Chloroplast Research in Arabidopsis: Methods and Protocols, Volume I (Methods in Molecular Biology 774)

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Chloroplast Research in Arabidopsis Methods and Protocols, Volume I

Edited by

R. Paul Jarvis Department of Biology, University of Leicester, Leicester, UK

Editor R. Paul Jarvis, PhD University of Leicester Department of Biology University Road LE1 7RH Leicester United Kingdom [email protected]

ISSN 1064-3745 e-ISSN 1940-6029 ISBN 978-1-61779-233-5 e-ISBN 978-1-61779-234-2 DOI 10.1007/978-1-61779-234-2 Springer New York Dordrecht Heidelberg London Library of Congress Control Number: 2011932678 © Springer Science+Business Media, LLC 2011 All rights reserved. This work may not be translated or copied in whole or in part without the written permission of the publisher (Humana Press, c/o Springer Science+Business Media, LLC, 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. Printed on acid-free paper Humana Press is part of Springer Science+Business Media (www.springer.com)

Preface Chloroplasts are green plastids found in land plants, algae, and some protists. They are the unique site for the reactions of photosynthesis in such cells, and thus chloroplasts are responsible for much of the world’s primary productivity. As photosynthesis is the only significant mechanism of energy-input into living cells, these organelles are essential for the survival of plants and animals alike. Consequently, agriculture is wholly dependent upon the photosynthesis that takes place in chloroplasts. Moreover, many other important cellular activities occur uniquely inside chloroplasts or in other non-photosynthetic types of plastid. These activities include the production of starch, amino acids, fatty acids, lipids, terpenoids, purine and pyrimidine bases, and colourful pigments in fruits, flowers, and leaves, as well as key aspects of nitrogen and sulphur metabolism. Many products of these biosynthetic processes are vital components of mammalian diets or offer opportunities for industrial exploitation. Advances in our understanding of plastid biogenesis will facilitate the manipulation and exploitation of these processes and aid improvements in the quantity or quality of the various products. Over the years, chloroplast biology has been studied in a variety of different organisms, based on technical considerations. Such work has undoubtedly led to major advances in the field, but has had the significant disadvantage that findings made using different experimental systems or species are not always directly cross-comparable. The relatively recent adoption of Arabidopsis thaliana as the model organism of choice for plant science research, across the globe, has led to its emergence as a pre-eminent system for research on chloroplasts and other types of plastid. The availability of genomic sequence resources and extensive germplasm collections for Arabidopsis, as well as its amenability to molecular genetic analysis, have all contributed to this change. This book (together with its partner, Volume II) aims to bring together in a single location some of the most important, modern techniques and approaches for chloroplast research, with the unifying theme of Arabidopsis as the model system. Within the confines of this remit, we have produced a book that is relatively broad in its scope, and which many Arabidopsis researchers and biotechnologists with a general interest in chloroplasts, plastids, or related processes might use as an aid to their work. In essence, it is a book for Arabidopsis integrative biologists with a general focus on chloroplast and plastid research. In spite of the central position afforded to Arabidopsis, many of the presented methods can be applied to other experimental organisms with minimal modification. Leicester, UK

R. Paul Jarvis

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

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Part I  Genetics, Cytology, and In Vivo Analysis   1 Screening or Selection for Chloroplast Biogenesis Mutants of Arabidopsis, Following Chemical or Insertional Mutagenesis . . . . . . . . . . . . . . . 3 Enrique López-Juez and Alison Hills   2 Analysis of Plastid Number, Size, and Distribution in Arabidopsis Plants by Light and Fluorescence Microscopy . . . . . . . . . . . . . . . . . 19 Kevin Pyke   3 Immunofluorescence Microscopy for Localization of Arabidopsis Chloroplast Proteins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 33 Stanislav Vitha and Katherine W. Osteryoung   4 Transient Expression and Analysis of Chloroplast Proteins in Arabidopsis Protoplasts . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 59 Dong Wook Lee and Inhwan Hwang   5 Visualisation of Stromules on Arabidopsis Plastids . . . . . . . . . . . . . . . . . . . . . . . . . 73 John C. Gray, James A. Sullivan, and Christine A. Newell   6 Analysis of Chloroplast Movement and Relocation in Arabidopsis . . . . . . . . . . . . . 87 Masamitsu Wada and Sam-Geun Kong   7 Studying Starch Content and Sedimentation of Amyloplast Statoliths in Arabidopsis Roots . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 103 John Stanga, Allison Strohm, and Patrick H. Masson   8 Studying Arabidopsis Chloroplast Structural Organisation Using Transmission Electron Microscopy . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 113 Stefan Hyman and R. Paul Jarvis   9 Transplastomics in Arabidopsis: Progress Toward Developing an Efficient Method . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 133 Kerry Ann Lutz, Arun Azhagiri, and Pal Maliga

Part II  Gene Expression and Protein Accumulation 10 Isolation, Quantification, and Analysis of Chloroplast DNA . . . . . . . . . . . . . . . . . . Beth A. Rowan and Arnold J. Bendich 11 Measurement of Transcription Rates in Arabidopsis Chloroplasts . . . . . . . . . . . . . . Yan O. Zubo, Thomas Börner, and Karsten Liere 12 Studying the Structure and Processing of Chloroplast Transcripts . . . . . . . . . . . . . . Alice Barkan 13 In Vitro RNA-Binding Assay for Studying Trans-Factors for RNA Editing in Chloroplasts . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Toshiharu Shikanai and Kenji Okuda

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14 Studying Translation in Arabidopsis Chloroplasts . . . . . . . . . . . . . . . . . . . . . . . . . . 209 Paolo Pesaresi 15 Studying Proteases and Protein Turnover in Arabidopsis Chloroplasts . . . . . . . . . . 225 Lars L.E. Sjögren and Adrian K. Clarke

Part III  Protein Transport, Localization, and Topology 16 In Silico Methods for Identifying Organellar and Suborganellar Targeting Peptides in Arabidopsis Chloroplast Proteins and for Predicting the Topology of Membrane Proteins . . . . . . . . . . . . . . . . . . . . . Sandra K. Tanz and Ian Small 17 Rapid Isolation of Arabidopsis Chloroplasts and Their Use for In Vitro Protein Import Assays . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Henrik Aronsson and R. Paul Jarvis 18 Energetic Manipulation of Chloroplast Protein Import and the Use of Chemical Cross-Linkers to Map Protein–Protein Interactions . . . . . . . . . . . . . . Hitoshi Inoue, Fei Wang, Takehito Inaba, and Danny J. Schnell 19 Isolation of Arabidopsis Thylakoid Membranes and Their Use for In Vitro Protein Insertion or Transport Assays . . . . . . . . . . . . . . . . . . . . . . . . . Thomas Bals and Danja Schünemann 20 Determining the Location of an Arabidopsis Chloroplast Protein Using In Vitro Import Followed by Fractionation and Alkaline Extraction . . . . . . . Chiung-Chih Chu and Hsou-min Li 21 Studying Arabidopsis Envelope Protein Localization and Topology Using Thermolysin and Trypsin Proteases . . . . . . . . . . . . . . . . . . . . John Froehlich

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Index . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 369

Contributors Henrik Aronsson  •  Department of Plant and Environmental Sciences, University of Gothenburg, Gothenburg, Sweden Arun Azhagiri  •  Waksman Institute of Microbiology Rutgers, The State University of New Jersey, Piscataway, NJ, USA Thomas Bals  •  Molecular Biology of Plant Organelles, Ruhr-University Bochum, Bochum, Germany Alice Barkan  •  Institute of Molecular Biology, University of Oregon, Eugene, OR, USA Arnold J. Bendich  •  Department of Biology, University of Washington, Seattle, WA, USA Thomas Börner  •  Institut für Biologie (Genetik), Humboldt-Universität zu Berlin, Berlin, Germany Chiung-Chih Chu  •  Institute of Molecular Biology, Academia Sinica, Taipei, Taiwan Adrian K. Clarke  •  Department of Plant and Environmental Sciences, Gothenburg University, Gothenburg, Sweden John Froehlich  •  MSU-DOE Plant Research Laboratory, Michigan State University, East Lansing, MI, USA John C. Gray  •  Department of Plant Sciences, University of Cambridge, Cambridge, UK Alison Hills  •  School of Biological Sciences, Royal Holloway, University of London, Egham, Surrey, UK Inhwan Hwang  •  Division of Integrative Bioscience and Biotechnology, Pohang University of Sciences and Technology, Pohang, Republic of Korea Stefan Hyman  •  Core Biotechnology Services Electron Microscopy Laboratory, University of Leicester, Leicester, UK Takehito Inaba  •  Department of Biochemistry and Molecular Biology, University of Massachusetts, Amherst, MA, USA Hitoshi Inoue  •  Department of Biochemistry and Molecular Biology, University of Massachusetts, Amherst, MA, USA R. Paul Jarvis  •  Department of Biology, University of Leicester, Leicester, UK Sam-Geun Kong  •  Department of Biology, Kyushu University, Higashi-ku, Fukuoka, Japan Dong Wook Lee  •  Department of Life Science, Pohang University of Science and Technology, Pohang, Republic of Korea Hsou-min Li  •  Institute of Molecular Biology, Academia Sinica, Taipei, Taiwan Karsten Liere  •  Institut für Biologie (Genetik), Humboldt-Universität zu Berlin, Berlin, Germany Enrique López-Juez  •  School of Biological Sciences, Royal Holloway, University of London, Egham, Surrey, UK ix

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Kerry Ann Lutz  •  Waksman Institute of Microbiology Rutgers, The State University of New Jersey, Piscataway, NJ, USA Pal Maliga  •  Waksman Institute of Microbiology Rutgers, The State University of New Jersey, Piscataway, NJ, USA Patrick H. Masson  •  Laboratory of Genetics, University of Wisconsin-Madison, Madison, WI, USA Christine A. Newell  •  Department of Plant Sciences, University of Cambridge, Cambridge, UK Kenji Okuda  •  Department of Life Science, Faculty of Science and Engineering, Chuo University, Tokyo, Japan Katherine W. Osteryoung  •  Department of Plant Biology, Michigan State University, East Lansing, MI, USA Paolo Pesaresi  •  Dipartimento di Scienze Biomolecolari e Biotecnologie, Università degli studi di Milano, Milano, Italy Kevin Pyke  •  Division of Plant and Crop Sciences, School of Biosciences, University of Nottingham, Leicestershire, UK Beth A. Rowan  •  Department of Biology, University of Washington, Seattle, WA, USA Danny J. Schnell  •  Department of Biochemistry and Molecular Biology, University of Massachusetts, Amherst, MA, USA Danja Schünemann  •  Molecular Biology of Plant Organelles, Ruhr-University Bochum, Bochum, Germany Toshiharu Shikanai  •  Department of Botany, Graduate School of Science, Kyoto University, Kyoto, Japan Lars L.E. Sjögren  •  Department of Plant and Environmental Sciences, Gothenburg University, Gothenburg, Sweden Ian Small  •  Australian Research Council Centre of Excellence in Plant Energy Biology, University of Western Australia, Crawley, WA, Australia John Stanga  •  Laboratory of Genetics, University of Wisconsin-Madison, Madison, WI, USA Allison Strohm  •  Laboratory of Genetics, University of Wisconsin-Madison, Madison, WI, USA James A. Sullivan  •  School of Biological and Chemical Sciences, Queen Mary, University of London, London, UK Sandra K. Tanz  •  Australian Research Council Centre of Excellence in Plant Energy Biology, University of Western Australia, Crawley, WA, Australia Stanislav Vitha  •  Microscopy and Imaging Center, Texas A&M University, College Station, TX, USA Masamitsu Wada  •  Department of Biology, Kyushu University, Higashi-ku, Fukuoka, Japan Fei Wang  •  Department of Biochemistry and Molecular Biology, University of Massachusetts, Amherst, MA, USA Yan O. Zubo  •  Institut für Biologie (Genetik), Humboldt-Universität zu Berlin, Berlin, Germany

Part I Genetics, Cytology, and In Vivo Analysis

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Chapter 1 Screening or Selection for Chloroplast Biogenesis Mutants of Arabidopsis, Following Chemical or Insertional Mutagenesis Enrique López-Juez and Alison Hills Abstract The power of Arabidopsis as a model organism lies in the depth and breadth of genetic tools available for its study. This also applies to the study of chloroplast biology. Although vast numbers of mutants have been identified in Arabidopsis, the continued use of forward-genetic screening approaches remains valuable for the isolation and study of previously overlooked mutants and novel mutations in sensitised backgrounds (i.e., suppressors or enhancers of previously known mutants). In addition, reverse-genetic collections of insertional mutants are now extensive and provide unique opportunities for gene function discovery. Here, we describe methods for the chemical mutagenesis of Arabidopsis, the screening of mutants visually, on the basis of gene-expression phenotypes (scored as reduced or enhanced activity of reporter genes), and the use of databases to select for existing mutations from historic collections or insertional mutagenesis programmes. Key words: Chloroplast, Plastid, Genetics, Mutant, Reporter gene, ADH, GFP

1. Introduction Two-thirds of primary productivity on the planet, and ultimately the majority of our food, depend on photosynthesis by land ­ecosystems, and of this the vast majority is carried out by higher plants. This makes studying not only photosynthesis, but also the biology of higher plant chloroplasts (the cellular organelles in which photosynthesis takes place) specifically such an important endeavour. Chloroplasts have been successfully isolated from many different organisms in the past. What makes Arabidopsis thaliana such a useful tool is not a particular high yield or ease of isolation of the organelles, but its power as a genetic model organism.

R. Paul Jarvis (ed.), Chloroplast Research in Arabidopsis: Methods and Protocols, Volume I, Methods in Molecular Biology, vol. 774, DOI 10.1007/978-1-61779-234-2_1, © Springer Science+Business Media, LLC 2011

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While  molecular genetic tools and resources for cereals such as maize and rice are rapidly being developed (1), at present the power of Arabidopsis genetics is unparalleled (2–4). With Arabidopsis, forward-genetic screens (the isolation of mutants altered in a chloroplast function based on their phenotype) can be carried out on Petri dishes or trays in growth rooms, rather than in fields, because of the plant’s small size. The phenotypes used for selection may be visual, or may be based on gene expression, using reporter gene technology, facilitated by the ease with which Arabidopsis transgenic lines can be generated (5). A large number of Arabidopsis mutants for chloroplast functions and development exist, making it possible to design and carry out more elaborate screens (e.g., for the identification of suppressor or enhancer mutations). This can potentially lead to the identification of functions that would otherwise go unnoticed in the absence of the sensitised background (6–8). Genome-wide and systems-based approaches are now identifying and cataloguing genes for proteins with a presumed chloroplast function, or which are predicted to be targeted to the organelle. The existence of large collections of Arabidopsis insertional mutants makes it possible to use a reverse-genetic approach, to identify lossof-function mutants in a particular gene of interest before characterising their phenotypes (9, 10). In this chapter, we first describe methods for the mutagenesis of Arabidopsis and the selection of mutants in forward-genetic approaches. Besides visual selection, two specific methods are described, for the isolation of mutants by selecting for (positive) or against (negative) expression of a reporter gene. We then briefly describe the identification of mutants in individual genes among existing collections, for reverse-genetic approaches. This last topic is complemented by a separate chapter in this book (see Chapter 9, Vol. 2), which covers in particular the systematic identification and characterisation of such mutants. Mutants identified by these different approaches may subsequently be characterised in detail using the many different techniques described in the other chapters in this book.

2. Materials 2.1. Forward Genetics: Chemical Mutagenesis and Visual Screening

1. All chemicals are from Sigma, Poole, UK, unless otherwise stated. 2. Seedling trays, half-trays, and propagator lids (PST-type, Desch Plantpak, Maldon, UK). 3. Levington M3 potting compost, John Innes No. 3 soil, and vermiculite, mixed in 6:6:1 proportions per volume (Scotts Professional, Ipswich, UK).

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4. KCl solution: 0.1% (w/v) KCl. 5. Ethyl methanesulfonate (EMS) solution: 0.1  M NaH2PO4, pH 5.0 (adjusted with H3PO4 as needed), 5% (v/v) dimethyl sulphoxide, and 75 mM EMS. EMS is a powerful carcinogen. See Subheading 3.1 for information on safe handling. 6. Sodium thiosulphate solution: 0.1 M Na2S2O3. 7. Solid sodium thiosulphate in a waste beaker. 8. Molten soft agar: 0.1% (w/v) Microagar (Duchefa, Melford Laboratories, Ipswich, UK), made molten by autoclaving. 9. Laminar flow hood (e.g., Heraeus, Newport Pagnell, UK), required for tissue culture-based screens. 10. Tissue culture Petri dishes: Cellstar 9-cm plates (see Note 1) (Greiner Bio-One, Stonehouse, UK). 11. Murashige and Skoog (MS) solid medium: 4.3 g MS salt mixture (Duchefa), 0.5 g 2-(N-morpholino)ethanesulphonic acid (MES), 10 g sucrose, pH 5.7, adjusted with KOH, and 8 g Microagar (Duchefa) (see Note 2). Autoclave at 121°C for 20 min. 2.2. Forward Genetics: Reporter Gene Negative Selection

1. Arabidopsis mutant R002 (ecotype Bensheim); Nottingham Arabidopsis Stock Centre (NASC) catalogue number N8102. 2. Arabidopsis transgenic line pOCA108 (11), harbouring LHCB1.2 promoter-driven alcohol dehydrogenase (ADH); NASC catalogue number N9400. This line is specific for the screen described in this chapter. 3. Cellstar six-well microtitre plates (Greiner). 4. Allyl alcohol (prop-2-en-1-ol) solution: 3 mM allyl alcohol in liquid MS medium.

2.3. Forward Genetics: Reporter Gene Positive Selection

1. Fluorescence-based molecular imager, designed primarily for gels, blots, or plates: FluorImager™ (Amersham Biosciences, GE Healthcare, Little Chalfont, UK), equipped with blue argon (488  nm) excitation laser (see Note 3 for details and alternatives). 2. Emission filters for FluorImager™ (Amersham Biosciences): 510–545 band-pass DF30 acrylic filter for green fluorescent protein (GFP) (12) and long-pass 610RG acrylic filter for chlorophyll. 3. ImageQuant image analysis software (Amersham Biosciences), or equivalent.

2.4. Forward Genetics: Survey of Existing Mutant Collections

Surveys of existing mutant collections would be carried out online, as described in Subheading 3.4 below. As such, the only material item required for identification of the mutant(s) is a personal computer with an Internet connection.

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2.5. Reverse Genetics: Diversity of Collections

1. Go-Taq DNA polymerase, buffer, and dNTPs (Promega, Southampton, UK). 2. Blue microcentrifuge tube pestles (see Note 4) (Anachem, Luton, UK). 3. DNA extraction buffer: 200 mM Tris (hydroxymethyl) aminomethane (Tris)–HCl, pH 7.5, 250 mM NaCl, 25 mM ethylenediaminetetraacetic acid disodium salt (EDTA), and 0.5% (w/v) sodium dodecyl sulphate (SDS). 4. Isopropyl alcohol. 5. 70% (v/v) ethanol. 6. TE: 10 mM Tris–HCl, pH 7.5, and 2 mM EDTA. 7. Standard reagents electrophoresis.

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3. Methods 3.1. Forward Genetics: Chemical Mutagenesis and Visual Screening

Mutations can be generated in Arabidopsis by treatment with EMS, a powerful chemical mutagen that introduces an ethyl group to a DNA base, causing mispairing and, in the subsequent round of DNA replication, a GC ® AT transition. Seeds treated with EMS will acquire mutations, in tissues including the embryonic shoot meristem cells, and will grow into plants designated as M1, with the mutations being present in heterozygous clones of cells in these plants. When such clones of cells contribute to the formation of flowers, seeds (M2) resulting from the self-fertilisation of those flowers will, with a frequency of 1/4, carry the mutation in a homozygous state, and therefore grow into mutant M2 plants. We have used the following protocol, which is adapted from the one devised by Leyser and Furner (13). As EMS is a dangerous, powerful mutagen, and is also volatile, all work should be carried out in a fume hood with appropriate protective clothing, and great care should be exercised to ensure complete inactivation, before disposal as general sink waste. All steps in the following protocol should be conducted at room temperature. 1. Collect 20,000–40,000 seeds (see Note 5) of Arabidopsis wild type (see Note 6) or, in the case of suppressor/enhancer screens, the mutant for which second-site modifiers are to be identified (see Note 7). 2. Place the seeds in a 50-mL polypropylene tube and pre-imbibe them overnight in KCl solution. 3. Remove the KCl solution by pipetting and soak the seeds for 3  h in 30  mL of EMS solution (see Note 8), with gentle shaking.

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4. Remove the EMS solution into a beaker containing solid sodium thiosulphate and add a further 30 ml of thiosulphate solution (see Note 9). 5. Wash the seeds (add the solution and allow the seeds to sediment before removing it) three times in 100 mM sodium thiosulphate for 15 min each time to destroy any traces of EMS. 6. Wash the seeds three times in sterile distilled H2O (dH2O) and resuspend in 50 mL of cold, molten, sterile 0.1% agar. 7. Store the seeds at 4°C for 3 days, to ensure rapid, synchronous germination. 8. Separate an approximate 1,000 seed aliquot (0.8–1.6 mL) into a sterile tube and keep to one side. Then, distribute 10 mL of the remaining solution with dispersed seeds into a total of five 50-mL tubes containing further molten agar (this will prevent the seeds from sedimenting readily), before dispersing with a Pasteur pipette onto standard seed trays with soil mixture, at a density of 150 seeds per tray. 9. Cover each of the trays with a propagator lid and transfer them to a growth room (21°C, 16 h day-length, at 150–200 mmol/ m2/s). Remove the propagator lids after 2–3 days. 10. Distribute the retained 1,000-seed aliquot onto five agar-­ solidified MS plates to assess mutagenesis. To do this, perform the following procedure under sterile conditions in a laminar flow hood. Place seeds in a 1.5-mL microcentrifuge tube, cover them in absolute ethanol, and mix. After 1  min, centrifuge briefly at low speed (~1,000 rpm) in a microfuge to sediment the seeds, remove the ethanol, then cover them in 25% (v/v) dilute household bleach, and mix as before. After 10 min, sediment the seeds, remove the bleach, wash five times in sterile dH2O, and then resuspend in cold, molten, sterile 0.1% agar. Distribute the seeds onto the surface of the plates, by collecting the soft agar solution containing the dispersed seeds with a 1,000-mL pipette, and pipetting slowly onto the surface of plates (see Note 10). Allow the liquid agar to dry by keeping the plates open in the laminar flow hood. Seal the plates with porous surgical tape and transfer them to a tissue culture incubator (21°C, 16  h day-length, at 150  mmol/m2/s). After approximately 7 days, score the frequency of albino seedlings. A frequency of albinos among the M1 between 0.1 and 1% will confirm successful mutagenesis, at a density sufficient to ensure saturating mutagenesis of the Arabidopsis genome, due to multiple mutations per M1 plant (see Note 7). 11. After 7–8 weeks of growth on soil, enclose plant flowering shoots in a cellophane or paper bag, and gradually reduce watering to allow seed maturation. Collect all seeds from each tray. The seed pool from each tray is considered an M2 pool (see Note 11). Allow for 1 week of after-ripening.

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12. Imbibe and stratify 1,500 seeds of each M2 pool (this will ensure on average 10 seeds per M1 plant, giving a >95% probability of including at least one homozygous mutant). To do this, place each pool of seeds in a microcentrifuge tube, cover them in ethanol, mix, centrifuge at low speed to sediment the seeds, remove the ethanol, wash twice in dH2O, and resuspend the seeds in cold, molten, 0.1% agar. Disperse the seeds onto MS medium or trays with soil (depending on the screening requirements; see below). 13. Subsequent analysis will depend on the approach to be used for screening for photosynthetic apparatus, chloroplast, or plastid biogenesis defects. Chloroplast development defects often result in seedlings that are, when grown on soil, either uniformly pale, green with pale interveinal regions, or virescent (pale newly emerging leaves that gradually become greener, hence leading to green rosettes with a pale centre, i.e., delayed greening) (11, 14–16). Early plastid development defects can result in albino seedlings, with very pale or white cotyledons, which will require growth on MS plates containing at least 1% sucrose. Photosynthetic defects often result in impaired electron transport, which leads to high chlorophyll fluorescence, or to decreased photosynthetic efficiency as measured as the reduced ratio of variable over maximum fluorescence of dark-adapted seedlings (see Note 12). Seedlings or plants identified by such screens will be putative mutants and will require confirmation of heritability of the phenotype in the M3 generation, before being considered true mutants. 3.2. Forward Genetics: Reporter Gene Negative Selection

A genetic approach can be taken one step further by searching for regulatory mutations. Such a search (e.g., in our case, for drivers of  plastid development) might identify mutants exhibiting mis-­ regulation of expression of genes necessary for plastid biogenesis. Negative screens might select for mutants in which a gene which would otherwise be expected to be expressed is not expressed or its expression is reduced; such mutants would harbour mutations in positive drivers of gene expression. Positive screens, on the contrary, would aim at identifying mutants with enhanced gene expression, and the mutated genes would be anticipated to be negative regulators of gene expression. Such screens can be carried out through the use of lines harbouring reporter genes under the control of the promoter of choice. In Subheading 3.3, a positive selection screen is presented, based on expression of GFP. Below we describe a negative selection screen. In the negative screen, normal expression of a reporter gene encoding an enzyme, ADH, driven by the promoter of a chloroplast-protein gene, causes conversion of relatively harmless allyl alcohol into the toxic aldehyde acrolein (propenal), and subsequent seedling lethality. Therefore, normal expression is selected

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against, and surviving mutants exhibit reduced reporter ­expression. The screen requires all the ADH activity present in the plants to be under the control of the promoter of choice, and so an Arabidopsis line devoid of endogenous ADH activity, the R002 mutant (17), is  used. The method, designed by Chory and collaborators, has been successfully used to identify defects in the expression of the ­light-harvesting chlorophyll-binding protein gene, LHCB1 (11, 15, 18). 1. The screen requires prior generation of a transgenic line harbouring a plastid- or chloroplast-protein gene promoter, driving expression of ADH, transformed into Arabidopsis mutant R002 (ecotype Bensheim). Such a line, harbouring LHCB1 promoter-driven ADH, is available from NASC. 2. Surface-sterilise and stratify 1,500 seeds per M2 mutagenised pool, as described in Subheading 3.1. 3. Distribute the seeds in batches of 300 in six-well microtitre plates (i.e., ~1,800 seeds per plate), with the wells containing 6 mL of liquid MS medium with 1% sucrose. Place the plates on a low-profile shaker in a plant tissue culture incubator under continuous light (150 mmol/m2/s, 21°C) for 5 days. Seedlings will grow into a single mass with cotyledons exposed to the surface. 4. Under aseptic conditions, aspirate the medium and dispose of as toxic waste. Replace it with 6 mL of MS medium containing allyl alcohol and then incubate for 1 h in the plant tissue culture incubator (see Note 13). 5. Wash the seedlings thrice in sterile, fresh MS and carefully distribute them with sterile forceps onto solid MS plates containing 1% sucrose. Avoid damaging the seedlings, but ensure that they are separate enough for the roots of all seedlings to be in contact with the medium. Transfer to the plant tissue culture incubator. 6. Over the following 2–7 days, cotyledons will bleach and most seedlings will die. Occasional surviving seedlings will bleach and appear to die, but will develop new green leaves. Allow growth for a further week before transferral to soil, as putative mutants. 3.3. Forward Genetics: Reporter Gene Positive Selection (see Note 14)

The protocol below describes a positive selection screen based on the one designed by Niwa and collaborators (12), for monitoring GFP expression in whole seedlings. The screen seeks elevated reporter expression under conditions, or in a mutagenised genetic background, in which it is normally not observed, or only observed at low levels.

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1. The screen requires prior generation of a transgenic line ­harbouring a plastid- or chloroplast-development associated gene promoter, driving expression of GFP, transformed into Arabidopsis. Use of a bright GFP with a high quantum efficiency, like the red-shifted, synthetic GFP (12), is essential. 2. Surface-sterilise and stratify pools of 1,500 seeds per M2 mutagenised pool, as described in Subheading 3.1. 3. Distribute the seeds in batches of approximately 200 in plates containing solid MS medium with 1% sucrose, and transfer the plates to appropriate growth or selection conditions. 4. After 7 days, subject these plates to analysis of fluorescence in a FluorImager™ or an equivalent imager (see Note 3). Use the blue laser as excitation light and emission filters corresponding to Fluorescein/GFP and to Texas Red/chlorophyll. Scan the plates and obtain two images per plate, one for each emission filter. 5. Examine images in the chlorophyll channel using ImageQuant software. Select bright areas (cotyledons), quantify the absolute fluorescence signal, copy and paste those areas onto the same image under the GFP channel, and then record the second absolute fluorescence signal. Using the two recorded values, calculate a ratio of GFP/chlorophyll (see Note 15). Seedlings in which the ratio is above a chosen threshold are selected as putative mutants for further genetic analysis. 6. Figure 1 provides examples of images obtained from seedling plates with the FluorImager (see Notes 3 and 16).

Fig. 1. In vivo reporter gene for positive selection screening. FluorImager™ images of seedlings of a light-hyposensitive mutant harbouring an LHCB1.2 promoter-driven GFP (two separate seed pools, upper half of each panel), or isogenic controls without the reporter gene (two pools, lower half of each panel). Seedlings were grown on solid MS medium under continuous white light of 25 mmol/m2/s irradiance for 7 days. The panels show images obtained under blue laser excitation and GFP emission filter (left ), chlorophyll emission filter (middle ), or overlay of both emissions (right ).

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3.4. Forward Genetics: Survey of Existing Mutant Collections

11

Plastid-related mutations often result in greening defects. The main international Arabidopsis stock centres, in Nottingham (UK), Ohio (USA), and Tsukuba/Sendai (Japan), have developed online resources to display searchable images of the visible phenotypes of available stocks. These stocks can be browsed by mutant image, or searched via keywords such as “pale” or “reticulate”. For example the Nottingham Arabidopsis Stock Centre (NASC) photograph collection carries 165 entries as “colour” mutants, as of October 2010. This approached has been successfully utilised in the past (14). More recently, two large-scale projects taking a systems biology approach, the Chloroplast 2010 project (19) (see Chapter 9, Vol. 2) and the Chloroplast Function Database project at RIKEN (20) (see Table  1), have examined and systematically catalogued phenotypes of insertional mutants for over 3,500 genes whose protein product is predicted to be targeted to the chloroplast. These data are available and searchable online using controlled vocabulary. Although intended as reverse-genetic resources, these can in theory also be used for forward screening. For example if one has identified an unusual phenotype associated with a mutation in a given gene, such a phenotype might then be sought in other mutants, the lesions in which could lie in genes of previously unsuspected relationship to the process under study. Table 1 summarises these available resources. Seeds can be ordered from the three stock centres, at Nottingham, Ohio, and Tsukuba, which also carry the stocks utilised and described by the large-scale chloroplast systems biology projects.

Table 1 Online databases of existing mutant collections containing potential chloroplast biogenesis mutants, and of genome-wide projects that have identified insertional mutants in genes for most proteins predicted to be targeted to the chloroplast Stock centre

Address for mutant search

Nottingham Arabidopsis Stock Centre (NASC)

http://www.arabidopsis.info/InfoPages?template=photopage; web_section=germplasm

Arabidopsis Biological Resource Centre (ABRC, Ohio)

http://arabidopsis.org/servlets/Order?state=catalog

RIKEN Bioresource Centre

http://www.brc.riken.jp/inf/en/index.shtml

Chloroplast 2010, University of Michigan

http://www.plastid.msu.edu/

Chloroplast Function Database, RIKEN

http://rarge.psc.riken.jp/chloroplast/

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3.5. Reverse Genetics: Diversity of Collections and Genotyping

Reverse genetics refers to the search for a mutant phenotype associated with a gene whose sequence is already known. While not without drawbacks, this is a powerful tool to understand gene function and is usually carried out by selecting already known, sequence-indexed insertional mutants at the locus under study. As of October 2010, seventeen different projects are listed in The Arabidopsis Information Resource (TAIR) as having generated collections of insertional mutants (see Note 17). The insertional mutagenesis generates in its first-generation heterozygous plants, from which sequence is obtained flanking the insertion, to catalogue the mutated gene (this is done by the organisation providing the mutant population). Often it is then necessary for the end-user to employ a genotyping assay to identify the homozygous mutants among the progeny of such plants, but for an increasing number of genes, such homozygous mutants have now been identified and deposited back in the stock centres (see Note 18). Both chloroplast-focused systems biology projects referred to in Subheading 3.4 have identified mutants for most genes encoding chloroplast-­ targeted proteins (when this is not prevented by lethality of the knockout mutation). However, such mutant lists are unlikely to be comprehensive yet, and in addition, mutants may be sought in other genes whose products are not targeted to the chloroplast, but have nevertheless a chloroplast biogenesis impact. It may, therefore, still be necessary to identify other insertional mutants. The following procedure seeks such mutants from online databases and briefly describes their genotypic characterisation. 1. Open the TAIR sequence viewer online (see Note 19). Search for the gene of interest. Note that for some genes the search may need to use a different database (see Note 20). 2. In the results page, select the gene in the five-chromosome panel, to open a close-up view. Display all gene models and T-DNA/transposon insertions, at an appropriately high resolution (20 kb). 3. Select an appropriate insertion line when available. The sequences shown (flanking sequence tags [FSTs]) typically read from the border of the insert and outwards into the affected gene; therefore, the beginning of the FST sequence is most likely to reflect the actual position of the insertion (see Note 21). 4. Go to the T-DNA primer design tool (see Note 22) of the Salk Institute Genomic Analysis Laboratory and obtain left primer (LP) and right primer (RP) sequences to genotype the line. The tool will provide predicted sizes for the amplicons resulting from polymerase chair reactions (PCRs) using LP and RP together, and using the border primer (BP) together with RP. Since different collections use either different T-DNA vectors,

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Table 2 Border primers (BP) needed to genotype mutants originating from different insertional mutant collections Line type

Name of primer

Sequence (5¢ to 3¢)

SALK

LBb1.3

ATTTTGCCGATTTCGGAAC

GABI-KAT

GK-TDNA

ATATTGACCATCATACTCATTGC

JIC Gene Trap

GT Ds3-1b

ACCCGACCGGATCGTATCGGT

Wisc DsLox

p745b (Ds-Lox)

GTCCGCAATGTGTTATTAAGTTG

JIC SM

Spm32b (SM)

CGAATAAGAGCGTCCATTTTAGAG

SAIL (GARLIC)

LB3b (SAIL)

CATCTGAATTTCATAACCAATCTCG

RIKEN (RATM)

RIKEN Ds5-2a

TCCGTTCCGTTTTCGTTTTTTAC

This information should be confirmed with the information for the individual resources available online, as some collections include more than one insertional vector, or have in the past had earlier primers replaced with improved alternatives

or transposons, different BPs will be required depending on the source of the line. Sequences of the most common BPs are provided in Table 2. 5. Order the insertion line from one of the stock centres, and grow at least 20 plants individually (see Note 23). 6. After 2–3 weeks on soil, collect one full leaf to obtain template genomic DNA from each individual plant. 7. Crush/grind the leaf by hand using a microfuge pestle in a microfuge tube, for a few seconds at room temperature, until the sample appears juicy. 8. Promptly add 0.5 mL of DNA extraction buffer and continue to grind to break up large clumps. 9. Vortex vigorously for 10 s (see Note 24). 10. Centrifuge for 5 min at full speed (13,000 rpm) in a microfuge at room temperature. Remove 450 mL of the supernatant to a new tube. 11. Add 450 mL of isopropyl alcohol and allow DNA precipitation to occur at room temperature for 10 min. 12. Centrifuge as in step 10 above (see Note 25). Pour off the supernatant. 13. Wash with 1 mL of ice-cold 70% ethanol. Thoroughly remove the ethanol.

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14. Dry the pellet by placing the tube at 65°C in a heat block with the tube lid open for 5 min. 15. Resuspend the pellet in 50 mL of TE. Use 1.5 mL of this DNA preparation per PCR. 16. Use the LP + RP and BP + RP primer combinations (see Note 26) in separate PCRs with each template genomic DNA. Use also wild-type DNA template as a positive control for the LP + RP reaction. 17. Resolve the amplicons by agarose gel electrophoresis. An amplicon of the expected size for LP + RP will indicate the presence of the wild-type allele in the template DNA. An amplicon for BP + RP will indicate the presence of the insertion, and therefore the mutant allele, within the target gene. Generation of both amplicons from the same template will indicate heterozygosity of the source plant. Plants producing only BP + RP amplicons are, therefore, homozygous knockout mutants.

4. Notes 1. Use these or other deep plates (1.5 cm high), to allow seedlings to grow without touching the plate lid. 2. Add microagar after pH adjustment. The agar will dissolve during autoclaving. High quality agar (e.g., Plant agar or Microagar, Duchefa) is essential for plant tissue culture. 3. The Storm™ or Typhoon™ (Amersham Biosciences) fluorescence molecular imagers may also be used. However, the Typhoon may not be recommended, as the image is obtained through confocal optics, making it highly sensitive during screening to the exact position of the seedling fluorescence source (the cotyledons) as described in Subheading 3.3, steps 4–6. This poses a difficulty if genotypes or growth conditions are used that result in seedling hypocotyl lengths (and consequently cotyledon positions) that are not highly consistent. Other fluorescence molecular imagers that use blue light ­excitation can also be used and are available from other manufacturers: e.g., Bio-Rad (Hercules, CA, USA) and Fujifilm (Tokyo, Japan). However, the Odyssey infra-red imaging ­system (Li-Cor, Lincoln, NE, USA) cannot be utilised as it uses longer wavelength light for both excitation and emission. 4. These pestles are best abraded with sand paper before first use. They can be reused after washing, incubating with 10% (v/v) household bleach, and autoclaving. 5. 1,000 Arabidopsis seeds weigh approximately 20–25 mg.

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6. If the mutant gene is to be identified by map-based cloning approaches, it is important to use an ecotype for which good information on genomic polymorphisms is available. The reference Arabidopsis sequence is derived from the Columbia-0 (Col-0) ecotype. A suitable choice would be to select for the mutant in Col-0, and to subsequently map it against the Landsberg erecta (La-er) ecotype. 7. The number of mutagenised M1 plants required for a saturating screen will depend on the average number of mutations carried in each M1 plant, but has been calculated to be in the range of 125,000 (21). The maximum number of mutations carried per M1 plant is limited by the accumulation of mutations that cause gametophytic (pollen or embryo sac) lethality, and therefore sterility. The recommendations given in this chapter have succeeded in isolating rare mutations from a population of 100,000 M2 seedlings. A more detailed discussion of the depth or saturation of chemical mutagenesis, as well as scoring alternatives to the identification of albinos, has been presented elsewhere (22). 8. Use of filter tips or plug-containing pipettes will minimise the risk of EMS exposure during and after handling. 9. Add 30 mL of 100 mM sodium thiosulphate to the used EMS solution (containing also thiosulphate crystals), and incubate at room temperature for 15 min to inactivate the EMS before discarding. 10. It is possible to pipette slowly by pressing the pipette plunger, or by turning the pipette graduation dial. Good dispersal of seeds facilitates the observation of albino seedlings. 11. Maintaining separate pools of progeny of the M1 plants serves two purposes: it facilitates subsequent genetic analysis, when mutants are known to be independent if isolated from different pools, and it allows re-screening of the relevant pool when a valuable putative mutant fails to survive. 12. Description of the screening of high chlorophyll fluorescence mutants is beyond the scope of this manual. For examples of outcomes, the reader is referred elsewhere (23). 13. A shaker’s rotation speed of 80 rpm, an allyl alcohol concentration of 3 mM, and incubation for 1 h may be appropriate, but should be optimised experimentally, as each of these parameters will have an effect on seedling growth and reporter expression, or on the ability of allyl alcohol to select against it. 14. Screening for active or enhanced gene expression is theoretically possible based on ADH expression, as ADH detoxifies ethanol and incorporates the product harmlessly into the ­respiratory chain. However, this screen is not feasible on

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Arabidopsis seedlings, as the amount of ethanol required to cause toxicity is sufficient to permeabilise membranes and cause physical damage to seedlings (our unpublished observations). 15. The chlorophyll fluorescence value is essential as an internal positive control, as not only the size and stage, but also the orientation of the seedlings will have an impact. Relatively intense fluorescence is exhibited even by chlorophyll-deficient mutants. However, it is possible that a sought mutation increases both GFP and chlorophyll fluorescence, and so an additional visual inspection of the plates, focusing on the seedlings identified as showing high absolute fluorescence values, is important. 16. It would be possible but more laborious to carry out a similar genetic screen with a hand-held UV lamp in a dark room (if using alternative GFPs, as the one utilised here is a S65T, redshifted type, and has blue-specific excitation). 17. http://www.arabidopsis.org/portals/mutants/findmutants.jsp 18. Stocks are available for homozygous insertional mutants individually or as large sets of lines: http://arabidopsis.info/ CollectionInfo?id=72 19. http://arabidopsis.org/servlets/sv 20. Mutants from some T-DNA collections are not listed in the TAIR database. Two alternatives are the database of the Salk Institute Genomic Analysis Laboratory (SIGnAL; http://signal. salk.edu/cgi-bin/tdnaexpress) and the NASC AtEnsemble genome browser (http://atensembl.arabidopsis.info/index. html; select graphical view). For example knockout mutants from the RIKEN transposon insertion collection are listed in NASC, but not in TAIR. 21. The sequence displayed in SeqViewer corresponds to the flanking sequence obtained from the insertional mutant. Such sequence can include the insert itself, and so may be used to determine the insertion site precisely. However, this is not always the case, as the sequence may start up to 300 bp downstream of the insertion site, and so it is important to choose the line that maximises the chances of the insertion being present within the gene’s open reading frame. 22. http://signal.salk.edu/tdnaprimers.2.html 23. Seeds of the different knockout collections can be sown on plates with media containing the appropriate selection antibiotic or herbicide, to eliminate homozygous wild-type seedlings directly. The SALK T-DNA insertion collection can in theory be grown on MS medium containing 50  mg/L kanamycin. However, the kanamycin resistance gene in this T-DNA suffers

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from frequent silencing, making this preliminary selection unreliable. 24. Six to ten extractions can be done in parallel. The individual extracts can be paused at room temperature after vortexing, until the set is complete. 25. A large, green-coloured pellet should be seen; this is normal. 26. BP + RP should be used to score for the presence of the insert, regardless of the orientation of the insertion. This is because the primer design tool referred to in Note 22 designates as RP the primer on the side of the genomic DNA closer to the T-DNA right border, which the BP targets. References 1. Stern, D. B., Hanson, M. R., and Barkan, A. (2004) Genetics and genomics of chloroplast biogenesis: maize as a model system. Trends Plant Sci. 9, 293–301. 2. Leister, D. (2003) Chloroplast research in the genomic age. Trends Genet. 19, 47–56. 3. Lopez-Juez, E. (2007) Plastid biogenesis, between light and shadows. J. Exp. Bot. 58, 11–26. 4. Sakamoto, W., Miyagishima, S.-y., and Jarvis, P. (2008) Chloroplast biogenesis: control of plastid development, protein import, division and inheritance. In, The Arabidopsis Book. American Society of Plant Biologists, Rockville, MD, USA, doi: 10.1199/tab.0110. 5. Desfeux, C., Clough, S. J., and Bent, A. F. (2000) Female reproductive tissues are the primary target of Agrobacterium-mediated transformation by the Arabidopsis floral-dip method. Plant Physiol. 123, 895–904. 6. Yu, F., Liu, X., Alsheikh, M., Park, S., and Rodermel, S. (2008) Mutations in SUPPRESSOR OF VARIEGATION1, a factor required for normal chloroplast translation, suppress var2-mediated leaf variegation in Arabidopsis. Plant Cell 20, 1786–1804. 7. Stanga, J. P., Boonsirichai, K., Sedbrook, J. C., Otegui, M. S., and Masson, P. H. (2009) A role for the TOC complex in Arabidopsis root gravitropism. Plant Physiol. 149, 1896–1905. 8. Meskauskiene, R., Wursch, M., Laloi, C., Vidi, P. A., Coll, N. S., Kessler, F., Baruah, A., Kim, C., and Apel, K. (2009) A mutation in the Arabidopsis mTERF-related plastid protein SOLDAT10 activates retrograde signaling and suppresses 1O2-induced cell death. Plant J. 60, 399–410. 9. Kuromori, T., Hirayama, T., Kiyosue, Y., Takabe, H., Mizukado, S., Sakurai, T., Akiyama,

K., Kamiya, A., Ito, T., and Shinozaki, K. (2004) A collection of 11 800 single-copy Ds transposon insertion lines in Arabidopsis. Plant J. 37, 897–905. 10. Alonso, J. M., Stepanova, A. N., Leisse, T. J., Kim, C. J., Chen, H., Shinn, P., Stevenson, D. K., Zimmerman, J., Barajas, P., Cheuk, R., Gadrinab, C., Heller, C., Jeske, A., Koesema, E., Meyers, C. C., Parker, H., Prednis, L., Ansari, Y., Choy, N., Deen, H., Geralt, M., Hazari, N., Hom, E., Karnes, M., Mulholland, C., Ndubaku, R., Schmidt, I., Guzman, P., Aguilar-Henonin, L., Schmid, M., Weigel, D., Carter, D. E., Marchand, T., Risseeuw, E., Brogden, D., Zeko, A., Crosby, W. L., Berry, C. C., and Ecker, J. R. (2003) Genome-wide insertional mutagenesis of Arabidopsis thaliana. Science 301, 653–657. 11. Li, H., Culligan, K., Dixon, R. A., and Chory, J. (1995) CUE1: a mesophyll cell-specific positive regulator of light-controlled gene expression in Arabidopsis. Plant Cell 7, 1599–1610. 12. Niwa, Y., Hirano, T., Yoshimoto, K., Shimizu, M., and Kobayashi, H. (1999) Non-invasive quantitative detection and applications of nontoxic, S65T-type green fluorescent protein in living plants. Plant J. 18, 455–463. 13. Leyser, H. M. O., and Furner, I. J. (2000) EMS mutagenesis of Arabidopsis. In, Arabidopsis - A Practical Approach (Wilson, Z. A., ed.), Oxford University Press, Oxford, UK, pp. 12–13. 14. Kinsman, E. A., and Pyke, K. A. (1998) Bundle sheath cells and cell-specific plastid development in Arabidopsis leaves. Development 125, 1815–1822. 15. Lopez-Juez, E., Jarvis, R. P., Takeuchi, A., Page, A. M., and Chory, J. (1998) New Arabidopsis cue mutants suggest a close connection between plastid- and phytochrome

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regulation of nuclear gene expression. Plant Physiol. 118, 803–815. 16. Chi, W., Ma, J., Zhang, D., Guo, J., Chen, F., Lu, C., and Zhang, L. (2008) The pentratricopeptide repeat protein DELAYED GREENING1 is involved in the regulation of early chloroplast development and chloroplast gene expression in Arabidopsis. Plant Physiol. 147, 573–584. 17. Jacobs, M., Dolferus, R., and Van den Bossche, D. (1988) Isolation and biochemical analysis of ethyl methanesulfonate-induced alcohol dehydrogenase null mutants of Arabidopsis thaliana (L.) Heynh. Biochem. Genet. 26, 105–122. 18. Chory, J., Li, H. M., and Mochizuki, N. (1995) Molecular methods for isolation of signal transduction pathway mutants. Methods Cell Biol. 49, 441–454. 19. Lu, Y., Savage, L. J., Ajjawi, I., Imre, K. M., Yoder, D. W., Benning, C., Dellapenna, D., Ohlrogge, J. B., Osteryoung, K. W., Weber, A. P., Wilkerson, C. G., and Last, R. L. (2008) New connections across pathways and cellular processes: industrialized mutant screening reveals novel associations between diverse

­ henotypes in Arabidopsis. Plant Physiol. 146, p 1482–1500. 20. Myouga, F., Akiyama, K., Motohashi, R., Kuromori, T., Ito, T., Iizumi, H., Ryusui, R., Sakurai, T., and Shinozaki, K. The Chloroplast Function Database: a large-scale collection of Arabidopsis Ds/Spm- or T-DNA-tagged homozygous lines for nuclear-encoded chloroplast proteins, and their systematic phenotype analysis. Plant J. 61, 529–542. 21. Jander, G., Baerson, S. R., Hudak, J. A., Gonzalez, K. A., Gruys, K. J., and Last, R. L. (2003) Ethylmethanesulfonate saturation mutagenesis in Arabidopsis to determine frequency of herbicide resistance. Plant Physiol. 131, 139–146. 22. Kim, Y., Schumaker, K. S., and Zhu, J.-K. (2005) EMS mutagenesis of Arabidopsis. In, Arabidopsis Protocols, 2nd edn. (Salinas, J. and Sanchez-Serrano, J. J., eds.), Humana Press, Totowa, NJ, USA, pp. 101–103. 23. Barkan, A., and Goldschmidt-Clermont, M. (2000) Participation of nuclear genes in chloroplast gene expression. Biochimie 82, 559–572.

Chapter 2 Analysis of Plastid Number, Size, and Distribution in Arabidopsis Plants by Light and Fluorescence Microscopy Kevin Pyke Abstract Methods are described which allow one to observe chloroplasts in mesophyll cells from leaves of Arabidopsis, determine their number per cell, measure their area, and determine a value for chloroplast coverage inside mesophyll cells. Non-green plastids can also be imaged either by using staining, or by exploiting fluorescent proteins targeted to the plastid in non-green parts of the plant, such as the roots, in transgenic Arabidopsis. Key words: Arabidopsis, Chloroplast, Non-green plastid, Roots, Fluorescent protein, Protoplasts

1. Introduction Observing chloroplasts and other types of plastids in plant cells is an important aspect of understanding plastid biology and function during plant development. In many respects, chloroplasts are relatively easy to observe since they are pigmented green with chlorophyll and they are relatively large organelles, in the range of 1–5 mm long. Consequently, with a standard light microscope using a ×20 or ×40 objective lens, it is possible to see individual chloroplasts in mesophyll cells in a leaf. However, leaves are complex cellular structures and are covered by a sheet of epidermal cells and thus observing intact leaves microscopically will not yield decent images of chloroplasts. Ripping of leaf tissue will probably yield some observable mesophyll cells, but in general a more specific preparation technique is required in order to view live chloroplasts in cells. The separation of leaf tissue into individual cells greatly improves the observation of chloroplasts within cells and enables information to be obtained about the size of chloroplast populations and R. Paul Jarvis (ed.), Chloroplast Research in Arabidopsis: Methods and Protocols, Volume I, Methods in Molecular Biology, vol. 774, DOI 10.1007/978-1-61779-234-2_2, © Springer Science+Business Media, LLC 2011

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c­ hloroplast sizes in individual cells. Intact leaf mesophyll cells can be separated by fixing them and weakening the middle lamella, which glues cell walls of adjoining cells together, thus enabling separation of individual intact cells. Chloroplasts are only one member of the family of organelles called plastids (1), different types of which reside in different types of cell within the Arabidopsis plant. In Arabidopsis, plastids other than chloroplasts are essentially colourless and lack pigment, thus making them difficult to image in non-green tissues. In order to image them, they need to be stained or tagged with molecules that can be easily visualised by light or fluorescence microscopy. Such techniques have greatly aided the study of non-green plastids in the last 15 years.

2. Materials General laboratory equipment in the form of pipettes, microscope slides, fine forceps, Eppendorf tubes, and a clicker counter are required for these methods, in addition to specific items mentioned below. A suitable light microscope with differential interference contrast (DIC) optics on it, such as a Nikon Optiphot microscope, is required, with a square-gridded graticule in one eyepiece. In order to image cells and chloroplasts, an image capture camera is also required to capture microscope images along with image analysis software, such as ImageJ (http://rsbweb.nih.gov/ij/). 2.1. Analysis of Chloroplast Number and Size in Arabidopsis Leaf Mesophyll Cells

1. 3.5% (v/v) glutaraldehyde in water. Glutaraldehyde normally comes as a concentrated stock solution and so will need diluting with water accordingly.

2.2. Analysis of Chloroplast Number and Size in Arabidopsis Leaf Mesophyll Protoplasts

1. Pre-plasmolysis buffer: 0.65 M sorbitol, 1 mM CaCl2, 5 mM 2-(N-morpholino)ethanesulphonic acid (MES)-KOH, pH 6.

2. 0.1  M sodium ethylenediaminetetraacetic acid (NaEDTA) at pH 9, titrated with NaOH.

2. Digestion medium: 1% (w/v) cellulase Onozuka R-10, 0.4% (w/v) macerozyme R-10, 0.2% (w/v) bovine serum albumin (BSA), 5 mM sodium ascorbate, 0.65 M sorbitol, 1 mM CaCl2, 0.25 mM EDTA, and 5 mM MES-KOH, pH 5.5. 3. Washing solution: 0.65  M sorbitol, 1  mM CaCl2, 0.25  mM EDTA, and 5 mM MES-KOH, pH 6.0. 4. Suspension medium: 0.65 M sorbitol, 1 mM CaCl2, 0.5 mM MgCl2, and 10  mM N-2 hydroxyethylpiperazine-N ¢2-ethanesulphonic acid (HEPES)-KOH, pH 7.0.

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2.3. Imaging and Analysing Non-green Plastids in Leaves and Roots

21

Item 1 below is for use when analysing non-green plastids in leaves (see Subheading 3.3). Items 2–5 below are for use when analysing non-green plastids in roots (see Subheading 3.4). 1. 1% (w/v) silver nitrate in water. 2. A mix of 1 × Murashige–Skoog medium dissolved in water containing 1% (w/v) sucrose and 1% (w/v) agar. Sterilise in an autoclave at 121°C for 20 min and pour into 9-cm standard Petri dishes. 3. 50% (v/v) commercial bleach (containing sodium hypochlorite), diluted in distilled water. 4. 10 mL of water containing a drop of Triton X-100 detergent. 5. Iodine (I2/KI) solution. This should consist of 0.5  g iodine (I2) and 1 g potassium iodide (KI) mixed with 8.5 mL distilled water. Alternatively, one can use Lugol’s solution, which is commercially available (Sigma).

3. Methods 3.1. Analysis of Chloroplast Number and Size in Arabidopsis Leaf Mesophyll Cells

1. Aliquot 1  mL of glutaraldehyde solution into a 1.5-mL Eppendorf tube; do this in a fume hood, since glutaraldehyde vapour is toxic. If several samples are to be processed, then 1 mL of glutaraldehyde in each of several Eppendorf tubes is required. 2. Harvest leaves or pieces of leaf tissue from growing Arabidopsis plants in which chloroplasts are to be analysed, submerge them in the glutaraldehyde solution, and close the lid. Small Arabidopsis leaves up to 0.5 cm2 can be fixed entirely but larger leaves are best cut into strips 1–5 mm wide with a scalpel or razor blade. It is important that all tissues are submerged in the fixative. Place in the dark at room temperature for 1 h. If required, samples can be kept in this glutaraldehyde fixative indefinitely before further processing in which case they are best stored in the dark at 4°C. However, the best images of cells and chloroplasts will be obtained from immediate processing of fresh tissues. 3. Pipette off the glutaraldehyde solution, wash once with water, and replace with the NaEDTA solution. This solution chelates calcium ions from the middle lamella of the cell wall, which joins together adjacent cells, and causes it to weaken, thereby facilitating cell separation. After 1 h, pipette off the glutaraldehyde solution in a fume hood and replace with distilled water. Shake tube gently, pipette off distilled water, and replace with the NaEDTA solution. Make sure that all tissues are submerged in the liquid.

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4. Heat the tube in a heating block at 60°C for 3  h. Let the samples cool and store at 4°C in the dark. Separation of cells on the microscope slide is normally better if samples are left overnight after heating before microscopic examination. Once processed in this way and stored in a fridge, samples are stable for many months, although the chlorophyll will slowly fade (see Note 1). 5. If larger amounts of leaf material are required, then sampling and processing would be more easily carried out in small glass vials, which can be heated in an oven, in a water bath, or on a heating plate. Batch sampling and processing of many individual leaf samples, for instance when screening mutant collections, may be more easily performed in 96-well plates rather than individual tubes. In this case, 96-well plates can be heated on a heating plate and sealed with Nescofilm prior to storage in a fridge. 6. To view chloroplasts in individual cells, remove a piece of tissue from the tube with fine forceps and lay on a microscope slide in a drop of water. If large leaf pieces were harvested, then small pieces can be cut from the processed tissue with a razor blade and the remainder returned to the tube for further storage. Using a blunt instrument, such as the handle end of forceps or scalpel handle, tap and macerate the tissue fairly vigorously. It should be soft and break up easily. If it remains hard and does not disintegrate, then the samples need heating for a longer period and can be reheated at 60°C in order to facilitate tissue break up. Trial and error at this point will be needed in order to get large numbers of individual isolated cells on the slide. Over zealous maceration will, of course, break cells. 7. Examine the macerated tissue by light microscopy, ideally using a ×10 or ×20 objective to start with. The preparations should contain both numbers of individual mesophyll cells, sometimes in small clumps, in addition to which there will be cell debris and larger lumps of tissue, most often associated with the vascular cells and sheets of epidermal cells which do not break apart readily. It should be appreciated that leaves contain several cell types in addition to chloroplast-containing mesophyll cells, including vascular cells and bundle sheath cells associated with the vascular as well as sheets of epidermal cells, containing stomata and trichomes. All of these cell types will be present in the cell preparation, although mesophyll and bundle sheath cells are the only ones that contain large numbers of green chloroplasts (see Note 2). 8. In order to image and count chloroplasts in mesophyll cells effectively, DIC/Nomarski optics are required on the microscope. This greatly facilitates focusing on the top and bottom

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Fig. 1. Analysing chloroplast populations in Arabidopsis leaf mesophyll cells. A clump of fixed separated mesophyll cells from an Arabidopsis leaf. Populations of individual chloroplasts can be seen in each cell. Bar = 20 mm.

surface of cells, enabling the chloroplasts on the upper and lower surfaces of the cell to be clearly observed, as the chloroplasts lying on the vertical edges of the cell. In the threedimensional mesophyll cells, chloroplasts form a monolayer in the cytosol, which is pressed tightly against the cell wall by the vacuole. Thus, an array of individual chloroplasts is seen in each cell, which spreads around the inner surface of the cell wall, covering up to 70% of the cell surface. Mesophyll cells are best observed using a ×40 objective lens. Adjust the DIC on the microscope so that the contrast between the background and the green chloroplasts is maximal and there is a shadow effect on the chloroplasts, making them appear three dimensional (Fig. 1). The author has used DIC optics on a Nikon Optiphot microscope, which is well suited to this method, but DIC optics on other microscopes should also be suitable. 9. Select an individual mesophyll cell and focus up and down through the cell. It should be possible to see all of the chloroplast in the cell, although those that reside on the side walls of the cell will be more difficult to identify as individual organelles. Counts of chloroplasts in individual cells should be possible. With a good eye, the number can simply be counted, probably with the aid of a clicker counter, as one focuses up and down through the cell. Counting, however, is greatly aided

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by using an eyepiece graticule in the microscope, consisting of a square grid, providing a frame of reference such that chloroplasts can be counted in individual squares in the grid, by focusing up and down. In order to avoid counting a chloroplast twice in a single cell, a rule of ignoring chloroplasts that touch the upper side or the right hand side of any graticule square, but counting all others, including those that impinge upon the lower and left hand edges of the graticule square, should help. In this way, a reasonably accurate estimate of the number of chloroplasts in a cell should be possible. This method does need some learning and development of consistency in counting. Mesophyll cells vary in the number of chloroplasts that they contain, normally in a range from 50 to 150 chloroplasts per cell in fully expanded cells. Chloroplasts in younger, expanding cells can also be counted, but smaller cells and smaller chloroplasts make for slightly more difficult counting. The number of chloroplasts in a mesophyll cell is related to the size of the cell. Although cell surface area would probably be the best index against which to relate chloroplast number per cell, the irregular three-dimensional shapes of mesophyll cells make such calculations fraught. Most studies have used the plan area of individual isolated cells as a measure of cell size, since it is easily measured from digital images captured on the microscope. Any standard image analysis package will be able to generate measurements of mesophyll cell plan area from digital images captured from the microscope (2). Whilst it might be possible to threshold individual cells in order to measure their plan area, it is probably easier to draw around the circumference of the cell image with the mouse or computer pen in order to define its edge. Images of cells in which chloroplasts have been counted can be captured and measured at a later date. Relationships between mesophyll cell plan area and chloroplast number per cell have been determined in Arabidopsis leaves (2) and in other plant species, for which this method also works well. Average Arabidopsis mesophyll cells from mature leaves will contain between 50 and 150 chloroplasts per cell. 10. The individual plan area of chloroplasts within cells can also be measured. This is best achieved by drawing around the outline of clearly defined chloroplasts on the upper surface or lower surface of the cell and determining their area using image analysis software. Obviously not all the chloroplasts within a cell can be measured but a sample of 10 or more from an individual cell should be sufficient to give an estimate of mean chloroplast size per cell. 11. There is a trade-off in mesophyll cells between chloroplast number and size; thus, a calculation of “chloroplast index”

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may prove a useful parameter with which to assess chloroplast population coverage of the cell surface of a mesophyll cell. This is calculated as follows: (chloroplast number per cell × mean chloroplast plan area per cell)/mesophyll cell plan area (3). Typical values for the chloroplast index for Arabidopsis mesophyll cells would be around 1.5. 3.2. Analysing Chloroplast Populations in Arabidopsis Leaf Protoplasts

In some situations, such as if very small cells are being observed or the tissue does not break up well, analysing fixed separated cells as described above may not be easy and an alternative approach might be required. One alternative approach is to make protoplasts from leaves or other tissues and observe and analyse the plastids within them. Such an approach makes counting chloroplasts or other plastids relatively straightforward, but such data cannot be related to cell size or shape, since the cell wall has been digested and has lost its defined morphology. However, imaging of mutant plastid morphologies or simple isolation of chloroplasts in order to image them is easily achieved by this method. There are several published protocols for protoplast isolation from leaves; in this instance, the method here is based on (4). 1. Harvest leaf material and if the leaves have prominent mid ribs, cut them out with a razor blade. 2. Peel the lower epidermis off the leaves using fine microdissection forceps. It is not necessary to remove all of the epidermis, but partially peeled leaves enable more efficient access by the digestion enzymes. 3. Cut the peeled leaf into small pieces, up to 1 cm wide, and float on 20 mL of pre-plasmolysis buffer in a Petri dish. Make sure that the peeled leaf surface is in contact with the liquid medium. 4. After 15 min, remove the pre-plasmolysis buffer from the dish and replace with ~20  mL of digestion medium. Place on an orbital shaker at 25°C for 40 min, shaking slowly at 30 shakes per min. 5. Carefully remove the digestion medium from the Petri dish and replace with ~20 mL of washing solution. Tap and swirl the Petri dish gently to release the protoplasts from the tissues. 6. All further operations should be performed at 4°C. Filter the suspension through a nylon filter with 60-mm pore size and centrifuge the filtrate at 100 × g for 3 min. Resuspend the pellet in suspension medium. 7. Prepare a microscope slide of the protoplast preparation using either a welled slide with a cover slip on top or use a normal slide with three square cover slips piled up on either side of a long cover slip, making a bridge with the liquid underneath.

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Fig. 2. Analysing chloroplast populations in Arabidopsis leaf protoplasts. (a) Protoplasts isolated from wild-type Arabidopsis leaves are spherical and contain populations of green chloroplasts within each. Intact protoplasts have a clearly defined edge, which is the plasma membrane. (b) Protoplasts from leaf mesophyll cells of the arc6 mutant of Arabidopsis (9) each contain one or two large green chloroplasts. Protoplasts lacking obvious green chloroplasts are derived from epidermal cells in the leaf. Bar = 40 mm.

This should prevent protoplasts being broken on the slide. Add the protoplast suspension to the slide using a large, 1-mL Gilson pipette tip or a plastic Pasteur pipette to prevent them being broken (see Note 3). 8. Observe the protoplasts as described in Subheading 3.1, step 8 onwards. Chloroplasts should be easily visible in the protoplasts derived from mesophyll cells and can be counted and analysed (Fig. 2) (see Note 4). 3.3. Imaging and Analysing Non-green Plastids in Leaves

Whilst green-pigmented chloroplasts in leaf mesophyll cells are relatively easy to image and analyse, other leaf cells contain plastids that have much lower levels of chlorophyll or none at all, making them difficult to visualise. One way of imaging such plastids is to use fluorescent markers (see Subheading  3.5). Alternatively they can be stained. A method that works well in staining plastids in the epidermal cells of Arabidopsis leaves uses silver nitrate, which is reduced in metabolically active plastids and results in them being stained dark brown. 1. Epidermal tissues to be examined can be left intact on the leaf, in which case small pieces of whole leaf can be mounted on a slide. Alternatively, it might be better to peel epidermal strips from leaves to enhance viewing of the stained plastids. In this case, using a pair of fine forceps, make a small hole in the leaf, and whilst holding it tight around a finger, pick up the edge of the lower epidermis and peel backwards away from you.

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Mature  Arabidopsis leaves should yield reasonable sheets of epidermal cells using this method. 2. Mount the tissue on a slide in the silver nitrate solution, cover with a cover slip, and observe microscopically, as described in Subheading 3.1. The chemical reaction that leads to reduction of silver and staining requires time to proceed, and also light and heat appear to assist this process. Thus, leaving the slide illuminated under the microscope for 15 min may enhance the staining (see Note 5). The chloroplasts in the stomatal guard cells stain darkly as do the diffuse populations of pale green plastids in the epidermal pavement cells (Fig. 3).

Fig. 3. Imaging and analysing plastids in epidermal cells and stomatal guard cells. (a, upper panel ) The staining of epidermal peels from Arabidopsis leaves reveals the distinct chloroplasts in the two guard cells forming a stoma, as well as the plastids in the surrounding epidermal pavement cells. (b, lower panel ) Excitation of chlorophyll in chloroplasts by fluorescence produces significant red fluorescence, here shown in two adjacent stomata, in which the guard cell chloroplasts fluoresce brightly. Bar = 10 mm.

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3.4. Imaging and Analysing Non-green Plastids in Roots

Plastids in roots cells are of major importance to root function but are difficult to visualise since they are small, highly variable in shape, and have no pigmentation. Whilst they can be imaged using fluorescent protein technology (see Subheading  3.5), the specialised plastids in the columella cells of roots, termed statoliths, contain significant quantities of starch and thus can be stained relatively easily and visualised. 1. Sterilise Arabidopsis seeds in 1 mL of 50% bleach solution for 7 min in an Eppendorf tube. 2. Replace the bleach solution with 1 mL of water containing a drop of Triton X-100. Shake and tip off, repeating the process twice. 3. Pour seeds out onto a filter paper in fume hood, wash with 70% ethanol, and leave to dry. 4. Sow seeds on agar plates, place in a suitable growth room, and leave to grow for the desired time, or until reasonable root growth has occurred, normally 7–10 days, if grown at 20°C. 5. Remove whole seedlings carefully from the agar plate and lay the seedling on a microscope slide in water. Cut off the root with a razor blade, remove the upper part of the seedling, cover with a coverslip, and examine the region of columella cells, distal of the meristem, and just behind the root cap cells, at the root tip. 6. Add a few drops of iodine solution to one edge of the coverslip and draw the liquid across the slide using a piece of filter paper placed at the opposite side of the coverslip. The starch in the statoliths in the columella cells will stain blue/black. If staining is not sufficiently intense, add more iodine solution. If the staining becomes too intense, draw water across the slide using a tissue. When the desired level of staining is attained, a filter paper can be used to absorb the remaining iodine solution on the ‘start’ side of the slide and root staining will not change further. This method allows a control of the intensity of starch staining and hence optimisation of the image captured (Fig. 4). In this way, images of distinct statoliths in columella cells in Arabidopsis roots can be obtained and the number of starch granules in each can also be seen. After optimisation of this staining technique, the number of columella cells in Arabidopsis roots, the number of statoliths in each, and the number of starch grains in each statolith can be determined.

3.5. Imaging Plastids with Fluorescent Markers Using Fluorescent and Confocal Microscopy

Whilst pigmented green chloroplasts are relatively easily imaged by light microscopy, the non-green plastids which lack pigment present a more difficult problem and this has hampered an understanding of their cell biology. In the last 15 years, however, the use of fluorescent proteins inside plastids has revolutionised the ability to

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Fig. 4. Imaging and analysing non-green plastids in roots. (a) An Arabidopsis root stained for starch reveals amyloplasts in the layers of columella cells at the root tip. (b) Higher magnification of the columella cells reveals distinct populations of amyloplasts within each columella cell. Darker stained patches within each amyloplast are individual grains of starch. (a) Bar = 40 mm. (b) Bar = 20 mm.

image them and learn more about their cell biology. Outstanding results have been obtained by genetically targeting fluorescent ­proteins to the plastid compartment in transgenic Arabidopsis using a variety of different plastid transit peptide sequences fused upstream of green fluorescent protein (GFP) or one of its coloured ­derivatives. The major advantage of such an approach is that if a powerful ­ectopic promoter is used to drive expression of the transgene, then, in theory, all plastids within the plant will contain fluorescent ­protein and can be imaged by fluorescence microscopy.

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Although conventional fluorescence microscopes can be used for such imaging, they suffer with problems of refraction and reflection of fluorescent light within tissues and of fluorescence bleaching after excitation. The method of choice now is to use a confocal laser scanning microscope, which produces high-quality fluorescent images, with the facility for optical sectioning of three-­ dimensional structures. Such an approach makes possible an effective comparison of different plastids types in different cell populations in the same transgenic Arabidopsis plant. In particular, such plants provide excellent material in which to image non-green plastids. However, imaging fluorescent proteins in normal green chloroplasts never appears to work so well due to the bright levels of fluorescence from the native chlorophyll. In some circumstances, chlorophyll fluorescence itself can be used as a useful marker, since plastids containing chlorophyll will fluoresce bright red (Fig. 3b). If one chooses to use transgenic plants containing plastid-­ targeted fluorescent protein in order to image plastids, then there are several ways in which to proceed. One can generate such transgenic material oneself using suitable vectors and an Arabidopsis transformation protocol (5). Alternatively, one could request seed of such lines from other laboratories in which the relevant transgenic Arabidopsis has been made (6–8) or from Arabidopsis stock centres. Indeed there is a particularly useful Arabidopsis line available from the ABRC stock centre (www.arabidopsis.org) in which plastids and several other organelles are genetically targeted with different coloured fluorescent proteins, enabling several different organelles to be imaged simultaneously in the same cell (­germplasm/ stock: CS16303). In this line, plastids are targeted with red fluorescent protein. 1. Sterilise seed of transgenic Arabidopsis containing a transgene encoding plastid-targeted fluorescent protein and plate on agar (as described in Subheading  3.4). Agar-grown Arabidopsis seedlings are a much more convenient source of plant material than those grown in compost, particularly if roots are to be imaged. 2. Place in a suitable growth room and when they have grown to a suitable size, remove a whole seedling carefully from the plate and mount in a drop of water on a slide and cover with a coverslip. 3. The exact protocol now depends somewhat on which type of confocal microscope is to be used. In general, however, to image GFP and chlorophyll in separate channels, one needs to use an excitation of 488  nm from an argon laser and collect emission signals between 495 and 526 nm, and between 631 and 729 nm for GFP and chlorophyll respectively. It is useful to false colour the two channels green and red respectively. Optical sections are normally collected at 1-mm intervals and

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Fig. 5. Imaging plastids with fluorescent markers. Plastids in Arabidopsis roots imaged by confocal microscopy, using fluorescence of a green fluorescent protein targeted to the plastid transgenically. Bar = 10 mm.

maximum projection images compiled using the confocal ­software. In this way, quality images of non-green plastids in non-green tissues such as roots, trichomes, and other tissues can be obtained (Fig.  5). Using such images, it should be ­possible to count the number of chloroplasts in a cell. In theory­, it would be easy to measure sizes of plastids from the digital confocal image by importing into suitable image analysis software, although one should be aware that the morphology of non-green plastids is often highly irregular and dynamic (see Chapter 5, Vol. 1), thus such measurements should be interpreted with care.

4. Notes 1. It is very important that samples are not left on the bench in bright sunlight for any length of time, since the chlorophyll will fade quickly and imaging of the chloroplasts will be poor. Samples should be stored in the fridge in the dark at all times. 2. In theory, this method can be used to separate and observe plastids in any tissues from the Arabidopsis plant, especially green tissues other than leaves. In practice, however, other tissues tend to have less air spaces between cells and hence the

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tissue is more densely packed, which makes it more difficult to get the fixed, heated tissue to disassociate into groups of observable cells. In some cases, such as cotyledons, the method works well, but for densely packed tissues such as roots, it will be more difficult to get effective separation. 3. Protoplasts are extremely delicate and are easily broken. Sucking them through a narrow orifice such as a yellow Gilson tip will break them, as will rough application of a coverslip on a slide. Use a blue 1-mL Gilson tip and be gentle with them at all times. 4. The protoplast preparation so made will consist of the entire population of cells from within a leaf. Although the majority will be mesophyll cells containing large populations of green chloroplasts, there will also be some protoplasts derived from epidermal cells, bundle sheath cells, and vascular cells, which will contain different types of plastids. This should be taken into account when analysing these protoplast populations. This method could be applied to making protoplasts in other Arabidopsis tissues, but digestion times may need to be increased for effective enzyme penetration into the tissue. 5. From experience, this method works better in young, fastgrowing tissues, such as cotyledons and young leaves, in which the plastids are metabolically active. Plastids in guard cells and epidermal cells can be stained and imaged effectively using this method (Fig. 3a) (9). References 1. Pyke, K. A. (2009) Plastid Biology, 1st edn. Cambridge University Press, Cambridge, UK. 2. Pyke, K. A., and Leech, R. M. (1991) A rapid image analysis screening procedure for identifying chloroplast number mutants in mesophyll cells of Arabidopsis thaliana (L.) Heynh. Plant Physiol. 96, 1193–1195. 3. Pyke, K. A., and Leech, R. M. (1987) The control of chloroplast number in wheat mesophyll cells. Planta 170, 416–420. 4. Riazunnisa, K., Padmavathi, L., Scheibe, R., and Raghavendra, A. (2007) Preparation of Arabidopsis mesophyll protoplasts with high rates of photosynthesis. Physiol. Plant. 129, 879–886. 5. Clough, S. J., and Bent, A. F. (1998) Floral dip: a simplified method for Agrobacterium-mediated transformation of Arabidopsis ­thaliana. Plant J. 16, 735–743.

6. Tirlapur, U. K., Dahse, I., Reiss, B., Meurer, J., and Oelmuller, R. (1999) Characterization of the activity of a plastid-targeted green fluorescent protein in Arabidopsis. Eur. J. Cell Biol. 78, 233–240. 7. Niwa, Y., Hirano, T., Yoshimoto, K., Shimizu, M., and Kobayashi, H. (1999) Non-invasive quantitative detection and applications of nontoxic, S65T-type green fluorescent protein in living plants. Plant J. 18, 455–463. 8. Kojo, K. H., Fujiwara, M. T., and Itoh, R. D. (2009) Involvement of AtMinE1 in plastid morphogenesis in various tissues of Arabidopsis thaliana. Biosci. Biotechnol. Biochem. 73, 2632–2639. 9. Robertson, E. J., Pyke, K. A., and Leech, R. M. (1995) arc6, a radical chloroplast division mutant of Arabidopsis also alters proplastid proliferation and morphology in shoot and root apices. J. Cell Sci. 108, 2937–2944.

Chapter 3 Immunofluorescence Microscopy for Localization of Arabidopsis Chloroplast Proteins Stanislav Vitha and Katherine W. Osteryoung Abstract Immunofluorescence microscopy reveals localization of proteins in cells and tissues by means of highly specific, fluorescently labeled antibodies. This technique is an important complement to localization methods that use genetically encoded fluorescent tags. This chapter describes the five stages of immunofluorescence localization of proteins in plant chloroplasts in sectioned leaf tissue: (1) fixation, (2) tissue embedding and sectioning, (3) treatment of sections prior to immunolabeling, (4) immunostaining, and (5) fluorescence microscopy and image capture. Protocols for both cryosectioning and sectioning of lowmelting-point wax-embedded samples are described. Immunofluorescence localization in chloroplasts is complicated by their intense autofluorescence background. Measures to suppress nonspecific background staining, confirm specificity of the fluorescence signal, and optimize imaging conditions are described. Key words: Aldehyde fixation, Autofluorescence, Antigen retrieval, Embedding, Cryosectioning, Anti-fade mounting medium

1. Introduction Subcellular localization of chloroplast proteins at the light microscopy level is commonly achieved using either genetically encoded fluorescent tags, such as green fluorescent protein (GFP), or using antibodies conjugated to fluorescent markers. Fluorescent ­proteins are invaluable for in vivo studies, especially of dynamic ­processes, and eliminate the need for fixation and extensive tissue processing. However, the presence of the fluorescent tag may alter the function or localization of the protein under study. Furthermore, fluorescent proteins may be more sensitive to pH than the synthetic fluorescent dyes, and less resistant to photobleaching (1). Another important consideration is that expression of tagged proteins may

R. Paul Jarvis (ed.), Chloroplast Research in Arabidopsis: Methods and Protocols, Volume I, Methods in Molecular Biology, vol. 774, DOI 10.1007/978-1-61779-234-2_3, © Springer Science+Business Media, LLC 2011

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be accompanied by overexpression of the protein under study, leading to localization artifacts. Immunolocalization of the ­endogenous protein thus remains an essential tool in cell biology and should be considered complementary to the use of genetically encoded fluorescent tags and biochemical methods based on cell fractionation (see Chapters 20 and 21, Vol. 1 and Chapters 10 and 11, Vol. 2) for defining the localization of chloroplast proteins. The immunofluorescence localization protocols described in this chapter are for use with sectioned tissue. While whole-mount immunofluorescence labeling is possible with some samples, such as embryos, root tips, hypocotyls, or entire young seedlings (2), the penetration of immunoreagents is severely hindered by the cell walls and hence permeabilization procedures must be used, which have the potential to extract some tissue components and compromise structural preservation of the tissue. Immunolocalization can also be performed on isolated intact chloroplasts, but that does not allow observation of the localization pattern within the context of the cell or tissue type. Additionally, the lengthy chloroplast isolation procedures may lead to redistribution or mislocalization of the protein of interest. For example, the isolation procedure used in our quantitative immunoblotting analysis of the Arabidopsis chloroplast division proteins FtsZ1 and FtsZ2 (3), which localize to a mid-plastid ring in fixed leaf tissue (4), resulted in loss of intact FtsZ rings (unpublished). Most plant tissues will need to be sectioned to facilitate effective immunostaining and fluorescence microscopy. This chapter presents two alternative methods for obtaining leaf sections: sectioning of frozen tissue and sectioning of waxembedded samples. The entire immunofluorescence localization protocol is divided into five stages: 1. Fixation 2. Tissue embedding and sectioning 3. Treatment of sections prior to immunolabeling 4. Immunostaining 5. Fluorescence microscopy and image capture These methods are not specific for Arabidopsis and have been directly applied to other plant species, including Pisum sativum, Zea mays, and Nicotiana tabacum (4) (unpublished observation). The principles and considerations for each stage are explained in the following subsections. 1.1. Fixation

The aim of fixation is to immobilize proteins in the same state and location as they were in the living sample at the instant of fixation, and minimize degradation of the sample during processing. Insufficient fixation may lead to localization artifacts, where the

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proteins of interest are degraded or diffuse away from their in vivo location. Different antigens may require different fixation for best results and thus finding the optimal fixative and fixation conditions through trial and error may be necessary. Two common fixatives are presented: a coagulating and weakly cross-linking fixative, FAA (formaldehyde–acetic acid–alcohol), and a strongly cross-linking fixative containing formaldehyde and a low concentration of glutaraldehyde, as well as dimethyl sulfoxide (DMSO) to aid penetration. Formaldehyde is the common cross-linker for immunofluorescence at the light microscope level. It penetrates the tissue quickly but acts very slowly, especially at low pH (5), requiring 4–16  h for complete fixation, and the cross-linking is somewhat reversible (6). Glutaraldehyde is a stronger cross-linker, but penetrates slowly and at higher concentrations causes tissue autofluorescence, which complicates immunofluorescence microscopy. Measures to quench the aldehyde-induced autofluorescence are given in Methods (Subheading 3.4). In recent years, microwaveassisted aldehyde fixation and immunolabeling protocols have gained popularity because of the improved structural preservation and speed of processing (7). However, we have observed that samples subjected to microwave-assisted fixation display higher autofluorescence background in chloroplasts, perhaps due to improved cross-linking and retention of some chloroplast pigments in the tissue. This is likely to be remedied by further optimization of the procedure, but for the protocol presented here, microwave irradiation is not used. 1.2. Tissue Embedding and Sectioning

Following fixation, the tissue must be embedded in a medium that will support the tissue for sectioning. For cryosectioning, the tissue may be infiltrated with a cryoprotectant, such as sucrose, to minimize ice crystal formation, and then frozen and sectioned using a cryostat (8–10). Such sections generally exhibit very good preservation of antigenicity, but frozen plant tissue is difficult to section and the resulting sections are often damaged. Wax embedding permits convenient sectioning with good structural preservation of plant tissues. The classical paraffin wax embedding medium requires relatively high temperatures for embedding (~60°C), which causes reduced or sometimes complete loss of antigenicity. Our preferred embedding medium is the Steedman’s wax, a mixture of polyethylene glycol distearate and hexadecanol (11). Steedman’s wax is hydrophilic, water-miscible, and soluble in ethanol, and has a low melting point of 37°C. This embedding medium has proven successful for immunodetection of numerous proteins in plants (12–15), including the chloroplast division proteins FtsZ1 and FtsZ2 (4), and in our hands, wax sectioning is much less time consuming than cryosectioning.

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1.3. Treatment of Sections Prior to Immunolabeling 1.3.1. D  ewaxing

Removal of the embedding medium ensures penetration of the antibodies and accessibility of the epitopes. Dewaxing of sections from Steedman’s wax-embedded samples is done in ethanol. The 95% standard grade ethanol is sufficient for complete dewaxing. Dewaxed sections are then gradually rehydrated and brought to phosphate-buffered saline (PBS).

1.3.2. Antigen Retrieval

It is well established that aldehyde fixation can result in a decrease or loss of reactivity for specific antibodies. This is caused by crosslinking of the target epitopes with nearby proteins, which then sterically block antibodies from binding (6). Chemical modification by dehydration and wax embedding may also contribute to epitope alteration and inhibit antibody binding (16). In such cases, antigen retrieval procedures can be employed on the tissue sections before immunostaining. Antigen retrieval involves breaking of ­protein cross-links introduced by chemical fixation and exposing antigen sites using chemical or physical means, such as proteinase digestion, denaturing agents such as sodium dodecyl sulfate and urea, or heating (17). Such treatments often enhance accessibility of the antibody to the antigen.

1.4. Immunolabeling

The specificity of a new primary antibody should be first confirmed using immunoblotting. A clean immunoblot with a specific band of expected molecular mass is often a good predictor of successful immunolabeling in situ. Highly specific antibodies, preferably affinity-purified, with minimal cross-reactivity to nontarget tissue components are extremely important for successful immunofluorescence labeling. Procedures for improving antibody specificity include blocking of nonspecific binding sites in the tissue prior to incubation with the antibody, optimization of the antibody concentration, and pre-absorption of the antibody with tissue from plants lacking the target epitope (such as a null mutant plant) (18–20). Using proper controls during immunolabeling of tissue sections is essential for correct interpretation of results. The following controls should be used to demonstrate the specificity of labeling: 1. No secondary antibody: indicates autofluorescence of the tissue. 2. No primary antibody: reveals nonspecific binding of the fluorescently conjugated secondary antibody. 3. Use of tissue lacking the protein of interest. Ideally, sections from a null mutant plant at the same developmental stage and grown under the same conditions as the test sample should be used. This control shows the specificity of the primary antibody and reveals its nonspecific binding to tissue components. If such a mutant is not available, additional independent confirmation, such as by analysis of a GFP fusion protein,

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is  recommended; antigen competition, where the primary antibody is pre-absorbed with a purified antigen, is a less useful approach (2). For convenient dual labeling, the primary antibodies should be raised in different species and the secondary antibody conjugates should have minimal cross-reactivity. The multiple labeling procedure should first be performed sequentially, i.e., neither the primary nor secondary antibodies should be mixed together, and thorough washing should be performed after each incubation. Controls for double labeling should include reverse order of labeling and omission of one of the primary antibodies to check for cross-reactivity of the reagents. Secondary antibody conjugates with traditional dyes such as fluorescein and rhodamine are outdated; these dyes have been ­surpassed by newer, much brighter and more ­photostable dyes. The cyanine dyes, especially Cy3 and Cy5 (Jackson Immuno­ research), remain a good choice for many applications. The list of modern fluorochromes includes Alexa Fluor (Invitrogen), DyLight (available, e.g., from KPL or Jackson Immunoresearch), Atto (Atto Tec), and Chromeo (Active Motif Chromeon). Finally, conjugates with quantum dots provide extreme photostability and may be advantageous in highly autofluorescent samples, where the autofluorescence can be bleached by prolonged UV irradiation without affecting the quantum dot signal (21). For multiple labeling, the fluorescence emission peaks of the fluorochromes should be well separated. For instance, dual labeling with Alexa Fluor 488 and Cy3 conjugates is not optimal because of their significant spectral overlap. Alexa Fluor 488 and Alexa Fluor 594, or Cy3 and Cy5, would be a much better choice. The light source of the microscope also needs to be considered when selecting fluorescent conjugates. If a confocal microscope with fixed laser wavelengths is going to be used, a conjugate with an absorption peak near the laser wavelength should be selected, for instance Alexa Fluor 633 or DyLight 647 for excitation with a 633-nm laser. The medium in which the sections are mounted after immunolabeling should preserve the fluorescence signal during microscopy and storage, and also provide an optimal optical environment for high-resolution microscopy. Some mounting media are incompatible with certain fluorescent dyes or with quantum dots, causing quenching or photobleaching. For instance, mountants containing p-phenylenediamine as an anti-fade reagent are not compatible with some cyanine dyes, especially Cy2 (22). Read product data sheets for potential incompatibilities. For best resolution, the refractive index of the mounting medium should be identical to the refractive index of the immersion liquid for the microscope objective. If this condition is not met, the resulting spherical

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aberration will degrade the performance of the microscope optics. Samples mounted in a buffer are best imaged with water immersion objectives. Hardening mounting media, such as Mowiol 4–88 or Prolong Gold (Invitrogen), eliminate the need to seal the cover glass and their refractive index after hardening is claimed to be close to that of oil and thus suited for imaging with oil immersion optics. Many non-hardening mounting media are glycerol based and should ideally be used with glycerol immersion objectives (23), but the refractive index mismatch-induced spherical aberrations are tolerable to some extent with oil immersion objectives (24). 2,2-Thiodiethanol (TDE) is a new mounting medium for highresolution microscopy (25). When used at 97% (v/v) (i.e., contains 3% water or buffer), its refractive index is 1.515, ideal for imaging with oil immersion objectives, but the mounting procedure is more labor intensive (see Note 1). For routine microscopy, the glycerolbased mounting media offer a good compromise between the convenience of use and optical properties. 1.5. Microscopy and Image Capture

Immunofluorescently labeled sections are viewed via a standard (wide-field) fluorescence microscope or by confocal fluorescence microscopy. Confocal microscopy offers optical sectioning and 3D reconstruction, but very weak signals may be difficult to detect. Relevant aspects of confocal and wide-field microscopy are explained in depth in the Handbook by Pawley (26). It is important to note that on microscopes equipped with differential interference contrast (a.k.a. Nomarski) optics, the Nomarski prism at the back focal plane of the objective degrades resolution of the fluorescent image, resulting in a less sharp or even doubled image. Therefore, this prism should be removed from the optical path for fluorescence imaging, and if a DIC image is desired, it should be acquired after the fluorescence images. Unfortunately, even microscopes marketed as fully motorized usually require manual removal of the DIC prism. In order to capture the image at the full resolution that the microscope optics provides, the smallest resolvable feature in the image should be represented at least by two pixels. This requirement is known as the Nyquist criterion (26, 27). The pixel size is freely adjustable by the amount of confocal zoom in point-­scanning confocal microscopes, but is usually fixed in wide-field and ­spinning-disk confocal microscopes, where it is determined by the magnification of the optics and the physical dimension of photosensing sites on the charge-coupled device (CCD) camera. In this respect, objectives with lower magnification and a relatively high resolution (high numerical aperture) pose a challenge for image capture on a CCD camera. However, a 100× oil immersion ­objective of 1.4 numerical aperture and a standard interline CCD camera with 6.45 mm pixels allow image recording at full optical resolution (see Note 2).

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When acquiring three-dimensional data (Z-stacks), the microscope should be equipped with motorized focus, and the Nyquist criterion should also be satisfied in the Z-dimension. Thus, the focal step between adjacent sections in the stack should be less than half of the axial resolution of the objective. If the Z-step is too large, information is lost, while a Z-step too small increases ­acquisition time and photobleaching of the sample without improving­the resolution. A convenient online tool for calculating the sampling step is the Nyquist calculator (http://support.svi.nl/ wiki/NyquistCalculator). Most acquisition software also has the ability to suggest the Z-step size. Image processing and image adjustment must be performed in accordance with the accepted ethical standards, and all image manipulations must be disclosed (28, 29). Raw data (original images) should be stored on media that cannot easily be altered or erased intentionally or unintentionally, such as CD or DVD.

2. Materials 2.1. Fixation

1. PBS 10× stock: 1.4 M NaCl, 27 mM KCl, 65 mM Na2HPO4, 15  mM KH2PO4, and 3.0  mM NaN3. Store at room temperature. Caution should be used to avoid exposure to NaN3. Prepare a 1× PBS working dilution as needed from the 10× stock. 2. Formaldehyde freshly prepared from paraformaldehyde, 30% (w/v): 3 g paraformaldehyde in a 15-mL conical tube is suspended in water to a final volume of 10 mL and heated in a water bath to ~60°C. Pellets of KOH are added and dissolved one by one, mixing by inverting the tube, until the solution clears and all paraformaldehyde is dissolved. Formaldehyde is used for preparation of the fixatives on the same day. Work in a chemical hood, avoid skin contact and inhalation. 3. (a) FAA fixative: 3% (w/v) formaldehyde, 5% (v/v) acetic acid, and 50% (v/v) ethanol. Work in a chemical hood, and avoid exposure. (b) Formaldehyde–glutaraldehyde (FG) fixative: 3% (w/v) formaldehyde, 0.1% (v/v) electron microscopy grade glutaraldehyde, and 1% (v/v) DMSO in PBS. Work in a chemical hood, and avoid skin contact and inhalation. 4. PBS with Tween-20 (PBST): 1 × PBS supplemented with 0.05% (v/v) Tween-20. Thoroughly mix; prepare on the day of use. 5. Dissecting and handling tools: fine-tip forceps and sharp razor blades.

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6. Rotator or a rocking platform for gentle agitation of samples. 7. Small glass containers (scintillation vials) and glass Pasteur pipettes (see Note 3). 2.2. Tissue Embedding and Sectioning

2.2.1. Infiltration with Sucrose, Freezing, and Cryosectioning

Supplies such as embedding molds and microtome knives can be obtained from vendors specializing in sample preparation for microscopy, e.g., Electron Microscopy Sciences (http://www. emsdiasum.com) or Ted Pella (http://www.tedpella.com). 1. Sucrose/PBS solution of increasing concentration: 8, 15, 30, and 60% (w/v) sucrose in PBS. 2. Cryostat (cryo-microtome) with specimen chucks and with a sharp disposable blade or a steel knife. 3. Poly-l-lysine-coated slides. These can be either purchased or prepared in the laboratory as follows. Plain, non-frosted glass slides (see Note 4) are thoroughly cleaned in water with detergent, rinsed, and dried. The slides are then submerged in 0.01% poly-l-lysine (e.g., Sigma) for 15 min in plastic staining jars, dried, and heated at ~55°C for 1  h. For best adhesion, only slides less than several weeks old should be used (see Note 5). 4. Silicone rubber embedding molds with 6-mm-deep cavities. 5. Tissue-freezing medium. 6. Fine-tip brush.

2.2.2. Steedman’s Wax Embedding and Sectioning

1. Steedman’s wax. This can be purchased pre-mixed (“Polyester Wax,” catalog number 19312, Electron Microscopy Sciences) or prepared in advance by melting 900 g polyethyelene glycol distearate (Aldrich) and 100 g 1-hexadecanol (Aldrich) at 65°C and stirring very thoroughly. The prepared wax is poured into 50-mL conical tubes and stored at room temperature for later use. 2. Ethanol, 95% (w/v). 3. Toluidine blue solution in ethanol: Toluidine blue O certified stain, approximately 0.1% (w/v) in ethanol. Stir well and then allow the non-dissolved residues to settle for several minutes. Use the supernatant for counterstaining of samples being embedded. The exact concentration of the dye is not critical. 4. Incubator set to 37°C. 5. Glass and plastic transfer pipettes (Pasteur pipettes) (see Note 3). 6. Silicone rubber embedding molds with 6-mm-deep cavities. 7. Two fine-point artist’s brushes. 8. Razor blades. 9. Rotary microtome with either a sharp steel knife or a disposable blade.

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10. Poly-l-lysine-coated slides (see Subheading 2.2.1). 11. Wooden stubs (blocks), approximately 15 × 25 × 25  mm (W × D × H) for mounting the wax blocks. Check that the stubs fit in the microtome specimen holder. 12. A gas or ethanol burner, or a hot plate. 13. Diamond scribe for marking the plain glass slides, or a pencil for marking frosted slides. 2.3. Treatment of Sections Prior to Immunolabeling (Dewaxing, Antigen Retrieval)

1. Glass slide staining jars (Coplin jars).

2.4. Immuno­ fluorescence Labeling

1. Sodium borohydride solution: 0.1% (w/v) NaBH4 in 1% (w/v) Na2HPO4. Prepare immediately before use. Sodium borohydride is very corrosive and a hazardous skin irritant. Use caution when handling the powder and the prepared solution.

2. Plastic (polypropylene) staining jars. 3. 95% Ethanol. 4. Tris–HCl buffer, 100 mM, pH >9.5. 5. Autoclave.

2. Blocking buffer: 2% (w/v) nonfat dry milk in PBST; this should be prepared and stirred for 1–2 h before use (see Note 6). 3. Parafilm, scissors, and fine-tip forceps. 4. Humid chamber, e.g., a large Petri dish lined with slightly wet filter paper. 5. Primary antibody and secondary antibodies, diluted in the blocking buffer before use, or, if necessary, pre-absorbed overnight with plant powder (see Note 7). 6. Toluidine blue O solution: Toluidine blue O certified stain, 0.01% (w/v) in PBS, filtered through a 0.2-mm syringe filter before use. 7. Mounting medium. Commercially available anti-fade mountants, such as ProLong, Slowfade (Invitrogen), and Vecta­ shield (Vector Labs), or prepare your own mountant with p-phenylenediamine (PPD) as follows. Place a small stir bar in a scintillation vial wrapped in aluminum foil (PPD is light sensitive), add 50 mg PPD and 5 mL of PBS, and stir until dissolved. PPD is toxic; therefore, wear gloves and avoid inhalation of the powder. Adjust the pH to 8.0 with carbonate/bicarbonate buffer (pH 9.2, prepared by mixing 4 mL of 0.2 M Na2CO3 and 46 mL of 0.2 M NaHCO3). Because of the small volume of the solution, use indicator paper to check the pH. The solution should be almost colorless or with a slight tint of pink (if it is of intense color, PPD is contaminated and should be discarded). Add the PPD solution to 45  mL of glycerol, and stir thoroughly. Aliquot to 1.5-mL microcentrifuge tubes and store protected from light at −20

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or −80°C. Do not use if the stored mounting medium turns brown. PPD anti-fade reagent may not be compatible with some cyanine dyes (22). 8. Cover glass #1.5, 22 × 40 mm (see Note 8). 9. Nail polish, clear or colored. 10. Slide storage trays or boxes. 2.5. Fluorescence Microscopy, Image Capture, and Processing

1. Wide-field or confocal fluorescence microscope equipped with high-resolution optics and acquisition software (see Note 9). In filter-based systems, the emission filters should be of bandpass type matched to the dyes used to facilitate specific detection and multicolor imaging. 2. Image processing and analysis software, such as the freeware ImageJ (http://rsb.info.nih.gov/ij/) with the bio-formats library for reading and writing life sciences image file formats (LOCI plugin, http://www.loci.wisc.edu/software/bio-formats) or commercial programs, e.g., Image Pro, Metamorph, SimplePCI, and SlideBook. 3. Software for general image editing, such as Photoshop (Adobe) or Gimp (freely distributed at www.gimp.org), and for assembling and annotating multi-panel figures, such as Canvas, CorelDraw, Illustrator, or Scribus (freely distributed at http:// www.scribus.net/) (see Note 10).

3. Methods Glass scintillation vials can be conveniently used for holding ­samples during fixation and subsequent steps. The volume of the solution should be in 10- to 100-fold excess relative to the volume of the tissue. Typically, 5–10  mL of liquid is sufficient for 10–20 leaf segments. 3.1. Fixation

Leaf tissue contains large amounts of air. Vacuum infiltration helps replace the air with the fixative and is essential for efficient fixation. The fixed tissue should not be stored in the fixative or in the buffer, since this would cause either over-fixation or reversal of fixation, respectively. 1. Cut a small piece of leaf tissue (less than 5 × 5 mm) with a sharp razor blade, and immediately immerse in the fixative. Samples should be collected within 5–10 min before proceeding to the next step. 2. Vacuum infiltrate the fixative by repeated vacuum cycles in a vacuum desiccator or using a 10-mL plastic syringe (see Note 11).

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3. Incubate at room temperature for 1.5 h. 4. Decant or aspirate the liquid with a glass Pasteur pipette (see Note 3). Wash FAA-fixed samples in 50% (v/v) ethanol in PBS. If the FG fixative was used, wash with PBS. Perform a total of three washes, for 20 min each, with occasional stirring. Samples should not be stored at this point; proceed to Subheading 3.2.1 or 3.2.2. 3.2. Tissue Embedding and Sectioning 3.2.1. Preparation of Frozen Sections

To improve cryosectioning, the fixed tissue is gradually infiltrated with sucrose as a cryoprotectant and embedded in tissue-freezing medium. The final concentration of sucrose may need to be optimized for best sectioning properties with a particular tissue. Sucrose concentrations between 15 and 60% have been used (9, 30, 31). Tissue fixed with the FAA fixative may not be stable enough, because the weak cross-linking by formaldehyde can be reversed during the lengthy infiltration protocol. Therefore, the use of the FG fixative is preferable. 1. FAA- or FG-fixed samples are transitioned from 50% ethanol to PBS in two steps: 20% ethanol/PBS and then PBS, 15 min each step. 2. Infiltrate the fixed and washed samples with sucrose/PBS solution of increasing concentration: 8, 15, 30, and 60% (w/v) sucrose in PBS, for 6–12 h each step at 4°C on a rotator or with occasional stirring. Perform the 60% step once more. The tissue should sink to the bottom when infiltrated. 3. Proceed to the next step, cryosectioning, or freeze and store the samples at −80°C. The frozen samples can be stored for at least several weeks and thawed on ice before use. 4. Set the cryostat chamber temperature to −25°C and allow the temperature to stabilize (9). 5. Put the specimen in a well of a silicone-embedding mold, removing all of the sucrose solution, and fill the mold cavity with tissue-freezing medium, making sure the surface is a little convex. Place a pre-chilled (−25°C) specimen chuck on top of the cavity (Fig. 1a) and put the assembly in the cryostat chamber to freeze completely. The medium will turn white and solidify and the block will adhere to the chuck. Remove the silicone mold and trim the frozen block to a pyramid shape with a razor blade (Fig. 1b and c). 6. Set the blade angle in the cryostat to approximately 2° above the clearance angle. Clearance angle is the angle of the bevel on the knife, and is usually marked on the cryostat knife holder as “0.” Clamp the specimen chuck in the specimen holder and adjust the tilt of the specimen so that the surface of the frozen block is parallel to the knife edge. Carefully approach the knife with the specimen, using the rapid advance control on the cryostat. Once the knife starts cutting into the specimen,

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Fig. 1. Cryosectioning of leaf segments. (a) Silicone-embedding mold cavities with samples are filled with tissue-freezing medium, and a chilled specimen chuck is inverted on top of each sample. (b) Frozen block on the specimen chuck. (c) The trimmed block mounted in the microtome specimen holder. (d) Sectioning with the use of an antiroll plate. A short ribbon of sections is formed. (e) Sections are lifted off by touching to a warm slide. (f) An alternative method of sectioning, without the antiroll plate. The front of the section is held down by the paintbrush to prevent curling. (g) One section on the blade edge, ready to be lifted off. (h) Microscope slide with several sections attached, ready to be dried and immunostained.

­ osition the antiroll plate over the knife. Cut cryosections of p 10-mm thickness and touch the sections on a warm (room temperature) lysine-coated slide (Fig. 1d and e) (see Note 12). 7. Dry the slides (Fig. 1h) at room temperature for 1 h. The slides may be stored before use at −20°C overnight. 8. Transfer the slides successively to staining jars containing sucrose/PBS of decreasing concentration, 60, 30, 15, and 8% (w/v), incubating for 5  min each step. Wash in PBS twice, 5 min for each step. Proceed immediately to immunolabeling (Subheading 4). 3.2.2. Steedman’s Wax Embedding and Sectioning

All dehydration steps should be carried out at room temperature. Infiltration with wax should be performed at 37°C. 1. Place containers with solidified wax in a 37°C incubator several hours in advance to melt. Melting of the wax can be accelerated by using a 37°C water bath, taking care not to contaminate the wax with water (see Note 13). 2. Dehydrate FG-fixed samples sequentially in ~10  mL of 10%, 25%, and 50% (v/v) ethanol in PBS, and then in 70% and 90% (v/v) ethanol in water, for 30 min at each step with occasional stirring. For samples fixed with FAA fixative, start dehydration

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with the 70% ethanol step, since the samples are already in 50% (v/v) ethanol. 3. Dehydrate in 95% ethanol for 30 min with occasional stirring. Repeat this step two more times. 4. Break point: samples can be left in 95% ethanol overnight. 5. Counterstain with 0.01% (w/v) solution of toluidine blue in ethanol for 30 min (see Note 14). 6. Rinse briefly with 95% (v/v) ethanol two to three times to remove excess stain. 7. Transfer the vials with specimens to a 37°C incubator and allow them to warm up for 30 min. All subsequent wax infiltration steps should be performed at ~37°C. 8. Remove most liquid from the sample, leaving approximately 5 mL. Add 1/3 volume (~1.5 mL) of wax and then swirl well to mix the wax and ethanol completely. Incubate for 1–2 h on a rotator or with occasional stirring. 9. Repeat step 8 once. 10. Remove all wax/ethanol mix and replace with ~5 mL of wax. Mix well and incubate for 2 h. 11. Repeat step 10 two more times. 12. Break point: Samples can be left in one of the pure wax changes overnight. 13. Pre-warm a heat block or a slide warmer to ~40°C. Pre-warm the silicone-embedding molds. 14. Pour the samples into a wide container, such as a plastic or aluminum weigh boat, and transfer each piece of tissue into a separate well of the embedding mold. Fill the wells with molten wax to form a slightly convex surface. Adjust the position and orientation of the tissue in the wells and allow the wax to harden on the benchtop overnight (Fig. 2a) (see Note 13). 15. Push the wax blocks out of the silicone mold and store in an air-conditioned room or in the refrigerator until needed for sectioning. Samples are stable for at least several months (see Note 15). 16. Mount the wax blocks on wooden stubs. The top of a new stub should be first soaked with molten wax. Warm a razor blade briefly over a flame or on a hot plate and lay it on top of the wooden block. Place the wax block on the blade and as its base starts to melt, slide it off the blade on the wood (Fig. 2b). Do not use too much heat or the whole wax block will melt. Allow the wax to harden at room temperature for about 30 min and then trim the sides of the wax block around the specimen with a razor blade, creating a pyramidal shape with straight edges (Fig.  2c). The top of the wax block should not be touched with one’s fingers because it would start melting.

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Fig. 2. Sectioning of wax-embedded samples and handling of the slides. (a) Silicone-embedding mold with solidified wax blocks. (b) Mounting the wax block on a wooden stub for sectioning. (c) Mounted and trimmed block. (d, e) Sectioning on a rotary microtome. The end of the ribbon is held by a paintbrush during sectioning. (f) The ribbon lying on a sheet of paper is being cut to smaller segments, lifted with the razor blade (g), and transferred on a slide (h). (i) A drop of water is added to stretch the sections. (j) Slides in a staining jar are arranged in a zig-zag manner to fit eight slides. The sections are facing toward the bottom of the image. (k) Slides are transferred between the staining jars using forceps, and excess liquid is wicked off on a paper towel (l). (m) 100 mL of antibody solution is applied on the sections and the area with sections is covered with a piece of Parafilm (n). (o) The finished slide; the coverslip is sealed with nail polish.

17. Sectioning is performed in a cool room. Ambient temperatures around 20°C permit sectioning at 5–15 mm. Thinner sections may be obtained by sectioning in a cold room (4°C) or in a cryostat set to 10°C. 18. Install a sharp steel knife or a new disposable blade in the microtome. Clamp the block into the specimen holder of a rotary microtome. Align the face of the wax block with the

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knife edge. Hold a paint brush ready in one hand and start cranking the microtome. As soon as the wax block starts cutting and a ribbon of sections begins to form, pick up the end of the ribbon with the brush and lift the ribbon so that it does not slide and stick to the knife surface (Fig. 2d). Ribbons over 30 cm long can be routinely cut this way (Fig. 2e). 19. Stop cutting and use a second paint brush to detach the ribbon from the knife edge. Transfer the ribbon on a clean sheet of paper. Cut the ribbon into shorter pieces with a razor blade (Fig. 2f) and place them on a glass slide (Fig. 2g and h) (see Note 16). 20. Holding the slide slightly tilted, add a drop of room temperature water to the upper end of the ribbons (Fig. 2i). The ribbons start to expand immediately. After the water reaches the bottom of the ribbon, blot the fluid thoroughly with a piece of filter paper. Let the slides dry at room temperature for several hours or overnight. The slides can then be used for immuno­ staining, or stored at 4°C for a day or two for later use. 3.3. Treatment of Sections Prior to Immunolabeling 3.3.1. Dewaxing of Wax-Embedded Sections

The wax needs to be removed from sections and the sections must be rehydrated prior to immunostaining. The hydrated sections may be then subjected to the optional antigen retrieval step if necessary (steps 4 and 5), in order to restore immunoreactivity lost during fixation and wax embedding. The sections should not dry at any point during the procedure. 1. Dewax the slides 3 × 10 min in a Coplin staining jar containing 95% ethanol. The staining jar can hold up to eight slides if they are arranged in a zig-zag manner (Fig. 2j–l) (see Note 17). 2. Rehydrate the slides in three steps, incubating in 90% (v/v) ethanol/water, 50% (v/v) ethanol/PBS, and PBS, for 5 min at each step. 3. If antigen retrieval is necessary, proceed to next section (Subheading  3.3.2); if not, proceed to immunolabeling (Subheading 3.4).

3.3.2. Antigen Retrieval (If Necessary)

1. Transfer the slides to a plastic (polyethylene) staining jar containing enough 100  mM Tris–HCl, pH > 9.5, to cover the slides completely. Place a lid loosely on the jar and then autoclave for 10 min, using liquid cycle settings. 2. Allow the jar to cool down at room temperature for 30 min and then transfer the slides to PBS for 10 min. 3. Proceed to immunolabeling (Subheading 3.4).

3.4. Immunoflu­o­ rescence Labeling

This section includes an optional sodium borohydride treatment for quenching glutaraldehyde-induced autofluorescence. Another effective measure to reduce autofluorescence is counterstaining the immunolabeled sections with toluidine blue prior to application of

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the mounting medium (19). Because toluidine blue has some red fluorescence, it is best suited for samples labeled with green- and orange-emitting dyes. The cryosectioned tissue retains chlorophyll and other pigments and exhibits intense fluorescence of chloroplasts in the red (peak emission around 680 nm). This is advantageous for imaging of chloroplasts by autofluorescence of chlorophyll, but also limits the choice of fluorescent dye conjugates mostly to those emitting in the green, yellow, and orange wavelengths, especially for detection of low-abundance proteins. Nonspecific binding of antibodies can be minimized by increased concentration of blocking agents (see Note 6) and by using antibodies at higher dilution. Pre-absorption of the polyclonal primary antibody with tissue powder lacking the protein of interest (e.g., from a null mutant plant), and pre-absorption of the secondary antibody with tissue powder from wild-type plants are other ways to minimize nonspecific binding and are described as optional steps (see Note 7). Optimal dilution of primary antibodies should be determined for each new antibody and new tissue. As a starting point, use a 10× lower dilution than what was determined as optimal for immunoblotting, and also prepare one or more higher and less-diluted antibody solutions. The commercially available secondary antibody conjugates have suggested dilution factors that can often be used without modification. Most of the processing steps are performed in slide staining jars (Coplin jars), except for the actual incubation with antibodies (see steps 4 and 7 in this section). Unless indicated otherwise, all steps should be performed at room temperature. Once the processing starts, the tissue sections should never be allowed to dry since this would cause nonspecific binding of antibodies and high background. 1. Optional: To pre-absorb the antibody, add 1 mL of the diluted antibody to a 0.2 mL of tissue powder (see Note 7) in a 1.5-mL microcentrifuge tube, vortex, and incubate at 4°C overnight. Vortex again and spin in a microcentrifuge at maximum speed (~14,000 × g) at 4°C for 5 min. Transfer the supernatant to a new tube and use for immunolabeling. 2. Optional: To quench glutaraldehyde-induced autofluorescence, immerse slides with sections in a staining jar containing sodium borohydride solution prepared immediately before use. Incubate for up to 30 min. Wash twice in PBS, for 5 min each. Sodium borohydride is very corrosive and a hazardous skin irritant. Use caution when handling the powder and the prepared solution. 3. Incubate sections in the blocking buffer for 1  h at room temperature. Set a small volume of the blocking buffer aside for diluting the antibodies.

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4. Prepare a dilution of the primary antibody in the blocking ­buffer, or use already diluted antibody that was pre-absorbed with a plant powder overnight (see Note 18). 5. Remove the slide out of the staining jar and blot off excess liquid by holding the slide vertically and touching the bottom edge on a paper towel for several seconds (Fig. 2l). Additionally, wipe the area around the sections using a tip of a folded paper towel. This helps keep the antibody from spreading over areas that do not have any sections. Apply 50–100 mL of the diluted primary antibody on the sections and rock the slide to distribute the antibody evenly (Fig. 2m). Cover the sections gently with a piece of Parafilm that is slightly narrower than the width of the slide (Fig. 2n). This will prevent drying of sections and help maintain a uniform layer of solution over the sections. Place the slides in a humid chamber (e.g., in a large Petri dish lined with wet filter paper) and incubate for 2–4  h at room temperature, or at 4°C overnight (see Note 19). 6. Remove and discard the Parafilm, and then wash the slides in PBST three times, for 10 min each time (see Note 20). 7. Prepare a dilution of the secondary antibody in the blocking buffer, or use the supernatant from pre-absorbed antibody, and spin down in a microcentrifuge for 5 min at ~14,000 × g to remove aggregates. Transfer the supernatant to a clean tube. 8. Apply the diluted secondary antibody in the same manner as described for the primary antibody (step 5 in this section). Incubate at room temperature for 1.5 h. 9. Remove and discard the Parafilm, and then wash the slides in PBST three times, for 10 min each time. 10. Optional: To reduce glutaraldehyde-induced autofluorescence, counterstain the tissue sections with 0.01% (w/v) toluidine blue in PBS for 10 min and then wash in PBS for 10 min. 11. Remove the slide from the staining jar and wick off excess liquid by holding the slide vertically and touching the bottom edge on a paper towel for several seconds (Fig. 2l). Put a small amount (20–30 mL) of mounting medium on the sections. Tilt the slide from side to side to distribute the medium evenly, and slowly apply a coverslip (see Note 21). Blot excess fluid off by touching a strip of filter paper along the edge of the coverslip and seal the coverslips with nail polish (Fig. 2o). Best image quality is usually achieved the next day after the mounting medium has evenly infiltrated the sections. 12. Store the sealed slides at −20°C for up to several weeks, or at −80°C for extended periods.

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3.5. Microscopy, Image Capture, and Processing

Specific operation of the microscope is hardware and software dependent, but a generalized protocol is presented. Fluorescence of the negative controls should be assessed and compared to that in samples of interest. The samples and the controls must be imaged using the same settings (exposure time, gain) to allow meaningful comparison. Chloroplasts contain large amounts of chlorophyll and other autofluorescent pigments and it is not trivial to distinguish the immunofluorescent signal from the background, especially in a cryosectioned tissue that retains most of the pigment. The peak emission from chlorophyll is at approximately 680 nm, which precludes the use of red-emitting fluorophores for immunolabeling, but the autofluorescence is very broad and is present regardless of excitation wavelength. The lowest amount of autofluorescence is found in the green part of the spectrum, and therefore the greenemitting fluorochromes, such as Alexa Fluor 488, Chromeo-488, or Atto 488, are the best options. In the wax-embedded tissue, most of the pigments are extracted by ethanol during dehydration, and the autofluorescence is less intense. In both wax and cryosections, fixation-induced autofluorescence may be significant in samples fixed with glutaraldehyde, and appropriate measures must be taken to quench this autofluorescence (see Subheading 3.4). It may also be possible to separate the signal of interest spectrally from the background using spectral scanning and unmixing, provided the microscope is equipped to do this (32). For multilabeled specimens, it is crucial to minimize cross talk between the channels by proper choice of fluorescent dyes for labeling, using specific filter sets with narrow band-pass emission filters, and in confocal imaging, performing sequential scanning if necessary. The amount of cross talk needs to be evaluated in control specimens where one of the secondary antibodies was omitted. Resolution and contrast in Z-stacks from both standard (widefield) and confocal microscopes can be improved and noise reduced by computational image restoration (deconvolution). Principles and algorithms as well as software options for deconvolution are detailed, e.g., in refs. (33–36). Bleaching (i.e., photodestruction or photoconversion of the fluorescent molecule to a nonfluorescent state) is most often encountered with high illumination intensity and in the presence of molecular oxygen (37, 38). One way to reduce bleaching is to choose secondary antibodies with the new-generation fluorophores that are much more resistant to photobleaching than the traditional dyes such as fluorescein or rhodamine. Additionally, mounting media containing anti-fade reagents greatly reduce bleaching (22, 38). During microscopy, use no more light than necessary and protect samples from light when they are not being imaged. Prepared slides may also undergo fading during storage. The slides may be stored at −20°C or even −80°C to extend their usability to weeks or months.

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1. Slides that were stored refrigerated or frozen should be allowed to warm up until the condensation on the slides evaporates. Previously viewed slides contaminated with immersion oil should be gently cleaned using cotton swabs and a glass cleaning solution. Oils of different brands should not be mixed. 2. Survey the slide using a dry, low-magnification objective and select an area of interest. 3. Switch to a suitable high-resolution immersion objective (glycerol or oil immersion), depending on the refractive index of the mounting medium. 4. If so equipped, set the objective’s correction collar to the coverslip thickness used (see Note 8). 5. Remove the Nomarski prism from the optical path. 6. Set the acquisition parameters (gain, exposure time) so that saturation of image pixels is avoided. To minimize photobleaching, lower the excitation intensity as much as possible, while maintaining reasonable exposure time. 7. Acquire the image or image stacks with sufficient Nyquist sampling in XY and Z planes (see Subheading 1.5 and Note 2) and save the original data in a native format of the imaging software (see Note 22). Record additional information regarding the imaging conditions as necessary. 8. Save the data to a reliable medium. Two copies of the data should be maintained and stored in separate locations. 9. Gray scale images from the original datasets may be pseudocolored using either the acquisition software or image analysis software of choice, e.g., the free ImageJ software. 10. Z-stacks can be reduced to 2D data by performing projections. Maximum intensity projections provide higher contrast, but the average intensity projection tends to preserve the lowercontrast detail. 11. Save processed images as TIFF, or other lossless image formats. Avoid using the JPG file format. The images may be further adjusted with image editing software (Gimp or Adobe Photoshop) (Fig. 3) (see Note 10).

4. Notes 1. It is not mandatory to use the highest purity grade 2,2-­thiodiethanol (T. Staudt, personal communication). The inexpensive grade (e.g., Aldrich, catalog number 166782) performs well for standard confocal imaging (39), but it is advisable to check for the level of autofluorescence in each batch.

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Fig. 3. Immunofluorescence localization of the chloroplast division protein FtsZ2 in Arabidopsis leaf mesophyll chloroplasts. Wide-field fluorescence images were acquired with a 100×/1.4 oil immersion objective. (a) FAA-fixed, Steedman’s waxembedded wild-type tissue was subjected to antigen retrieval before immunostaining. (b) Cryosectioned wild-type tissue did not require antigen retrieval. (c) FAA-fixed, Steedman’s wax-embedded tissue from the arc6 chloroplast division mutant (42), immunostained after antigen retrieval. Note the fragmented FtsZ filaments in the grossly enlarged chloroplast of the mutant. Chloroplast shape is marked with the dashed line. Arrows in (a) and (b) indicate FtsZ rings at the chloroplast division site. Images in (a) and (b) were pseudo-colored for clarity. Green = FtsZ2 signal; red = autofluorescence of chlorophyll in the chloroplast. Scale bar = 5 mm.

2. Under optimal conditions, an oil immersion objective of 1.4 numerical aperture resolves objects separated by approximately 0.2 mm. In the image projected by a 100× objective onto the CCD camera, this minimal separation corresponds to 100 × 0.2 mm, i.e., 20 mm. This is more than double the CCD pixel size in a typical interline CCD camera (6.45 mm pixels). Thus, Nyquist criterion is satisfied and the image can be captured at the full resolution afforded by the objective. Similar calculation shows that a 60× objective of the same optical resolution would result in under-sampling and loss of resolution in the recorded image. 3. Plant leaf tissue often adheres to plastic utensils and containers. The use of glass containers for fixation and dehydration, and glass Pasteur pipettes for aspirating liquid from sample vials is recommended. For handling the molten wax during wax embedding, either glass or plastic Pasteur pipettes (transfer pipettes) can be used. 4. Slides without a frosted writing surface are preferred, since the frosted area tends to wick away the antibody solutions applied for immunostaining, causing some sections to dry. 5. Besides poly-l-lysine-coated slides, silanized or specially treated, positively charged Superfrost Plus slides can also be used for good adherence of tissue sections. These are available from many vendors. 6. Various blocking agents can be used instead of nonfat dry milk, such as bovine serum albumin (BSA) at 2–5% (w/v). BSA usually contains a significant amount of fatty acids, which may interfere with immunolabeling. Therefore, fatty acid-free BSA

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is recommended. Dry milk and most commercial sources of BSA contain bovine IgG, which interferes with the use of most anti-goat or anti-sheep IgG fluorescent conjugates, unless these were pre-absorbed against bovine IgG. If this is a problem, cold water fish skin gelatin at 0.5 % (w/v) in PBST can be used as a blocking agent. 7. In order to eliminate nonspecific binding, the diluted antibodies can be pre-absorbed with plant tissue powders. For preabsorption of primary antibodies, the powder is prepared from a null mutant plant not containing the target protein (if available). For pre-absorption of secondary antibodies, the same genotype and tissue type as the one under investigation are used. Fix and wash a sufficient amount of tissue, blot the tissue dry on a filter paper, and using a mortar and pestle grind the tissue in liquid nitrogen. Transfer the frozen powder to a suitable container, such as a 15-mL polypropylene conical tube, and store at −80°C. 8. Most microscope objectives are designed to be used with a 0.17-mm-thick cover glass. The standard #1.5 coverslips have a nominal thickness range of 0.16–0.19  mm and should be used for routine work. Nevertheless, even a 10-mm deviation of the coverslip thickness can lead to 50% loss of resolution in high numerical aperture dry and water immersion objectives. Even oil immersion lenses perform optimally only with a coverslip thickness of 0.17 mm (40). For critical applications, one should either measure and handpick individual coverslips, or purchase coverslips with narrow tolerances (Carl Zeiss, catalog number 474030–9000). Some microscope objectives are equipped with an iris, permitting an increase in contrast by decreasing the numerical aperture of the objectives. For image capture at highest resolution, the maximum aperture should be used, since both axial and lateral resolution depend on numerical aperture (40). 9. Many cameras and wide-field microscopes can be controlled by the freeware MicroManager (41) (http://www.micro-­manager. org). 10. Our typical workflow involves importing of the original image files into ImageJ, applying filters or other processing if necessary, and generating RGB pseudo-color images or image stacks by assigning each original dataset into the R, G, or B channel of an RGB image (ImageJ menu commands “Image-ColorMerge Channel”). If necessary, maximum or average intensity projections are generated from image stacks. The resulting images are then saved in TIFF format and multi-panel figures are created in software that can handle both raster and vector graphics, such as Scribus (freeware, http://www.scribus.net) or commercial software, e.g., Canvas (ACD Systems International

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Inc.), Adobe Illustrator (Adobe Systems Incorporated), or CorelDraw (Corel Corporation). 11. Vacuum infiltration in a syringe is often faster and more effective than using a vacuum desiccator. Remove the piston from a 10-mL plastic syringe, stop the outlet of the syringe with a piece of Parafilm and a gloved finger, and pour in several milliliters of the fixative with pieces of tissue. Insert the piston and invert the syringe so that the outlet is upward. Purge all air and excess fixative from the syringe, leaving only about 2  mL of liquid. Stop the outlet with the finger and pull the piston back. This will create a vacuum. Many small bubbles appear on the surface of the samples, as the air trapped inside the tissue expands and escapes. Shake the syringe vigorously, still holding the piston back and keeping the finger on the outlet. The goal is to shake the bubbles off the leaf surface so that the air does not return when the vacuum is released. Samples will often get stuck on the syringe walls. Check this occasionally and keep shaking. When all specimens are in the fixative, release the piston. The pressure inside reverts to normal and the fixative is pushed inside the tissue, replacing the air that escaped. Leaves that are infiltrated with the fixative turn darker green and are semi-translucent. Repeat the infiltration two to three times. Finally, remove the piston and transfer the samples into fresh fixative. 12. Cryosectioning is usually performed with the use of an antiroll plate positioned parallel to the knife to flatten the sections and keep them from curling. The sections should slide between the antiroll plate and the knife, and form short ribbons (Fig. 1d) that can be touched to a warm slide (Fig. 1e) or lifted with a brush and transferred to a chilled slide, which is later transferred to room temperature. Setting the antiroll plate correctly may be difficult, and some researchers prefer to remove the antiroll device and use a brush to hold the newly formed section down to prevent it from curling (Fig. 1f and g). 13. Melt only the amount of wax that will be needed for embedding; prolonged storage of wax in the molten stage changes its sectioning properties. Avoid overheating the wax; do not use microwave oven for melting. 14. Counterstaining helps make the leaf segment visible in the solidified, white wax block at the end of the embedding procedure. Instead of toluidine blue, basic fuchsin or other dyes for microscopy can be used, provided they are not highly fluorescent. Nevertheless, most of the dye is washed away during the wax infiltration procedure and leaf tissue in particular is often barely visible in the final wax block. Tissues with young, lessvacuolated cells, such as shoot apices or siliques, retain the

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stain better. It is helpful to be consistent in positioning all samples in the same way in the embedding mold, so that trimming of the wax block and microtome sectioning can be performed more or less blindly, without seeing the tissue. 15. We have successfully immunostained leaf sections from wax blocks that have been stored at room temperature for over 4 years. 16. Tilt the razor blade so that the edge is pointing away from you, and cut the ribbon of sections. The short ribbon will adhere to the razor blade and can be transferred to the slide. Use the paint brush to detach the ribbon gently from the razor blade. 17. When handling slides, use forceps to transfer slides from one staining jar to the next solution in another jar, rather than pouring the liquid out and in. Gently pull the slide out of the jar, blot off excess liquid by touching the bottom edge of the slide on a paper towel, and place the slide in the next solution without delay. 18. Repeated freezing and thawing of antibodies should be avoided. The concentrated primary antibody is divided into small ­aliquots and stored at −80°C. When needed, one tube is thawed and kept at 4°C for making dilutions. Secondary antibodies can be stored in a similar manner, or can be mixed with an equal volume of glycerol and stored in smaller aliquots at  −20°C. At this temperature, the antibody does not freeze and the required amount can be immediately pipetted out and the tube returned to the freezer. Since the antibody in glycerol is half the original concentration, use twice the amount when making dilutions. 19. The 2-h antibody incubation may not be sufficient for good signals due to slow penetration; overnight incubation may be required. In this case, sodium azide should be added to 0.1% (w/v) to prevent microbial growth (add 1 mL of 10% (w/v) NaN3 per 100 mL of diluted antibody). Overnight incubation is often performed at 4°C, but incubation at room temperature allows faster diffusion of the antibody. It is possible to incubate for several hours at room temperature and then overnight at 4°C. 20. If more thorough washing is necessary to reduce background staining, increase the number of washing steps. Alternatively, instead of using the staining jars, washing efficiency can be improved by laying the slides flat in a container (Petri dish or a lid from a pipette tip box) with a 10-mm layer of PBST. Agitate the tray very gently on an orbital shaker for 10 min each step. Sections that do not adhere well to the slide may be lost using this procedure.

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21. Lay one edge of the coverslip down on the slide and support the other edge with forceps or a dissecting needle. Slowly lower the coverslip onto the slide and avoid trapping bubbles underneath. 22. The native data formats for imaging software typically include not only images, but also the associated metadata with acquisition parameters. These primary data files should always be saved and archived. If necessary, the images may be exported to a different format for processing or sharing, such as TIFF. Conversion to a lossy compression format, such as JPEG, should be avoided if the images are to be used for processing and analysis.

Acknowledgments Preparation of this manuscript was supported by grants from the National Science Foundation and US Dept. of Energy to K.W.O. References 1. Johnson, I. D. (2006) Practical considerations in the selection and application of fluorescent probes. In, Handbook of Biological Confocal Microscopy, 3rd edn. (Pawley, J., ed.) Springer, New York, USA, pp. 353–367. 2. Sauer, M., Paciorek, T., Benkova, E., and Friml, J. (2006) Immunocytochemical techniques for whole-mount in situ protein localization in plants. Nat. Protocols 1, 98–103. 3. McAndrew, R. S., Olson, B. J. S. C., KadirjanKalbach, D. K., Chi-Ham, C. L., Vitha, S., Froehlich, J. E., and Osteryoung, K. W. (2008) In vivo quantitative relationship between plastid division proteins FtsZ1 and FtsZ2 and identification of ARC6 and ARC3 in a native FtsZ complex. Biochem. J. 412, 367–378. 4. Vitha, S., McAndrew, R. S., and Osteryoung, K. W. (2001) FtsZ ring formation at the chloroplast division site in plants. J. Cell Biol. 153, 111–119. 5. Wick, S. M., and Duniec, J. (1986) Effects of various fixatives on the reactivity of plant cell tubulin and calmodulin in immunofluorescence microscopy. Protoplasma 133, 1–18. 6. Sompuram, S. R., Vani, K., Messana, E., and Bogen, S. A. (2004) A molecular mechanism of formalin fixation and antigen retrieval. Am. J. Clin. Pathol. 121, 190–199. 7. Ferris, A. M., Giberson, R. T., Sanders, M. A., and Day, J. R. (2009) Advanced laboratory

techniques for sample processing and immunolabeling using microwave radiation. J. Neurosci. Meth. 182, 157–164. 8. Nakazono, M., Qiu, F., Borsuk, L. A., and Schnable, P. S. (2003) Laser-capture microdissection, a tool for the global analysis of gene expression in specific plant cell types: identification of genes expressed differentially in epidermal cells or vascular tissues of maize. Plant Cell 15, 583–596. 9. Feltlová, M. (2000) Preparing plant tissue cryosections for light microscopy – a little improvement. Biol. Listy 65, 270–272. 10. Slot, J. W., and Geuze, H. J. (2007) Cryosectioning and immunolabeling. Nat. Protocols 2, 2480–2491. 11. Steedman, H. F. (1957) A new ribboning embedding medium for histology. Nature 179, 1345. 12. Vitha, S., Baluška, F., Mews, M., and Volkmann, D. (1997) Immunofluorescence detection of F-actin on low melting point wax sections from plant tissues. J. Histochem. Cytochem. 45, 89–95. 13. McCartney, L., Marcus, S. E., and Knox, J. P. (2005) Monoclonal antibodies to plant cell wall xylans and arabinoxylans. J. Histochem. Cytochem. 53, 543–546. 14. Samaj, J., Ovecka, M., Hlavacka, A., Lecourieux, F., Meskiene, I., Lichtscheidl, I., Lenart, P.,

3  Immunofluorescence Microscopy for Localization of Arabidopsis… Salaj, J., Volkmann, D., Bogre, L., Baluska, F., and Hirt, H. (2002) Involvement of the mitogenactivated protein kinase SIMK in regulation of root hair tip growth. EMBO J. 21, 3296–3306. 15. Paciorek, T., Sauer, M., Balla, J., Wisniewska, J., and Friml, J. (2006) Immunocytochemical technique for protein localization in sections of plant tissues. Nat. Protocols 1, 104–107. 16. Otali, D., Stockard, C. R., Oelschlager, D. K., Wan, W., Manne, U., Watts, S. A., and Grizzle, W. E. (2009) Combined effects of formalin fixation and tissue processing on immunorecognition. Biotech. Histochem. 84, 223–247. 17. D’Amico, F., Skarmoutsou, E., and Stivala, F. (2009) State of the art in antigen retrieval for immunohistochemistry. J. Immunol. Methods 341, 1–18. 18. Gong, H. Q., Peng, Y. B., Zou, C., Wang, D. H., Xu, Z. H., and Bai, S. N. (2006) A simple treatment to significantly increase signal specificity in immunohistochemistry. Plant Mol. Biol. Rep. 24, 93–101. 19. Vitha, S., Baluška, F., Jasik, J., Volkmann, D., and Barlow, P. (2000) Steedman’s wax for F-actin visualization. In, Actin: a Dynamic Framework for Multiple Plant Cell Functions (Staiger, C. J., Baluška, F., Volkmann, D., and Barlow, P., eds.) Kluwer, Dordrecht, The Netherlands, pp. 619–636. 20. Finney, M. (1998) Nonradioactive methods for visualization of protein blots. In, Immunochemical Protocols (Pound, J. D., ed.) Humana Press, Totowa, NJ, USA, pp. 207–216. 21. Orcutt, K. M., Ren, S. S., and Gundersen, K. (2009) Detecting proteins in highly autofluorescent cells using quantum dot antibody conjugates. Sensors-Basel 9, 7540–7549. 22. Collins, T. Mounting Media and Antifade Reagents, Collation of information from the Confocal listserver archives and the Histonet archives as well as other web-resources. http:// www.uhnres.utoronto.ca/facilities/wcif/ PDF/Mountants.pdf. 23. Martini, N., Bewersdorf, J., and Hell, S. W. (2002) A new high-aperture glycerol immersion objective lens and its application to 3D-fluorescence microscopy, J. Microsc. 206, 146–151. 24. Egner, A., and Hell, S. W. (2006) Aberrations in confocal and multi-photon fluorescence microscopy induced by refractive index mismatch. In, Handbook of Biological Confocal Microscopy, 3rd edn. (Pawley, J., ed.) Springer, New York, USA, pp. 404–413. 25. Staudt, T., Lang, M. C., Medda, R., Engelhardt, J., and Hell, S. W. (2007) 2,2’-thiodiethanol: a new water soluble mounting medium for high

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resolution optical microscopy. Microsc. Res. Techn. 70, 1–9. 26. Pawley, J. (2006) Handbook of Biological Confocal Microscopy, 3rd edn. Springer, New York, USA. 27. Wayne, R. (2009) Light and Video Microscopy, 1st edn. Academic Press, New York, USA. 28. Mackenzie, J. M., Burke, M. G., Carvalho, T., and Eades, A. (2006) Ethics and digital imaging. Microsc. Today 14, 40–41. 29. Cromey, D. (2010) Avoiding twisted pixels: ethical guidelines for the appropriate use and manipulation of scientific digital images. Sci. Eng. Ethics doi:10.1007/s11948-010-9201-y. 30. Tirichine, L., Andrey, P., Biot, E., Maurin, Y., and Gaudin, V. (2009) 3D fluorescent in situ hybridization using Arabidopsis leaf cryosections and isolated nuclei. Plant Methods 5, 11. 31. Koiwai, H., Nakaminami, K., Seo, M., Mitsuhashi, W., Toyomasu, T., and Koshiba, T. (2004) Tissue-specific localization of an abscisic acid biosynthetic enzyme, AAO3, in Arabidopsis. Plant Physiol. 134, 1697–1707. 32. Moreno, N., Bougourd, S., Haseloff, J., and Feijó, J. A. (2006) Imaging plant cells. In, Handbook of Biological Confocal Microscopy, 3rd edn. (Pawley, J., ed.) Springer, New York, USA, pp. 769–787. 33. Murray, J. M., Appleton, P. L., Swedlow, J. R., and Waters, J. C. (2007) Evaluating performance in three-dimensional fluorescence microscopy. J. Microsc. 228, 390–405. 34. Cannell, M. B., McMorland, A., and Soeller, C. (2006) Image enhancement by deconvolution. In, Handbook of Biological Confocal Microscopy, 3rd ed. (Pawley, J., ed.) Springer, New York, USA, pp. 488–500. 35. Y. Sun, Davis, P., Kosmacek, E. A., Ianzini, F., and Mackey, M. A. (2009) An open-source deconvolution software package for 3-D ­quantitative fluorescence microscopy imaging. J. Microsc. 236, 180–193. 36. Biggs, D. S. (2010) 3D deconvolution microscopy. Curr. Protoc. Cytom. 52, 12.19.1112.19.20. 37. Lichtman, J. W., and Conchello, J.-A. (2005) Fluorescence microscopy. Nat. Methods 2, 910–919. 38. Ono, M., Murakami, T., Kudo, A., Isshiki, M., Sawada, H., and Segawa, A. (2001) Quantitative comparison of anti-fading mounting media for confocal laser scanning microscopy. J. Histochem. Cytochem. 49, 305–312. 39. Vitha, S., Bryant, V. M., Zwa, A., and Holzenburg, A. (2010) 3D confocal imaging of pollen. Microsc. Today 18, 26–28. 40. Keller, H. E. (2006) Objective lenses for confocal microscopy. In, Handbook of Biological

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Confocal Microscopy, 3rd edn. (Pawley, J., ed.) Springer, New York, USA, pp. 145–161. 41. Stuurman, N., Amodaj N., and Vale, R. D. (2007) Micro-Manager: open source software for light microscope imaging, Microsc. Today 15, 42–43.

42. Vitha, S., Froehlich, J. E., Koksharova, O., Pyke, K. A., van Erp, H., and Osteryoung, K. W. (2003) ARC6 Is a J-domain plastid division protein and an evolutionary descendant of the cyanobacterial cell division protein Ftn2. Plant Cell 15, 1918–1933.

Chapter 4 Transient Expression and Analysis of Chloroplast Proteins in Arabidopsis Protoplasts Dong Wook Lee and Inhwan Hwang Abstract Although chloroplasts have their own genome, most chloroplast proteins are encoded in the nuclear genome and are targeted to chloroplasts posttranslationally. In vitro import studies with isolated chloroplasts have been widely used and have helped to elucidate the complex mechanisms involved in protein targeting to chloroplasts. Recently, an in vivo targeting method using protoplasts emerged as an alternative method to investigate protein targeting into chloroplasts. The present study describes a set of principles and methods, including polyethylene glycol-mediated reporter plasmid transformation, fluorescence microscopy, immunocytochemistry, and Western blotting, for studying chloroplast interior and envelope membrane protein targeting using protoplasts isolated from Arabidopsis thaliana leaf tissues. Key words: Chloroplast, Protoplast, Transit peptide, Green fluorescent protein, Polyethylene glycol

1. Introduction Although chloroplasts and mitochondria have their own genomes, over 90% of their proteins are encoded in the nucleus and are transported to their target organelles posttranslationally. The N-terminal cleavable targeting signal, designated the transit peptide, is necessary and sufficient for protein targeting of chloroplast interior ­proteins (1–3). Unlike targeting signals for proteins destined to the endomembrane systems, nucleus, or peroxisome, transit peptides are dissimilar among chloroplast proteins and have very long lengths (13–146 amino acids), hinting at the complexity of protein targeting to chloroplasts (1, 4). Multiple steps are required for chloroplast targeting of chloroplast interior proteins, including navigation through the cytosol to chloroplasts after translation,

R. Paul Jarvis (ed.), Chloroplast Research in Arabidopsis: Methods and Protocols, Volume I, Methods in Molecular Biology, vol. 774, DOI 10.1007/978-1-61779-234-2_4, © Springer Science+Business Media, LLC 2011

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chloroplast binding, translocation across the envelope, and ­maturation (1, 2). To investigate the mechanisms involved in these events in detail, in vitro import assays using isolated chloroplasts have been used widely (5, 6) in which chloroplast precursor proteins are in  vitro translated in wheat germ extracts or rabbit reticulocyte lysate in the presence of isotope-labeled amino acids. The importation of in vitro translation protein products into isolated chloroplasts is qualitatively and quantitatively analyzed by changing the ATP concentration, temperature, and light/dark conditions. The interaction of in vitro translation products with translocons of the outer (TOC) and inner (TIC) chloroplast envelope machineries during protein import can be analyzed through cross-linking followed by immunoprecipitation. As an alternative method for investigating the targeting mechanisms of chloroplast proteins, an in vivo targeting assay using isolated protoplasts has emerged as a powerful tool (2, 3, 7–9). In this assay, isolated protoplasts are transformed with plasmids encoding chloroplast-targeted reporter proteins. After incubation, chloroplast targeting of preproteins in transformed cells can be observed in  vivo by fluorescence microscopy or analyzed by immunoblotting. In some cases, chloroplast targeting can also be examined through immunocytochemistry using specific antibodies. One of the major advantages of the protoplast system over the in  vitro import system is that the cytosolic regulator steps for chloroplast targeting can be studied. In addition, protein targeting to chloroplasts can be examined even in seedling lethal mutants that have severely defective chloroplasts (e.g., plastid protein import 2 [ppi2], tic20-I, and tic21) (8, 10). Here, we present methods regarding protoplast isolation, poly­ethylene glycol (PEG)-mediated reporter plasmid transfor­mation, fluorescence microscopy, immunoblotting, and immunocytochemistry with transformed protoplasts.

2. Materials 2.1. Plants and Media

1. Wild-type Arabidopsis (Arabidopsis thaliana) should be grown on B5 medium plates (see item 3 below) at 22°C in a growth chamber with a 16/8 h light/dark cycle. Leaf tissues harvested from 2-week-old plants are used immediately for protoplast manipulation (see Note 1). 2. For albino mutants such as ppi2, tic20-I, or tic21, leaf tissues are harvested from 3-week-old plants grown as described in item 1 above and used immediately for protoplast manipulation (10).

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3. B5 medium plates: 3.2 g/L Gamborg’s B5 medium (Duchefa Biochemie, Haarlem, The Netherlands), 2% (w/v) sucrose, 0.05% (w/v) 2-(N-morpholino)ethanesulfonic acid (MES) free-acid monohydrate, and 0.8% (w/v) agar. The medium should be autoclaved for 15  min at 121°C and then poured into Petri dishes. 2.2. Buffers and Reagents

1. Enzyme solution: 0.25% (w/v) macerozyme R-10 (Yakult Honsha Co. Ltd., Tokyo, Japan), 1.0% (w/v) cellulase R-10 (Yakult Honsha Co. Ltd.), 400 mM mannitol, 8 mM CaCl2, and 5 mM MES-KOH, pH 5.6. 2. 21% (w/v) sucrose solution. 3. W5 solution: 154  mM NaCl, 125  mM CaCl2, 5  mM KCl, 5 mM glucose, and 1.5 mM MES-KOH, pH 5.6. 4. MaMg solution: 400 mM mannitol, 15 mM MgCl2, and 5 mM MES-KOH, pH 5.6. 5. PEG solution: 400  mM mannitol, 100  mM Ca(NO3)2, and 40% (w/v) PEG 8,000. 6. 16% (w/v) paraformaldehyde solution. 7. TSW buffer: 10  mM Tris–HCl, pH 7.4, 0.9% (w/v) NaCl, 0.25% (w/v) gelatin, 0.02% (w/v) sodium dodecyl sulfate (SDS), and 0.1% (w/v) Triton X-100. 8. Monoclonal anti-GFP antibody raised in mouse (Clontech Co., Mountain View, CA, USA). This is suitable for both immunocytochemistry and immunoblotting. 9. FITC-labeled goat anti-mouse IgG (Roche Ltd., Basel, Switzerland). This should be diluted in TSW buffer at a titer of 1:300 and can be used for immunocytochemistry. 10. Sonication buffer: 20 mM Tris–HCl, pH 7.4, 2.5 mM MgCl2, 2  mM ethylene glycol-bis(2-aminoethylether)-N,N,N¢,N¢tetraacetic acid (EGTA), 1  mM ethylenediaminetetraacetic acid (EDTA), 160  mM NaCl, 1% (v/v) Triton X-100, and protease inhibitor cocktail (Roche Ltd.). 11. 6× SDS sample buffer: 0.375 M Tris–HCl, pH 6.8, 12% (w/v) SDS, 60% (w/v) glycerol, 0.6  M dithiothreitol (DTT), and 0.06% (w/v) bromophenol blue. 12. 30% (w/v) monomer solution (200 mL): 60 g of acrylamide (FW 71.08) and 1.6 g of bisacrylamide (FW 154.2). 13. 4× Separating gel buffer: 1.5 M Tris–HCl, pH 8.8. 14. 4× Stacking gel buffer: 0.5 M Tris–HCl, pH 6.8. 15. 10% (w/v) SDS. 16. Double distilled water (DDW). 17. 10% (w/v) ammonium persulfate.

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18. N,N,N¢,N¢-tetramethylethylenediamine (TEMED). 19. Tank buffer: 0.025% (w/v) Tris, 0.192 M glycine, and 0.1% (w/v) SDS. 20. Western transfer buffer: 39 mM glycine, 48 mM Tris, 0.037% (w/v) SDS, and 20% (v/v) methanol. 21. Poly(vinylidene fluoride) (PVDF) membrane (Millipore Corporation, Billerica, MA, USA). 22. 10× Tris-buffered saline with Tween 20 (TBS-T): 1.37  M NaCl, 250 mM Tris–HCl, pH 7.5, and 1% (v/v) Tween 20. 23. Blocking solution: 6% (w/v) nonfat dry milk in 1× TBS-T buffer. 24. Primary antibody buffer: 3% (w/v) nonfat dry milk in 1× TBS-T buffer. Dilute a primary antibody in this buffer for immunoblotting. The optimal dilution factor depends on the titer of the particular antibody to be used. The mouse antiGFP antibody listed above can be used at a titer of 1,000:1. 25. Secondary antibody buffer: 3% (w/v) nonfat dry milk in 1× TBS-T buffer. Dilute anti-IgG (the secondary antibody) conjugated to horseradish peroxidase (HRP) in this buffer for Western blotting. The optimal dilution factor may vary depending on the manufacturer’s specification; in addition, the nature of the secondary antibody that should be used depends on the organism (e.g., mouse and rabbit) used to raise the primary antibody. The HRP-conjugated anti-mouse IgG antibody listed below can be used at a titer of 5,000:1. 26. HRP-conjugated goat anti-mouse IgG antibody (Amersham Biosciences Corp., Uppsala, Sweden). 27. Enhanced chemiluminescent (ECL) reagents (Amersham Biosciences Corp.). 2.3. Plasmid DNA

The plasmid that will be used in any given protoplast transfection experiment will depend to a large extent on the specific biological question that is to be addressed. In the example experiments that we have conducted, as illustrated in the presented figures, the plasmids described in items 1–3 below were used. 1. All reporter proteins were encoded in plasmid DNA derived from a pUC-based vector (11). 2. Reporter construct for protein import into chloroplasts: RbcSnt:GFP, the N-terminal 79 amino acids of rubisco small subunit are fused to GFP (2); Cab-nt:GFP, the N-terminal 67 amino acids of chlorophyll a/b binding protein are fused to GFP (3). 3. Reporter construct for protein targeting to outer envelope membrane: OEP7:GFP, full-length OEP7 fused to the

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N-terminus of GFP at the C-terminus; AtToc64 (1–29):GFP, the N-terminal 29 amino acids of AtToc64 are fused to GFP; T7:AtHsc70-4, a small epitope T7 is tagged at the N-terminus of AtHsc70-4 (9, 12–14). 4. Other vectors encoding different transit peptide sequences or different passenger/fluorescent proteins may also be used. 5. Plasmid DNA should be purified using Qiagen Plasmid Maxi Prep columns (Qiagen, Cologne, Germany), or equivalent columns. 2.4. Equipment and Software

1. Mesh (140  mm of pore size). For example, cell dissociation sieve CD1 (Sigma–Aldrich, Saint Louis, MI, USA). 2. Tabletop centrifuge and swinging-bucket rotor to accommodate 15-mL Falcon tubes. For example, Eppendorf 5702 (Eppendorf North America, Hauppauge, NY, USA). This is used throughout the protoplast isolation and transformation procedures. 3. Hemacytometer (Cole-Parmer, London, UK), and a microscope equipped for phase-contrast optics. 4. Fluorescence microscope, such as a Zeiss Axioplan (Jena, Germany), with the following filter sets: XF116 (exciter, 474AF20; dichroic, 500DRLP; emitter, 510AF23), XF33/E (exciter, 535DF35; dichroic, 570DRLP; emitter, 605DF50), and XF137 (exciter, 540AF30; dichroic, 570DRLP; emitter, 585ALP) (Omega Inc., Brattleboro, VT, USA) for green fluorescent protein (GFP), red fluorescent protein (RFP), and chlorophyll autofluorescence, respectively. The microscope should be equipped with a cooled charge-coupled device (CCD) camera. 5. Adobe Photoshop software (Mountain View, CA, USA). 6. Positively charged microscope slides (Fisher Scientific, Pittsburgh, PA, USA). 7. Sonicator. For example, Sonic Dismembrator Model 100 (Fisher Scientific). 8. Rotating shaker. For example, Orbital Shaker SH30 (FINEPCR Co., Seoul, Korea). 9. Electrophoresis system for SDS-polyacrylamide gel electrophoresis (SDS-PAGE). For example, the Mini Protean 3 system (Bio-Rad, Hercules, CA, USA), including the supplied gel casting cassette, is suitable for the procedures outlined in this chapter. 10. Power supply for electrophoresis and blotting. For example, power supply model SP-250 (Seoulin Co., Seoul, Korea).

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11. Semi-dry transfer apparatus. For example, Semiphor Transfer Unit (Amersham Biosciences Corp.). 12. LAS-3000 luminescent image analyzer (Fujifilm, Tokyo, Japan).

3. Methods All of the steps should be performed at room temperature, unless otherwise specified. 3.1. Isolation of Protoplasts from Arabidopsis Plants

1. Grow the plants for 2–3 weeks on a B5 plate. 2. Harvest leaf tissues (150 plants for 10 transformation experiments) of Arabidopsis plants from the plate using a new scalpel. 3. Dip the leaves into ~20 mL of enzyme solution. At this step, the amount of enzyme solution should be barely enough to soak all the tissues. 4. Incubate for ~8–12  h with gentle agitation (very slowly, at ~22–23°C, in the dark). After incubation, the solution should display a strong green color throughout. If the incubation time is too long, the protoplasts will be stressed. 5. Pass the protoplast solution through the 100-mm mesh to remove debris. 6. Load the mixture onto ~30 mL of 21% sucrose solution in a 50-mL Falcon tube. 7. Centrifuge at 730 rpm (98 × g) for 10 min in a swinging-bucket rotor. After centrifugation, the Falcon tube contains top and bottom fractions that consist of enzyme solution and 21% sucrose solution, respectively. Protoplasts are observed in four different locations: (1) in the top fraction containing enzyme solution, (2) at the interface between top and bottom fractions, (3) on the wall of the Falcon tube, and (4) at the bottom of the tube. Only protoplasts in the top fraction and at the interface should be used because the other fractions contain broken protoplasts. 8. Transfer intact protoplasts into ~30  mL of W5 solution in a 50-mL Falcon tube. At this step, it is preferable to use a Pasteur pipette. Be careful not to touch the 21% sucrose because sucrose usually prevents intact protoplasts from being pelleted to the bottom in the next step. 9. Centrifuge the mixture at 530  rpm (51 × g) for 6  min in a swinging-bucket rotor. 10. Discard the supernatant completely.

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11. Add fresh W5 solution (~15 mL). 12. Mix gently and completely. 13. Store at 4°C. Wait until all of the protoplasts sink to the bottom before use (~2  h) or centrifuge at 500  rpm (46 × g) for ~2–3 min in a swinging-bucket rotor. 3.2. PEG-Mediated Transformation

1. Before PEG-mediated transformation, aliquot the plasmid DNA (5–30 mg) into a 15-mL tube. It is recommended that the plasmid DNA concentration be adjusted to 1 mg/mL before transformation. 2. Pellet the protoplasts to the bottom of the Falcon tube at 4°. For this, just leave it at 4°C for 2 h, or centrifuge at 500 rpm (46 × g) for 2–3 min, as described in Subheading 3.1. 3. Discard the W5 solution (the supernatant from step 13 of Subheading 3.1). 4. Suspend the protoplasts in ~3 mL MaMg solution at a density of 5 × 106 protoplasts/mL. The number of protoplasts is counted using a hemacytometer and a phase-contrast microscope. 5. Resuspend protoplasts gently and completely by rotating the Falcon tube by hand. 6. Add 300  mL of protoplasts to each 15-mL tube containing ~5–30 mg of plasmid DNA, and mix gently and completely by rotating the tubes several times at an almost-horizontal position by hand. This step is very important because it will prevent protoplasts from being coagulated during the next two steps. 7. Add 300  mL of PEG solution to each tube. Mix gently and completely by rotating the tubes at an almost-horizontal position by hand. Incubate for 30 min. 8. Add 1 mL of W5 solution. Mix gently and completely by rotating the tubes at an almost-horizontal position by hand, and incubate for 10  min. Repeat this step two further times for three times in total. 9. Add 1.5  mL of W5 solution (750  mL × 2 times). Mix completely after each addition, as described above. The total volume will be ~5.1 mL. 10. Centrifuge for 4 min at 500 rpm (46 × g) in a swinging-bucket rotor. Discard the supernatant. 11. Add 2 mL of W5 solution, mix well, and incubate at ~22–23°C in a dark incubator.

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3.3. Fluorescence Microscopy

GFP or RFP fusion proteins can be observed directly in the ­transformed protoplasts. Best results are obtained approximately 8–24  h after completion of the transfection procedure in Subheading 3.2. 1. Capture images with a cooled CCD camera using a fluorescence microscope. 2. Process data using Adobe Photoshop software for pseudocolor images. 3. Protein imported into chloroplasts will show close overlap with autofluorescence of chlorophyll. In the case of proteins targeted to the outer envelope membrane, a ring pattern surrounding the chlorophyll autofluorescence can be observed (Fig. 1).

Fig. 1. In vivo localization of GFP-tagged reporter proteins. RbcS-nt:GFP (a) or AtToc64 (1–29):GFP together with AtOEP7: RFP (b) were transformed into protoplasts and localization of reporter proteins was examined by a fluorescent microscope. Bar = 20 mm.

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Subcellular localization of nonfluorescent chloroplast proteins can be determined by immunohistochemistry using an antibody raised against the passenger protein. In certain cases, localization of GFP or RFP fusion proteins also needs to be detected by immunohistochemistry using anti-GFP or anti-RFP antibodies, respectively (see Note 2). 1. Transform protoplasts with various constructs (see Subheading 3.2) and then finally resuspend them in 300 mL of W5 solution (to do this, the cells can be pelleted as described in Subheading 3.2, step 2). 2. Spread the cell suspension onto positively charged microscope slides. 3. Fix the cells by adding 70  mL of 16% paraformaldehyde and incubating for 1.5 h. 4. Permeabilize the cells by washing three times with 300 mL of TSW buffer for 30 min (10 min for each wash) on the slide. No mixing or shaking is required during the wash steps; surface tension prevents the buffer from leaving the slide. 5. After the final wash with TSW, stain the cells with primary antibodies diluted in 300 mL of TSW buffer (e.g., anti-GFP at a dilution of 1:300) for 16 h at 4°C. Again, no mixing or shaking is required. The dilution factor largely depends on the titer of the particular antibody used. 6. Wash the cells three times with 300  mL of TSW buffer, for 10 min each wash, with no shaking. 7. Stain the cells with secondary antibody diluted in 300 mL of TSW buffer (e.g., FITC-labeled goat anti-mouse IgG at a titer of 1:300) for 16 h at 4°C. The dilution factor largely depends on the titer of the particular antibody used. 8. Wash the cells three times with 300  mL of TSW buffer, for 10 min each wash, with no shaking. 9. Examine the samples by fluorescence microscopy, as described in Subheading 3.3 and previously (7) (see Note 2). 10. Protein localized to the stroma produces as a disc pattern that closely overlaps with autofluorescence of chlorophyll. In contrast, proteins localized to the inner or outer envelope membrane produce a ring pattern that surrounds the chloroplast (Fig. 2).

3.5. Preparing Total Protein Extracts from Transformed Protoplasts for Immunoblotting

1. Before preparing total protein extracts, remove W5 solution from each 15-mL tube containing transformed protoplasts (to do this, the cells can be pelleted as described in Subheading  3.2, step 2) and resuspend each sample with 300 mL of sonication buffer.

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Fig. 2. Localization of reporter proteins by immunohistochemistry. T7:AtHSC70-4 was detec­ted by immunohistochemistry using anti-T7 antibody followed by FITC-labeled anti-mouse IgG as the secondary antibody. Bar = 10 mm.

2. Sonicate the transformed protoplasts with using a sonicator at output amplitude 1 for 10 s. 3. Incubate at 4°C for 15  min with agitation in a rotating shaker. 4. Centrifuge in a microfuge at 3,000 × g at 4°C for 10 min and harvest the supernatant. 5. Add 50 mL of 6× SDS sample buffer to the supernatant, mix well, and boil for 5 min. 6. Store at −20°C before use. 7. Use 50 mL per sample for immunoblotting. 3.6. Immunoblotting

1. Prepare the separating gel solution (10% SDS-PAGE, 30 mL) as follows: 12.1 mL of DDW, 10 mL of 30% (w/v) monomer solution, 7.5 mL of 4 × separating gel buffer, 300 mL of 10% (w/v) SDS, 150 mL of 10% (w/v) ammonium persulfate, and 10 mL of TEMED. Do not add the last two components until you are ready to pour the separating gel. 2. Prepare the stacking gel solution as follows: 12 mL of DDW, 2.66 mL of 30% (w/v) monomer solution, 5 mL of 4× stacking gel buffer, 200  mL of 10% (w/v) SDS, 100  mL of 10% (w/v) ammonium persulfate, and 10 mL of TEMED. Do not add the last two components until you are ready to pour the stacking gel. 3. Prepare an SDS polyacrylamide gel using the aforementioned separating gel and stacking gel solutions. Set up a mini-gel

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casting cassette. First pour 10  mL of separating gel solution into the mini-gel casting cassette and then add immediately 0.5 mL of normal butanol to the top of the separating gel to make the surface of the separating gel even. Let the separating gel polymerize at room temperature for at least 20 min. After polymerization of the separating gel, thoroughly remove the butanol by washing with DDW and then pour the stacking gel solution (approximately 1.5–2.0 mL per gel) to fill the plate; immediately insert the comb. Let the stacking gel polymerize at room temperature for at least 5 min. 4. Place the gel in the electrophoresis apparatus and fill both reservoirs with tank buffer. Load 30–50 mL of each sample to the gel. Run the gel at 65 V through the stacking gel and at 120 V through the separating gel using a power supply. 5. After SDS-PAGE, transfer proteins from the gel to a PVDF membrane using a semi-dry transfer machine at a constant current of 60 mA for each gel, for 1 h 30 min. Blotting should be conducted in Western transfer buffer. 6. After transfer, incubate the membrane in 40 mL of blocking solution with shaking on a rotating shaker for 30 min. 7. Incubate the membrane in 5 mL of primary antibody buffer at 4°C overnight on a rotating shaker. 8. Wash three times (15  min each) in 1× TBS-T on a rotating shaker. 9. Incubate the membrane in 5 mL of secondary antibody buffer at 4°C overnight, or at room temperature for 1.5 h, on a rotating shaker. 10. Wash three times (15  min each) in 1× TBS-T on a rotating shaker. 11. Immerse the membrane with ECL reagents. 12. Capture the chemiluminescence images with an LAS-3000 luminescent image analyzer, or a similar device. 13. Protein that has been imported into chloroplasts migrates slightly faster than the precursor form on SDS-PAGE gels, due to proteolytic processing of the N-terminal transit peptide (see Fig. 3).

4. Notes 1. The plant growth stage is critical for a successful transformation efficiency. We recommend that the growth stage be between the 4- and 10-rosette leaves stage for protoplast isolation and transformation (15).

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Fig. 3. Western blot analysis of proteins imported into chloroplasts. Total protein extracts from protoplasts that had been transformed with RbcS-nt:GFP or RbcS[T1A+3S]:GFP (the latter is like the former, but it has a transit peptide mutation) were analyzed by immuno­ blotting using anti-GFP antibody. Pre, precursor form; Pro, processed form. RbcS-nt, RbcS-nt:GFP; T1A+3S, RbcS[T1A+3S]:GFP.

2. Chloroplast interior proteins are unfolded during translocation across the chloroplast envelope membranes. Therefore, for some preproteins with mutant transit peptides that are defective in translocation across the chloroplast membrane (e.g., GFP-fused RbcS-nt [T4A/T7A] or Cab [T4A]), the GFP ­signal cannot be observed by fluorescence microscopy (2, 3). In such cases, the mutant preproteins can be visualized using immunocytochemistry with an anti-GFP antibody.

Acknowledgments This work was supported in part by grants from the National Research Foundation of Korea (NRF) (20100000737), World Class University Program (Project No. R31-2008-000-10105-0) of Ministry of Education, Science and Technology, and Technology Development Program (609004-05-1-SB210) for Agriculture and Forestry, Ministry for Food, Agriculture, Forestry and Fisheries (Republic of Korea).

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References 1. Li, H. M., and Chiu, C. C. (2010) Protein transport into chloroplasts. Annu. Rev. Plant Biol. 61, 157–180. 2. Lee, D. W., Lee, S., Lee, G. J., Lee, K. H., Kim, S., Cheong, G. W., and Hwang, I. (2006) Functional characterization of sequence motifs in the transit peptide of Arabidopsis small subunit of rubisco. Plant Physiol. 140, 466–483. 3. Lee, D. W., Kim, J. K., Lee, S., Choi, S., Kim, S., and Hwang, I. (2008) Arabidopsis nuclearencoded plastid transit peptides contain multiple sequence subgroups with distinctive chloroplast-targeting sequence motifs. Plant Cell 20, 1603–1622. 4. Zhang, X. P., and Glaser, E. (2002) Interaction of plant mitochondrial and chloroplast signal peptides with the Hsp70 molecular chaperone. Trends Plant Sci. 7, 14–21. 5. Perry, S. E., Li, H. M., and Keegstra, K. (1991) In vitro reconstitution of protein transport into chloroplasts. Methods Cell Biol. 34, 327–344. 6. Smith, M. D., Schnell, D. J., Fitzpatrick, L., and Keegstra, K. (2003) In vitro analysis of chloroplast protein import. Curr. Protoc. Cell Biol. Chapter 11, Unit 11.16. 7. Jin, J. B., Kim, Y. A., Kim, S. J., Lee, S. H., Kim, D. H., Cheong, G. W., and Hwang, I. (2001) A new dynamin-like protein, ADL6, is involved in trafficking from the trans-Golgi network to the central vacuole in Arabidopsis. Plant Cell 13, 1511–1526. 8. Lee, D. W., Lee, S., Oh, Y. J., and Hwang, I. (2009) Multiple sequence motifs in the rubisco small subunit transit peptide independently contribute to Toc159-dependent import of proteins into chloroplasts. Plant Physiol. 151, 129–141. 9. Lee, S., Lee, D. W., Lee, Y., Mayer, U., Stierhof, Y. D., Lee, S., Jürgens, G., and Hwang, I.

(2009) Heat shock protein cognate 70–4 and an E3 ubiquitin ligase, CHIP, mediate plastiddestined precursor degradation through the ubiquitin-26S proteasome system in Arabidopsis. Plant Cell 21, 3984–4001. 10. Kikuchi, S., Oishi, M., Hirabayashi, Y., Lee, D. W., Hwang, I., and Nakai, M. (2009) A 1-megadalton translocation complex containing Tic20 and Tic21 mediates chloroplast protein import at the inner envelope membrane. Plant Cell 21, 1781–1797. 11. Kim, D. H., Eu, Y. J., Yoo, C. M., Kim, Y. W., Pih, K. T., Jin, J. B., Kim, S. J., Stenmark, H., and Hwang, I. (2001) Trafficking of phosphatidylinositol 3-phosphate from the trans-Golgi network to the lumen of the central vacuole in plant cells. Plant Cell 13, 287–301. 12. Lee, Y. J., Kim, D. H., Kim, Y. W., and Hwang, I. (2001) Identification of a signal that distinguishes between the chloroplast outer envelope membrane and the endomembrane system in vivo. Plant Cell 13, 2175–2190. 13. Lee, Y. J., Sohn, E. J., Lee, K. H., Lee, D. W., and Hwang, I. (2004) The transmembrane domain of AtToc64 and its C-terminal lysinerich flanking region are targeting signals to the chloroplast outer envelope membrane. Mol. Cells 17, 281–291. 14. Bae, W., Lee, Y. J., Kim, D. H., Lee, J., Kim, S., Sohn, E. J., and Hwang, I. (2008) AKR2Amediated import of chloroplast outer membrane proteins is essential for chloroplast biogenesis. Nat. Cell Biol. 10, 220–227. 15. Boyes, D. C., Zayed, A. M., Ascenzi, R., McCaskill, A. J., Hoffman, N. E., Davis, K. R., and Görlach, J. (2001) Growth stage-based phenotypic analysis of Arabidopsis: a model for high throughput functional genomics in plants. Plant Cell 13, 1499–1510.

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Chapter 5 Visualisation of Stromules on Arabidopsis Plastids John C. Gray, James A. Sullivan, and Christine A. Newell Abstract Stromules are thin stroma-filled tubules that extend from all plastid types in all multicellular plants examined. They are most easily visualised by epifluorescence or confocal microscopy of plastids containing green fluorescent protein (GFP) or other fluorescent proteins. Transient expression of gene constructs encoding plastidtargeted GFP following bombardment of whole plants or organs of Arabidopsis with gold or tungsten particles coated with plasmid DNA is a relatively rapid and simple means of producing material for observation of stromules. Key words: Stromule, Chloroplast, Plastid, GFP, Particle bombardment

1. Introduction Stromules are stroma-filled tubules extending from the surface of plastids (1–3). They are 0.35–0.85 mm in diameter and may extend up to 200 mm in some cells (2–5). Stromules are bounded by the two plastid envelope membranes and contain soluble proteins, including the 550-kDa major stromal protein Rubisco, but not internal thylakoid membranes (2, 5). Stromules have been visualised on all plastid types (including chloroplasts, etioplasts, amyloplasts, leucoplasts, chromoplasts, and elaioplasts) (6, 7), although the frequency with which they occur depends on the cell type, the stage of plant or organ development, and environmental factors (4–7). In general, stromules are more abundant on nongreen plastids in cells with a low plastid density (4). Mesophyll cells, with many large, closely packed chloroplasts, generally contain none or only a few relatively short (8% available chlorine) and 0.1% (v/v) Tween 20. 7. Sterile deionised, or distilled, water. 8. Murashige–Skoog (MS) medium: 4.3  g MS salts (Duchefa Biochemie BV, Haarlem, The Netherlands), 30.0  g sucrose, and 7.0  g Phytoagar (Duchefa Biochemie) dispersed in ~900 mL deionised water, adjusted to pH 6.0 with 1 M NaOH, and made up to a final volume of 1  L. Autoclave at 15  psi (1.0 × 105 Pa) for 15 min. 9. 9-cm diameter Petri dishes. 10. Parafilm.

2.2. Particle Bombardment

1. Tungsten M-10 (~0.7  mm) or 1.0-mm gold microcarriers ­(Bio-Rad Laboratories, Hercules, CA, USA) (see Note 2). 2. 1.5-mL microfuge tubes. 3. Ethanol, 70% (v/v), freshly prepared. 4. Vortex. 5. Microfuge. 6. Sterile deionised, or distilled, water. 7. Glycerol, 50% (v/v). 8. Plasmid DNA encoding plastid-targeted fluorescent protein (1 mg/mL in water) (see Note 3). 9. Calcium chloride solution, 2.5 M, made up freshly, or stored frozen in 500 mL aliquots at −20°C. 10. Spermidine (Sigma–Aldrich S0266): dissolve complete contents of bottle in deionised water to give 0.1 M solution and store frozen in 200 mL aliquots at −20°C (see Note 4). 11. Ethanol, 100%. 12. Macrocarrier discs and rupture discs (1,100 psi) for Bio-Rad PDS-1000/He particle delivery system. 13. Bio-Rad PDS-1000/He particle delivery system. 14. Parafilm.

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1. Forceps. 2. Scalpel. 3. Glass slides and coverslips. 4. Deionised, or distilled, water. 5. Epifluorescence or confocal microscope fitted with appropriate filters for GFP and other fluorescent proteins (see Note 5).

3. Methods Stromules can be observed by epifluorescence or confocal laser scanning microscopy of plant material containing plastid-located fluorescent proteins. The plant material may be stable Arabidopsis transformants, as described in Subheading 1.2, or cells or tissues transiently expressing plastid-targeted fluorescent proteins. This section describes methods for the bombardment of Arabidopsis plants and organs with gold or tungsten particles coated with plasmid DNA encoding plastid-targeted GFP, and for observation of the resulting plant material. 3.1. Preparation of Plant Material

1. Surface-sterilise 1 mg (~50) of dry Arabidopsis seeds in a 1.5-mL microfuge tube by adding 1 mL of 70% ethanol and vortexing vigorously for 1 min. After 2 min, pellet the seeds by centrifugation in a microfuge for a few seconds (allow microfuge to reach top speed and then turn off) and pour off the supernatant. 2. Add 1 mL of 10% sodium hypochlorite and 0.1% Tween 20, vortex for 1  min, leave for 13  min, and then vortex again for  1  min. Collect the seeds by centrifugation. Pour off the supernatant. 3. Wash the seeds with 1 mL of sterile deionised water, vortex for 1 min, pellet by centrifugation, and then pour off the supernatant. Repeat four times. 4. Evenly space 4–6 seeds on the surface of MS medium in 9-cm diameter Petri dishes, seal the dishes with Parafilm, and place at 4°C for 48 h for stratification. 5. Transfer the dishes to a growth room or growth cabinet at 20°C with a photon flux density of 100 mmol photons/m2/s for a photoperiod of 16 h light, 8 h dark and allow the plants to grow for 21 days.

3.2. Preparation of Particles for Bombardment

1. Weigh 30 mg of gold or tungsten particles (see Note 2) in a 1.5mL microfuge tube, add 1 mL of freshly prepared 70% (v/v) ethanol, and vortex for 3 min. Leave at room temperature for 15 min. Pellet the particles by centrifugation in a microfuge for

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a few seconds (allow the microfuge to reach top speed and then turn off). Pour off the supernatant and discard. 2. Add 1 mL of sterile water, vortex for 1 min, leave for 1 min, centrifuge for a few seconds, and then pour off the supernatant and discard. Repeat three times. 3. Resuspend washed particles in 500 mL of 50% (v/v) glycerol (see Note 6). 4. Remove 50  mL to a new 1.5-mL microfuge tube and, while vortexing, add in the following order: 5 mL DNA (1 mg/mL), 50 mL CaCl2 (2.5 M), and 20 mL spermidine (0.1 M). Continue to vortex for 1 min, allow the particles to settle for 1 min, and then pellet by rapid centrifugation (allow microfuge to reach half top speed and then turn off). Pour off and discard the supernatant. 5. Carefully, add 140 mL of 70% ethanol to wash the pellet (do not disturb the pellet). Pour off the liquid and discard. 6. Add 100 mL of 100% ethanol and gently resuspend the particles by flicking the tube. 7. Pipette 10 mL into the centre of the macrocarrier disc and allow to dry. Place in the microcarrier launch assembly of a Bio-Rad PDS-1000/He particle delivery system. 3.3. Bombardment of Plant Material

1. Place a Petri dish containing 21-day-old Arabidopsis plants (see Note 7) on the second shelf from the base (to give a distance of ~7 cm from the stopping screen to the surface of the leaves) of the PDS-1000/He system, fitted with an 1,100-psi rupture disc. 2. Evacuate the chamber to at least 28 in Hg and turn on the flow of helium gas (see Note 8). 3. Following bombardment, remove the Petri dish, seal with Parafilm, and place in a growth cabinet at 20°C with a photon flux density of 100 mmol photons/m2/s for a photoperiod of 16 h light, 8 h dark for up to 4 days.

3.4. Fluorescence Microscopy

1. Cut 0.5 × 0.5 cm sections of plant tissue adjacent to any areas badly damaged by the bombardment and place on a glass slide. Add water and cover with a glass coverslip. 2. Examine at low magnification (×10 objective) in an epifluorescence or confocal microscope to identify areas with plastids containing fluorescent proteins (see Note 9). 3. Examine under higher magnification (×40 and ×100 oil immersion, or ×63 water immersion objectives) to detect stromules. Stromules marked with a fluorescent protein will extend beyond the central region of chloroplasts, which is defined by its chlorophyll fluorescence (see Note 10).

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4. With a confocal laser scanning microscope, take serial optical sections at 1.0-mm intervals along the vertical (z) axis and combine to follow the length of the stromule. Digital images can be processed in Adobe Photoshop (see Note 11).

4. Notes 1. We have successfully visualised stromules following bombardment of Columbia-0 and Wassilewskija ecotypes, and do not see any reasons why the protocols cannot be used for other ecotypes. However, successful visualisation of stromules may depend on genetic factors and optimal growth conditions, which may differ between ecotypes. 2. We have routinely used 1-mm gold particles for bombarding Arabidopsis, but recent batches from Bio-Rad have produced unacceptable levels of particle agglomeration, using the protocol described in Subheading 3.2. Tungsten M-10 particles, which are considerably cheaper, can also be used. 3. We have used plasmids encoding GFP fused to the plastid-­ targeting signal (transit peptide) of tobacco RbcS (2) or to the complete precursor protein of Arabidopsis InfA (29), and YFP fused to the full-length outer envelope protein OEP14 (Fig. 2) to visualise stromules. However, transit peptides from other plastid proteins, such as RecA, PAC, or FtsZ1, which have been used to target GFP to plastids in stable Arabidopsis transformants (7), should also be successful. Different methods of preparing plasmid DNA have not been evaluated with respect to the visualisation of stromules. However, we have successfully visualised stromules following bombardment with DNA prepared by Qiagen plasmid mini, midi, and maxi kits (Qiagen, Crawley, UK). 4. Spermidine is very deliquescent. We recommend dissolving the complete contents of the bottle in deionised water to a concentration of 0.1 M and then storing at −20°C in small aliquots. Spermidine spontaneously deaminates over time, so extended storage is not recommended. Spermidine solutions should not be autoclaved. 5. Dedicated filter sets or filter cubes for GFP and other fluorescent proteins are available for most microscopes. GFP is normally excited with the 488-nm line of an argon laser and emitted light collected in the fluorescein isothiocyanate (FITC) channel (535 ± 20  nm). YFP can be excited with the 514-nm line of an  argon laser and emitted light collected at 515–560  nm. CFP  can be excited at 425–445  nm and light collected at 460–510 nm.

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6. Washed gold or tungsten particles in 50% (v/v) glycerol may be stored at −20°C. Tungsten particles should not be stored for more than 1 month because of surface oxidation, which may have deleterious effects on the recipient cells. After storage, it is important to vortex thoroughly for 5  min to resuspend and disrupt agglomerated particles. 7. Instead of 21-day-old Arabidopsis plants, it is possible to bombard excised individual organs (e.g., leaves) or whole seedlings. Excised organs can be arranged on a sterile 8.5-cm Whatman No.1 filter paper wetted with 2 mL of MS liquid medium (MS medium without agar) in a sterile Petri dish. Three or four excised rosette leaves can be placed, adaxial (top) surface down, on the filter paper around the middle of the Petri dish and bombarded under the conditions given in Subheading  3.3. Following bombardment, the bombarded leaves are turned over so that the adaxial surface is uppermost and the end of the petiole is cut off. The leaves are then positioned with the cut end of the petiole against one side of the Petri dish so that, when the dish is tilted on its side, the base of the petiole is sitting in a pool of MS liquid medium. The dishes are then sealed with Parafilm and left standing on end in a slanted position with the ends of the petioles in liquid. 8. The chamber vacuum should be at least 28 in Hg. A vacuum of 30 in Hg gives a higher proportion of expressing cells, whereas a vacuum of 26 in Hg is ineffective for expression. 9. This initial screening for areas containing plastids marked with a fluorescent protein is important for determining whether the bombardment has been successful. Bombardment of Arabidopsis plants or leaves is generally less efficient than bombardment of tobacco leaves for expression of plastid-targeted fluorescent proteins, and if expression is not obtained following bombardment of Arabidopsis tissue, we would recommend that the plasmid preparation is used for bombardment of tobacco leaves, using the method outlined in Note 7. If novel chimeric gene constructs are being used for bombardment for the first time, we would recommend that a positive control construct, known to target a fluorescent protein to Arabidopsis plastids following bombardment, is also used. The following gene constructs, all under the control of the CaMV 35S promoter, have been used successfully for bombardment of Arabidopsis leaves, leading to observation of stromules: rice Rpl12-1 transit peptide-GFP (28), pea RbcS transit peptide-GFP (2), arabidopsis InfA-GFP (29), and pea OEP14-YFP (see Fig. 2). 10. The bombardment conditions have been optimised for expression in epidermal pavement cells, which are more likely than other leaf cell types to display easily visible stromules. The plastids in these epidermal cells are chloroplasts, although

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they contain lower amounts of chlorophyll than mesophyll chloroplasts, and are likely to be widely spaced, facilitating the observation of long stromules. Stromules are more difficult to observe in mesophyll cells because of the large, closely packed chloroplasts and the lower abundance of stromules. Stromules may also be observed in stomatal guard cells, although at a lower frequency than in epidermal pavement cells. If stromules are not observed on epidermal cell plastids clearly marked with fluorescent proteins, we would recommend comparison with leaves bombarded with a positive control construct known to visualise stromules (see Note 9). If the chimeric gene construct encodes the complete fulllength protein (not just the transit peptide) fused to a fluorescent protein, stromules may not be detected if the protein is excluded from stromules (for example, if it is associated with the thylakoid membrane or other large macromolecular complexes whose mobility in the stroma may be restricted). 11. The extent of image and data processing depends on the aims of the investigation. Single-channel images may be satisfactory for documenting the presence of stromules, but images combining fluorescent protein and chlorophyll fluorescence provide a clearer distinction between the stromal material and the thylakoid membranes in the plastid body. For quantitative assessment of stromule abundance, we usually examine at least 10 cells, ideally in at least 3 separate bombarded leaves, and express the number of plastids with stromules as a percentage of the total number of plastids in a cell. The statistical analysis of results expressed as percentages requires the use of arc-sine transformation of the data (4).

Acknowledgments We would like to thank Julian Hibberd, Michael Hansen, and Senthil Natesan for developing methods for the observation of stromules. This work was supported by grants from BBSRC and the European Commission. References 1. Köhler, R.H., Cao, J., Zipfel, W.R., Webb, W.W., and Hanson, M.R. (1997) Exchange of protein molecules through connections between higher plant plastids. Science 276, 2039–2042. 2. Gray, J.C., Sullivan, J.A., Hibberd, J.M., and Hansen, M.R. (2001) Stromules: mobile

protrusions and interconnections between plastids. Plant Biol. 3, 223–233. 3. Kwok, E.Y. and Hanson, M.R. (2004) Stromules and the dynamic nature of plastid morphology. J. Microsc. 214, 124–137. 4. Waters, M.T., Fray, R.G., and Pyke, K.A. (2004) Stromule formation is dependent upon

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plastid size, plastid differentiation status and the density of plastids within the cell. Plant J. 39, 655–667. 5. Hanson, M.R. and Sattarzadeh, A. (2008) Dynamic morphology of plastids and stromules in angiosperm plants. Plant Cell Env. 31, 646–657. 6. Köhler, R.H. and Hanson, M.R. (2000) Plastid tubules of higher plants are tissue-specific and developmentally regulated. J. Cell Sci. 113, 81–89. 7. Natesan, S.K.A., Sullivan, J.A., and Gray, J.C. (2005) Stromules: a characteristic cell-specific feature of plastid morphology. J. Exp. Bot. 56, 787–797. 8. Wildman, S.G., Hongladarom, T., and Honda, S.I. (1962) Chloroplasts and mitochondria in  living plant cells: cinephotomicrographic studies. Science 138, 434–436. 9. Gunning, B.E.S. (2005) Plastid stromules: video microscopy of their outgrowth, retraction, tensioning, anchoring, branching, bridging, and tip-shedding. Protoplasma 225, 33–42. 10. Kwok, E.Y. and Hanson, M.R. (2003) Microfilaments and microtubules control the morphology and movement of non-green plastids and stromules in Nicotiana tabacum. Plant J. 35, 16–26. 11. Kwok, E.Y. and Hanson, M.R. (2004) In vivo analysis of interactions between GFP-labeled microfilaments and plastid stromules. BMC Plant Biol. 4, 2. 12. Natesan, S.K.A., Sullivan, J.A., and Gray, J.C. (2009) Myosin XI is required for actin-associated movement of plastid stromules. Mol. Plant 2, 1262–1272. 13. Sattarzadeh, A., Krahmer, J., Germain, A.D., and Hanson, M.R. (2009) A myosin XI tail domain homologous to the yeast myosin ­vacuole-binding domain interacts with plastids and stromules in Nicotiana benthamiana. Mol. Plant 2, 1351–1358. 14. Tirlapur, U.K., Dahse, I., Reiss, B., Meurer, J., and Oelmüller, R. (1999) Characterization of the activity of a plastid-targeted green fluorescent protein in Arabidopsis. Eur. J. Cell Biol. 78, 233–240. 15. Fester, T., Strack, D., and Hause, B. (2001) Reorganization of tobacco root plastids during arbuscule development. Planta 213, 864–868. 16. Fester, T., Lohse, S., and Halfmann, K. (2007) “Chromoplast” development in arbuscular mycorrhizal roots. Phytochemistry 68, 92–100. 17. Kwok, E.Y. and Hanson, M.R. (2004) Plastids and stromules interact with the nucleus and cell

membrane in vascular plants. Plant Cell Rep. 23, 188–195. 18. Huang, J., Taylor, J.P., Chen, J-G, Uhrig, J.F., Schnell, D.J., Nakagawa, T., Korth, K.L., and Jones, A.M. (2006) The plastid protein THYLAKOID FORMATON1 and the plasma membrane G-protein GPA1 interact in a novel sugar-signaling mechanism in Arabidopsis. Plant Cell 18, 1226–1238. 19. Langeveld, S.M.J., van Wijk, R., Stuurman, N., Kijne, J.W., and de Pater, S. (2000) B-type granule containing protrusions and interconnections between amyloplasts in developing wheat endosperm revealed by transmission electron microscopy and GFP expression. J. Exp. Bot. 51, 1357–1361. 20. Bechtel, D.B. and Wilson, J.D. (2003) Amyloplast formation and starch granule development in hard red winter wheat. Cereal Chem. 80, 175–183. 21. Tirlapur, U.K. and König, K. (2001) Femtosecond near-infrared lasers as a novel tool for non-invasive real-time high-resolution time-lapse imaging of chloroplast division in living bundle sheath cells of Arabidopsis. Planta 214, 1–10. 22. Gremillon, L., Kiessling, J., Hause, B., Decker, E.L., Reski, R., and Sarnighausen, E. (2007) Filamentous temperature-sensitive Z (FtsZ) isoforms specifically interact in the chloroplasts and in the cytosol of Physcomitrella patens. New Phytol. 176, 299–310. 23. Barrero-Gil, J., Rodriguez-Navarro, A., and Benito, B. (2007) Cloning of the PpNHAD1 transporter of Physcomitrella patens, a chloroplast transporter highly conserved in photosynthetic eukaryotic organisms. J. Exp. Bot. 58, 2839–2849. 24. Escobar, N.M., Haupt, S., Thow, G., Boevink, P., Chapman, S., and Oparka, K. (2003) Highthroughput viral expression of cDNA-green fluorescent protein fusions reveals novel subcellular addresses and identifies unique proteins that interact with plasmodesmata. Plant Cell 15, 1507–1523. 25. Shiina, T., Hayashi, K., Ishii, N., Morikawa, K., and Toyoshima, Y. (2000) Chloroplast tubules visualized in transplastomic plants expressing green fluorescent protein. Plant Cell Physiol. 41, 367–373. 26. Reed, M.L., Wilson, S.K., Sutton, C.A., and Hanson, M.R. (2001) High-level expression of a synthetic red-shifted GFP coding region incorporated into transgenic chloroplasts. Plant J. 27, 257–265. 27. Okumura, S., Sawada, M., Park, Y.W., Hayashi, T., Shimamura, M., Takase, H., and Tomizawa, K.-I. (2006) Transformation of poplar

5  Visualisation of Stromules on Arabidopsis Plastids (Populus alba) plastids and expression of foreign ­proteins in tree chloroplasts. Transgenic Res. 15, 637–646. 28. Arimura, S., Hirai, A., and Tsutsumi, N. (2001) Numerous and highly developed tubular projections from plastids observed in tobacco epidermal cells. Plant Sci. 160, 449–454. 29. Millen, R.S., Olmstead, R.G., Adams, K.L., Palmer, J.D., Lao, N.T., Heggie, L., Kavanagh, T.A., Hibberd, J.M., Gray, J.C., Morden, C.W., Calie, P.J., Jermiin, L.S., and Wolfe, K.H. (2001) Many parallel losses of infA from chloroplast DNA during angiosperm evolution with multiple independent transfers to the nucleus. Plant Cell 13, 645–658. 30. Vitha, S., McAndrew, R.S., and Osteryoung, K.W. (2001) FtsZ ring formation at the chloroplast division site in plants. J. Cell Biol. 153, 111–119. 31. Holzinger, A., Kwok, E.Y., and Hanson, M.R. (2008) Effects of arc3, arc5 and arc6 mutations on plastid morphology and stromule formation in green and nongreen tissues of Arabidopsis thaliana. Photochem. Photobiol. 84, 1324–1335.

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32. Itoh, R.D., Yamasaki, H., Septiana, A., Yoshida, S., and Fujiwara, M.T (2010) Chemical induction of rapid and reversible plastid filamentation in Arabidopsis thaliana roots. Physiol. Plant. 139, 144–158. 33. Raab, S., Toth, Z., de Groot, C., Stamminger, T., and Hoth, S. (2006) ABA-responsive RNAbinding proteins are involved in chloroplast and stromule function in Arabidopsis seedlings. Planta 224, 900–914. 34. Kojo, K.H., Fujiwara, M.T., and Itoh, R.D. (2009) Involvement of AtMinE1 in plastid morphogenesis in various tissues of Arabidopsis thaliana. Biosci. Biotechnol. Biochem. 73, 2632–2639. 35. Holzinger, A., Buchner, O., Lütz, C., and Hanson, M.R. (2007) Temperature-sensitive formation of chloroplast protrusions and stromules in mesophyll cells of Arabidopsis thaliana. Protoplasma 230, 23–30. 36. Stokes, K.D., McAndrew, R.S., Figueroa, R., Vitha, S., and Osteryoung, K.W. (2000) Chloroplast division and morphology are differentially affected by overexpression of FtsZ1 and FtsZ2 genes in Arabidopsis. Plant Physiol. 124, 1668–1677.

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Chapter 6 Analysis of Chloroplast Movement and Relocation in Arabidopsis Masamitsu Wada and Sam-Geun Kong Abstract Chloroplast photorelocation movement is essential for the sessile plant survival and plays a role for efficient photosynthesis and avoiding photodamage of chloroplasts. There are several ways to observe or detect chloroplast movement directly or indirectly. Here, techniques for the induction of chloroplast movement and how to detect the responses, as well as various points of attention and advice for the experiments, are described. Key words: Accumulation, Arabidopsis, Avoidance, Blue light, Chloroplast, Light, Microbeam, Movement, Movie, Phototropin

1. Introduction Chloroplast movement is ubiquitous in almost all terrestrial sessile plants tested, including algae (but not in swimming algae), mosses, ferns, and seed plants, and has been known since the nineteenth century (1), although there are some plants reported not to show chloroplast movement, such as rice leaves (2). Chloroplasts in mesophyll cells show different distribution patterns under different light conditions (Fig. 1) (3–5). Chloroplast photorelocation movement is crucial for plant survival (6). Chloroplasts move away from strong light (avoidance response) to avoid photodamage of chloroplasts followed by cell death, and move toward weak light (accumulation response) to receive more light for photosynthesis. Chloroplast movement is crucial for the plants living under a canopy where strong light and shadow of upper leaves are frequently repeated. In this situation, chloroplasts should move every time up

R. Paul Jarvis (ed.), Chloroplast Research in Arabidopsis: Methods and Protocols, Volume I, Methods in Molecular Biology, vol. 774, DOI 10.1007/978-1-61779-234-2_6, © Springer Science+Business Media, LLC 2011

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Fig. 1. Distribution patterns of chloroplasts under different light conditions. The patterns shown are for darkness (a), and weak (b) and strong (c) light. Chloroplast distribution in a palisade cell is shown schematically as a side view of its mid part, and as top views with three different levels (i.e., top, middle, and bottom of the cell). The cell was irradiated from the leaf surface, namely, from the top.

to the cell surface or down to the cell side when the light ­conditions change. Under the circumstances where strong light is continuously shining, such as in open fields or at the top of the canopy, chloroplasts may always show an avoidance response and do not need to move. Similarly, under circumstances where weak light conditions occur throughout the daytime, chloroplasts must be at the mesophyll cell surface showing an accumulation response to absorb more light, so that they do not need to move. Even in the same species, an individual plant has to live under either one of these circumstances, so chloroplasts show different responses depending on the conditions where the plants live. Because of these reasons, pre-culture conditions are very important when performing experiments. Chloroplasts in vacuolated cells localize between the tonoplast and the plasma membrane. Probably they stick to the plasma membrane (7, 8) with their concave side (Fig. 2), although we do not know whether it occurs directly or indirectly, and how. Hence, they move along the plasma membrane but do not go through the midst of a vacuole. Chloroplasts can move in any direction by sliding without turning or rotating but not rolling (9, 10). They have a top and bottom but not a head and tail, at least from the point of view of their movement. Recently, short actin filaments were found around the chloroplast periphery, between the chloroplast and the plasma membrane, and were named chloroplast actin (cp-actin) filaments (11). The actin filaments may play a role in attaching chloroplasts onto the plasma membrane when chloroplasts do not

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Fig. 2. Electron microphotograph of chloroplasts attached to the plasma membrane with their peripheral region. Chloroplasts in a dark-adapted mesophyll cell of Arabidopsis thaliana are shown.

Fig. 3. Cp-actin filaments in an Arabidopsis thaliana palisade cell. Actin filaments were visualized by using a GFP-mouse talin fusion protein (a). A part of the cell shown in a rectangular dotted line was illuminated with a strong blue light continuously. The numbers at the upper-left corners show the time after the start of light irradiation. The corresponding chloroplasts were shown by chlorophyll autofluorescence (b). See the text for detail.

move and stay in one place. When chloroplast movement is induced, more cp-actin filaments appear at the front side of moving chloroplasts (Fig. 3) (11). Chloroplast unusual positioning 1 (CHUP1) is one of the factors controlling the cp-actin filament dynamics (11). Kinesin-like protein for actin-based chloroplast movement 1 (KAC1) and KAC2 also control the maintenance of cp-actin ­filaments (12).

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Not only the living circumstances, but also leaf age, leaf t­ hickness (layers of mesophyll cells), cell size, and other factors of leaf condition must be different from leaf to leaf. This lack of ­uniformity influences the level of chloroplast response, especially when detected by sensitive ways such as light transmittance. The best way to avoid such artifacts is direct observation of chloroplast movement induced by experimental conditions under microscopy. Here, we describe several methods for how to induce chloroplast movement and how to detect and analyze the movement. 1.1. Detection of Chloroplast Movement by Naked Eyes

In natural circumstances, both avoidance and accumulation responses occur under strong and weak light, respectively. But we cannot detect the movement with our naked eyes. Partial leaf irradiation through a slit is a simple and quick way to detect chloroplast movement without special machine(s) but by our own eyes. An area irradiated with strong light through a slit can easily be detected by a different light transmittance (as a pale green band over a green background), exaggerated by a sharp contrast at the border between the two areas. This method can be applied for both accumulation and avoidance responses (see below). We developed this method to screen for mutant plants defective in chloroplast movements (band assay) (Fig. 4) (8, 12, 13).

Fig. 4. Screening methods for mutants deficient in chloroplast movements (band assay). To screen for mutants deficient in the avoidance response, the middle part of mutagenized leaves placed on an agar plate was irradiated through a slit to induce an avoidance response (a). To screen for mutants deficient in the accumulation response, whole leaves were irradiated with strong light, followed by partial irradiation with weak light through a slit (b). See the text for detail.

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Chloroplast movement can be detected by a sensitive photosensor as transmission of monitoring beam light through leaf cells. If the sensitivity of photosensors is high enough, even a time course of small changes in light absorbance or transmittance caused by chloroplast movement can be detected continuously (Fig.  5). Inoue and Shibata detected successfully, for the first time, chloroplast photorelocation movement of terrestrial plants by absorption spectrophotometry (2). Any kind of photometer can be used if stimulating light for chloroplast movement induction is given without interference with the measuring beam. Excitation light must be blue or white light, but the measuring beam is preferably weak red light (around 660 nm) or longer wavelength. Numerical analysis is easily performed using the chart data obtained by spectrophoto­ metry (Fig. 5c). Moreover, when we use a microplate reader equipped with a 96-well plate, 96 different samples can be sorted and all the data from each sample are detected at once using a multi-detector system; this is very powerful to analyze many samples at the same time and under the same conditions (Fig. 5).

Fig. 5. Induction and detection of chloroplast movements by a commercially available 96-well microplate reader with some modifications. The whole system is shown with a photograph on the left (a). This system is composed of a microplate reader, a blue LED, a PC, a power supply, and a temperature-controlled incubator. A representative data chart taken with the system is shown on the right (c). The transmittance was reduced under accumulation response and increased under avoidance response. Chloroplast movement was induced in leaves of wild-type and phototropin mutants by applying different intensities of blue light using an LED (peak wavelength at 470 nm). The values shown are means ± SD derived from eight leaves. Chloroplast distribution patterns under each light condition are shown with the photographs above the chart (b).

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But we should be aware that the data obtained with this method are not guaranteed to reflect real chloroplast movement itself. The change in leaf transparency could be caused not only by the change in chloroplast distribution pattern from periclinal to anticlinal walls or vice versa, but also by chloroplast shape or direction changes and/or alterations of chlorophyll content, although the main change might be depending upon chloroplast movement. 1.3. Observation of Moving Chloroplasts Under Microscopy

This is the most reliable method to detect chloroplast movement. The behavior of each chloroplast before, during, and even after the irradiation can be observed under a microscope and/or recorded as a time-lapse movie. Precise analyses of chloroplast behavior can be made afterward using the movies. For more precise analysis of each chloroplast behavior, a part of a cell (Fig. 6a, b) (9) or even

Fig. 6. Chloroplast movements induced by two sequential blue microbeam irradiations. To induce an accumulation response, a part of the dark-adapted Arabidopsis palisade cell was irradiated with the first blue microbeam (100 mmol/m2/s) continuously (a). In turn, when the chloroplast reached position 2, the second blue microbeam was given to a different part of the cell (b). Similarly, to induce an avoidance response, a half portion of a chloroplast was irradiated with the first blue microbeam (100 mmol/m2/s) continuously (c), and then when the chloroplast reached position 2, the second blue microbeam was given to a different part of the chloroplast (d). The lines in (a) to (d) show the paths of moving chloroplast from positions 1 to 2 or 2 to 3 for 12 min (a), 38 min (b), 5 min (c), and 5 min (d), respectively. The microbeam was 7 mm in diameter in (a) and (b), or a 10 × 10 mm square in (c) and (d).

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Fig.  7. Structure of a microbeam irradiator remodeled from an ordinary microscope. A new light source and a light path were added. The samples can be observed with the eye and/or by taking a photograph or using a monitor (Ph). Bs, beam splitter; C-ob, ­condenser objective; Cs, 15% (w/v) CuSO4 solution (light path, ca. 6 mm); D, diaphragm; Fs, focusing stage; Hf, heat-cut filter; If, interference filter; Is, tungsten-halogen lamp; Is-ob, incandescent lamp for observation (30 W); mV, mV meter; Oc, ocular; Ob, objective lens; Pf, plastic filter; S, stage; Sp, silicon photocell; Sh, shutter; St, voltage stabilizer. Modified from Fig. 1 of Kadota and Furuya (14).

a  part of a chloroplast (Fig.  6c, d) (10) can be irradiated with a microbeam of various fluence rates with different size, different color, or different duration. In this case, a remodeled microscope or a custom-made microscope is needed (Figs. 7 and 8). We have made five microbeam irradiators so far, when needed for special purposes, with different structure and functions. The first generation was the most simple version, remodeling an old microscope by inserting a small diaphragm at a focal plane of the light path and adding another light source for observation (Fig. 7) (14). The second one was custom-made specifically for microbeam irradiation, and is still functioning after 30 years (Fig. 8) (15). Our confocal microscope (Leica SP5) has also been used as a microbeam irradiator. Microbeam irradiation is easily applicable within a small area with different wavelengths under live data mode. It is quite useful for the induction of the avoidance response, but the laser beam is too strong to induce an accumulation response. Without a costly microbeam irradiator, chloroplast movement can also be observed under an ordinary microscope by setting a sample preparation on a stage and keeping the observation light through the sample leaves strong enough, adjusted by the condenser lens. An inverted microscope for epifluorescence microscopy is ideal (see Note 1).

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Fig. 8. Structure of an order-made microbeam irradiator based on an inverted microscope (Biophot, Nikon, Tokyo). It has two light sources: one for microbeam illumination from above and another for observation with infrared light from below. Observation (Obs) of the samples and the microbeam can be done with the eye and/or by taking a photograph or using a monitor (Ph). C, cut-off filter; D, diaphragm; Dp, depolarizer; Fd, field diaphragm; H, heat-cut filter; HM, half mirror; Id, iris diaphragm; If, interference filter; IR, infrared light filter; L, tungsten-halogen lamp (12 V, 100 W); M, mirror; Nd, neutral density filter; Pl, polarizer; S, shutter; W, Wollaston prism. Modified from Fig. 1 of Yatsuhashi and Wada (15).

It takes more than 30 min for chloroplasts to accomplish either accumulation or avoidance movement. For continuous observation of these responses under microscopy, the sample leaf should neither move nor be dried up during the period. For this purpose, we use a custom-made cuvette for sealing a detached leaf (Fig. 9). The speed of chloroplast movement is rather slow, at about 1 mm/min on average (9, 10), so that the real-time detection of chloroplast movement under a microscope is almost impossible and the recorded data at real time are essentially meaningless. Hence, time-lapse recording of the phenomenon every 1  min and replaying it about 600 times faster than the normal speed are preferable, making it easy to analyze the precise behavior of chloroplasts.

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Fig. 9. Structure of a cuvette system used for chloroplast movement studies. Schematic drawings of the cuvette (a) and the ring holder (b) are shown. The whole system is shown with a photograph in (c). See the text for detail.

2. Materials 2.1. Detection of Chloroplast Movement by Naked Eyes

1. Plant material: young and fully expanded Arabidopsis leaves cultured under a light (16  h)/dark (8  h) photoperiod using white light (80–100 mmol/m2/s) at 23°C (see Note 2). 2. Light source: (a) For the accumulation response under weak light, use a fluorescence tube (e.g., FLR40SW, Mitsubishi Electric Corp., Japan); (b) For the avoidance response under strong light (higher than 100 mmol/m2/s), use an incandescent bulb (halogen lamp; e.g., Focusline 12V-100W HAL, Philips Lighting, The Netherlands) or a blue light-emitting diode (LED) illuminator (e.g., peak wavelength at 470  nm, half-­ bandwidth = 30 nm; LED-mB, Eyela, Japan) (see Note 3); (c) For mutant screening with many leaves at once, use an overhead projector with a wide irradiation space and strong light emission (Fig. 10). 3. A non-light-transmitting board with a slit for partial leaf irradiation (see Note 4). 4. A plastic plate with solidified agar (see Note 5). 5. A transparent sheet fitted to the plastic plate.

2.2. Detection of Light Transmittance Using Photosensors

1. Plant material; the same as described in Subheading 2.1, step 1. 2. Microplate reader (e.g., VersaMax, Molecular Devices) (see Note 6). 3. 96-Well plates.

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Fig. 10. Structure of the band assay system, providing illumination through a slit, used for chloroplast movement studies. The whole system is shown with a schematic diagram (a) and a photograph (b). See the text for detail.

4. Murashige and Skoog (MS) medium. 5. Gellan gum (see Note 7). 6. Sealing film for 96-well PCR plates. 7. Forceps or a needle. 8. Blue LED illuminating system (Fig. 5a). 2.3. Observation of Moving Chloroplasts Under Microscopy

1. Plant material; the same as described in Subheading 2.1, step 1. 2. A syringe (10 mL). 3. Cuvette (preparation): a custom-made cuvette is composed of two steel rings mating each other with complementary threads, and two round cover glasses and a silicon ring in between. The outer ring has an inner thread and a stopper at the bottom, and the inner ring is just a ring with a thread outside (Fig. 9). 4. Microbeam irradiator or an ordinary microscope equipped with a camera (Figs. 7 and 8) (see Note 1). 5. Time-lapse recording system (see Note 8).

3. Methods 3.1. Detection of Chloroplast Movement by Naked Eyes (see Fig. 10) 3.1.1. Avoidance Response

1. Cut off a rosette leaf from a plant incubated under darkness for a couple of hours (or under weak light conditions) and put the leaf, adaxial side up, on the solidified surface of 0.8% agar medium poured into a plastic square dish (e.g., sterile no. 2 square scale, Eiken Chemical Co., Ltd., Japan) (see Note 5). 2. Cover the leaf with a transparent film and then a slit board (or else, e.g., negative film with high contrast) such that the slit should cross on the middle part of the leaf (Fig. 4). 3. Put the agar plate under strong light (e.g., 50 W halogen lamp or blue LED) to irradiate the sample leaf through the slit. The fluence rate should be more than 100 mmol/m2/s of white light or 30 mmol/m2/s of blue light, respectively (see Notes 9 and 10).

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4. After irradiation for 30 min or longer, take off the slit board and see whether the irradiated part of the leaf becomes yellowish green (we call it a “white band”) (8, 13, 16) (see Note 11). 3.1.2. Accumulation Response

1. Almost all procedures are the same as described in Subheading 3.1, step 1 for the avoidance response except a few steps below. 2. A whole leaf put on an agar plate should be irradiated first with strong white light (800 mmol/m2/s) for 1 h, before the irradiation through a slit with weak light as described below (see Note 12). 3. Set a board with a slit on the leaf. 4. Irradiate the leaf with weak white light (47  mmol/m2/s) through the slit for 30 min. 5. Take off the slit board and observe the “green band.”

3.2. Detection of Light Transmittance Using Photosensors (see Fig. 5)

1. Pour 300  mL of 0.5% (w/v) gellan gum melted in quarterstrength MS medium into each well and leave it until it becomes solid (see Note 13). 2. Place detached leaves, adaxial side up, at the center of the agar surface of each well, one leaf for each well. 3. Seal the plate with a transparent sealing film (e.g., a sealing film for 96-well PCR plates) and then make two small holes over each well with a needle or sharp forceps (see Note 14). 4. Keep the sample in the dark for several hours to overnight at room temperature. 5. Set the sample plate on an automated plate reader under darkness or dim light conditions. 6. Record transmittance changes automatically using a 660-nm measuring light. 7. Irradiate the sample plate with blue light of appropriate intensity during the period to induce chloroplast movements using a blue LED illuminator (see Note 15). 8. Analyze the recorded date using analysis software such as Excel (Microsoft).

3.3. Observation of Moving Chloroplasts Under Microscopy

1. Pretreat the plants to see clearer responses (see Note 16). (a) For the avoidance response, cultivate plants under weak light conditions and then incubate them in darkness for a couple of hours. (b) For the accumulation response, incubate plants overnight (or at least 6  h) in the dark to induce dark positioning where almost all chloroplasts move from periclinal to anticlinal walls.

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2. Detach a rosette leaf and evacuate it gently in a syringe filled with deionized water (see Note 17). 3. Seal an excised and evacuated leaf, adaxial side up, in a cuvette or between a glass slide and coverslip (Fig.  9c) (see Notes 18 and 19). 4. Observe chloroplasts under a microscope or a microbeam irradiator with infrared light (or dim red light) and decide on target chloroplasts to induce either accumulation or avoidance responses (see Note 20). 5. Adjust a microbeam spot of infrared light with intensity higher than that of the background infrared light. For observation under an ordinary microscope, skip this step. (a) For the accumulation response, adjust the microbeam onto an area near the target chloroplast but not on the chloroplast itself (Fig. 6a and b). (b) For the avoidance response, adjust the infrared microbeam spot on the whole of the target chloroplast or a part of chloroplast under infrared light conditions (Fig. 6c and d). 6. Change the microbeam light from infrared to blue light with low fluence rates (1–10  mmol/m2/s for 40–60  min) for the induction of an accumulation response, or high fluence rates (30 mmol/m2/s or higher for 30 min) for an avoidance response (17) (see Note 21). 7. Record the chloroplast behavior during movement by using a time-lapse video system for further analyses of chloroplast movement. 8. Analyze the recorded responses (see Note 22).

4. Notes 1. Partial cell irradiation with a microbeam is a much easier way to observe and analyze chloroplast movement precisely. For this purpose, a diaphragm must be inserted in a focal plane of the light path of the microscope, and the figure of diaphragm should be focused on a part of the sample cell set on the sample stage. However, finding a focal plane is not easy; even if a focal plane position is found, usually there is no hole or slit on a microscope to insert a diaphragm in that position. Some methods for partial cell irradiation are still possible using an ordinary microscope. The easier one is the use of field stop. Close the field stop so that the fins can be seen in the field, set a leaf across the edge of the fins, and observe after some time. If the light is strong enough, chloroplasts move out of the light; if the light is weak, chloroplasts move into the light-irradiated

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area. However, this method is not suitable for precise analyses of chloroplast movement. Hence, custom-made microbeam irradiators are more suitable for precise analysis. 2. The length of photoperiod is not so important, but plants should be grown under photoperiodic conditions. In addition, aged leaves are not recommended because of their low sensitivity to light. Cotyledons can also be used. Dark adaptation at least for several hours before use makes leaves more sensitive. Chloroplast movement in other seed plant leaves, such as Coleus (18) or Perilla (Satoru Tokutomi, personal communication; easily purchased as a herb at supermarkets in Japan), fern gametophytes (19), and moss (20) could also be detected. 3. If incandescent bulbs or halogen lamps are used, heat-cut filters (such as HS 50, Hoya Glass Co., Tokyo) and/or a thick water layer (more than 10 cm) with 15% (w/v) CuSO4 solution should be inserted between the light source and samples (Fig. 10a). 4. When leaves are to be irradiated partially through a slit or a hole or other figures, any kind of screening device made of non-light-transparent material can be used. If you cannot find an appropriate item, make it by yourself using aluminum sheet. Cut out a slit of 1–2 mm in width from a piece of aluminum sheet. Stick the aluminum piece with a slit firmly onto a transparent sheet, such as a film for an overhead projector, with double-sided sticky tape. Even a black-and-white drawing or a negative film of a photograph with high contrast is useful; see cover of Science, 16th March issue, 2001; and Fig.  3 of Suetsugu et al. (12). 5. Wet soft paper can be used instead of an agar plate. However, a transparent agar plate is preferable to detect weak responses. 6. A custom-made spectroradiometer remodeled for measuring weak light transmittance is also useful (21, 22). We assembled such a piece of equipment by ourselves from a LED (GL5UR3K; Sharp Corp., Osaka, Japan), a photodiode (S1227-66BR; Hamamatsu Photonics K.K., Hamamatsu, Japan), a voltmeter, and a power supply (6). 7. Gellan gum is better than ordinary agar because of its higher transparency. 8. Any kind of PC that can control time-lapse acquisition of photographs taken with an infrared light-sensitive camera is recommended for use. 9. We use an overhead projector for the light source in which incident light comes from the bottom, so that the plate is put upside down onto the glass plate of the projector (Fig. 10b). In this case, a kind of supporter like a transparent, thick plastic plate should be inserted into the dish to prevent the slit board

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from coming off. If the light source emits too much heat, a heat-cut filter or a layer of 15% (w/v) CuSO4 solution (e.g., 10  cm in thickness) should be inserted between the sample dish and the light source. Blowing cool air is also effective (Fig. 10a). 10. Chloroplast movement is induced by blue light, but red light enhances the response, so that using white light is more preferable than using blue light. 11. It can be seen even by naked eyes, but if observed under a fluorescent microscope, the contrast of brilliant red and darker red chlorophyll fluorescence between the irradiated and non-­ irradiated area is much more clearly seen. 12. This procedure induces an avoidance response in the whole area of the leaf and makes it easy to observe the accumulation response as a “green band” at the area irradiated through a slit (13). 13. The importance of this procedure is to fix the leaf so that it does not move during the repeated measurements of transmittance from the same position; thus, not only gellan gum but also solidified normal agar is usable. The most important part is the hardness of the supporting gel. MS medium is also not essential; distilled water or even tap water can be substituted. 14. This procedure is necessary to keep cut leaves in sufficient moisture, avoiding dehydration, and to prevent condensation of water on the film during measurement. But the holes should not be in the center where the measuring beam passes. 15. Appropriate fluence rates needed for accumulation and avoidance responses are easily obtained by adjusting a power supply or inserting neutral-density filters under the LED illuminator as shown in Fig. 5a. 16. We recommend this procedure optionally to induce chloroplast movement more efficiently. Even if you skip these procedures, the responses are detectable. 17. Air spaces in a leaf are a serious obstruction to the observation of chloroplasts clearly under a microscope, and so the air in air spaces should be eliminated by evacuation using a syringe. Dip a leaf under distilled water in the tube of the syringe, push out any remaining air in the syringe through the tip-hole (normally used for applying a needle) by pushing up the piston, then seal the tip-hole with a finger, and pull and release the piston five to ten times repeatedly, to take out the air in the leaf, until the leaf becomes transparent. 18. Chloroplasts staying on the periclinal wall of the top layer of palisade cells, namely, the chloroplasts just under the epidermal cells of the adaxial side, should be observed. All procedures

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had better be done under safe light conditions, i.e., dim green light or dim red light, because chloroplast movement is induced by blue light. 19. A sample leaf is sealed with water between two round cover glasses supported by a ring-shaped silicon-rubber spacer. Then the cover glasses with the sample is put on the bottom of the outer ring of the cuvette and then tightened with the inner ring (Fig. 9c). An easy way to seal leaves and keep them moistened is to put them with water between a slide and cover glass sealed with double-sided sticky tape. 20. To see chloroplasts under infrared light, an infrared-sensitive camera connected to a microscope or a microbeam irradiator and a monitor screen should be used. 21. Red background light irradiation for whole view field is better to be used, because red light enhances chloroplast movement but has no influence on the directional movement (23). 22. Analytical studies will be required for the description and/or evaluation of chloroplast movement in scientific papers. The path and time course of chloroplast movement, a lag time before the start of movement, movement speed and direction, and all other behaviors can be calculated from the movie frames recorded. The analyses are easily done using the image analysis software package “ImageJ,” which is freely downloaded (http://rsbweb.nih.gov/ij/).

Acknowledgments The authors thank Mr. Hidenori Tsuboi for time-lapse images of chloroplast movement induced by microbeam irradiation and Dr. Isao Uemura (Tokyo Metropolitan University) for his support on electron microscopy. This work was supported in part by Grantsin-Aid for scientific research from the Ministry of Education, Culture, Sports, Science and Technology of Japan (MEXT) (Grants 17084006 to M.W. and 21770050 to S.-G.K.) and the Japan Society for the Promotion of Science (JSPS) (Grant 20227001 to M.W.). References 1. Senn, G. (1908) Die Gestalts- und Lageverän­ derung der Pflanzen-Chromatophoren., Leipzig, Germany. 2. Inoue, Y., and Shibata, K. (1973) Light-induced chloroplast rearrangements and their action

spectra as measured by absorption spectrophotometry. Planta 114, 341–358. 3. Wada, M., Kagawa, T., and Sato, Y. (2003) Chloroplast movement. Annu. Rev. Plant Biol. 54, 455–468.

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4. Suetsugu, N., and Wada, M. (2007) Chloroplast photorelocation movement mediated by phototropin family proteins in green plants. Biol. Chem. 388, 927–935. 5. Suetsugu, N., and Wada, M. (2009) Chloroplast photorelocation movement. In, The Chloroplast: Interactions with the Environment (Sandelius, A. S., and Aronsson, H., eds.), Springer, Berlin / Heidelberg, Germany, pp. 235–266. 6. Kasahara, M., Kagawa, T., Oikawa, K., Suetsugu, N., Miyao, M., and Wada, M. (2002) Chloroplast avoidance movement reduces photodamage in plants. Nature 420, 829–832. 7. Oikawa, K., Yamasato, A., Kong, S.G., Kasahara, M., Nakai, M., Takahashi, F., Ogura, Y., Kagawa, T., and Wada, M. (2008) Chloroplast outer envelope protein CHUP1 is essential for chloroplast anchorage to the plasma membrane and chloroplast movement. Plant Physiol. 148, 829–842. 8. Oikawa, K., Kasahara, M., Kiyosue, T., Kagawa, T., Suetsugu, N., Takahashi, F., Kanegae, T., Niwa, Y., Kadota, A., and Wada, M. (2003) Chloroplast unusual positioning1 is essential for proper chloroplast positioning. Plant Cell 15, 2805–2815. 9. Tsuboi, H., Yamashita, H., and Wada, M. (2009) Chloroplasts do not have a polarity for light-induced accumulation movement. J. Plant Res. 122, 131–140. 10. Tsuboi, H., and Wada, M. (2011) Chloroplasts can move in any direction to avoid strong light. J. Plant Res. 124, 201–210. 11. Kadota, A., Yamada, N., Suetsugu, N., Hirose, M., Saito, C., Shoda, K., Ichikawa, S., Kagawa, T., Nakano, A., and Wada, M. (2009) Short actin-based mechanism for light-directed chloroplast movement in Arabidopsis. Proc. Natl. Acad. Sci. USA 106, 13106–13111. 12. Suetsugu, N., Yamada, N., Kagawa, T., Yonekura, H., Uyeda, T. Q., Kadota, A., and Wada, M. (2010) Two kinesin-like proteins mediate actin-based chloroplast movement in Arabidopsis thaliana. Proc. Natl. Acad. Sci. USA 107, 8860–8865. 13. Suetsugu, N., Kagawa, T., and Wada, M. (2005) An auxilin-like J-domain protein, JAC1,

regulates phototropin-mediated chloroplast movement in Arabidopsis. Plant Physiol. 139, 151–162. 14. Kadota, A., and Furuya, M. (1977) Apical growth of protonemata in Adiantum capillusveneris. I. Red far-red reversible effect on growth cessation in the dark. Dev. Growth Differ. 19, 357–365. 15. Yatsuhashi, H., and Wada, M. (1990) Highfluence rate responses in the light-oriented chloroplast movement in Adiantum protonemata. Plant Sci. 68, 87–94. 16. Kagawa, T., Sakai, T., Suetsugu, N., Oikawa, K., Ishiguro, S., Kato, T., Tabata, S., Okada, K., and Wada, M. (2001) Arabidopsis NPL1: a phototropin homolog controlling the chloroplast high-light avoidance response. Science 291, 2138–2141. 17. Sakai, T., Kagawa, T., Kasahara, M., Swartz, T. E., Christie, J. M., Briggs, W. R., Wada, M., and Okada, K. (2001) Arabidopsis nph1 and npl1: blue light receptors that mediate both phototropism and chloroplast relocation. Proc. Natl. Acad. Sci. USA 98, 6969–6974. 18. Hangarter, R. P., and Gest, H. (2004) Pictorial demonstrations of photosynthesis. Photosynth. Res. 80, 421–425. 19. Wada, M. (2007) The fern as a model system to study photomorphogenesis. J. Plant Res. 120, 3–16. 20. Kasahara, M., Kagawa, T., Sato, Y., Kiyosue, T., and Wada, M. (2004) Phototropins mediate blue and red light-induced chloroplast movements in Physcomitrella patens. Plant Physiol. 135, 1388–1397. 21. DeBlasio, S. L., Luesse, D. L., and Hangarter, R. P. (2005) A plant-specific protein essential for blue-light-induced chloroplast movements. Plant Physiol. 139, 101–114. 22. Gabryś, H., and Walczak, T. (1980) Photometric study of chloroplast phototranslocation in leaves of land plant. Acta Physiol. Plant. 2, 281–290. 23. Kagawa, T., and Wada, M. (2000) Blue lightinduced chloroplast relocation in Arabidopsis thaliana as analyzed by microbeam irradiation. Plant Cell Physiol. 41, 84–93.

Chapter 7 Studying Starch Content and Sedimentation of Amyloplast Statoliths in Arabidopsis Roots John Stanga, Allison Strohm, and Patrick H. Masson Abstract Amyloplasts, organelles responsible for the synthesis and storage of starch, are of critical importance to gravitropism in higher plants. We discuss two methods that are useful for describing the histology and behavior of amyloplasts. First, because mutants with little or no plastidic starch accumulation are defective in their gravitropic response, we review a method to observe starch accumulation quickly in plant tissue. Second, we discuss a method for measuring amyloplast sedimentation in the dynamic environment of Arabidopsis root columella cells, which is thought to provide a directional cue to a reoriented plant. Key words: Amyloplast, Sedimentation, Statolith, Gravitropism, Starch staining

1. Introduction Plant gravitropism is a research subject with a long and rich history (for reviews, see refs. 1, 2). Amyloplasts are starch-filled plastids whose movements are necessary for a normal gravitropic response (3). Because the sensitivity of the gravitropic response directly correlates with starch accumulation (4, 5), we first briefly revisit a commonly used starch staining method that can be used to assess quickly and qualitatively the presence or absence of starch in root statocytes. Upon reorientation of the plant, the amyloplasts sediment toward the new bottom of the gravity-sensing cells, called statocytes. Because both the sedimentation and saltation of amyloplasts have been proposed to contribute to gravitropism, it is important to understand the nature of amyloplast movement (6, 7). Much research into the positioning of amyloplasts in the moments following gravistimulation is based on tissues that have been fixed,

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sectioned, and analyzed (e.g., ref. 8). While these experiments do allow for very detailed measurements, they do not always provide adequate insight into the dynamic environment of a gravistimulated statocyte, as each individual root can only represent a single time point. There is ongoing interest in using in  vivo imaging experiments to describe amyloplast movements in both roots and shoots of various plants (e.g., refs. 9, 10). Here, we describe a method for observing and quantifying the bulk sedimentation of amyloplasts immediately following gravistimulation in a living Arabidopsis root (11). Because much of the current work on gravitropism uses Arabidopsis, these protocols were developed specifically for this model organism. They are particularly useful for characterizing gravitropic mutants to determine if their amyloplasts contain normal amounts of starch and sediment with normal kinetics. These protocols are also likely to work well for studies on other plant species with minimal modifications.

2. Materials 2.1. Iodine Staining for Starch Content

1. Seed sterilization solution: 20% (v/v) bleach and 0.01% (w/v) sodium dodecyl sulfate (SDS). 2. Growth medium (GM): half-strength Linsmaier and Skoog salts with macro- and micronutrients, vitamins, and 30  g/L sucrose, pH 5.7 (Caisson Laboratories, UT, USA), and 1% (w/v) type E agar (see Note 1). 3. Lugol stain: 10% (w/v) potassium iodide (KI) and 5% (w/v) iodine (I). 4. Chloral hydrate, 2.5 g/mL (see Note 2). 5. Other supplies and equipment: laminar flow hood, Petri plates, micropore tape (3M), aluminum foil, growth chamber, microscope slides, coverslips, and light microscope.

2.2. In Vivo Analysis of Amyloplast Sedimentation

1. Seed sterilization solution (see Subheading 2.1). 2. Growth medium (see Subheading  2.1), containing 0.75% (w/v) agar. 3. Microscope cartridges (see Note 3). 4. Light microscope equipped with a vertical rotatable stage and a camera (see Note 4). An example setup is shown in Fig. 1. 5. Computer with image capture software and image analysis software (see Note 5). 6. Other supplies and equipment: laminar flow hood, sterile 60 × 24 mm rectangular #1 or #1 1/2 coverslips (see Note 6), Petri plates, Parafilm, vacuum grease, and timer.

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Fig. 1. Example horizontal microscope setup. Our setup consists of a light microscope positioned on its side so that the rotatable stage is vertical. The microscope is connected to a camera and a computer for image acquisition.

3. Methods 3.1. Iodine Staining for Starch Content (see Note 7)

1. Pour molten growth medium into the Petri plates in a laminar flow hood and allow it to harden. Sterilize the Arabidopsis seeds for 10  min in the seed sterilization solution and then rinse them four times with water. Then place individual seeds on the surface of the growth medium. Seal the plates with micropore tape, wrap them in aluminum foil, and place them at 4°C in the dark for 48 h. 2. Remove the foil and place the plates upright in a growth chamber. Allow the seedlings to grow to the desired age (about 5 days). 3. Mount the roots on a slide with water and place a coverslip over the roots. Stain the roots briefly (about 1 min) with 1% Lugol solution (see Note 8). 4. Clear the sample with a drop of chloral hydrate (see Note 9). Observe the roots under a light microscope. Starch grains in amyloplasts will appear dark blue over a light brown background. An example of typical results is shown in Fig. 2.

3.2. In Vivo Analysis of Amyloplast Sedimentation

1. Sterilize the Arabidopsis seeds for 10 min in the seed sterilization solution and then rinse them four times with water. Keep the seeds in water in the dark at 4°C for 2–3 days (see Note 10).

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Fig.  2. Typical images illustrating starch staining of amyloplasts. Dark blue staining indicates the presence of starch in an Arabidopsis root tip that was stained with 1% Lugol solution for about 1  min. The arrow points to one of the stained columella cells. This method allows the observer to analyze quickly and qualitatively the presence or absence of starch in the columella cells, the approximate number of amyloplasts in a cell, and which cells contain amyloplasts.

2. In a laminar flow hood, add about 500 mL of molten growth medium to the center of each of several sterile coverslips. Once the growth medium has solidified, place about five seeds in a single row across each coverslip width-wise. The seeds should be placed directly on the coverslip and touch the edge of the growth medium. Place the coverslips in a Petri plate, seal the plate with Parafilm, and place the plate upright in a growth chamber (see Note 11). The plate should be positioned so that the growth medium is below the seeds, allowing the roots to grow downward between the coverslip and the medium. Let the seedlings grow for 4–7 days. 3. Prepare the microscope cartridges by adding a thin layer of vacuum grease to the perimeter of the opening and by adhering a blank coverslip to one side. In the same way, adhere a coverslip with 4–7-day-old roots to the other side so that the seedlings are within the hollow space. Press gently around the perimeter to create an airtight seal. Sterile conditions are not necessary for this step. The setup is shown in Fig. 3.

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Fig. 3. Cartridge assembly used for measurement of amyloplast sedimentation kinetics. Seeds are placed on a coverslip so that they touch a layer of growth medium. After 4–7 days of growth, a root that is growing underneath the growth medium and directly on top of the coverslip is chosen for analysis. The coverslip carrying the seedling is placed on one side of the cartridge so that the seedling’s root is in the hollow space formed by the cartridge and the coverslip. A blank coverslip is placed on the other side of the cartridge, and the apparatus is sealed with vacuum grease.

4. Identify a root that is growing underneath the growth medium and against the coverslip, and orient it vertically on the microscope stage. Allow it to remain vertical for 20–30 min before gravistimulation (see Note 12). 5. Set the camera to take a picture every 10 s for at least 10 min. As quickly as possible, begin a timer, rotate the stage 90°, find the root in the field of vision, refocus the microscope if necessary, and begin sequential image acquisition (see Note 13). Example images are shown in Fig. 4. 6. Combine each set of images into a single stack. Use NIH ImageJ software, including the object tracker and click forward macros, to measure a landmark (such as the corner of the columella cell being measured) that is visible throughout the stack and can be used as a reference point (see Note 14). The landmark measured is shown in Fig. 4. 7. Repeat the measurement as in step 6 for the leading edge of the amyloplast farthest from the cell membrane in each of the x and y dimensions (see Note 15). The amyloplast coordinates measured are shown in Fig. 4. 8. Subtract the individual amyloplast coordinates and the landmark coordinates for each frame of the stack to provide a relative distance in pixels. Convert the distance to microns by measuring an object of known length, such as a microscope calibration slide, in pixels. 9. Use a Student’s t-test at each time point to compare the experimental data with the controls (see Note 16).

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Fig. 4. Example images and analysis of amyloplast sedimentation. (a) Following 90° reorientation of a seedling, the amyloplasts within a central S2 columella cell sediment over time. In this example, amyloplasts from wild-type (Ws) and mar2-1 arg1-2 plants sediment with similar kinetics, while ACG20 amyloplasts sediment more slowly. (b) The distance from the former cell bottom to the leading plastid was measured for each time point on the x-axis. (c) The distance from the new cell bottom to the top-most plastid was measured for each time point on the y-axis. Error bars represent the standard error, and stars and crosses represent significant t-test values (P 8, then repeat the neutralization step. 5. Cross-link the DNA to the membrane using UV exposure. 6. Label a cpDNA probe (see Note 6) using a commercially available alkaline phosphatase direct or indirect labeling system and chemiluminescence detection. 7. Detect the hybridization signal by exposing the hybridized blot to X-ray film. We recommend making several different exposures. Choose the exposure where the weakest signal can be detected without overexposing the strongest signal. A densitometer or imaging software program can be used to quantify the hybridization signal. See Fig. 2 for an image of A. thaliana cpDNA after PFGE and blot hybridization. 8. The signals from each band should appear as distinct peaks, although sometimes the peaks corresponding to the genomic oligomers are partially obscured by the background smear of cpDNA fragments for Arabidopsis. 3.3. The Analysis of cpDNA as a Fraction of Total Tissue DNA

The methods described in this section provide information about the relative proportion of total cellular DNA that is represented by cpDNA. Because these methods do not require prior chloroplast isolation, they can be performed quickly and easily. However, this also limits the type of information that one can obtain because the measured proportion of cpDNA is an average of different cell types and only some of the cells within a tissue contain plastids that develop into chloroplasts. Another limitation of the method described in Subheading 3.3.3 is that the proportion of cpDNA measured can be artifactually inflated due to the presence of chloroplast-derived sequences that are integrated into the nuclear genome. Additionally, both of these methods provide only the ratio of chloroplast DNA to nuclear DNA, which can change as the result of a change in the amount of either one of these DNAs. If the nuclear DNA amount is constant from cell to cell and throughout development, then a change in the ratio can be ascribed to a change in the amount of cpDNA and these methods are reliable indicators of the amount of cpDNA. Otherwise, these

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methods can only report the relative proportion of total DNA represented by cpDNA. It is important to consider this difference if the goal of the experiment is to measure the amount of cpDNA. For Arabidopsis researchers who want to measure the amount of cpDNA, we recommend using one or more of the techniques described in Subheadings 3.1 and 3.2, instead of the techniques described in the following section, because Arabidopsis is a species for which nuclear DNA amounts are not constant during leaf development. For species with a constant amount of nuclear DNA, this method can be used to assess the total amount of cpDNA (9, 10). 3.3.1. CTAB Method for Isolating Total Tissue DNA from Plants

1. Place 50–100  mg of leaf tissue in a 1.5-mL microcentrifuge tube. The tissue can be frozen in liquid nitrogen and disrupted by grinding the tissue in the microcentrifuge tube using a small pestle. 2. Add 250 mL of CTAB solution. Mix by inverting the tube several times and incubate at 60–65°C for 30 min. 3. Add 250 mL of 24:1 chloroform:isoamyl alcohol and mix by inverting the tube several times. 4. Centrifuge at 12,000–16,000 × g for 5–10  min in a microcentrifuge. 5. Transfer 220 mL of the upper phase to a new microcentrifuge tube containing 150 mL of isopropanol. 6. Incubate for 5 min at room temperature. 7. Centrifuge at 12,000–16,000 × g for 10 min. 8. Remove supernatant and wash the pellet with 300 mL of 70% ethanol. Centrifuge at 12,000–16,000 × g for 10 min. 9. Air-dry the DNA pellet for 10 min. 10. Resuspend the DNA pellet in 50  mL of water or TE, pH 8. Store the DNA temporarily at 4°C or at −20°C for longer periods of time.

3.3.2. Quantifying cpDNA by Blot Hybridization of Restriction-Digested Total Tissue DNA

1. Isolate total tissue DNA using a commercially available kit or by the CTAB method provided in Subheading 3.3.1. 2. Digest the DNA using a restriction enzyme that will generate a fragment that will hybridize to a cpDNA probe of your choice and that is of the appropriate size for analysis by conventional agarose gel electrophoresis. You should choose a cpDNA probe that will bind to a fragment size between 300 and 1,200 bp for optimal resolution on an agarose gel. 3. Perform agarose gel electrophoresis and blot hybridization (see Subheading 3.2.4) using a probe that is designed to hybridize to a cpDNA (or nuclear DNA) fragment of a known length.

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4. Take several different X-ray film exposures of the blot. Choose the exposure time that can detect the weakest signal without overexposing the strongest signal. 5. Use an imaging software program or densitometer to compare the hybridization signal from DNA samples on the same blot. 3.3.3. Measuring the Ratio of cpDNA to Nuclear DNA Using Real-Time Quantitative PCR

1. Design primers to amplify a 100–200-bp fragment of cpDNA. Test the primers to make sure that they amplify with at least 90% efficiency (see Note 5). Design and test primers that amplify a fragment of similar size from the nuclear genome. 2. Isolate total tissue DNA using a commercially available kit or by the CTAB method (see Subheading 3.3.1). 3. Perform PCRs in 20–25 mL volume with total tissue DNA as template and containing 0.12  mM primers, 0.2  mM dNTPs, 4.5 mM MgCl2, 0.25 mg/mL SYBR Green, and 1 unit of Taq polymerase or a commercially available SYBR Green PCR mastermix (see Note 7). 4. Set the background subtraction and establish the threshold manually. Ideally, the threshold should intersect the reaction amplification curves during the log-linear phase of the reaction. 5. Record the cycle at which the reaction amplification curve crosses the threshold (Ct). 6. Calculate an average of the Ct values for the nuclear reactions for each sample. Subtract the chloroplast Ct value for each replicate for each sample from the average nuclear Ct value (x). Calculate the ratio of chloroplast copies to the number of nuclear DNA copies (R) using the equation R = 2x. Then, obtain the mean and standard deviation among the replicates for each sample.

4. Notes 1. We find no difference in DNA amount for chloroplasts prepared using the step gradient and those prepared using only the 70% Percoll solution. 2. Glutaraldehyde can cause headaches, dizziness, and severe irritation of the lungs, eyes, nose, and throat. Wear gloves and work in a fume hood when using this fixative. 3. Calculation of DNA contents of chloroplasts using DAPI and virus particles of known DNA content. We illustrate the process here with Vaccinia virus. The number of chloroplast genome equivalents per plastid is calculated using the following equation: chloroplast genome equivalents = 1.33 V (where V = the DAPI-DNA Rfl of the plastid divided by the mean Rfl

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of Vaccinia virus particles). The value 1.33 is a constant that accounts for the differences between the size and base composition between the Arabidopsis chloroplast genome and the Vaccinia virus genome, and is determined as (% AT content of Vaccinia virus genome/% AT content of Arabidopsis chloroplast genome) × (number of bp of Vaccinia virus genome/ number of bp of Arabidopsis chloroplast genome), where % AT for Vaccinia (Copenhagen strain) is 66.6, % AT for Arabidopsis cpDNA is 64%, number of bp for Vaccinia DNA is 197,361, and number of bp for Arabidopsis cpDNA is 154,361. 4. We recommend at least three technical replicates per run for each of the standards and each of the samples. Running a replicate plate is desirable in order to adjust for run-to-run variability and plate position effects. Similarly, for relative quantification, reactions with chloroplast and nuclear primers should each be performed in triplicate for each run and each triplicate sample should be run at least twice. We recommend including a set of biological replicates to adjust for subtle variations in growth conditions that may affect the results. 5. In order to determine the efficiency of amplification, plot the log of the concentration of DNA as a function of the Ct value and fit a linear regression to the points. If the reactions are 90% efficient (meaning that the amplification results in at least a 1.9-fold increase in the amount of DNA per cycle) and the replicates are consistent, then the R2 value should be >0.99 and the slope should be between −0.278 and −0.301. If you do not know the absolute concentration of the DNA to be analyzed (e.g., with the relative quantification method described in Subheading 2.3.3), then prepare a twofold dilution series of DNA. You can then use the Ct values in a similar manner, using mock concentrations in twofold increments to generate a similar plot and linear regression. 6. There are many available commercial kits for labeling DNA to be used as probes. We recommend following the manufacturer’s directions to label and hybridize the probe to the membrane. 7. If preparing your own SYBR Green PCR amplification mixture, we recommend preparing a mastermix for use among experiments.

Acknowledgments The authors thank Dr. Delene J. Oldenburg for her contributions toward developing many of the techniques described in this chapter and for critically reading the manuscript.

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References 1. Lopez-Juez E., and Pyke K. A. (2005) Plastids unleashed: their development and their integration in plant development. Int. J. Dev. Biol. 49, 557–577. 2. Waters M. T., and Langdale J. A. (2009) The making of a chloroplast. EMBO J. 28, 2861–2873. 3. Fujie M., Kuroiwa H., Kawano S., Mutoh S., and Kuroiwa T. (1994) Behavior of organelles and their nucleoids in the shoot apical meristem during leaf development in Arabidopsis thaliana L. Planta 194, 395–405. 4. Kuroiwa T, Suszuki T, Ogawa K, and Kawano S. (1981) The chloroplast nucleus: distribution, number, size, and shape, and a model for the multiplication of the chloroplast genome during chloroplast development. Plant Cell Physiol. 22, 381–396. 5. Oldenburg D. J., and Bendich A. J. (2004) Changes in the structure of DNA molecules and the amount of DNA per plastid during chloroplast development in maize. J. Mol. Biol. 344, 1311–1330.

6. Rowan B. A., Oldenburg D. J., and Bendich A. J. (2004) The demise of chloroplast DNA in Arabidopsis. Curr. Genet. 46, 176–181. 7. Rowan B. A., Oldenburg D. J., and Bendich A. J. (2009) A multiple-method approach reveals a declining amount of chloroplast DNA during development in Arabidopsis. BMC Plant Biol. 9, 3. 8. Oldenburg D. J., and Bendich A. J. (2009) Chloroplasts. In, Molecular Genetic Approaches to Maize Improvement (Kriz A. L., and Larkins B. A., eds.) Springer-Verlag, Heidelberg, Germany, pp. 325–343. 9. Oldenburg D. J., Rowan B. A., Zhao L., Walcher C. L., Schleh M., and Bendich A. J. (2006) Loss or retention of chloroplast DNA in maize seedlings is affected by both light and genotype. Planta 225, 41–55. 10. Shaver J. M., Oldenburg D. J., and Bendich A. J. (2008) The structure of chloroplast DNA molecules and the effects of light on the amount of chloroplast DNA during development in Medicago truncatula. Plant Physiol. 146, 1064–1074.

Chapter 11 Measurement of Transcription Rates in Arabidopsis Chloroplasts Yan O. Zubo, Thomas Börner, and Karsten Liere Abstract The regulation of gene expression is still one of the major issues in modern plant molecular biology. The amount of RNA in a cell is regulated by both transcriptional and posttranscriptional events. Methods to determine these steady-state levels of RNAs, such as Northern analysis, ribonuclease protection assay (RPA), and quantitative real-time PCR, do not discriminate between regulation by de novo RNA synthesis and the influence by degradation or stabilization. To assess the rate of transcription of individual genes, run-on transcription is utilized. To this end, isolated chloroplasts are used in brief in vitro transcription reactions in the presence of radiolabeled nucleotides, with a subsequent hybridization of the isolated RNA with DNA fragments spotted on membranes. Here, we describe a protocol for run-on transcription in chloroplasts isolated from Arabidopsis leaves and present data on the transcriptional activity of several plastid genes in detached leaves of different Arabidopsis ecotypes. Key words: Arabidopsis thaliana, Chloroplast, Gene expression, Run-on assay, Transcription rate, Plastid isolation, DNA/RNA hybridization

1. Introduction Chloroplasts originate from cyanobacteria that were engulfed as endosymbionts by a host cell, the ancestor of algae and plants. During evolution, most genes of the ancestral cyanobacterial genome have been lost or transferred into the nucleus. In case of higher plants, the extant plastid genome (plastome) harbors only about 120–140 genes for products that function primarily in photosynthesis and gene expression (1). However, the core subunits of a eubacteria-type RNA polymerase are still encoded by the plastid rpoA, rpoB, rpoC1, and C2 genes on the plastid genome. Nuclearencoded s-factors complement this plastid-encoded plastid RNA polymerase (PEP) to the functional holoenzyme (2, 3). Moreover, R. Paul Jarvis (ed.), Chloroplast Research in Arabidopsis: Methods and Protocols, Volume I, Methods in Molecular Biology, vol. 774, DOI 10.1007/978-1-61779-234-2_11, © Springer Science+Business Media, LLC 2011

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several PEP-associated factors were isolated that are thought to be involved in regulating plastid transcription (4–7). Although resulting in an albino phenotype and impaired photosynthesis, lack of the PEP still allows for heterotrophic growth and transcription of plastid genes (8–10). This is explained by the existence of a second, plastid-localized, nuclear-encoded transcription activity (nuclearencoded plastid RNA polymerase, NEP) that supplements PEP to transcribe the genes encoded in the plastome fully (11, 12). The NEP activity is represented by nuclear-encoded, phage-type RNA polymerases (13–20). Aside from mitochondrial targeting (RpoTm), it was shown that a second RNA polymerase of bacteriophage T3/T7 type (RpoT) enzyme is targeted into plastids both in monocots and dicots (RpoTp) (14, 15, 21–27). Furthermore, a third RpoT polymerase found exclusively in eudicots (RpoTmp) has been shown to be dually targeted into both mitochondria and plastids of Arabidopsis and Nicotiana (21, 25, 28). A broad set of methods used for the examination of RNA steady-state levels allow one to estimate the length and abundance of transcripts (Northern hybridization), quantify low abundant transcripts (real-time qPCR), determine 3¢- or 5¢-end positions (3¢- or 5¢-RACE, RNase protection assay, primer extension analysis), determine transcription initiation sites (in  vitro capping assay; tobacco acid pyrophosphatase-based 5¢-RACE) (29), and provide information on the entire transcriptome (microarray) (30, 31). However, to evaluate the rate of individual gene transcription directly, run-on transcription is the method of choice: transcripts are labeled by adding radiolabeled nucleotides during a brief time of incubation and subsequently analyzed by dot-blot hybridization (30). This technique applies to all genome-containing compartments such as bacterial cells, nuclei, mitochondria, and chloroplasts. In general, chloroplast isolation is a simple and quick procedure which does not require special equipment (32, 33). Since the transcription of most chloroplast genes is comparatively strong and the plastome codes only for about 120–140 genes (34), it is feasible even to determine the transcription rates of all plastid genes in one single experiment (10). However, this method also has some limitations. In contrast to more expensive single-stranded DNA oligonucleotides, the commonly used double-stranded PCR probes are able to hybridize with both sense and antisense transcripts. This is especially significant for transcription of chloroplast genes, since antisense transcripts (of unknown size and function) are generated during plastome transcription (10, 35). Various ecotypes of Arabidopsis thaliana are found in a broad geographic distribution around the world. To demonstrate the method of run-on-transcription and to test if there are differences in the rate of plastid transcription among the Arabidopsis ecotypes Columbia (Col-0), Landsberg erecta (Ler-0), and Wassilewskija (Ws), we performed run-on analyses with chloroplasts isolated from plants grown under the same conditions.

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2. Materials 2.1. Dot- or Slot-Blot Preparation

1. 10 M NaOH. 2. Nylon membrane Hybond N+ (Amersham Pharmacia Biotech, UK) and 3MM chromatography paper (Whatman, Maidstone, UK). 3. 2× SSC: 0.03 M sodium citrate and 0.3 M NaCl, pH 7.0. Store at +4°C. 4. Dot-blot apparatus (e.g., Bio-Dot™ apparatus, Bio-Rad, USA). 5. Ultraviolet chamber (e.g., GS Gene Linker™, Bio-Rad).

2.2. Chloroplast Isolation

1. Homogenization buffer: 0.33  M sorbitol, 50  mM TricineKOH, pH 8.0, 2 mM ethylenediaminetetraacetic acid (EDTA), and 5  mM 2-mercaptoethanol. Freshly prepare the buffer before starting the experiment. 2. Percoll buffer: Homogenization buffer plus 3% (w/v) polyethylene glycol, 0.5% (w/v) bovine serum albumin (BSA), and 0.5% (w/v) Ficoll 400. Store at −20°C. 3. RNase inhibitor (e.g., RiboBlock; Fermentas, Lithuania). 4. Miracloth (Calbiochem-Behring, USA). 5. Plant material: Grow Arabidopsis seedlings for 12 days under long-day conditions (16 h/8 h) at 22°C.

2.3. In Vitro Transcription, Hybridization, and Washing of Membrane

1. Transcription buffer: 50  mM Tris–HCl, pH 8.0, 10  mM MgCl2, 0.2  mM CTP, GTP and ATP, 0.01  mM UTP, and 10 mM 2-mercaptoethanol. Store at −20°C. 2. Resuspension buffer: 50  mM Tris–HCl, pH 7.0, 10  mM MgCl2, 10 mM KCl, and 4 mM 2-mercaptoethanol. Store at −20°C. 3. [a-32P]-UTP (3,000  Ci/mmol; Perkin-Elmer LAS GmbH, Germany). 4. Stop buffer: 5% (w/v) Na-lauroylsarcosine (Sigma, USA), 50  mM Tris–HCl, pH 8.0, and 25  mM EDTA. Store at −20°C. 5. Aqua-Roti-Phenol (pH 4.5–5) for RNA isolation (Carl Roth GmbH, Germany) and chloroform (Carl Roth GmbH). 6. 3 M sodium acetate, pH 6.0. 7. Yeast tRNA 10 mg/mL (Ambion, USA). 8. 96% and 75% (v/v) ethanol. 9. Hybridization buffer: 250 mM Na2HPO4, pH 7.2, 7% (w/v) sodium dodecyl sulfate (SDS), and 2.5 mM EDTA, pH 8.0. Store at room temperature.

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10. Washing solutions: High-salt buffer (0.5× SSC and 0.1% [w/v] SDS) and low-salt buffer (0.2× SSC and 0.1% [w/v] SDS). Store at room temperature. 2.4. Signal Detection and Stripping of Dot-Blots

1. Saran Wrap (ScienceLab, USA). 2. Amersham Hyperfilm™ MP autoradiography film (Amersham Pharmacia Biotech) or Imaging Screen K and Molecular Imager® FX system (Bio-Rad). 3. Stripping solution: 0.1% (w/v) SDS and 5  mM EDTA, pH 8.0.

3. Methods Run-on assays consist of four steps: (1) dot-blot preparation, (2) chloroplast isolation, (3) in vitro transcription and hybridization to the dot-blot membrane, and (4) signal detection and subsequent stripping of the dot-blots. The dot-blot membranes can be stored for a long time and reused after stripping for up to ten times. After extraction, chloroplasts can be stored in homogenization buffer at 4°C overnight. However, the transcription rates will decline after such storage. Therefore, it is preferable to perform the run-on transcription on the same day as the chloroplast extraction. Most transcripts are already initiated and partially elongated before the chloroplast isolation and are further elongated or finished during the transcription reaction (36). Although initiation of transcription takes place in vitro as well, the impact of these transcripts on the overall transcriptional rates is lesser than that of elongated RNAs. A limited period of time of the transcription reaction is necessary to minimize the influence of processes, leading to transcript degradation and/or stability (36). To detect the freshly transcribed RNAs, radioactively labeled UTP ([a-32P]-UTP) is routinely used. At present, there is no chance to use nonradioactive labeling techniques such as fluorescence-labeled nucleotides, since the endogenous plastid RNA polymerases are not capable of processing these nucleotides. 3.1. Preparation of Dot-Blot Membranes

1. Produce DNA fragments of genes of your interest by PCR. At least two dots per gene should be spotted on the membrane, with each dot containing 1  mg DNA (see Note 1). Although it is not essential to remove the components of the PCR before spotting the DNA fragments onto the membrane, further purification of the DNA fragments results in more defined and sharper spots. The size of the PCR fragments may differ in most cases (see Note 2), but should be in the size range between 200 and 1,000 bp.

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2. DNA fragment denaturation. The DNA fragments are denaturated by NaOH (final concentration is 0.5 M) and heating (95°C for 10 min). The volume of the denaturated DNA solution should be adjusted with distilled water to 100 mL per dot if using the Bio-Dot™ apparatus. After heating, immediately transfer the tubes into ice to prevent DNA renaturation. Briefly centrifuge the tubes in a microfuge after cooling to collect any condensate. 3. Prepare the dot-blot apparatus according to the manufacturer’s protocol and load the DNA fragments onto the Nylon membrane according to a previously designed scheme (Fig. 1a). 4. After drying the membrane at room temperature for 5  min, cross-link DNA and membrane by UV irradiation (37). a accD

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Fig. 1. Transcription activities of several chloroplast genes in different Arabidopsis ecotypes. Chloroplasts were isolated from 12-day-old Arabidopsis rosette leaves of the ecotypes Landsberg erecta (b), Wassilewskija (c), and Columbia (d) and used for run-on transcription assays. The [32P]-labeled transcripts were isolated and hybridized to plastid gene probes blotted on Nylon membrane according to the scheme presented in (a). The transcription activity of plastid genes of the ecotypes tested is under these conditions highly similar. Therefore, the differences in the nuclear backgrounds of these ecotypes do not influence the transcription activity of their plastid genes. Note: representative autoradiograms are shown in (b), (c), and (d). To assess data, multiple experiments of at least two biological and two technical repetitions should be carried out.

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5. For neutralization and removal of components of the PCR (if applicable), wash the membrane for 5 min at room temperature in 2× SSC buffer. Dried membranes can be store at room temperature for months until being used in hybridization experiments. 3.2. Chloroplast Isolation

1. Before isolating the chloroplasts, Percoll step gradients (40/70%) should be prepared in, e.g., 15-mL Falcon or similar transparent tubes (Corex). A volume of 3  mL of the 40% Percoll solution should be gently and accurately loaded on top of 70% Percoll solution (3 mL; see Note 3). Gradients should be precooled before loading chloroplast suspension at 4°C. 2. Homogenize the Arabidopsis leaf material (3–10  g) briefly (2–5 s) in a Warring blender in 80-mL ice-cold homogenization buffer (see Note 4). 3. To remove any cell debris, the homogenate is filtered through two layers of Miracloth into a fresh beaker and centrifuged at 3,500 × g for 5 min (see Note 5). 4. Resuspend the pellet carefully in a small volume of homogenization buffer (1  mL buffer per up to 10  g of initial plant material) by swirling or with the help of a small, round- or rigger-tip paintbrush. 5. Load 3–5 mL of the chloroplast suspension onto Percoll step gradients (40/70%) and centrifuge in a swing-out rotor at 6,000 × g for 30  min (break off). After centrifugation, intact chloroplasts are found at the border of the 40% and 70% Percoll layers, while broken chloroplasts are retained on top of the 40% Percoll layer. 6. Remove the liquid above the band of intact chloroplasts. Transfer the chloroplast band to a clean beaker using a 1-mL Gilson pipette tip and cut at the end (wide bore). 7. To remove the Percoll, add 50 mL of homogenization buffer to the isolated chloroplasts and then invert the tube carefully one time to mix and wash off the Percoll. Centrifuge the chloroplasts in a swing-out rotor at 3,500 × g for 5 min (break on). 8. Decant and discard the supernatant, and subsequently resuspend the chloroplast pellet carefully in 1 mL of homogenization buffer by swirling or with the help of a small, round- or rigger-tip paintbrush. Transfer the homogenate into a clean 2-mL reaction tube. Finally, determine the number of isolated chloroplasts microscopically using a Rosenthal–Fuchs hemocytometer (see Note 6).

3.3. Run-On Transcription and Hybridization to Dot-Blot Membranes

1. Transfer aliquots containing 106–107 chloroplasts to a new 1.5-mL reaction tube and sediment by centrifugation at 3,500 × g for 4 min. Carefully remove the supernatant and add 50 mL of chloroplast resuspension buffer (see Note 7).

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2. To start the in  vitro transcription reaction, add 20  units of RNase inhibitor, 50 mL of transcription buffer, and 50 mCi of [a-32P]-UTP. Mix the reaction by vortexing or pipetting up and down (see Note 8). 3. Immediately transfer the tubes into a preheated water bath or incubation block and incubate at 25°C for 3–5 min (see Note 9). 4. Transfer the reaction tubes back on ice and add 1 volume of stop buffer (100  mL) to stop the transcription reaction. Mix the reaction by vortexing. 5. To perform deproteinization, add 200 mL of phenol (pH 4.5–5) to the reaction and mix by vortexing for 30 s. Centrifuge at maximum speed for 5  min and subsequently transfer the watery phase into a fresh tube. 6. Repeat step 5 followed by chloroform extraction by adding 1 volume of chloroform (200 mL). Centrifuge at maximum speed for 5  min and subsequently transfer the watery phase into a fresh reaction tube. 7. Add 1/10 volume 3 M sodium acetate and 2.5 volume of 96% ethanol. The addition of 2 mL of yeast tRNA (10 mg/mL) will increase the precipitation efficiency (see Note 10). Mix the reaction by vortexing and incubate for 30 min at −20°C. 8. Centrifuge the tubes at maximum speed for 30 min at 4°C. 9. During the centrifugation step, prepare the membrane for hybridization. Before hybridizing the dot-blots with RNA extracted from the in  vitro transcription reaction, a prehybridization is performed. Transfer the membrane into a hybridization tube, add an appropriate amount of hybridization buffer (see Note 11), and incubate with rotation in a hybridization oven for at least 1 h at 58°C. 10. Discard the supernatant from the precipitation reaction (steps 7 and 8) and wash the RNA pellet by adding 1 mL of 75–80% ethanol. Centrifuge the reaction tube at maximum speed for 5 min at 4°C. 11. Subsequently discard the supernatant, briefly air-dry the RNA pellet (5 min, do not over dry), and redissolve in 100 mL of RNase-free water. 12. Denature the RNA solution by heating at 95°C for 3 min and stop the denaturation reaction by immediately transferring the reaction tubes onto ice. 13. Now discard the pre-hybridization solution from the hybridization tubes and add an appropriate amount of fresh, prewarmed hybridization solution (58°C; see Note 12). Pipette the denatured RNA solution into the hybridization tube and continue to incubate with rotation overnight at 58°C.

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14. After hybridization, wash the membrane twice for 10 min with pre-warmed (58°C) high-salt washing buffer at 58°C, and at least once for 10 min with pre-warmed low-salt washing buffer (each with about 300–500 mL). Monitor the radioactivity of the membrane after each washing step and stop if the membrane counts are significantly, but not exceedingly higher than the counts of the environmental background. 3.4. Signal Detection and Reusing of Membrane

1. Wrap the membrane with Saran wrap or put it in a plastic transparent but water-impermeable bag, making sure that there are no wrinkles. Expose to autoradiography film overnight or to a Molecular Imager screen for 1 h. However, the exposure time is dependent on the intensity of radioactive signals and may take from a few hours until several days (usually overnight exposure is convenient). Adjust the exposure time accordingly. Analysis and quantification of the signals are performed by available software (e.g., Quantity One, Bio-Rad; see Note 13). Examples of obtained results are shown in Fig. 1b–d. 2. The membrane can be reused (usually about ten times). However, be aware that the stripping procedure has the potential to denature sensitive nucleic acids, rendering them unrecognizable by the probe. Furthermore, any physical defects of a membrane will be increased when stripped and reprobed. Boil the membrane in stripping solution for 20  min. The volume of the solution is dependent on the size and amount of membranes. About 500 mL of stripping solution should be used per up to 5 membranes 8 × 12 cm in size. To remove all bound RNA, repeat boiling of the membrane up to four times. Test for stripping efficiency by exposing to autoradiography film or Imaging Screen.

4. Notes 1. To compare the transcript levels of all genes and not to be limited during hybridization by the amount of the DNA loaded, it is necessary to have an excess of DNA molecules on the membrane. Initial experiments with various amounts of DNA fragments of strongly expressed genes such as psbA, rbcL, and rrn16 will show the limits of saturation of the system. 2. Ideally, length and/or G/C content of the gene fragments used in dot-blots should be very similar. However, DNA fragments of different sizes are still suitable to investigate the differences in transcriptional regulation of individual genes while comparing different treatments or mutants. 3. Intact chloroplasts from mature green leaves band between roplasts appear between the extraction buffer and 40%

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Percoll layer. To isolate plastids from etiolated or pigment-­ deficient mutant Arabidopsis plants, the percentage of the upper Percoll layer is lowered to 30 or even 20%. Note that lower percentages lead to contamination by cell debris including nuclei. 4. To perform run-on assays with chloroplasts from plant species other than Arabidopsis, some parameters of the chloroplast isolation method need to be adjusted. As mentioned for etiolated or pigment-deficient plastids in Note 3, the percentages of the Percoll gradient may need some changes to accommodate possible physical differences of the plastids isolated from other plant species. Similarly, the centrifugation forces used during chloroplast isolation may need to be optimized. 5. Brief (2–5 s) rounds of homogenization give an optimal yield of intact chloroplasts. If some leaves are not completely homogenized during the first blending step, it is possible to collect the filter cake from the Miracloth and to repeat the homogenization using 80 mL ice-cold homogenization buffer. 6. Counting the number of chloroplasts using a microscope and a Rosenthal–Fuchs hemocytometer is the most usual way to estimate the amount of organelles in a given preparation. However, if the number and size of chloroplasts significantly vary in the examined tissues, determination of the DNA or chlorophyll content of the chloroplasts may provide more accurate results. 7. Although placed on ice, some transcription may still occur if the necessary components are present. Since the osmotic power of the homogenization buffer leads to disruption of the chloroplasts, the addition of transcription buffer and [a-32P]-UTP immediately starts RNA synthesis (38). However, while the non-osmotic chloroplast resuspension buffer contains some components of the transcription system, the absence of ribonucleotides prevents RNA synthesis. Thus, isolated chloroplasts may be stored in resuspension buffer for about half an hour without any effect on transcriptional activity. 8. To prevent contamination of gloves and equipment, try to exclude contact between the radioactive solution and the lid of the reaction tube. Furthermore, using a pair of pliers to open the lid carefully helps to prevent the risk of contaminating the gloves by residual radioactive liquid on the lid’s inside. 9. Although the incubation time may be increased, one should keep in mind that the influence of RNA degradation or stabilization on the results will increase as well. Therefore, the time of incubation should not be longer than 10 min. 10. In case of contamination with nuclear DNA, the precipitate appears to be gel-like. Large amounts of nuclear DNA lead to strong nonspecific background on the membrane after hybridization. To remove the nuclear DNA contamination, perform

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an additional treatment with DNase I. To remove the degraded DNA and DNase I subsequently, deproteinize the sample by phenol/chloroform treatment. Even if a gel-like precipitate appears just in one sample, the DNase treatment needs to be performed with all samples. Contamination by chloroplast DNA does not cause problems during hybridization. 11. Although several protocols convert the pre-hybridization buffer into hybridization buffer by simply adding the probe, we recommend a complete buffer change. To remove components on the dot-blot possibly interfering with the hybridization efficiently, the membrane is washed with an excess of hybridization buffer during pre-hybridization. In the case of a 200-mL tube, use at least 50 mL of hybridization buffer. It is possible to pre-hybridize several membranes in one tube. 12. The volume of the hybridization tube and the amount of hybridization buffer depend on the size of the membranes used. In the case of using a membrane of 8 × 12 cm in size, a 100-mL tube and 7 mL of hybridization buffer are optimal. If smaller membranes are used, it is possible to perform the hybridization in Falcon™ tubes. Between 1 and 1.5  mL of hybridization buffer is added to a 15-mL tube, and 4–5 mL into a 50-mL tube. The Falcon™ tubes are subsequently placed into the standard hybridization tubes. The membranes are oriented with the DNA-side up inside the tube. They may overlap, however, only once. 13. Make sure to prevent drying-out of the membranes. There is no reasonable chance of successfully stripping dried membranes.

Acknowledgments This work was funded by Deutsche Forschungsgemeinschaft (SFB 429 to T.B. and K/L.). We thank Liliana Borsellino, Dr. Victor V. Kusnetsov, and Dr. Maria V. Yamburenko for their helpful discussions. References 1. Martin, W. (2003) Gene transfer from organelles to the nucleus: frequent and in big chunks. Proc. Natl. Acad. Sci. USA 100, 8612–8614. 2. Shiina, T., Tsunoyama, Y., Nakahira, Y., and Khan, M. S. (2005) Plastid RNA polymerases, promoters, and transcription regulators in higher plants. Int. Rev. Cytol. 244, 1–68.

3. Liere, K., and Börner, T. (2007) Transcription of plastid genes. In, Regulation of Transcription in Plants (Grasser, K. D., ed.) Blackwell Publishing, Oxford, UK, pp. 184–224. 4. Suzuki, J. Y., Jimmy Ytterberg, A., Beardslee, T. A., Allison, L. A., Wijk, K. J., and Maliga, P. (2004) Affinity purification of the tobacco

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Chapter 12 Studying the Structure and Processing of Chloroplast Transcripts Alice Barkan Abstract Most chloroplast genes in land plants are represented by multiple transcript isoforms that arise via differential splicing, endo- and exo-nucleolytic processing, and/or RNA editing. Exploration of the functional significance and mechanisms of these processing events is an active area of current research. This chapter focuses on methods that can be used to define the termini of chloroplast RNAs, quantify the relative levels of alternative processed RNA isoforms, and identify the binding sites of proteins that mediate chloroplast RNA processing. Various approaches for defining the sequence specificity of chloroplast RNA binding proteins are discussed, as are the parameters to consider in designing in vitro assays for RNA binding activities. A protocol is provided for a poisoned-primer extension assay for quantifying different splice isoforms. Key words: Chloroplast, Plastid, RNA processing, RNA binding protein, RNA binding assay

1. Introduction RNA populations in land plant chloroplasts are characterized by a remarkable complexity that was not foreshadowed in the chloroplast’s cyanobacterial ancestor (reviewed in ref. (1)). Most chloroplast genes are represented by many different transcript isoforms that arise through the differential processing of one or several primary transcripts. These processing events include the trimming of sequences at 5¢ and 3¢ termini, endonucleolytic cleavage, group I and group II intron splicing, and RNA editing. The mechanistic basis and functional significance of these processing events have been longstanding questions. The past decade has witnessed enormous progress in the identification of the protein factors that mediate various aspects of

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chloroplast RNA maturation. Advances have been achieved through genetic screens for mutants with defects in chloroplast gene expression, biochemical purification of RNA processing activities, and reverse genetic analysis of candidate genes. These approaches have revealed the chimeric nature of the chloroplast RNA processing machineries: conserved proteins of bacterial ancestry function in parallel with diverse RNA binding protein classes that emerged in the context of nuclear-organellar coevolution (1–4). The field is now poised to address the mechanisms employed by the many hundreds of RNA binding proteins in chloroplasts, and the roles of the RNA processing events in chloroplast gene expression and its regulation. This chapter provides an overview of techniques that can be used to map chloroplast transcript termini, quantify distinct transcript isoforms, and identify the binding sites of RNA binding proteins that mediate the processing events. Where possible, the reader is referred to protocols that have been published previously. A detailed protocol is provided for a poisoned-primer extension assay that is particularly useful for monitoring chloroplast RNA splicing. Methods for studying RNA editing are the focus of a different chapter in this book (see Chapter 13, Vol. 1) and are not addressed here. 1.1. Overview of Methods for Mapping and Quantifying Chloroplast Transcript Isoforms 1.1.1. N  orthern Blotting

1.1.2. Primer Extension

Northern blotting remains one of the most useful methods for displaying different transcript isoforms derived from a particular chloroplast gene. A detailed protocol for Northern blotting was published previously (5). Highly resolved, sharp bands can be obtained routinely if care is taken to use RNA that shows no sign of degradation, to minimize the volume and mass of RNA applied to the gel (