 cells) and mix carefully. The cells are collected by two centrifugation steps. First for 10 min at 400 g; the supernatant is discarded cautiously without agitating the protoplasts (sediment). Transfer protoplasts carefully to a 1.5-ml reaction tube and centrifuge a second time for 30 s at 400 g. The majority of buffer is removed, leaving an almost dry pellet. It can be frozen in liquid nitrogen and stored at −80°C until use (see Note 7). 2. To extract the GUS enzyme, The protoplasts must be lysed and completely disrupted. Often, this is accomplished by

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freezing in liquid nitrogen. Additionally, rapidly applying 800 µl extraction buffer to the frozen protoplasts and vortexing thoroughly for 30 s helps (2). Keep on ice afterwards or snap freeze in liquid nitrogen (see Note 8). 3. The extraction is then cleared of cellular debris by centrifugation at 14,000 g for 10 min at 4°C (see Note 9). 4. The fluorescence of 4-MU is detected by excitation at 365 nm and measuring emission at 455 nm. Use any suitable cuvette or plate fluorimeter. It is necessary to calibrate each experiment to a freshly prepared MU standard (purchased from a commercial manufacturer), for example, in 100 nM, 1 µM, and 10 µM 4-MU concentrations in extraction buffer (see Note 10). 5. The reaction buffer is recommended to contain a final concentration of 1 mM 4-MUG in extraction buffer (2, 6). Prepare a reaction solution of 2 mM 4-MUG in extraction buffer and mix this 1:1 with the cleared protein extract. 6. Add 100 µl protein extract to 100 µl 4-MUG reaction solution (2, 12). The reaction is run typically for 60 min and time points are taken at regular intervals (i.e., every 10 min, including the zero time point). 7. Stop the reaction by transferring 20 µl reaction mixture to a tube (or well) containing 180 µl Na2(CO3) solution (roughly a 1:10 volume ratio) (see Note 11). 8. Determine the protein concentration of the plant extracts using the method of Bradford (1976) (13). The GUS specific activity must be given as pmol 4-MU per second per mg total protein (4-MU/s/mg) to normalize between samples (see Note 12). 9. Calculate the linear increase in 4-MU over time using normalized values (pmol 4-MU mg protein/time) (Fig. 20.1b) (see Note 13). Alternatively, one can normalize the GUS activity to the amount of DNA or relative LUC activity if an additional vector has been transformed simultaneously as internal control (see Note 12, 13). 3.2. Computer Aided Sequence Analysis – Searching for Cis-Regulatory Elements (CREs)

The previous section should provide a guideline for the functional analysis of regulatory sequence in planta. Subsequently, many investigators take their regulatory sequence and search for the presence of known or yet unknown DNA motifs with the help of computer programs. Major advances in computer aided regulatory sequence analysis have been made during the past years. To date, there are several databases for eukaryote cis-regulatory elements – some specialized on plant cis-elements – and there are tools for analysis based on various algorithms. With several eukaryote genomes fully sequenced, many existing analysis programs have been trained on several genome sequences and thus

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317

have been improved by the pace of the vastly growing genomic era. On the other hand, the number of well-characterized regulatory elements is comparably low. Many motifs have been computationally deduced from the analysis of co-regulated genes of expression profile experiments using microarrays. Although there is a significant probability that those motifs harbor a certain regulatory function, their contribution to gene expression has often not been experimentally verified. This is a problem as databases for transcription factor binding sites exist, which include these computationally predicted motifs in their list of functionally characterized elements. Therefore, one has to say that there is no standardized workflow that ensures a “true” result. Moreover, the abundant numbers of databases compose the same cisregulatory element in different ways, to suit their specific needs for the algorithm used. Similarly, there are different calculation methods that deduce significant motifs by different means. On the following pages we provide an overview over both, the cis-regulatory element databases and the available analysis tools, starting with a brief introduction into cis-regulatory elements in plant regulatory sequences. 3.2.1. Functional Classification of Regulatory DNA Motifs

Generally, there are three different ways in which the presence of a certain DNA pattern can affect gene expression: 1. A DNA motif constitutes a binding site for regulatory proteins that have an indirect or direct effect on the transcription machinery. The nature of the resulting gene expression changes might be positive or negative, that is, more or less gene-transcripts, respectively. From what we know so far, a single protein might function as an enhancer under a certain condition and might act as a repressor of gene expression in a different context. 2. In eukaryotes, certain DNA patterns can be targets for in vivo methylation by methyltransferases at cytosine nucleotides. Although the function of 5-methylcytosines in plant regulatory sequences is not yet fully understood, their effect on transcription, silencing, and the positioning of nucleosomes has been shown several times. While symmetric methylation at cytosines in CpG or CpNpG patterns has been associated with the expression strength, asymmetric methylation at CpNpN-sites is assumed to control expression throughout different developmental stages (14). 3. There is a possibility that nucleotide composition affects gene expression changes via alterations in the biophysical properties of DNA, which results in conformational changes. For example, it has been shown for some simple sequence repeats that a Z-form DNA has a high probability to form also under conditions inside a cell. Other motifs have been

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described to mediate DNA bending (15). Similarly, the formation of hairpins in extended palindromic sequences is known from prokaryotes to abort gene expression. It has not been ruled out that similar conformational changes in DNA might function in eukaryotes as well. 3.2.2. Describing Transcription Factor Binding Sites

When analyzing a DNA motif that might be a functional transcription factor binding site with different bioinformatics tools, we require the information on how the respective binding site is recognized by the program (16). As our goal is to propose a workflow for the analysis of a single regulatory DNA sequence, we have chosen 1.5-kb of the flagellin-responsive receptor kinase promoter (SIRK/FRK; At2g19190) from Arabidopsis thaliana as a representative sequence to demonstrate general principles of the computational algorithms. The WRKY transcription factor binding sites, the W-boxes are present in this promoter 15 times and have been functionally characterized (3, 4). Here, they serve as examples for a typical transcription factor binding site in a regulatory sequence.

3.2.2.1. Strict Motif Consensus

Some of the analysis programs or databases use perfectly matching DNA-consensus sequences as input or output. They are generally unambiguous and consist of only one strict sequence. In cases where a transcription factor binds to more than only one related motif, each of the consensus sequences has to be entered independently. As a consequence, only one strict consensus will be returned if a database lists only one of the possible sequences. In case of our example, there exist two related W-box core motifs that are bound by WRKY transcription factors, TGACT and TGACC (Fig. 20.2a). Both are of about the same frequency in the SIRK/FRK promoter. In databases with strict motif consensus option, only one of the two possible motifs might be listed, which reduces the number of detected W-boxes by 50%.

3.2.2.2. IUPAC Motif Consensus

The IUPAC (International Union of Pure and Applied Chemistry; http://www.iupac.org/) recognizes not only the four bases A, C, G, and T as nucleotides, but 11 additional letters to describe all possible sequences. Hence, variable or ambiguous nucleotides can be expressed using the degenerate IUPAC code. In case of our example, the two strict W-box consensus sequences can be summarized with TTGACY for the sense orientation (Fig. 20.2b). In antisense orientation, this would be displayed as RGTCAA. Here, Y is C or T and R is A or G.

3.2.2.3. Position Weight Matrix (PWM)

Position weight matrices (PWM) are derived from sequence alignments and describe a matrix of the proportion of each nucleotide at each position in a given motif (Fig. 20.2c). There are cis-regulatory element databases that use the PWMs to search for motifs. PWMs have an advantage over an IUPAC-based sequence

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319

Fig. 20.2. Describing transcription factor binding sites. The W-box motifs from the SIRK/FRK promoter sequence (3) have been chosen as an example to explain the different ways of describing a cis-regulatory element (CRE). (A) Strict consensus sequences of the two possible W-box sequences. (B) Degenerate IUPAC consensus representing both possible versions of the W-box, where Y stands for C or T. (C) Building a position weight matrix (PWM) on the basis of the 15 W-box motifs present in the SIRK/FRK promoter sequence. The number of nucleotides per position is recorded in the matrix and can be used to deduce an extended IUPAC-consensus. (D) A different way to display the same information on the W-box sequences is to use a sequence logo (http://weblogo.berkeley.edu//). Here, the sizes of the nucleotide displays are proportional to their frequency at a certain position. Using the W-boxes from the SIRK/FRK promoter with a TTGAC pentamere instead of the conserved TGAC core-motif changes the frequencies of the neighboring nucleotides.

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code, because it has a weighted score for the nucleotide usage at each position. In a degenerate IUPAC-code all contributing nucleotides are of equal weight. Finally, the number of nucleotides per position is recorded in the matrix and can be used to deduce an extended motif-consensus (Fig. 20.2c) in IUPAC code. In case of our example, the W-box penta-nucleotide TGACY is present 15 times in the SIRK/FRK promoter. Hence, each of the motifs contributes 1/15 to the PWM. While the TGAC core is invariant (each position has a weight of 15/15 in the matrix), the Y is composed of a C with a weight of 8/15 and a T with a weight of 7/15. Hence, in a PWM-based investigation the C is considered to have a slightly higher probability to occur in a sequence than the T. Using this approach, an extended consensus for functional W-boxes can be deduced (T N N T T G A C Y A W A W) on the basis of only the SIRK/FRK promoter. 3.2.2.4. Sequence Logo

Sequence Logos are a versatile way to display PWM-based DNA motifs. Some organizations provide free online tools to establish sequence logos on the basis of an alignment (e.g. http:// weblogo.berkeley.edu/). The sizes of the displayed nucleotides are proportional to their frequency at the respective position. Although the intention of sequence logos is to display a DNA motif in a weighted manner, it also allows a preliminary investigation of neighboring positions flanking a motif. In case of our example, the invariant TGAC core is located at positions 5–8 (Fig. 20.2d, upper panel). About the same number of C or T nucleotides are at position 9. It is worth noting that the four nucleotides flanking the TGACY consensus at the 3’ end are significantly enriched for A/T, while the position immediately 5’ of the core favors a T at position 4. To get a closer insight into how the sequence environment changes with respect to the consensus used, we show the same analysis with an invariant TTGAC core as an example (Fig. 20.2d, lower logo). When comparing the sequence logos with TGAC or TTGAC core motifs, the proportion of C/T at position 9 has not changed. In contrast, there are notable alterations in the frequency at other positions: The occurrence of T now dominates position 1 and A is of higher frequency at position 10.

3.2.3. Bioinformatic Tools for CRE Analysis of Plant Genomes

On the following pages we provide information on several tools for the analysis of plant regulatory sequence analysis. These analysis programs have been tested for their suitability and the results are summarized in accompanying tables. A general workflow for the iterative process of regulatory sequence analysis is shown in Fig. 20.3. The regulatory sequence of interest is retrieved from a database or by sequencing. There are two possible ways for the computational analysis of a regulatory sequence; they are searching either for CREs of known function

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Fig. 20.3. Proposed workflow for the analysis of a single regulatory DNA sequence. The two possible ways for the analysis of a regulatory sequence by computational means are shown. Either sequences of known cis-regulatory function (CRE) can be identified with the help of databases, or one can search the sequence for unknown motifs on the basis of various significance models. Both ways of analysis will identify motifs that have to be tested for functionality in a biological system. This again can be performed with the transient expression assay in protoplast.

or for motifs of yet unknown function. In either way, the sequence motifs identified so far lack any evidence of importance for regulatory capacity in the native context. Interestingly, the number of CREs is proportional to the size of the regulatory sequence and to the number of responsive conditions under which a gene is expressed (17). However, two recent reports could demonstrate by computational analysis that functional CREs have a tendency to be localized closer to the gene sequence (9, 17). Hence, an identified motif located closer to the start of a gene has a higher probability to be of regulatory importance. Therefore, additional functional assays are indispensable. To test for functionality of the motifs identified, the transient protoplast assay is a suitable method for rapid investigation of mutated motifs in the native regulatory sequence or as multimerized sequences. Moreover, direct binding of a transcription factor or the deletion of the

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motif by single nucleotide exchanges gives further insight into its role in regulating gene expression. 3.2.3.1. Plant Cis-Regulatory Element (CRE) Databases

After you have identified regulatory regions of interest, it is possible to search for transcription factor binding sites (TFBSs) and CREs electronically primarily using web-based bioinformatic tools. We have summarized several databases in Table 20.1 that catalogue known plant TFBSs. These databases allow the retrieval of published binding sites from plants. They are typically recorded as DNA sequences using only one side of the strand and occasionally contain IUPAC letters or are described as a position weight matrix (PWM). As most computer groups are trying to continually develop and expand their analysis tools, it is advisable to watch for updates and modifications, including URL changes to any of these web-based tools. The tables include Unique Identifiers [uid] or PubMed Unique Identifiers [PMID] (http://www.ncbi.nlm.nih.gov/entrez/ query/Pmc/pmchelp.html) for rapid access to the referenced papers in PubMed (http://www.pubmed.gov). The Plant Cisacting Regulatory DNA Elements (PlaCE) (18) database allows one to search for and retrieve information on individual TFBS, and they also allow one to look for homologous TFBS using their Homology Search tool. TRANSFAC (19) is primarily a commercial analysis suite, but they have several programs or algorithms available for non-profit public, such as an extensive database for TFBSs of all species; The public programs and datasets are available after a free registration. Finally, PlantCARE (20) attempts to track plant transcription sites and provides the Query CARE tool allowing one to query through their database and retrieve the CREs and known target genes.

3.2.3.2. Regulatory Sequence Retrieval

Retrieving a known promoter (or regulatory sequence) can be a simple or complicated matter and is usually dependent on the number of sequences one wishes to analyze. A regulatory region is defined as any stretch of a sequence that has been mapped

Table 20.1 Cis-regulatory element databases Database

PMID [Ref.]

URL

PlaCE

9847208

http://www.dna.affrc.go.jp/PLACE/

PlantCare

11752327

web site moved*, http://bioinformatics.psb.ugent.be/ webtools/plantcare/html/

TRANSFAC

17118134

http://www.gene-regulation.com/pub/databases.html

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323

by functional experimentation, and it can be a specific part of a promoter region, a specific intron, a 3¢UTR or any other part of a gene shown to be functionally significant for its regulation. Once this region is known, it can be extracted from any electronic sequence file by simply highlighting the sequence from an electronic file and copying it to a new text file. Generally, when one searches for CREs, one should first focus on those that are most likely responsible for the observed responses. For example, if you are interested in cold-induced genes, then searching for coldresponsive elements in your upstream promoter region might bring immediate results. “Promoters” are sometimes regarded to be upstream of the coding region (ATG) or the transcription start site (TSS). The ATG is frequently used as reference when no or little information for the TSS is available. While it is possible to extract promoters by hand from genomic sequences (e.g. BACs, YACs, chromosomal pseudomolecules), it is particularly time-consuming when one needs promoters from many genes. Fortunately, bioinformatic tools that automatically do this for us already exist. These programs typically extract the promoter before continuing with other parts of their analysis. Here, we call this useful automated function dynamic promoter extraction (dPE). Programs that do not dynamically extract promoters require that the user provides the sequence data for them. Sequence retrieval tools exist that extract the desired sequences from a database (usually returned in FASTA format) to download to your own computer. Once one has obtained the sequences from such programs, they are available for further analysis, such as input into other programs. Three programs that can extract promoter sequences (and other sequence bits as well) are mentioned in Table 20.2. Regulatory Sequence Analysis (RSA) Tools (21) is an excellent analysis suite for searching for known and unknown motifs using various approaches. Additionally, RSA Tools provide one of the few program tools that can extract promoters from many annotated genomes by using the Sequence Retrieval tool. This is

Table 20.2 Regulatory sequence retrieval Database - Tool – Task

PMID [Ref.]

URL

RSA tools - sequence retreival

10641039

http://rsat.scmbb.ulb.ac.be/rsat/

Motif mapper - creating your own database

17137509

http://www2.mpiz-koeln.mpg.de/coupland/ coupland/mm3/html/

TAIR - sequence bulk download and analysis

12519987

http://www.arabidopsis.org/tools/bulk/ sequences/index.jsp

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easily done by choosing the correct genome, entering the start and stop positions, and choosing from which the genes (all or a selection) for which one wishes to retrieve promoters. Although originally designed for yeast, RSA offers many tools and implements a variety of several downstream analysis programs that are useful for the investigation of any kind of DNA sequence. The Arabidopsis Information Resource (TAIR) (22) maintains a database of genetic and molecular biology data for the model higher plant Arabidopsis thaliana. TAIR offers many tools concerning sequence retrieval from Arabidopsis for the public, and in addition, they offer a large variety of tools and information concerning research on Arabidopsis. The TAIR tool Sequence Bulk Download and Analysis allows a user to download promoter (and other sequence-type) datasets by entering Arabidopsis Genome Identifiers (AGI) (23). The sequences can be returned in FASTA or tab-delimited formats. TAIR is strictly limited to Arabidopsis thaliana. Finally, we mention a non-web-based analysis suite, which has certain advantages and disadvantages. The Motif Mapper analysis suite is a collection of open source visual basic scripts, which are useful for analyzing single and multiple promoters for IUPAC compatible word motifs both individually and collectively (9). The principle advantage of Motif Mapper is that it can process large datasets, and this is done independently of the internet. The major disadvantage of Motif Mapper is that its scripts are command line, meaning that the user enters data by a series of question prompts and requires a steep learning curve. Similar to RSA Tools (21), it provides a tool for extracting promoters from various organisms. In the case of Motif Mapper, it can extract sequences from any GenBank (24) flat file. A flat file is a text file containing GenBank annotations followed by the sequence. GenBank formatted data are seen when browsing sequences using a web browser (e.g., Firefox, Internet Explorer, Opera, etc.) at The National Center for Biotechnology Information (NCBI; http://www.ncbi.nlm.nih.gov/). To save a GenBank file as text and not an HTML file, one must either copy and paste the GenBank file data into a text file program (e.g., Editor, Notepad, Wordpad, Textedit, Vi, etc.) or when viewing the sequence, change the Display option to GenBank and click on the SendTo option and choose Text. Thereafter, the file can be saved directly as text. After this is done, the user must follow the instructions in Creating your own database from the Motif Mapper website, which explains how to prepare the GenBank files for extraction and analysis by the suite. 3.2.4. Regulatory Sequence Analysis Programs

Tables 20.3 and 20.4 summarize some programs for analyzing promoter sequences for known or unknown cis-regulatory elements. The analysis programs have been divided into those that can be used to search for CREs in a single promoter (Table 20.3,

•







•

•

•

•

•




AthaMap - gene analysis

Agris - cisDB

DoOP - plant dataset

TAIR - motif finder

TAIR - Patmatch •




Promomer

PlaCE

PlantCare

•

•

•

•

•

•




•










•







•




























•

•

Predefined clusters Arabidopsis only

Word matching Word matching Word matching

•

•

•

•




One sequence at a time 11752327

Word matching

9847208

12519987

12519987

15608291

15733318

17148485

Vascular plants only

Arabidopsis only

Arabidopsis only

15960624

17137509

Word matching

Word matching

Arabidopsis only

Word matching

•

Arabidopsis only

Enumeration

GenBank flat file format

•




10641039

web site moved*, http://bioinformatics.psb.ugent.be/webtools/plantcare/ html/

http://www.dna.affrc.go.jp/PLACE/

http://www.arabidopsis.org/cgi-bin/ patmatch/nph-patmatch.pl

http://www.arabidopsis.org/tools/bulk/ motiffinder/index.jsp

http://doop.abc.hu/

http://arabidopsis.med.ohio-state.edu/ AtcisDB/

http://www.athamap.de/search_gene.php

http://bbc.botany.utoronto.ca/ntools/ cgi-bin/BAR_Promomer.cgi

http://www2.mpiz-koeln.mpg.de/ coupland/coupland/mm3/html/

http://www.gene-regulation.com/pub/ programs.html

http://rsat.scmbb.ulb.ac.be/rsat/

PMID [Ref.] URL

Commerical, with pub- 17118134 lic releases

Incoporated genomes

Word matching

Various

Various

•

•

•

Restrictions

dPE (dynamically extracted promoters), SE (statistical evaluation), UDM (user defined motifs), IUPAC (IUPAC wobble DNA code), MP (motif prediction), IGBM (identify genes by motifs)




•







•

•

•




•




Motif mapper

•

•

•




TRANSFAC

•

•

•

RSA - tools

UDM IUPAC MP IGBM Method(s)

•

dPE SE

Algorithm/suite

Table 20.3 Regulatory sequence analysis programs for single sequence inputs

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Sect. 3.2.4.1) or into those that require more than one promoter sequence as input (Table 20.4, Sect. 3.2.4.2). Although one may want to identify TFBS that are defined as “statistically significant” from an analysis and hence, disregard all others, the reader must keep in mind that all results are based on probabilistic methods. Moreover, the analysis programs vary in their background models, which may or may not differ from each other, and what one program may consider significant, another may not. Furthermore, it could be the case that an element not considered important by a statistical appraisal is the very element important for the regulation you observe. Thus, we have noted which programs provide a statistical evaluation (SE) and reassuringly, many of these programs also return the “non-significant motifs” for the user to muse over. Likewise, it is important to know which input sequences are accepted. We note that some programs only allow querying sequences against their defined set of TFBSs, whereas other programs allow the user to enter their own user defined motifs (UDM). In addition to searching a sequence for motifs, some sites offer tools that predict unknown CREs (motif prediction, MP). Finally, querying the promoter dataset for the presence of a CRE can help the researcher to narrow down the dataset to promoters that possess an interesting element (identify genes by motifs, IGBM). Typically, a promoter is depicted in the 5¢ to 3¢ orientation of transcription. Most programs do not just return the identified TFBSs, but also give the position of the TFBSs with respect to the input sequence. We have also noted some of the major limitations (restrictions) imposed by each analysis tool. 3.2.4.1. Motif Analysis Based on a Single Sequence Input

As mentioned, RSA Tools can dynamically extract promoter sequences (Sect. 3.2.3.2). Additionally, RSA Tools can locate user defined motifs (UDM) or predicted motifs (MP) with various programs using primarily over-representation statistics (21). TRANSFAC (19) has a wide variety of useful programs and analysis tools, which are too numerous to iterate here. It is highly recommended to peruse their web site carefully. Motif Mapper (9) can search through any list (unlimited) of user defined FASTA or RAW sequences for an unlimited list of query motifs; however, searching for novel motifs is not possible. The analysis suite Athena (25) is not recommended for the analysis of one promoter, but could use it as cross-reference for other programs mentioned here. The PlaCE database (18) provides the Signal Scan Search tool for scanning one sequence (FASTA format, only the first sequence is processed) against CREs in their database. The PlaCE tool PLACE Web Signal Scan accepts an uploaded text file containing multiple FASTA sequences for CRE analysis and returns the results by e-mail. Promomer (26) searches for overrepresented elements, but also can identify genes with a queried




•

•




•




MEME

Athena

Cis-Element searcher

Weeder

RSA - Tools

MDscan







•




•

•










•

•

•

•




•

•

•




•




Oryza only

Arabidopsis only

Version dependent

FASTA format

FASTA format, 50kb

Incoporated genomes Job limits

Various Word enumeration and PWMs

Word enumeraMotifs ³ 6 bp, locator tion and PWMs tool

Word matching, MEME

Hypergeometric probability distribution

•

•

MEME

Gibbs motif sampling

Gibbs motif sampling










Restrictions

12101404

10641039

16845071

NO PMID

16223790

16845028

12015892

10698627

PMID [Ref.]

http://ai.stanford.edu/~xsliu/ MDscan/

http://rsat.scmbb.ulb.ac.be/ rsat/

http://159.149.109.16:8080/ weederWeb/

http://hpc.irri.cgiar.org/tool/ nias/ces

http://www.bioinformatics2. wsu.edu/cgi-bin/Athena/ cgi/home.pl

various web sites; http:// meme.nbcr.net

http://homes.esat.kuleuven. be/~thijs/Work/MotifSampler.html

http://atlas.med.harvard.edu/

URL

dPE (dynamically extracted promoters), SE (statistical evaluation), UDM (user defined motifs), IUPAC (IUPAC wobble DNA code), MP (motif prediction), IGBM (identify genes by motifs)

•

•

•

•

•

•

•

•

•







•




MotifSampler

•




•




AlignACE

UDM IUPAC MP IGBM Method(s)

SE

Algorithm/suite dPE

Table 20.4 Regulatory sequence analysis programs for multiple sequence inputs

Bioinformatic Analysis of Promoter Elements 327

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motif in various sets of dynamically extracted promoters (limited to Arabidopsis). The TAIR tool Motif Finder (22) returns over-represented hexamers using a background model of 500 or 1000 bps from the ATG (Arabidopsis only). The TAIR tool Patmatch searches TAIR sequence datasets with IUPAC compatible motifs allowing the researcher to identify promoters that carry the sought motif. One major drawback is that it is not yet possible to restrict datasets by gene identifiers and one gets every gene identified; To circumvent that, one can download the entire output file and sort for the genes of interest. The AthaMap’s Gene Analysis (27) tool extracts promoters dynamically using an AGI input list and identifies CREs in their database; this algorithm is one of the few that allow mismatches. The Arabidopsis Gene Regulatory Information Server (AGRIS) (28) carries extensive data on transcription factor families in Arabidopsis and is a good source for initiating research with transcription factors. The AGRIS tool AtcisDB (29) accepts an AGI input list and returns a list of promoters, but only if they have already been mapped and recorded in their database. Databases of Orthologous Promoters (DoOP) (30) analyzes defined clusters of promoters, yet the user query is a single promoter. One can identify which CREs the clusters may have in common with other co-regulated genes (and promoters) or from homologous genes. Hence, DoOP is not restricted to Arabidopsis and it allows the user to download all of the promoter sequences for further analysis. PlantCARE (20) can identify TFBSs listed in their database in a single FASTA formatted sequence using their Search for CARE tool. 3.2.4.2. Motif Analysis Based on a Multiple Sequence Input

Investigation with multiple sequence input is based on the idea that the input sequences have something in common that has to be unveiled by the analysis program. For example, genes with similar expression profiles in microarray experiments might be regulated by the same cis-regulatory elements. Thus, the motif shared in most of the regulatory sequences might be detected using one of the following analysis tools. The programs Aligns Nucleic Acid Conserved Elements (AlignACE) (31), MotifSampler (32), and MDscan (33) require a set of sequences to calculate novel motifs (MP). The results can vary from one run to the next, so it is advised to make several analyses using the same and different input parameters. The algorithm termed Multiple Expectation maximization for Motif Elicitation is commonly referred to as MEME (34, 35). MEME calculations are implemented in many analysis suites, and it is not uncommon to find that many web-based sites are including a version of MEME for novel motif prediction. The link provided in Table 20.4 is therefore only one example of an implemented MEME; it is advised to search the web for others. Weeder (36) builds PWM for TFBS based on a promoter set

Bioinformatic Analysis of Promoter Elements

329

(co-regulated or homologues genes) using promoter/enhancer and 5’UTR regions as a background model. For larger jobs, there is a stand alone version one can download and run on your own computer. Additionally, Weeder also provides the Locator tool, which can find motifs that must be at least 6 nucleotides long and allow mismatches, in a single sequence. RSA Tools also allows the analysis of promoter clusters with a variety of methods: oligoanalysis (words), dyad-analysis (spaced pairs), position-analysis, and consensus (matrices), just to name a few. We highly recommend visiting RSA Tools to learn all that they have to offer (21). Cis-Element Searcher (34, 35) is limited to Oryza sativa, but it carries a wide set of valuable analysis tools. Cis-Element Searcher dynamically extracts promoters, it allows user defined motifs, has a database of known CREs, and has implemented a MEME to predict CREs. Athena (25) is restricted to Arabidopsis and is best used to search a cluster of promoters for functionally characterized motifs that are statistically enriched in the entire cluster. Additionally, Athena provides a statistical evaluation for enriched GO annotations, maps predicted CpG islands, and has a MEME integration in the works. 3.2.4.3. Special Analysis Suites

Finally, we would like to mention other analysis suites that are useful for identifying CREs (see Table 20.5). TAIR maintains a webpage entitled Plant Promoter and Regulatory Element Resources, which lists invaluable analysis tools for CRE research for all plant life; keep this page bookmarked and make an occasional visit on your searches through the World Wide Web. The FLAGdb++ (37) application is a program that works at the genomic scale, aiming at investigating genome evolution and the relationships between the structure and function of genes. FLAGdb++ approaches

Table 20.5 Special analysis suites Database

PMID [Ref.]

URL

FLAGdb++

14681431

http://urgv.evry.inra.fr/projects/FLAGdb++/ HTML/index.shtml

The SIGnAL Arabidopsis Methylome Mapping Tool

16949657

http://signal.salk.edu/cgi-bin/methylome

TAIR - Plant Promoter and Regulatory Element Resources

12519987

http://www.arabidopsis.org/portals/genAnnotation/genome_annotation_tools/cis_element.jsp

UCSC Genome Browser

16949657

http://epigenomics.mcdb.ucla.edu/ DNAmeth/project.html

330

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this goal by exploring the genomes of Arabidopsis thaliana, Oryza sativa, Vitis vinifera and Populus trichocarpa. FLAGdb++ provides a means of interactively viewing and scrolling through genomic annotations on chromosome pseudomolecules, and displays transcripts, coding regions, GO annotations, transcription factor families, Affymetrix probes, and much more. Furthermore, FLAGdb++ has the Search Pattern tool, which maps IUPAC compatible motifs to the entire genome and a Blast tool option (38), which maps blast hits (with a chosen cutoff) to all positions on the chromosomes. The SIGnAL Arabidopsis Methylome Mapping Tool allows one access to DNA methylation sites in Arabidopsis, which were obtained by a functional mapping experiment at the genomic scale (14). A genome-wide mapping of DNA methylation sites in Arabidopsis (14) has been deposited at the UCSC Genome Browser (http://epigenomics.mcdb.ucla.edu/DNAmeth/) (39). The Arabidopsis UCSC Genome Browser carries not only information on methylation in wild type plants but also in methylation deficient mutants. Additionally, the Arabidopsis UCSC Genome Browser also shows nucleosome density plots, the location of small RNAs, and TF binding sites from AtcisDB against TAIR gene models along all chromosomes. 3.2.4.4. Concluding Remarks

Here, we have provided a short summary of programs that we find useful and reliable for routine investigations that aims at functional validation of CREs in the laboratory. We cannot stress enough how imperative it is that one reviews each website carefully before starting the analysis. In most cases, help files and related literature are available, which provide an insight into what each tool or program can do. As every program will give you some “results”, the wise use of the results will lead to additional validation in the laboratory.

4. Notes 1. During the protoplast preparation (Schütze et al., this volume) you may have some debris or non-perfect cells. You may wish to be stringent on the cell quality or tolerant; however, the cell count should only be based on the number of cells with a round and healthy appearance. 2. You may wish to use less (down to 10 µg) of DNA, and a good rule of thumb is 30 µg. Be aware that you should always maintain the total volume “DNA solution” at a minimum of 20 µl and at a maximum of 30 µl. In case you are not sure if your construct is expressing well, some protocols have

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found that linear plasmid gives better transient expression (40), while others have found that linearized plasmids have no benefit (5). Every plasmid may have different transient expression levels due to DNA topology (40), which cannot be entirely avoided except by using the same plasmid backbones for co-transformation experiments. Furthermore, some researchers have found better results when using dam− strains of Escherichia coli (strain GM2163) (41) for plasmid amplification. The purity of plasmid DNA can play a large role in efficiency, and it is suggested to purify DNA with CsCl or silica-matrix column chromatography. Often however, an RNAse digestion and protein removal with phenol/chloroform/ isoamylalcohol followed by an isopropanol precipitation also works well in most cases. 3. The efficiency of the co-transformation will be dependent on the total transformation efficiency. For comparing experiments it is necessary to have a standardization plasmid. Some protocols use a separate plasmid, e.g., a plasmid expressing Luciferase (2), to estimate the total transformation efficiency. Of course, it would be better to have the standardization gene on the same plasmid; it could be based on luminescence (LUC), fluorescence (GFP), or simply the protein amount (detection with an antibody through Western blotting). We have commonly observed that 10% of all transformations are co-expressing when two plasmids are used. Fluorogenic detection of GUS activity is the most sensitive method for estimating the amount of GUS protein present. 4. The final concentration of PEG is 20%, which has been found to be the optimal concentration for the majority of protoplasts from various plant species (5, 40). The MMM solution contains Mg2+, which has been found to be critical and most effective for transformation of various plant species protoplasts (40), including Arabidopsis thaliana Columbia (5, 42), although Ca2+ occasionally may also be more helpful (40). 5. Any protoplast aggregates visible after addition of PEGsolution should resolve during this step. There are two likely causes for aggregates that do not dissolve: On the one hand the protoplasts were damaged or were not properly washed after the enzymatic digestion step. On the other hand, the plasmid DNA used harbors impurities or is concentrated too high. 6. The incubation duration is dependent on which proteins are being expressed and by which promoter. Promoters that require induction should be quantitatively tested for activation and reporter activity. The proper folding of fluorophores (i.e., green fluorescent protein (GFP), red fluorescent protein

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(RFP)) will take time (2–24 hours) depending on the protein and any modifications that have been made to the protein sequence. The highest transient expression is seen after 2–24 h (1), yet as long as the cells are in K3 medium, some will continue to express protein for up to 7 days or longer. 7. The protoplasts float in K3 medium and will not pellet, not even if they are centrifuged. Therefore, it is necessary to exchange the medium to one where the cells have the proper osmotic pressure, yet sink. MMM or W5 medium is sufficient for this. 8. Different methods are suggested, likely depending on the amount of material, the age of the protoplasts, and method of choice. It has been reported that the FastPrep System (Savant Instruments, model FP120) can yield improved results (43); however, as mentioned freezing is often sufficient. If desired, one can grind the material by using a small pestil (with or without sand) or by using glass beads and vortexing (6). 9. Extracts can be stored at −80°C for at least two months (6). 10. Fluorescence is linear and product sensitivity is dependent on the machine being used; therefore, it is possible to measure down to less than 1 nM 4-MU dependent on machine sensitivity (6). 11. The volumes are adjusted for protein extractions from 500,000 protoplasts. The original protocol from Jefferson et al. (1987) (6) ran a total reaction of 1 ml (presumably by adding 500 µl protein extract in 500 µl of 4-MUG reaction solution) and removing aliquots of 200–800 µl of Na2(CO3) stop solution. This was adapted to a 200-µl microtiter welled fluorimeter plates (2, 12). 12. One way to proceed is taking the slope between each data point and averaging them. Better would be calculating the slope using least-squares regression analysis. 13. The data points should be nearly linear without manipulation. If not, it is quite likely that the reaction was not properly mixed leading to unequal 4-MUG turnover rates. There are also reports that successfully use DNA content for normalization. The total DNA amount might reflect the number of cells that were lysed, assuming that the ploidy content or cell cycle divisions are equally distributed between samples. Two methods for calculating the DNA amount are measuring Horchst 33258 fluorescence (6) or SybrGreen I fluorescence (12). Moreover, one can quantify the DNA amount of the reporter-plasmid or standardization-plasmid by qRT-PCR for normalization. In addition, if one wishes to use LUC as a standardization measure, the reader is referred to the method described by Hartmann et al. (2).

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Acknowledgements The authors would like to thank Joachim Kilian and Friederike Ladwig for their critical comments and Sabine Hummel for technical assistance. References 1. Abel, S. and Theologis, A. (1994) Transient transformation of Arabidopsis leaf protoplasts: a versatile experimental system to study gene expression. Plant J. 5, 421– 427. 2. Hartmann, U., Valentine, W. J., Christie, J. M., Hays, J., Jenkins, G. I. and Weisshaar, B. (1998) Identification of UV/blue lightresponse elements in the Arabidopsis thaliana chalcone synthase promoter using a homologous protoplast transient expression system. Plant Mol Biol. 36, 741–754. 3. Robatzek, S. and Somssich, I. E. (2002) Targets of AtWRKY6 regulation during plant senescence and pathogen defense. Genes Dev. 16, 1139–1149. 4. Miao, Y., Laun, T., Zimmermann, P. and Zentgraf, U. (2004) Targets of the WRKY53 transcription factor and its role during leaf senescence in Arabidopsis. Plant Mol. Biol. 55, 853–867. 5. Damm, B., Schmidt, R. and Willmitzer, L. (1989) Efficient transformation of Arabidopsis thaliana using direct gene transfer to protoplasts. Mol. Gen. Genet. 217, 6–12. 6. Jefferson, R. A., Kavanagh, T. A. and Bevan, M. W. (1987) GUS fusions: betaglucuronidase as a sensitive and versatile gene fusion marker in higher plants. EMBO J. 6, 3901–3907. 7. Mathur, J. and Koncz, C. (1998) PEG-mediated protoplast transformation with naked DNA. Methods Mol. Biol. 82, 267–276. 8. Rushton, P. J., Reinstadler, A., Lipka, V., Lippok, B. and Somssich, I. E. (2002) Synthetic plant promoters containing defined regulatory elements provide novel insights into pathogen- and wound-induced signalling. Plant Cell 14, 749–762. 9. Berendzen, K. W., Stueber, K., Harter, K. and Wanke, D. (2006) Cis-motifs upstream of the transcription and translation initiation sites are effectively revealed by their positional disequilibrium in eukaryote genomes using frequency distribution curves. BMC Bioinformatics 7, 522.

10. Stoeber, F. (1961) Thèse de Docteur des Sciences, Paris. 11. Dangl, J. L., Hauffe, K. D., Lipphardt, S., Hahlbrock, K. and Scheel, D. (1987) Parsley protoplasts retain differential responsiveness to UV light and fungal elicitor. EMBO J. 6, 2551–2556. 12. Coté, C. and Rutledge, R. G. (2003) An improved MUG fluorescent assay for the determination of GUS activity within transgenic tissue of woody plants. Plant Cell Rep. 21, 619–624. 13. Ausubel, F. M., Brent, R., Kingston, R. E., Moore, D. D., Seidman, J. G., Smith, J. A. and Struhl, K. (1995) Current Protocols in Molecular Biology, John Wiley & Sons. 14. Zhang, X., Yazaki, J., Sundaresan, A., Cokus, S., Chan, S. W., Chen, H., Henderson, I. R., Shinn, P., Pellegrini, M., Jacobsen, S. E. and Ecker, J. R. (2006) Genome-wide high-resolution mapping and functional analysis of DNA methylation in Arabidopsis. Cell 126, 1189–1201. 15. Asayama, M. (2006) Regulatory System for Light-Responsive Gene Expression in Photosynthesizing Bacteria: Cis-Elements and Trans-Acting Factors in Transcription and Post-Transcription. Biosci. Biotechnol. Biochem. 70, 565–573. 16. D’haeseleer, P. (2006) How does DNA sequence motif discovery work? Nat Biotechnol. 24, 959–961. 17. Walther D., Brunnemann R. and Selbig J. (2007) The Regulatory Code for Transcriptional Response Diversity and Its Relation to Genome Structural Properties in A. thaliana. PLoS Genet. 3, e11 18. Higo, K., Ugawa, Y., Iwamoto, M. and Korenaga, T. (1999) Plant cis-acting regulatory DNA elements (PLACE) database Nucl. Acids Res. 27, 297–300. 19. Kel, A., Voss, N., Jauregui, R., Kel-Margoulis, O. and Wingender, E. (2006) Beyond microarrays: Finding key transcription factors controlling signal transduction pathways. BMC Bioinformatics 7(Suppl. 2), S13.

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20. Lescot, M., Déhais, P., Thijs, G., Marchal, K., Moreau, Y., Van de Peer, Y., Rouzé, P. and Rombauts, S. (2002) PlantCARE, a database of plant cis-acting regulatory elements and a portal to tools for in silico analysis of promoter sequences. Nucl. Acids Res. 30(1), 325–327. 21. Van Helden, J., Andre, B., Collado-Vides, J. (2000) A web site for the computational analysis of yeast regulatory sequences. Yeast 16, 177–187. 22. Rhee, S. Y., Beavis, W., Berardini, T. Z., Chen, G., Dixon, D., Doyle, A., GarciaHernandez, M., Huala, E., Lander, G., Montoya, M., Miller, N., Mueller, L, A., Mundodi, S., Reiser, L., Tacklind, J., Weems, D. C., Wu, Y., Xu, I., Yoo, D., Yoon, J and . Zhang, P. (2003) The Arabidopsis Information Resource (TAIR): a model organism database providing a centralized, curated gateway to Arabidopsis biology, research materials and community. Nucl. Acids Res. 31, 224. 23. Arabidopsis Genome Initiative (2000) Analysis of the genome sequence of the flowering plant Arabidopsis thaliana. Nature 408, 796–815. 24. Benson, D. A., Karsch-Mizrachi, I., Lipman, D. J., Ostell, J. and Wheeler, D. L. (2006) GenBank. Nucl. Acids Res. 34(Database issue), D16–20. 25. O’Connor, T. R., Dyreson, C., and Wyrick, J. J. (2005) Athena: a resource for rapid visualization and systematic analysis of Arabidopsis promoter sequences. Bioinformatics 21, 4411–4413. 26. Toufighi, K., Brady, S. M., Austin, R., Ly, E. and Provart, N. J. (2005) The botany array Resource: e-northerns, expression angling, and promoter analyses. Plant J. 43, 153–163. 27. Galuschka, C., Schindler, M., Bulow, L. and Hehl, R. (2007) AthaMap web tools for the analysis and identification of co-regulated genes. Nucl. Acids Res. 35(Database issue), D857–862. 28. Palaniswamy, S. K., James, S., Sun, H., Lamb, R. S., Davuluri, R. V. and Grotewold, E. (2006) AGRIS and AtRegNet: A platform to link cis-regulatory elements and transcription factors into regulatory networks. Plant Physiol. 140, 818–829. 29. Molina, C., and Grotewold, E. (2005) Genome wide analysis of Arabidopsis core promoters. BMC Genomics 6, 25. 30. Barta, E., Sebestyen, E., Palfy, T. B., Toth, G., Ortutay, C. P. and Patthy, L. (2005) DoOP: Databases of orthologous

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promoters, collections of clusters of orthologous upstream sequences from chordates and plants. Nucl. Acids Res. 33(Database issue), D86-D90. Hughes, J. D., Estep, P. W., Tavazoie, S. and Church, G. M. (2000) Computational identification of cis-regulatory elements associated with groups of functionally related genes in Saccharomyces cerevisiae. J. Mol. Biol. 296, 1205–1214. Thijs, G., Marchal, K., Lescot, M., Rombauts, S., De Moor, B., Rouzé, P. and Moreau, Y. (2002) A Gibbs sampling method to detect overrepresented motifs in the upstream regions of coexpressed genes. J. Comput. Biol. 9, 447–464. Liu, X. S., Brutlag, D. L. and Liu, J. S. (2002) An algorithm for finding proteinDNA binding sites with applications to chromatin immunoprecipitation microarray experiments. Nat. Biotechnol. 20, 835–839. Bailey, T.L. and Elkan C. (1994) Fitting a mixture model by expectation maximization to discover motifs in biopolymers. Proc. Int. Conf. Intell. Syst. Mol. Biol. 2, 28–36. Bailey, T. L., Williams, N. Misleh, C., and Li. W. W. (2006) MEME: discovering and analyzing DNA and protein sequence motifs. Nucl. Acids Res. 34(Web Server issue), W369–373. Pavesi, G., Mereghetti, P., Zambelli, F., Stefani, M., Mauri, G. and Pesole, G. (2006) MoD Tools: Regulatory motif discovery in nucleotide sequences from co-regulated or homologous genes. Nucl. Acids Res. 34, W566-W570. Samson, F., Brunaud, V., Duchene, S., De Oliveira, Y., Caboche, M., Lecharny, A. and Aubourg, S. (2004) FLAGdb++: a database for the functional analysis of the Arabidopsis genome. Nucl. Acids Res. 32(Database issue), D347–350. Altschul, S. F., Gish, W., Miller, W., Myers, E. W. and Lipman, D. J. (1990) Basic local alignment search tool. J. Mol. Biol. 215, 403–410. Hinrichs, A. S., Karolchik, D., Baertsch, R., Barber, G. P., Bejerano, G., Clawson, H., Diekhans, M., Furey, T. S., Harte, R. A., Hsu, F., Hillman-Jackson, J., Kuhn, R. M., Pedersen, J. S., Pohl, A., Raney, B. J., Rosenbloom, K. R., Siepel, A., Smith, K. E., Sugnet, C. W., Sultan-Qurraie, A., Thomas, D. J., Trumbower, H., Weber, R. J., Weirauch, M., Zweig, A. S., Haussler, D and . Kent, W. J. (2006) The UCSC Genome Browser Database: update 2006. Nucl. Acids Res. 34(Database issue), D590–598.

Bioinformatic Analysis of Promoter Elements 40. Negrutiu, I., Shillito, R., Potrykus, I., Biasini, G. and Sala, F. (1987) Hybrid genes in the analysis of transformation conditions. I. Setting up a simple method for direct gene transfer in plant protoplasts. Plant Mol. Biol. 8, 363–373. 41. Tover Torres, J., Block, A., Hahlbrock, K. and Somssich, I. E. (1993) Influence of bacterial strain genotype on transient expression of plasmid DNA in plant protoplasts. Plant J. 4, 587–592.

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Chapter 21 RNA-Protein Interaction Mediating Post-Transcriptional Regulation in the Circadian System Jan C. Schöning and Dorothee Staiger Abstract Post-transcriptional control makes an important contribution to shaping transcript profiles of circadianly regulated genes. In Arabidopsis thaliana, the clock-regulated glycine-rich RNA-binding protein ATGRP7 oscillates with a 24-h rhythm and transmits the rhythmicity generated by the central oscillator within the cell. ATGRP7 negatively auto-regulates its own expression at the post-transcriptional level. In response to an elevated protein level, a shift to a cryptic 5′ splice site within the intron occurs, leading to an unproductively spliced transcript that rapidly vanishes due to its short half-life. This feedback regulation relies on direct binding of the RNA-binding protein to its own RNA. Here we describe the analysis of RNA-protein interaction in vitro employing recombinant RNA-binding protein and 32P-labelled in vitro transcripts or synthetic RNA oligoribonucleotides comprising the binding site under study. Key words: RNA-protein interaction, EMSA, synthetic oligoribonucleotide, in vitro transcription.

1. Introduction Higher plants, like most organisms, employ an endogenous timer, the so-called circadian clock, to coordinate physiological, biochemical and developmental processes with the environmental cycle of day and night and associated temperature changes (1,2). It is well established that the circadian clock operates at the level of single cells (3, 4). Clock proteins build an auto-regulatory feedback loop and generate their own 24-h rhythm through interfering with the transcription of their own genes. The Arabidopsis core clockwork comprises the Myb-type transcription factors CCA1 (circadian clock associated) and LHY (late elongated T. Pfannschmidt (ed.), Plant Signal Transduction, vol. 479 Humana Press, a part of Springer Science + Business Media, LLC 2009 DOI: 10.1007/978-1-59745-289-2_21

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hypocotyl), and TOC1 (timing of CAB expression), which cycle in antiphase and reciprocally regulate each other (5–8). Additional feedback circuits interlock into this basic loop (9–11). Apart from regulating their own expression, clock proteins signal rhythmicity to downstream targets to produce overt rhythms. They may accomplish this by direct control of downstream genes or via cascades of oscillating gene products (12, 13). In particular, post-transcriptional control mechanisms have been implicated in shaping rhythmic transcript profiles (14–16). So far, only a limited number of RNA-binding proteins have been shown to be under clock control. Drosophila LARK functions in clock output leading to rhythms in adult eclosion (17). The ortholog in mouse, LARK, has been implicated in the regulation of the clock protein mPERIOD1 (18). LARK binds to the 3′ UTR of the mPER1 mRNA in vitro and causes elevated mPER1 protein levels presumably by translational regulation. In Chlamydomonas reinhardtii, CHLAMY1 is a heterodimer that binds to UG repeats in the 3′ UTRs of several mRNAs and mediates translational control (19, 20). In Arabidopsis, ATGRP7 (Arabidopsis thaliana glycine-rich RNA-binding protein) represents an RNA-binding protein that operates within an output pathway from the CCA1-LHY-TOC1 clockwork (21). ATGRP7 shows a circadian expression pattern with the highest mRNA and protein abundance at the end of the day (22). Ectopic overexpression of ATGRP7 in transgenic plants leads to negative auto-regulation of its own mRNA. Binding of the protein to intronic sequences and the 3′ untranslated region of the ATGRP7 pre-mRNA is required for this negative feedback (23, 24). Moreover, ATGRP7 also binds to the ATGRP8 transcript encoding a related RNA-binding protein to regulate alternative splicing and steady-state abundance of this ATGRP7 target transcript (24). Here we describe techniques to analyse the interaction of bacterially expressed recombinant RNA-binding proteins with candidate target sites, using radiolabelled in vitro transcripts or synthetic oligoribonucleotides as binding substrates. Further, we describe the determination of binding specificity via competition assays and the determination of equilibrium dissociation constants. Additional methods to study various aspects of RNA-protein interaction have been assembled in an excellent and detailed manual (25).

2. Materials 2.1. Bacterial Growth

1. Luria-Bertani (LB) medium: 10 g tryptone/peptone from casein, 5 g yeast extract, 10 g NaCl, adjust to pH 7.4 with NaOH, autoclave. 2. Ampicillin 100 mg/ml. Dissolve in sterile water and store at −20°C in small aliquots.

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3. Sterile 1-l shaking flasks with breathable plugs. 4. IPTG 0.1 M. Dissolve in sterile water and store at −20°C in small aliquots. 5. Escherichia coli cells harbouring the pGEX-RNA-binding protein fusion (see Note 1). 2.2. Affinity Chromatography for Purification of Recombinant Protein

1. 10 X PBS: 1.37 M NaCl, 27 mM KCl, 100 mM Na2HPO4, 18 mM KH2PO4, adjust to pH 7.4 with HCl. 2. 1 X PBS-T: Dilute 100 ml of 10 X PBS with 1 ml Tween-20 in 899 ml H2O. 3. Protease inhibitor tablets complete protease inhibitor (Roche, Mannheim, Germany). Use as indicated by the supplier. 4. Glutathione Sepharose 4B (GE Healthcare, Buckinghamshire, UK). 5. Poly-prep empty column (Bio-Rad, Munich, Germany). 6. Elution buffer: 20 mM glutathione (reduced form) in 1 X PBS. 7. Centricon microconcentrator centrifugal filter devices (Millipore, Billerica, MA, USA). 8. Bio-Rad protein assay (Bio-Rad, Munich, Germany). 9. HEPES buffer: 25 mM HEPES-KOH pH 7.5, 0.1 M NaCI, 1 mM MgCl2, 1 mM DTT.

2.3. SDSPolyacrylamide Gel Electrophoresis (SDS-PAGE)

1. Forty percent acrylamide/bis solution (39:1) (this is a neurotoxin when unpolymerized and should be handled with care). Store at 4°C. 2. Ammonium persulfate (APS): prepare 10% solution in water and store at 4°C for a few weeks. 3. 10 X Laemmli running buffer: 250 mM Tris, 1.92 M glycine, 1% SDS. Store at room temperature. 4. 2 X SDS sample buffer: 250 mM Tris, 40% glycerol, 20% β-mercaptoethanol, 8% SDS, 0.004% bromophenol blue (BPB). Store in small aliquots at −20°C. 5. Coomassie staining solution: Dissolve 0.5 g Coomassie Brilliant Blue in 500 ml methanol. Add 100 ml glacial acetic acid and make up to 1 l with H2O. 6. Destaining solution: 50% methanol, 10% glacial acetic acid.

2.4. In Vitro Transcription to Prepare Radiolabelled RNA-Binding Substrates

1. In vitro transcription kit: MegaScript T7 Kit (Ambion Inc., Austin, TX, USA). 2. [α-32P] UTP: 30 TBq/mmol.

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2.5. Denaturing Polyacrylamide–Urea Gel Electrophoresis

1. 10 X TBE: 890 mM Tris, 890 mM boric acid, 20 mM Na2EDTA, adjust to pH 8.0 with NaOH. 2. Stop solution: 95% formamide, 10 mM Na2EDTA pH 8.0, 0.02% bromophenol blue. Store at 4°C. 3. Elution buffer: 2 M ammonium acetate, 0.1% SDS, 30 µg/ml tRNA from baker’s yeast (add directly before use).

2.6. Electrophoretic Mobility Shift Assay

1. 10 X binding buffer: 200 mM HEPES-KOH pH 7.5, 1 M NaCl, 10 mM MgCl2, 0.1% NP-40). Store in small aliquots at −20°C. 2. 10 X TAE buffer: 400 mM Tris, 18 mM Na2EDTA, adjust to pH 7.8 with acetic acid. 3. Ribolock RNase-inhibitor (Fermentas, St. Leon-Rot, Germany). 4. 6 X native loading dye: 6 X TAE, 40% glycerol. Store at room temperature.

2.7. Preparation of 5 ′ Radiolabelled Oligoribonucleotide

1. Synthetic oligoribonucleotide covering the binding site (e.g., from Biomers, Ulm, Germany) (see Note 2). 2. 10 X denaturing buffer: 0.25 M Tris pH 9.5, 6 mM Na2EDTA pH 8.0, 25 mM spermidine. 3. [γ-32P] ATP: 185 TBq/mmol. 4. T4 polynucleotide kinase (PNK) (Fermentas, St. Leon-Rot, Germany). 5. Mini Quick Spin RNA columns (Roche, Mannheim, Germany).

3. Methods 3.1. Induction of Recombinant Protein Expression in Escherichia coli

1. An overnight starter culture is prepared in 10 ml LB containing 100 µg/ml ampicillin by inoculating a single colony of E. coli expressing the glutathione S-transferase (GST) RNAbinding protein fusion. 2. Inoculate 500 ml LB (100 µg/ml ampicillin) in a 1-l shaking flask with 5 ml overnight culture. 3. Grow on a rotary shaker with 160 rpm at 37°C until an OD600 of 0.5 is reached. Remove 1 mL culture (uninduced control). Centrifuge 3 min at 16,000× g, remove the supernatant and resuspend the pellet in 20 µl SDS sample buffer. Store at −20°C for analysis by SDS PAGE (cf. Sect. 3.3) (see Note 3). 4. Induce recombinant protein by adding 500 µl 0.1 M IPTG (0.1 mM final concentration) and shake culture for 24 h at 18°C.

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5. Remove 1 ml culture (induced sample) and treat it like the uninduced control (cf. Sect. 3.1.3). 3.2. Purification of GST-Tagged Recombinant Protein

1. The induced E. coli culture is harvested by centrifugation for 15 min at 5,000× g, and the cells are resuspended in 20 ml PBS buffer supplemented with complete protease inhibitor (Roche, Mannheim, Germany). 2. Cells are disrupted by passing twice through a French pressure cell at 1,000 psig. 3. The lysate is clarified by centrifugation at 4°C and 16,000× g for ten minutes. All subsequent steps are performed at 4°C. 4. Equilibrate 1 ml of glutathione Sepharose 4B in PBS: Sediment the beads by centrifugation (2 min, 500× g), remove the supernatant, and resuspend the beads in 1 ml PBS. Repeat four times. 5. About 1 ml glutathione Sepharose 4B (50% slurry in PBS) is added to the clarified E. coli lysate. Batch absorption is performed in 50-ml screw capped tubes for 1 h by end-over-end rotation. 6. Sediment the beads by centrifugation (5 min, 500× g) and remove the supernatant. Take a 15-µl aliquot of the supernatant and store to analyse the degree of absorption by SDS PAGE (cf. Sect. 3.3). 7. Resuspend the beads in PBS-T and fill into a Poly-prep column (Bio-Rad). Wash with 4 × 5 ml PBS-T. Remove a 15-µl aliquot from each wash fraction. 8. Elute the protein by adding 1 mL of elution buffer. Repeat the elution steps four times. Take a 5-µl sample from each elution fraction for final analysis (cf. Sect. 3.3). 9. Store samples at −70°C (see Note 4).

3.3. Analysis of Protein Induction and Purification on SDS-PAGE

1. These instructions assume the use of a Mini Protean Gel System (Bio-Rad). 2. Prepare a 12% polyacrylamide SDS gel. Mix 2.4 ml 40% acrylamide/bis (39:1) solution, 2.4 ml 1.5 M Tris-HCl pH 8.8, 80 µl 10% SDS, 3.3 ml H2O, 80 µl 10% APS and 10 µl TEMED and pour the separation gel. Leave some space for the stacking gel and cover the separation gel with 500 µl of 2-propanol. The gel should polymerize within 40 min. Pour off the 2-propanol and rinse the surface of the separation gel with water. 3. Prepare the stacking gel by mixing 560 µl of 40% acrylamide/bis (39:1) solution, 560 µl 1 M Tris-HCl pH 6.8, 45 µl 10% SDS, 3.26 ml H2O, 60 µl 10% APS and 6 µl TEMED.

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Pour the gel and insert the comb. The gel should polymerize in about 20 min. 4. Prepare the 1 X Laemmli running buffer by diluting 100 ml 10 X Laemmli running buffer with 900 ml of H2O in a measuring cylinder. Cover with parafilm and invert to mix. 5. Assemble the gel system and fill the buffer chambers with the 1 X running buffer. Remove the comb carefully and rinse the pockets with running buffer using a 5-ml syringe fitted with a 21-gauge needle. 6. Prepare the samples taken during the purification process by adding an equal amount of 2 X SDS sample buffer and heat for 5 min at 95°C. In parallel, denature the uninduced and induced control samples (cf. Sects. 3.1.3 and 3.1.5). Load the samples and a molecular weight standard onto the gel and run the gel at 80 V (const.) until the BPB front has reached the end of the gel. 7. Disassemble the glass plates and stain the gel in Coomassie staining solution for 2 h at room temperature on a rotary shaker. Remove the staining solution and incubate the gel in destaining solution until the protein bands are visible against a clear background (e.g., overnight). 8. Identify the fractions containing the eluted fusion protein. 9. Remove glutathione in the eluate fractions containing the fusion protein by centrifugation through centricon YM-30. Centrifuge at 4°C and 5,000× g until the sample volume has been reduced to one-fifth of the initial volume. Fill the retentate cup with HEPES buffer and centrifuge again. Repeat this procedure a second time. Elute the protein from the filter by placing the retentate cup over the sample reservoir, inverting the unit and centrifuging at 1,000× g for two minutes to transfer concentrate into retentate cup. Store fusion protein in small aliquots at −70°C (see Note 4). 10. Determine the protein concentrations of the eluate fractions by Bradford protein assay (Bio-Rad). Prepare a dye working solution by diluting the Bio-Rad dye concentrate 1:5 with H2O. Mix 20 µl of protein eluate with 1 ml of the diluted Bio-Rad dye working solution and incubate for 5 min at room temperature. Do the same for the calibration standard, but add 20-µl samples with increasing amounts (0.2–1.5 mg/ml) of bovine serum albumin (BSA) instead of the eluate. Determine the absorption at 595 nm in a spectrophotometer. Plot the A595 of the standard samples against the BSA concentration and perform a linear regression. Calculate the protein concentration of the eluate fractions on the basis of the standard curve.

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3.4. In Vitro Transcription to Prepare Radiolabelled Binding Substrates Containing the Binding Site

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1. Plasmids harbouring the binding site under study downstream of a T7 promoter (e.g., pBSK- (Stratagene)) serve as templates (Fig. 21.1). Plasmid DNA is isolated according to standard procedures (26, 27) or with commercial plasmid isolation kits (DNeasy, Qiagen; Invisorb Spin Plasmid Mini Two, Invitek). 2. To prevent read-through of the RNA polymerase, the recombinant plasmid is linearized by restriction enzyme digestion using a suitable restriction site downstream of the binding site (see Notes 5 and 6). 3. In vitro transcription is performed using the MegaScript T7 Kit. About 1 µg of linearized template DNA is transcribed in 2 µl 10 X T7 buffer, 1 µl 10 mM ATP, 10 mM GTP, 10 mM CTP each (0.5 mM final concentration), 1 µl 25 µM UTP and H2O ad 16 µl. Add 2 µl 0.74 MBq [ -32P] UTP (12.5 µM final concentration) and 2 µl T7 enzyme mix. 4. Incubate for 1 h at 37°C.

3.5. Purification of 32 P-Labelled In Vitro Transcripts on Denaturing Polyacrylamide–Urea Gels

1. Prepare 10% polyacrylamide–urea gel. Dissolve 3.6 g urea in 1.05 ml 40% acrylamide/bis (39:1) solution, 0.75 ml 10 X TBE and 4 mL H2O by gentle warming. Make up to 7.5 ml with H2O.

Fig. 21.1. Example of a vector for in vitro transcription. The pBSK-Vector (Stratagene) harbours a T7 promoter site upstream of a multiple cloning site (MCS). The ATGRP7 3′ UTR was cloned into the MCS using BamHl and XbaI (23).

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2. Add 60 µl APS solution and 10 µl TEMED. Pour the gel and insert the comb. The gel should polymerize in about 30 min. 3. Prepare the 1 X TBE running buffer from the 10 X TBE stock solution. 4. Carefully remove the comb. Use a 5-ml syringe fitted with a 21-gauge needle to flush the wells with 1 X TBE running buffer. 5. Pre-run the gel at 300 V for 1 h. 6. Denature the samples for 10 min at 94°C. Before loading, again thoroughly flush the wells with 1 X TBE to completely remove the urea. 7. Load the samples and run the gel until the BPB front has run about three-fourth of the gel. Check regularly that the gel maintains a temperature of about 50°C to ensure that the in vitro transcripts remain denatured (see Note 7). 8. Remove the upper glass plate, cover the wet gel with SARAN wrap and expose to an x-ray film for 2 min. 9. Align the autoradiogram with the gel and excise the 32P-labelled band using a scalpel. 10. Transfer the gel piece to a reaction tube containing 400 µl elution buffer. The radioactively labelled in vitro transcript is recovered from the gel by incubation at 25°C overnight under moderate shaking. Subsequently precipitate the RNA from the supernatant with 800 µl ethanol for 30 min at −20°C. Pellet the RNA by centrifugation at 16,000× g for 15 min. Remove the supernatant and dissolve the pellet in 30 µl nuclease-free water. 11. Check the yield of the radioactively labelled fragment by measuring Ćerenkov counts (see Note 8). Adjust to 2,500 cpm/µl using nuclease-free water. 3.6. Electrophoretic Mobility Shift Assay Using In Vitro Transcripts as Binding Substrates

1. Prepare a 6% native polyacrylamide gel by mixing 1.05 ml 40% acrylamide/bis (39:1) solution, 0.75 ml 10 X TAE, 5.2 ml H2O, 60 µl APS solution and 10 µl TEMED. Pour the gel and insert the comb. The gel should polymerize in about 30 min. Prepare the 1 X TAE running buffer from the 10 X TAE stock solution. Carefully remove the comb. Use a 5-ml syringe fitted with a 21-gauge needle to flush the wells with 1 X TAE. Assemble the gel system and fill the buffer chambers with 1 X TAE. Pre-run the gel for 15 min at 20 mA. 2. Denature the labelled in vitro transcripts briefly by heating to 80°C for 3 min. Allow RNA to refold by cooling down slowly to room temperature. 3. Prepare the binding assays containing 0.5 µg recombinant GST fusion protein, 1 µl 10 X binding buffer and 0.25 µl Ribolock

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and bring to a volume of 7 µl with H2O. Add 1 µl of tRNA from baker’s yeast in concentrations varying between 0.01 and 10 mg/ml. Start the binding reaction by addition of 2 µl 32 P-labelled in vitro transcripts = 5,000 cpm. 4. Prepare a control sample omitting the recombinant GST fusion protein to visualize the unbound RNA. 5. Incubate at 20°C for 30 min. 6. Add 2 µl 6 X loading dye, load the samples onto the gel and run the gel at 17 mA (const.) until the BPB front has reached the end of the gel. 7. Dry the gel and expose to PhosphorImager Screen or x-ray film. 8. Binding of the recombinant GST fusion protein is indicated by the appearance of additional bands of lower electrophoretic mobility than the unbound RNA. Non-specific interactions tend to disappear at higher concentrations of tRNA (see Note 9). 3.7. Preparation of 5 ′ 32 P-Labelled Oligoribonucleotides

1. The commercially obtained synthetic oligoribonucleotide (ORN) is dissolved in RNase-free water by heating to 65°C for 10 min. OD260 is measured and the concentration is adjusted to 250 pmol/µl. 2. The ORN is denatured immediately before the labelling reaction. To 1 µl (= 250 pmol) ORN, 0.5 µl denaturation buffer and 3.5 µl H2O are added. The sample is incubated at 70°C and then placed on ice. 3. The ORN is end-labelled using T4 polynucleotide kinase and [γ-32P] ATP. Add to a micro-centrifuge tube 13 µl H2O, 2 µl 10 X T4 PNK buffer, 1 µl denatured ORN (50 pmol), 1 µl Ribolock, 2 µl [γ-32P] ATP and 1 µl T4 PNK (10 U/µl). Incubate at 37°C for 60 min. Stop the reaction by incubation at 70°C for 10 min and add 20 µl of H2O. 4. Unincorporated nucleotide triphosphate is removed by spin column chromatography: Mini Quick Spin RNA columns are inverted to resuspend the matrix. The upper and lower caps are removed. Spin the column at 1,000× g for 1 min to remove the buffer. Carefully apply the sample to the centre of the column bed. Centrifuge for 4 min at 1,000× g. Recover the eluate containing the ORN. 5. The labelling is checked by measuring Cerenkov counts.

3.8. Electrophoretic Mobility Shift Assay Using Synthetic Oligoribonucleotides as Binding Substrates

1. Prepare 6% native polyacrylamide gel (cf. Sect. 3.6.1). 2. Prepare binding assays containing 0.5 µg recombinant GST fusion protein in 1 µl 10 X binding buffer, 0.25 µl Ribolock, 1 µl tRNA (1 mg/ml); bring to a volume of 8 µl with H2O and start the binding reaction by adding 2 µl 32P-labelled ORN = 50 fmol.

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3. Prepare a control sample omitting the recombinant GST fusion protein to visualize the unbound ORN. 4. Incubate at 20°C for 30 min. 5. Add 2 µl 6 X loading dye and run the native gel as described (cf. Sect. 3.6.5) until the BPB front has migrated about three-fourth of the gel. 6. Dry the gel and expose to PhosphorImager Screen or x-ray film. 3.9. Determination of Equilibrium Dissociation Constants (Kd Values)

1. Prepare 6% native polyacrylamide gel (cf. Sect. 3.6.1). 2. Prepare binding assays containing 0, 0.1, 0.5, 1, 2.5, 5, 10, 25, 50 and 100 pmol recombinant protein in 1 µl 10 X binding buffer, 0.25 µl Ribolock, 1 µl tRNA (1 mg/ml), bring to a volume of 8 µl with H2O and start the binding reaction by adding 2 µl 32P-labelled ORN = 50 fmol. 3. Incubate at 20°C for 30 min. 4. Add 2 µl 6 X loading dye and run native gel as described (cf. Sect. 3.8.4). 5. Dry gel and expose to PhosphorImager Screen. 6. Determine the signal intensity of the retarded complex and free ORN for each sample (e.g., with the ImageQuant software, Amersham). 7. The Kd value is calculated by plotting the log (complexed/free probe) of the mean curve against the log (protein concentration). The log (Kd) is obtained as x-intercept (Fig. 21.2).

3.10. Competition Assay to Determine Binding Specificity

1. To check for the specificity of the retarded complex, perform competition experiments. Increasing amounts of homologous or heterologous unlabelled cold competitor RNA are added to the binding reactions (cf. Sect. 4.8). 2. Prepare nine binding assays each containing 50 pmol recombinant protein in 1 µl 10 X binding buffer, 0.25 µl Ribolock, 1 µl tRNA (1 mg/ml) and bring to a volume of 7 µl with H2O. 3. Start the binding reaction by adding 2 µl = 50 fmol of the 32 P-labelled ORN. At the same time add 1 µl of unlabelled homologous ORN to four reactions in increasing concentrations (e.g., 1, 10, 100 and 1000 pmol). Add 1 µl of unlabelled heterologous ORN of the same concentrations to another four samples (see Note 9). Prepare two control samples: one without competitor ORN and another without protein and competitor ORN. 4. Incubate at 20°C for 30 min.

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Fig. 21.2. Use of EMSA to determine the binding affinity of recombinant ATGRP7 for an oligoribonucleotide derived from the 3′ UTR of ATGRP7. About 50 fmoles of labelled ORN is incubated with increasing concentrations of recombinant protein, as indicated, in the presence of 1 µg tRNA (A) The PhosphorImager scan obtained after gel electrophoresis is shown. The amount of radioactivity in the retarded complex and the free ORN are quantified for each lane using ImageQuant (GE Healthcare). (B) The percentage of bound ORN is plotted against the concentration of recombinant protein. The Kd corresponds to the protein concentration that shifts 50% of the input ORN. (C) Plot of the log (complexed/free probe) of the curve against the log (protein concentration). The x-intercept yields log (Kd).

5. Add 2 µl 6 X loading dye and run native gel as described (cf. Sect. 3.8.4). 6. Dry gel and expose to PhosphorImager screen. 7. Determine the signal intensity of the retarded complex and free ORN for each sample (e.g., with the ImageQuant software, Amersham). If the binding is specific, a loss of complex formation should be observed with increasing concentration of homologous competitor but no major changes should be visible in the case of the heterologous competitor (Fig. 21.3).

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Fig. 21.3. Competition experiment to determine the specificity of complex formation between recombinant ATGRP7 and an oligoribonucleotide derived from the ATGRP8 3 ′ UTR. Recombinant ATGRP7 protein is incubated with 50 fmol labelled ORN in the presence of 1 µg tRNA and 2, 20, 100 and 250 pmoles of unlabelled ORN or 2, 20, 100 and 250 pmoles of the same oligoribonucleotide carrying mutations. First lane: Free labelled ORN without protein; second lane: labelled ORN with ATGRP7 protein and tRNA but without competitor.

4. Notes 1. Conventional peptide tags including His-tags are used to purify recombinant RNA-binding proteins from E. coli. Here the use of the glutathione-S-transferase system based on the vector pGEX-6P (GE Healthcare) is described. This system allows for the purification of the entire fusion protein by elution with glutathione or for recovery of the RNA-binding moiety by cleavage with PreScission protease (see the PreScission protease protocol from GE Healthcare). 2. The length of the oligoribonucleotide depends on the binding sequence under study. For ATGRP7, 27mers work well, but shorter oligoribonucleotides may be sufficient. The RNA sequence actually bound by the RNA-binding protein is likely much shorter. Flanking bases may contribute to complex stability. 3. The optimal yield of intact fusion protein depends on the duration of induction of the IPTG concentration and the temperature and should be determined experimentally for each new fusion protein. 4. Conditions under which the fusion protein is most stable during storage (4°C, −20°C, −70°C) vary between proteins and should be determined experimentally for each protein under study. 5. To linearize the template DNA downstream of the binding site for in vitro transcription, restriction enzymes generating

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5′ protruding ends should be favoured over enzymes generating blunt ends. Avoid 3′ protruding ends. 6. As an alternative to subclone binding sites into T7 promoter vectors, templates generated by PCR can be used. In this case, the T7 promoter is incorporated during the PCR reaction using an upstream primer with a T7 promoter at its 5′ end. Alternative promoters such as T3 and SP6 for which RNA polymerases are available can also be used. 7. This gel serves to remove unincorporated nucleotide triphosphates and prematurely terminated in vitro transcript from the full-length transcript. 8. For the quantification of 32P labelled molecules, the measurement of Cerenkov radiation can be used. This method is based on the emission of electro-magnetic radiation from charged particles with high energies (85% of 32P β-particles have an E > 0.265 MeV) in colourless solutions. The emission occurs mainly in the blue light and UV-spectrum and can be detected in standard scintillation analysers. 9. If the competition experiment is performed using synthetic ORNs as binding substrates and competitor, the heterologous ORN should be of the same size but of a different sequence. To determine the bases important for binding, synthetic ORNs of the same sequence with defined point mutations are employed. If the competition experiment is performed using in vitro transcripts as binding substrates and competitor, the heterologous in vitro transcript has to be transcribed from a plasmid containing a fragment of similar size downstream of a T7 promoter. To obtain unlabelled in vitro transcripts, [α-32P] UTP is omitted from the reaction (cf. Sect. 3.4) and the concentration of unlabelled UTP is raised to 0.5 mM. The fragments are purified by phenol extraction and ethanol precipitation.

Acknowledgements This work was supported by the DFG (STA 653/2 and SFB 613). J.C.S. is a fellow of the German National Academic Foundation. References 1. Schöning, J. C., Streitner, C., and Staiger, D. (2006) Clockwork Green - the circadian oscillator in Arabidopsis. Biological Rhythm Res. 37, 335–352.

2. McClung, C. R. (2006) Plant circadian rhythms. Plant Cell 18, 792–803. 3. Nagoshi, E., Saini, C., Bauer, C., Laroche, T., Naef, F., and Schibler, U. (2004) Circadian

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Schöning and Staiger gene expression in individual fibroblasts; cellautonomous and self-sustained oscillators pass time to daughter cells. Cell 119, 693–705. Dunlap, J. C. (1999) Molecular bases for circadian clocks. Cell 96, 271–290. Wang, Z. Y., and Tobin, E. M. (1998) Constitutive expression of the CIRCADIAN CLOCK ASSOCIATED1 (CCA1) gene disrupts circadian rhythms and suppresses its own expression. Cell 93, 1207–1217. Strayer, C., Oyama, T., Schultz, T. F., Raman, R., Somers, D. E., Mas, P., Panda, S., Kreps, J. A., et-al. (2000) Cloning of the Arabidopsis clock gene TOC1, an autoregulatory response regulator homolog. Science 289, 768–771. Alabadi, D., Oyama, T., Yanovsky, M. J., Harmon, F. G., Mas, P., and Kay, S. A. (2001) Reciprocal regulation between TOC1 and LHY/CCA1 within the Arabidopsis circadian clock. Science 293, 880–883. Schaffer, R., Ramsay, N., Samach, A., Putterill, J., Carre, I. A., and Coupland, G. (1998) The late elongated hypocotyl mutation of Arabidopsis disrupts circadian rhythms and the photoperiodic control of flowering. Cell 93, 1219–1229. Mizuno, T., and Nakamichi, N. (2005) Pseudo Response Regulators (PRR) or True Oscillator Components (TOC). Plant Cell Physiol. 46, 677–685. Locke, J. C., Southern, M. M., KozmaBognar, L., Hibberd, V., Brown, P. E., Turner, M. S., and Millar, A. J. (2005) Extension of a genetic network model by iterative experimentation and mathematical analysis. Mol. Syst. Biol. 1, http://dx.doi. org/10.1038/msb4100018. Salome, P. A., and McClung, C. R. (2005) PSEUDO-RESPONSE REGULATOR 7 and 9 are partially redundant genes essential for the temperature responsiveness of the Arabidopsis circadian clock. Plant Cell 17, 791–803. Brown, S. A., and Schibler, U. (1999) The ins and outs of circadian timekeeping. Curr. Opin. Genet. Dev. 9, 588–594. Staiger, D., Streitner, C., Rudolf, F., and Huang, X. (2006) Multiple and slave oscillators in Endogenous Plant Rhythms, Hall, A. and McWatters, H., eds., Blackwell Publishers, pp. 57–83. So, W. V., and Rosbash, M. (1997) Posttranscriptional regulation contributes to Drosophila clock gene mRNA cycling. EMBO J. 16, 7146–7155.

15. Edery, I. (1999) Role of posttranscriptional regulation in circadian clocks: lessons from Drosophila. Chronobiol. Int. 16, 377–414. 16. Mittag, M. (2003) The function of circadian RNA-binding proteins and their cisacting elements in microalgae. Chronobiol. Int. 20, 529–541. 17. McNeil, G. P., Zhang, X., Genova, G., and Jackson, F. R. (1998) A molecular rhythm mediating circadian clock output in Drosophila. Neuron 20, 297–303. 18. Kojima, S., Matsumoto, K., Hirose, M., Shimada, M., Nagano, M., Shigeyoshi, Y., Hoshino, S., Ui-Tei, K., et-al. (2007) LARK activates posttranscriptional expression of an essential mammalian clock protein, PERIOD1. Proc. Natl. Acad. Sci. USA 104, 1859–1864. 19. Zhao, B., Schneid, C., Iliev, D., Schmidt, E. M., Wagner, V., Wollnik, F., and Mittag, M. (2004) The circadian RNA-binding protein CHLAMY 1 represents a novel type heteromer of RNA recognition motif and lysine homology domain-containing subunits. Eukaryot. Cell 3, 815–825. 20. Iliev, D., Voytsekh, O., Schmidt, E. M., Fiedler, M., Nykytenko, A., and Mittag, M. (2006) A heteromeric RNA-binding protein is involved in maintaining acrophase and period of the circadian clock. Plant Physiol. 142, 797–806. 21. Rudolf, F., Wehrle, F., and Staiger, D. (2004) Slave to the rhythm. The Biochemist 26, 11–13. 22. Heintzen, C., Nater, M., Apel, K., and Staiger, D. (1997) AtGRP7, a nuclear RNA-binding protein as a component of a circadian-regulated negative feedback loop in Arabidopsis thaliana. Proc. Natl. Acad. Sci. USA 94, 8515–8520. 23. Staiger, D., Zecca, L., Wieczorek Kirk, D. A., Apel, K., and Eckstein, L. (2003) The circadian clock regulated RNA-binding protein AtGRP7 autoregulates its expression by influencing alternative splicing of its own pre-mRNA. Plant J. 33, 361–371. 24. Schöning, J. C., Streitner, C., Page, D. R., Hennig, S., Uchida, K., Wolf, E., Furuya, M., and Staiger, D. (2007) Autoregulation of the circadian slave oscillator component ATGRP7 and regulation of its targets is impaired by a single RNA recognition motif point mutation. Plant J. 52, 1119–1130. 25. Hartmann, R. K., Bindereif, A., Schön, A., and Westhof, E. (2005) Handbook of RNA Biochemistry, WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim.

RNA-Protein Interaction 26. Sambrook, J. and Russel, D. W. (2001) Molecular Cloning: A Laboratory Manual, Cold Spring Harbor Laboratory, New York.

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27. Ausubel, F. M., Brent, R., Kingston, R. E., Moore, D., Seidman, J. G., Smith, J. A., and Struhl, K. (2005) Current Protocols in Molecular Biology, John Wiley & Sons, Inc.

INDEX A abiotic additives ............................................................... 43 abscisic acid ..................................................................... 42 acousto-optical tunable filters (AOTF) ......................... 102 ade2 mutation ................................................................ 304 ADE2 and HIS3 receptor genes ................................... 230 aequorin ........................................................................... 80 side effects on activity ................................................ 87 affinity chromatography ................................................ 339 Agrobacterium tumefaciens....................................... 193, 196 amplex-red..................................................................... 111 antiphase........................................................................ 338 Arabidopsis suspension culture ............................... 251, 253 ARGONOUTE.............................................................. 10 ATGRP7 ....................................................................... 338 AUX/IAA repressor ...................................................... 160 auxin...... ...................................................................... 5, 44 auxotrophy marker ......................................................... 301

B back-crossing ................................................................... 19 bait................................................................................. 219 fusion ....................................................................... 225 binding affinity .............................................................. 347 bioinfomatics analysis .................................................... 312 bioinformatic tools for CRE element analysis ............... 320 biomolecular fluorescence complementation (BiFC)..... 189 brassinosteroids ............................................................... 44 branching pattern ............................................................ 29 bridge assay............................................................ 219, 225 bZIP factors .................................................................. 191

C calcium .............................................................................. 9 dyes ............................................................................ 84 intracellular variation of ............................................. 79 signatures ................................................................... 79 calcium measurement ...................................................... 81 with tobacco cells ....................................................... 86 with seedlings ............................................................ 86 calibration of fluorescence ............................................... 98 casein... .....................................................................256 casein dephosphorylation .............................................. 256 CCA1... ......................................................................... 337

cDNA synthesis .................................................................... 69 library of Arabidopsis ................................................ 236 cell division ...................................................................... 45 epidermal ................................................................... 27 guard .......................................................................... 27 cell culture ............................................................. 179, 205 of Chlamydomonas reinhardtii ................................... 179 circadian rhythm............................................................ 173 chemical stressors ............................................................ 37 chemical transformation ................................................ 197 chlorophyll determination ............................................................ 28 extraction ................................................................. 112 concentration ........................................................... 138 chromatin immunoprecipitation (ChIP) ........................... 261, 263 extraction ................................................................. 263 preparation of .......................................................... 266 chromatography ....................................................... 276 circadian system ............................................................. 337 cis-regulatory DNA sequences ....................................... 311 cis- regulatory element (CRE) ....................................... 292 data basis ................................................................. 322 CLAVATA3 ...................................................................... 3 cloning into BiFC vectors ............................................. 194 CNBr.... ................................................................. 125, 287 colonies .......................................................................... 300 colorimetric ONPG assay.............................................. 302 column packing ............................................................. 282 competition assay........................................................... 346 complex formation......................................................... 348 computer aided sequence analysis .................................. 316 confocal laser scanning microscopy (CLSM) ............ 80, 82 microscope settings .............................................. 82, 83 confocal settings ............................................................ 206 CONSTANS (CO) ........................................................... 7 CONSTITUTIVE PHOTOMORPHOGENESIS 1 (COP1) .................................................................. 6 Coomassie R 250, 284 COP9-signalosome ....................................................... 156 cotransfection ................................................................ 315 cotyledons ........................................................................ 23 cross-linking of proteins ........................................ 262, 264

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cross talk ........................................................................ 209 C-terminal half of ubiquitin (Cub)................................ 218 cullin-RING-ligases ...................................................... 149 curve fitting ................................................................... 208 cytokinins ........................................................................ 45

excitation spectrum ........................................................ 100 experimental design ......................................................... 69 expression library ........................................................... 298 ex vivo kinase assay ................................................ 251, 254 ex vivo phosphatase assay ...................................... 252, 256

D

F

data evaluation................................................................... 73 processing .................................................................. 73 analysis of FCCS ..................................................... 210 DAPI.... ......................................................................... 271 de-etiolation .................................................................... 54 degradation substrates ........................................... 149, 156 DELLA protein ............................................................ 161 denaturing polyacrylamid urea gel ......................... 340, 343 2D-EMSA .....................................................277, 283, 284 principle of............................................................... 284 2-DE-PAGE ......................................................... 177, 181 detection assays.............................................................. 223 di-amino benzidine........................................................ 111 dicer endonuclease ........................................................... 10 diffusion-independent fluctuation ................................. 212 DNA.................................................................................. 9 affinity chromatography .................................. 277, 282 affinity column media .............................................. 281 binding motif ........................................................... 292 calculation tool .......................................................... 66 ligand ....................................................................... 281 quantification of ...................................................... 267 protein complexes .................................................... 274 purification of .......................................................... 269 DNase treatment ............................................................. 69 differential isotope labelling .......................................... 144 driver plasmid ................................................................ 295

F-box protein substrate interactions .............................. 160 FCCS application to plant cells ..................................... 209 FCCS in nuclei.............................................................. 211 FCCS measurement ...................................................... 207 fEMSA .......................................................................... 278 filter lift assay................................................................. 302 FLOWERING LOCUS C (FLC) .................................................................... 11 D (FD) ........................................................................ 4 T (FT) ......................................................................... 3 flowering time ................................................................. 29 fluorescence cross-correlation spectroscopy (FCCS) ..... 203 fluorescence DNA probe ....................................... 275, 279 fluorescence image ....................................................................... 208 intensity ................................................................... 208 EMSA ..................................................................... 276 fluorophor...................................................................... 205 FRET..............................................................190, 204, 218 feedback circuits ............................................................ 338 functional classification 317 functional complementation .......................................... 235 hog1.......................................................................... 242 function-not-known ........................................................ 17 fusion tags...................................................................... 158

E E. coli RNA plymerase ................................................... 284 electrophoretic mobility shift assay (EMSA)................. 273 electroporation............................................................... 196 EMSA.. ..........................................................274, 340, 344 principle of............................................................... 274 setup ........................................................................ 280 environment ...................................................................... 2 enzymes metabolic ..................................................................... 6 epitope. .......................................................................... 159 equilibrium dissociation constant .................................. 346 ESI-MS................................................................. 140, 142 esterification of isolated peptides ................................... 141 ethylene ........................................................................... 46 ethylene receptor ETR1 ................................................ 231 E3-ubiquitin-ligase ....................................................... 148 mutants .............................................150, 151, 152, 153 interactions .............................................................. 160

G β-galactosidase activity .................................................. 301 Gateway®-compatible mbSUS vectors .......................... 221 gene internal control ........................................................ 63 genome tiling arrays....................................................... 262 genotype .......................................................................... 19 germination assay ............................................................ 22 gibberellins (GA)....................................................... 23, 47 GIBBERELLIN INSENSITIVE DWARF 1 (GID1) ............................................... 5 glutathione-S-transferase system ................................... 348 gravity assay ..................................................................... 25 growth condition ............................................................. 20 GST-tagged recombinant protein ................................. 341 GUS enzyme activity..................................................... 315 GUS expression assay .................................................... 312

H HCF164 target proteins ................................................ 128 Hela cell ........................................................................ 207 heparin-affinity chromotography .......................... 174, 180 heparin-bound-proteins ................................................ 175

PLANT SIGNAL TRANSDUCTION 355 Index heparin-sepharose .......................................................... 276 chromatography ....................................................... 278 herbicides......................................................................... 53 hexokinase ..................................................................... 1, 6 homozygous lines ............................................................ 19 hormones table of ....................................................................... 37 hydrogen peroxide ......................................................... 109 hypocotyls length............................................................. 23

I IMAC.................................................................... 140, 141 immunodetection........................................................... 199 immunoprecipitation ..................................................... 254 infiltration ..................................................................... 198 infrared image................................................................ 279 interacting hybrid-protein ............................................. 303 interaction of MAPK ............................................ 248, 250 in vitro phosphorylation ................................................ 255 in vitro transcription .............................................. 339, 343 IPG-strip ....................................................................... 181 isolation of nuclei .................................................. 262. 265 IUPAC code .................................................................. 320 IUPAC motif consensus ................................................ 318 interaction model of..................................................................... 13

K

mating ........................................................................... 227 mating-based Split-Ubiquitin System (mbSUS) ............................................................ 217 mCherry ........................................................................ 211 MEL1 reporter gene...................................................... 304 methyl jasmonate ............................................................. 48 microRNA (miRNA) ...................................................... 10 molecular interaction ..................................................... 204 morphology flower and fruit .......................................................... 30 siliques ....................................................................... 30 seed .............................................................................. 3 motif consensus ............................................................. 318 multi-cellular organism.................................................... 12 multiple sequence input ................................................. 328 MUG............................................................................. 316

N native polyacrylamid gel ................................................ 280 negativ regulation .......................................................... 249 network topology............................................................. 12 Nicotiana benthamiana ........................................... 194, 197 nitro blue tetra solium ................................................... 110 non-denaturing gel ........................................................ 274 N-terminal half of ubiquintin (Nab) ............................. 218

O ONPG-assay ................................................................. 227

knock-out mutation .................................................................... 18 Krypton/Argon laser........................................................ 89

L lac-repressor ................................................................... 274 laser scanning microscope.............................................. 207 LC-ESI-MS...................................................176, 179, 184 leaves color ........................................................................... 28 expansion ................................................................... 45 rosette ........................................................................ 26 true ............................................................................ 20 vein pattern of............................................................ 27 LEDs..... .......................................................................... 24 lexA....... ......................................................................... 219 light spectra........................................................................ 24 quality ........................................................................ 24 luminometer .................................................................... 81

M MAP kinase mutants ..................................................... 242 MAP kinase pathway .................................................... 236 mass spectrometry ......................................................... 134 Matchmaker system ...................................................... 295

P PCR................................................................262, 269, 270 pipetting scheme of ................................................. 270 real time ................................................................... 269 real time quantitative ................................................. 61 program ..................................................................... 70 PDI-like protein ............................................................ 176 PEG-mediated transfection .................................. 195, 314 pENTR-TOPO vector .................................................. 305 perfusion of dyes .............................................................. 88 perturbation ..................................................................... 12 phenotypic screen ...................................................... 20, 41 streamlining of ........................................................... 21 flow chart ................................................................... 22 phenotype .................................................................. 18, 19 developmental............................................................ 18 recording of ............................................................... 21 variations ................................................................... 35 phosphatase activities .................................................... 247 phophorimager .............................................................. 346 phosphopeptide enrichment .......................................... 136 photomultiplier tube ...................................................... 102 photoperiod ..................................................................... 29 phototrophic assay ........................................................... 26 phototropism ................................................................... 26

LANT SIGNAL TRANSDUCTION 356 PIndex

phyllotaxis........................................................................ 30 PHYTOCHROME AND FLOWERING TIME (PFT1)..................................................... 11 Place..... ......................................................................... 326 plant genome ...................................................................... 17 plant compatible BiFC vectors .............................. 191, 195 plasmid rescue ................................................237, 239, 241 positive clones................................................................ 300 poly(dIdC) ..................................................................... 287 position weight matrix ................................................... 319 PP2C phosphatase................................................. 248, 250 prey....... ......................................................................... 219 primer design of .................................................................... 65 mastermix .................................................................. 68 optimization .............................................................. 69 probe walking ................................................................ 276 Pro-Q diamond phosphostain ........................134, 136, 139 proteins DELLA ....................................................................... 5 induction ................................................................. 341 stability .................................................................... 147 quantitive analysis of................................................ 183 protein phosphorylation ........................................ 133, 143 relative quantification of .......................................... 143 protein-A sepharose beads............................................. 268 protein-DNA complexes ................................................................ 267 interaction................................................................ 311 protein-protein interaction .................................... 189, 217 protoplasts ......................................................195, 198, 251 isolation ................................................................... 253 transformation ......................................................... 254 pseudo-colour Look Up Table ....................................... 103 p-Thr-antibody ...................................................... 134, 138

R radiolabelled oligoribonucleotide................................... 340 radiolabelled probe ........................................................ 275 radiolabelled transcript, purification of .......................... 343 ratiometric imaging ..........................................98, 102, 104 reactive oxygen species (ROS) ............................53, 94, 109 in vivo imaging of ............................................ 109, 113 feeding of probes ..................................................... 111 receptors light ............................................................................. 5 multiplicity .................................................................. 7 reconstitution of fluorescent YFP .................................. 190 redox-sensitive green fluorescent protein (roGFP) .......... 93 calibration of ............................................................ 103 characterisation of.............................................. 97, 100 imaging of fluorescence ............................................. 96 properties of ............................................................... 94

purification of ............................................................ 99 recombinant ............................................................... 96 stable expression of ............................................ 98, 101 transient expression of ....................................... 97, 100 variants of .................................................................. 95 redox state........................................................................ 93 regulatory DNA sequence ............................................. 321 regulatory sequence retrieval.................................. 322, 323 regulatory sequence analysis programmes.............. 324, 325 replica plating ................................................................ 226 reporter plasmid..................................................... 295, 313 responses to abiotic stress .......................................................... 35 to hormones ............................................................... 35 triple .......................................................................... 46 reversible phosphorylation ............................................. 134 RING-finger-substrate interactions .............................. 160 RNA binding proteins....................................................... 338 isolation of ................................................................. 67 optimization of concentration ................................... 67 quantification of ........................................................ 61 protein interaction ................................................... 337 root development .............................................................. 25 lateral ......................................................................... 25 simultaneous growth assay ......................................... 42 RTqPCR verification of quality ........................................... 71, 72

S 19S cap complex ............................................................ 153 20S central cylinder ....................................................... 153 SDS-PAGE .................................... 135, 194, 198, 255, 339 second messenger .............................................................. 9 senescence........................................................................ 31 sequence logo................................................................. 320 sequencing of phosphopeptides ..................................... 142 signalling network nodes ......................................................................... 11 links ........................................................................... 11 hubes ......................................................................... 11 signals light ............................................................................. 2 long distance ................................................................ 3 target of ....................................................................... 3 silver staining ......................................................... 178, 183 singlet oxygen ................................................................ 109 imaging of................................................................ 114 singlet oxygen sensor green............................................ 110 SLY1.............................................................................. 161 small interfering RNA (siRNA) ...................................... 10 soil...........................................................................................20 special analysis suites ..................................................... 329

PLANT SIGNAL TRANSDUCTION 357 Index spinach Trx-m ............................................................... 125 26S proteasome ..................................................... 148, 153 stem length ...................................................................... 29 sterilization of seeds......................................................... 18 by liquid ..................................................................... 19 by vapour ................................................................... 19 stratification..................................................................... 20 stress cold ............................................................................ 48 heat ............................................................................ 49 influence of ................................................................ 48 light ........................................................................... 53 osmotic ...................................................................... 51 oxidative..................................................................... 53 salt. ................................................................................51 stroma fraction............................................................... 124 substrate labelling .......................................................... 256 superoxide ...................................................................... 109 supershift ....................................................................... 275 SYBR green I .......................................................... 64, 270 synthetic oligoribonucleotide......................................... 345

T target DNA multimerization of ................................................... 296 target genes ........................................................................ 262 promoter .................................................................. 265 transcript.................................................................. 338 temperature ....................................................................... 2 The Arabidopsis Information Resource (TAIR)............................................................... 324 TGA2, 265 thiol-reactive proteins .................................................... 127 thioredoxin (Trx) ........................................................... 117 affinity chromatography .......................................... 118 binding specificity of ............................................... 123 cystein residues ........................................................ 118 disulfide bond .......................................................... 118 efficiency of purification .......................................... 121 isoforms ................................................................... 118 purification strategy ................................................. 120 screening of targets .......................................... 122, 126 target proteins .......................................................... 117 three-dimensional structure ..................................... 119 thylakoid membranes isolation of ............................................................... 137 TILLING ....................................................................... 18 TIR1.............................................................................. 162 tissue culture .................................................................... 18 tobacco BY2 cells..................................................... 81, 212 T7 promoter .................................................................. 343

transcription factor ................................................ 274, 292 bHLH ......................................................................... 8 MADS-box ............................................................... 11 binding sites............................................................. 318 TRANS FAC ................................................................ 326 TRANSPORT INHIBITOR RESPONSE 1 (TIR1) .......................................... 5 trichomes ......................................................................... 27 Trx-immobilised resin ................................................... 125 tryptic digestion ............................................................. 184

U UV-B irradiation ............................................................. 55 ubiquitilation ................................................................. 148 ubiquitilation assays ....................................................... 162 ubiquitin-ligase-pathway mutants ......................... 154, 155 ubiquitin-proteasome-system ........................................ 147 targets of .................................................................. 157 uidA gene ....................................................................... 312 unspecific competitor..................................................... 287 URA3 gene user defined motifs ........................................................ 326 UV-cross-linking ............................................275, 278, 285 UV-light ........................................................................ 285

V vector...........................................................................220 for transient protoplast transfection ......................... 192 for stable transformation.......................................... 192 VERNALIZATION 1 (VRN1) .................................................................. 11 2 (VRN2) .................................................................. 11 VERNALIZATION INSENSITIVE 3 (VIN3) ........... 11

W water potential ................................................................. 52 western blot ....................................................135, 228, 255 western-analysis ..................................................... 194, 198 wounding ....................................................................... 249

X XPD plates .................................................................... 293

Y yeast-one-hybrid-assay .................................................. 291 yeast reporter strain ....................................................... 297 yeast-strains ........................................................... 220, 293 yeast transformation and mating .............222, 224,238, 240 yeast-two-hybrid ........................................................... 218 YEB medium................................................................. 197 YPD medium ................................................................ 238