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 (<10  mm) stromules. In contrast, stromules may be observed on a high proportion of plastids in epidermal cells of leaves, petals, roots, or hypocotyls, which contain fewer smaller plastids than mesophyll cells (5–7 and Fig. 1). 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_5, © Springer Science+Business Media, LLC 2011

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Fig.  1. Stromules in different tissue and cell types. Confocal images of plastids visualised with fluorescent proteins. (a) Plastids in a leaf epidermal cell of a transgenic tobacco plant expressing a chimeric gene encoding GFP fused to the RbcS transit peptide under the control of the CaMV 35S promoter; (b) plastids in a petal epidermal cell of transplastomic tobacco containing a GFP gene under the control of the tobacco plastid ribosomal RNA operon (rrn) promoter; (c) plastids in a leaf trichome of a transgenic tobacco plant expressing a pea OEP14-YFP gene fusion under the control of the CaMV 35S promoter. Scale bars = 5 mm.

Stromules are highly dynamic and they may rapidly change shape over a period of seconds to minutes (8, 9). They can be observed to extend and retract, to branch, and, very rarely, for the end of the stromule to detach, creating an apparently independent plastid vesicle (8, 9). Stromule movement is dependent on the actin cytoskeleton (2, 10, 11) and is powered by the ATPase activity of class XI myosins (2, 12, 13). The functions of stromules are not yet fully established (5, 7). In some cells, they interconnect two, or more, plastids and enable the transfer of stromal material between the interconnected plastids (1, 14). However, in many cells, plastids interconnected by stromules are rarely observed, suggesting that stromules have other functions. The formation of stromules is likely to lead to an increase in the plastid surface area (2, 4, 5) and this may facilitate transport processes into, or out of, the plastids. A large increase in stromules accompanies arbuscular mycorrhizal infection of root cortical cells, leading to a functional symbiosis that requires transfer of plastid metabolic products to the mycorrhizae (15, 16). Stromules have also been observed in close proximity to other cell organelles, including nuclei, mitochondria, endoplasmic reticulum, and plasma membrane (3, 9, 14, 17, 18), leading to suggestions this may facilitate signalling or transfer of materials between organelles. Stromules appear to have a specialised function in starch deposition in wheat endosperm, where small starch granules have been observed within stromules protruding from amyloplasts (19, 20). The description of stromule function and identification of the molecular mechanisms involved are likely to be facilitated by studies with a genetically amenable plant, such as Arabidopsis thaliana.

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1.1. Visualisation of Stromules

Plastid structures that can now be identified as stromules were observed using bright-field light microscopy over 100 years ago (2, 3). However, stromules are more clearly visible by differential interference contrast (DIC or Nomarski) microscopy under white light (1, 8, 9), or by using confocal microscopy with a near-infrared femtosecond laser (21). These methods enable the examination of virtually any plant material, without the need for introducing a visible marker into the plastids. However, stromules are most easily seen if the plastids are marked with a fluorescent protein, such as green fluorescent protein (GFP), yellow fluorescent protein (YFP), or cyan fluorescent protein (CFP) (1–3). Stromules visualised by confocal microscopy of tobacco and petunia plants expressing plastid-targeted GFP were first reported by Köhler et  al. (1). They examined stable transgenic lines containing a nuclear transgene encoding GFP fused to the plastid-targeting signal (transit peptide) of the Arabidopsis chloroplast RecA protein, under the control of the cauliflower mosaic virus (CaMV) 35S promoter (1). Various plastid transit peptides have subsequently been used to direct nucleus-encoded GFP to the stroma, or to chloroplast envelope membranes, in stable transgenic plants for the visualisation of stromules (7). YFP targeted to plastids using the complete outer envelope protein OEP14 is shown in Figs. 1 and 2. The expression of the OEP14-YFP fusion protein produces intricate patterns of interconnected plastids, with some very long stromules (Figs. 1 and 2). Chimeric gene constructs with plastid transit peptides have also been used for transient expression of plastid-targeted GFP following microprojectile bombardment of leaves or other plant organs (7) (as described in this chapter), or by polyethylene glycol (PEG)mediated transfection of protoplasts (22, 23) (see Chapter 4, Vol. 1). In addition, a transient expression system using a tobacco mosaic virus vector, introduced for the high-throughput screening of subcellular compartmentation of GFP fusion proteins, has resulted in the identification of several chloroplast proteins present in stromules (24). Stable transplastomic lines of tobacco and poplar, with the GFP gene integrated directly in plastid DNA, can produce very high levels of GFP, facilitating the observation of stromules in many cell types (7, 25–27), including those in which expression from the CaMV 35S promoter is weak (7). The stromules can be observed by epifluorescence microscopy (10, 17, 18, 27) or  by confocal laser-scanning microscopy (1–3, 5, 7).

1.2. Stromules in Arabidopsis thaliana

Stromules were first observed in stomatal guard cells of cotyledons or young leaves of Arabidopsis seedlings transformed with a chimeric gene construct encoding the PALE CRESS (PAC) protein fused at its C-terminus to GFP, under the control of the PAC promoter (14). GFP was also observed in stromules in stomatal guard cells of dark-grown seedlings, and of seedlings treated with norflurazon, to bleach the chlorophyll (14). However, in this study, stromules

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Fig.  2. Stromules in Arabidopsis. Confocal images of plastids visualised with YFP fused to full-length pea OEP14. The ­chimeric gene construct was under the control of the CaMV 35S promoter in stable transgenic plants (a and d) or in bombarded leaves (b and c). (a) Plastids in a root epidermal cell of a stable transgenic line containing the CaMV 35S::OEP14-YFP construct; (b) plastids in a leaf epidermal cell following particle bombardment; (c) plastids in a leaf epidermal cell of arc6 following particle bombardment; (d) plastids in a hypocotyl epidermal cell of arc6 seedlings, obtained by crossing arc6 with a stable transgenic line containing the CaMV 35S::OEP14-YFP construct. In b and d, the red fluorescence is from chlorophyll in the underlying untransformed mesophyll cells. Scale bars = 5 mm.

were not observed in root tissue or in epidermal pavement cells of leaves or cotyledons, most probably due to the lack of expression from the PAC promoter in these tissues, rather than the absence of stromules in these tissues. Subsequent studies using plastid-targeted GFP, YFP, or CFP constructs under the control of the CaMV 35S promoter detected stromules in a wide range of other cell types, including epidermal cells of leaves (2, 28–30), roots (17, 18, 31, 32) and seedling hypocotyls (5, 31, 33) and cotyledons (34), leaf mesophyll cells (35), root tips and root hairs (17, 18, 31, 34), and stamen filaments and petals (31, 34). Stromules have also been detected in leaf mesophyll and hypocotyl epidermal cells by DIC microscopy (17, 36), and in leaf epidermis, mesophyll, bundle sheath, and stomatal guard cells by nearinfrared laser microscopy (21). The abundance and/or length of stromules is altered in some chloroplast division mutants (2, 29, 31, 34). Very long stromules,

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up to 65 mm long, have been detected in epidermal cells of leaves, roots, and hypocotyls of the arc6 (accumulation and replication of chloroplasts6) mutant (2, 31 and Fig.  2), and similar phenotypes have been detected in mutants created by overexpression of AtFtsZ1-1 (30) or AtMinE1 (34). Longer and more abundant stromules were also detected in epidermal cells of roots, hypocotyls, and stamen filaments of the arc3 mutant, and it was possible to visualise stromules emerging from chloroplasts in mesophyll cells by electron microscopy (31). These studies led to the conclusion that plastid division proteins play a role, either directly or indirectly, in the regulation of stromule development (31). Stromule formation in Arabidopsis also appears to be affected by environmental conditions, such as temperature. Increasing temperature increased the proportion of leaf mesophyll chloroplasts with stromules and protrusions, and the length and number of protrusions per chloroplast (35). In leaf mesophyll cells at temperatures above 20°C, stromules up to 20  mm were occasionally observed when there were large spaces between chloroplasts. However, at 5°C, very few chloroplasts (~5%) showed stromules or protrusions, whereas at 45°C, the number of chloroplasts with protrusions had increased more than tenfold. These chloroplasts had as many as five short beak-like protrusions (35). Antimycin A, an inhibitor of the cytochrome bc1 complex of the mitochondrial respiratory electron transfer chain, dramatically affected the morphology of root plastids, producing long filamentous plastids resembling stromules (32). Morphological changes were initiated within 5 min and complete in 1 h, and appeared not to be dependent on new protein synthesis, as neither chloramphenicol nor cycloheximide affected antimycin A-induced plastid filamentation (32). The morphological effects of antimycin A could be blocked by salicylhydroxamic acid (SHAM), a known inhibitor of the mitochondrial alternative oxidase, suggesting the involvement of an antimycin-activated alternative oxidase-mediated signalling pathway (32). The studies outlined above indicate that there is much to learn about the formation, movement, and functions of stromules. Arabidopsis thaliana offers an unsurpassed set of genetic resources for future studies. However, the observation of stromules is facilitated by the availability of plants or tissues containing plastidlocated fluorescent proteins that can be easily detected by confocal laser scanning microscopy. Stable transgenic lines containing chimeric genes encoding plastid-targeted GFP, YFP, or CFP have been used for most of the studies on Arabidopsis stromules reported above, and the lines should be available from the relevant laboratories. However, transient expression of plastid-targeted fluorescent proteins offers a means of detecting stromules in a wide range of Arabidopsis mutants, without the need for producing stable transgenic lines, and will be described in this chapter.

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2. Materials 2.1. Plant Material

1. Seeds of Arabidopsis thaliana (see Note 1). 2. 1.5-mL microfuge tubes. 3. Ethanol, 70% (v/v). 4. Vortex. 5. Microfuge. 6. 10% (v/v) sodium hypochlorite (>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 < 0.05) compared with Ws at each time point. This figure is published in Stanga et al. (11).

4. Notes 1. Seedlings may be grown for analysis using any growth conditions desired, e.g., varying agar or sucrose concentrations. These conditions are standard for our laboratory. 2. Chloral hydrate is classified as a Schedule IV controlled substance in the USA. Acquiring chloral hydrate at your institution may require a Drug Enforcement Administration (DEA) license, or equivalent. Check with your institution’s Environmental Health and Safety Department. Furthermore, chloral hydrate is

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toxic and used as a sedative. Wear gloves when handling it, work with it in a well-ventilated area, and wear a mask. 3. These cartridges are not commercially available. They are hard plastic and measure 75 × 30 mm with a rectangular hole in the center measuring 35 × 20 mm. The hole should be large enough to allow the roots to grow within the hollow space, yet small enough to allow room for the perimeter of a full-sized coverslip to adhere to the face of the cartridge. If the hole is slightly recessed around the perimeter to match the dimensions of a coverslip (60 × 24 mm), it will prevent the coverslip from sliding during the experiment, but this is not an essential feature. 4. In our setup, we used an upright microscope (Nikon Universal DIC Phase Condenser with Achr-Apl 1.4 lens, Japan; 40× objective) lain on its side such that the normally horizontal rotatable stage is vertically oriented. A rotatable stage is not standard for this model. Our setup consists of a round piece of aluminum machined to fit over the round stage and mounted with a screw. Any stage that is small enough to fit under the objectives and can rotate 360° can be used. A CCD camera (Spot RT-Slider Diagnostic Instruments Inc., MI, USA) is used for image capture. Bright-field images are sufficient for our purposes, but researchers seeking higher resolution images should consider using differential interference contrast (DIC) microscopy. The setup is shown in Fig. 1. 5. The software is required to capture images at predetermined intervals. Most cameras come with software capable of this function (e.g., SPOT RT, Diagnostic Instruments). For image analysis, the publicly available NIH ImageJ software package (http://rsbweb.nih.gov/ij/) can be used to convert the images to a stack, adjust thresholds, and track objects. 6. Coverslips must be thin enough to allow the microscope’s focal plane to lie within the root. 7. This method for starch staining is qualitative and is best used to assess quickly the presence or absence of starch in the columella cells, the approximate number of amyloplasts present in a cell, and which cells within the root cap contain amyloplasts. Starchless pgm mutants (12) are effective negative controls. Crude comparative assays may be done, but care should be taken to expose the roots to Lugol and chloral hydrate for the same duration because over time, Lugol staining intensifies, creating a high background signal, while chloral hydrate can quickly deteriorate the colored stain. Researchers seeking a quantitative comparison should seek an alternative protocol (13). 8. Mount the root(s) on one slide with water and a coverslip, and then apply a drop of 1% Lugol to the edge of the coverslip until the starch appears stained. Then add a drop of chloral hydrate

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(Subheading  3.1, step 4). Capillary action should draw the solutions through the roots, but a small kimwipe placed at the opposite end of the coverslip may help by serving as a wick to draw away excess moisture. This approach allows visualization of the root during the procedure. Alternatively, the solutions may be placed in small vials or culture dishes, and the seedlings may be moved from one solution to the next with tweezers before being mounted for microscopy. 9. Chloral hydrate is effective at removing background staining. If chloral hydrate proves too difficult to obtain, try using a more dilute Lugol stain to reduce or delay the appearance of background staining. 10. Seeds may keep for a few days longer, but over time they will begin to germinate in the dark and should not be used. 11. A small bottle of growth medium kept in a warm (55°C) incubator will keep for a few days and may be used to plate multiple batches of seeds on successive/alternate days to maintain a steady supply of seedlings ready for microscopy. If the drop of growth medium extends to the edges of the coverslip, trim the excess with the edge of another coverslip. Trimming the top edge of the medium so that it is flat will make plating easier and enable more seeds to grow under the medium and along the coverslip. If working with a mutant whose roots grow randomly, a droplet of seeds should be placed at the center of a coverslip, and a solidified drop of growth medium should be carefully overlain using another coverslip. Plates must be sealed with Parafilm to prevent the medium from drying out. 12. Stage clips should be tight enough so that the cartridge does not slip during the experiment. Roots must be kept vertical to allow the amyloplasts to sediment to the bottoms of the cells after disruptive handling. 13. Prior to reorientation, check to make sure that the camera and the eyepiece are parfocal: what may appear to be in focus to the user may not be in focus to the camera. It may help to mark a 90° angle before reorientation so that the stage can be quickly rotated to the proper angle. Rotating the root will likely move the tip out of the field of vision when using a 40× objective. Switch to a lowpower objective, then reorient the stage, find the root with the 40× objective, focus, and begin the image acquisition. The root will grow significantly over this time course, so begin with the columella cells positioned near the upper edge of the camera’s field of vision so that the root may grow into the field. 14. The central second-story columella cells of the root are of primary interest because of their strong contribution to gravitropism (14). Before making measurements, verify that the root is in focus throughout the stack. To speed the processing time,

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crop the stack to a smaller size. The object tracker macro will give coordinates for the chosen landmark only if it is present in every frame. If the macro has difficulty distinguishing the created mark from random background pixels, adjust the threshold levels so that the created mark is 100% black and anything lighter is white. 15. Due to the dynamic environment of the columella cells, it is extremely difficult to track an individual amyloplast over the course of an experiment. Therefore, a measurement is used that describes the movement of the amyloplasts en masse (15). 16. Under our growth conditions, mutants with intermediate amounts of starch (4) were suitable negative controls because their amyloplasts sediment slightly slower than wild-type amyloplasts. References 1. Morita, M. (2010) Directional gravity sensing in gravitropism. Ann. Rev. Plant Biol. 61, 705–720. 2. Blancaflor, E. and Masson, P. (2003) Plant gravitropism. Unraveling the ups and downs of a complex process. Plant Physiol. 133, 1677–1690. 3. Kiss, J., Hertel, R., and Sack, F. (1989) Amyloplasts are necessary for full gravitropic sensitivity in roots of Arabidopsis thaliana. Planta 177, 198–206. 4. Kiss, J., Guisinger, M., Miller, A., and Stackhouse, K. (1997) Reduced gravitropism in hypocotyls of starch-deficient mutants of Arabidopsis. Plant Cell Physiol. 38, 518–525. 5. Vitha, S., Yang, M., Sack, F., and Kiss, J. (2007) Gravitropism in the starch excess mutant of Arabidopsis thaliana. Am. J. Bot. 94, 590–598. 6. Evans, M.L. and Ishikawa, H. (1997) Cellular specificity of the gravitropic motor response in roots. Planta Suppl. 203, S115–S122. 7. Chen, R., Rosen, E., and Masson, P. (1999) Gravitropism in higher plants. Plant Physiol. 120, 343–350. 8. MacCleery, S. and Kiss, J. (1999) Plastid sedimentation kinetics in roots of wild-type and starch-deficient mutants of Arabidopsis. Plant Physiol. 120, 183–192.

9. Sack, F.D., Suyemoto, M.M., and Leopold, A.C. (1986) Amyloplast sedimentation and organelle saltation in living corn columella cells. Am. J. Bot. 73, 1692–1698. 10. Saito, C., Morita, M., Kato, T., and Tasaka, M. (2005) Amyloplasts and vacuolar membrane dynamics in the living graviperceptive cell of the Arabidopsis inflorescence stem. Plant Cell 17, 548–558. 11. 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. 12. Caspar, T. and Pickard, B. (1989) Gravitropism in a starchless mutant of Arabidopsis: implications for the starch-statolith theory of gravity sensing. Planta 177, 185–197. 13. Smith, A.M. and Zeeman, S.C. (2006) Quantification of starch in plant tissues. Nat. Protoc. 1, 1342–1345. 14. Blancaflor, E., Fasano, J., and Gilroy, S. (1998) Mapping the functional roles of cap cells in the response of Arabidopsis primary roots to gravity. Plant Physiol. 116, 213–222. 15. Leitz, G., Kang B.H., Schoenwaelder, M.E.A., and Staehelin L.A. (2009) Statolith sedimentation kinetics and force transduction to the cortical endoplasmic reticulum in gravity-sensing Arabidopsis columella cells. Plant Cell 21, 843–860.

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Chapter 8 Studying Arabidopsis Chloroplast Structural Organisation Using Transmission Electron Microscopy Stefan Hyman and R. Paul Jarvis Abstract Chloroplasts, as well as other, non-photosynthetic types of plastid, are characteristic structures within plant cells. They are relatively large organelles (typically 1–5 mm in diameter), and so can readily be analysed by electron microscopy. Chloroplast structure is remarkably complex, comprising at least six distinct suborganellar compartments, and is sensitive to developmental changes, environmental effects, and genetic lesions. Transmission electron microscopy (TEM), therefore, represents a powerful technique for monitoring the effects of various changing parameters or treatments on the development and differentiation of these important organelles. We describe a method for the analysis of Arabidopsis plant material by TEM, primarily for the assessment of plastid ultrastructure. Key words: Arabidopsis thaliana, Chloroplast ultrastructure, Plastid development, TEM, Trans­ mission electron microscopy

1. Introduction Plastids are a functionally and structurally diverse group of related organelles, including photosynthetic chloroplasts and a variety of non-photosynthetic variants such as leucoplasts, amyloplasts, and chromoplasts (1, 2). They are characteristic components of plant cells and have been subjected to extensive ultrastructural analysis by electron microscopy (EM) over many years. Such analysis is invaluable, as organellar structure is influenced by a variety of factors including developmental, environmental, and genetic parameters (3–5). Thus, ultrastructural analysis of plastids enables the consequences of various treatments and conditions for organelle structure and development to be analysed and quantified.

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_8, © Springer Science+Business Media, LLC 2011

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Electron microscopy has many variants; even a focused approach to a single specimen has many options that may be explored. We have aimed to describe procedures based on access to the expertise and not insignificant physical resources of an established electron microscopy laboratory. We provide a basic methodology for the collection, preservation (fixation), and embedding of the sample. Brief descriptions of the processes of ultramicrotomy and section staining, and use of the transmission electron microscope are also provided, but detailed instructions for these are beyond the scope of this text and are described in many other publications, for example, Kuo (6). The described techniques are those we have successfully employed, and they should be readily reproducible in any electron microscopy laboratory. By restricting ourselves to these conventional methods, we avoid the need for specialised preparative apparatus, such as that needed for high pressure freezing and freeze substitution techniques as utilised by Pfeiffer and Krupinska (7). It should be noted that the described methods are far from exhaustive and indeed are offered as a starting point from which modifications may be made to suit more “local” applications.

2. Materials 2.1. Major Equipment

All of the following items are essential to these procedures. However, similar items should already be present in any biological sciences EM laboratory. This summary is based upon what we use and does not imply superiority of one brand or manufacturer over another. 1. A ducted fume cupboard, suitable for handling flammable and carcinogenic materials (i.e., it should be spark proof and not of the portable, re-circulating variety). 2. Spark-proof refrigerator or cold room, set at 4°C. 3. Stereo dissecting microscope (e.g., Olympus SZ51, Olympus UK Ltd., Southend-on-Sea, Essex, UK). 4. Ultramicrotome (e.g., Leica EM UC7, Leica Microsystems (UK) Ltd., Milton Keynes, UK). 5. Light Microscope (e.g., Olympus BX Series, Olympus UK Ltd.). 6. Transmission electron microscope. We have a JEOL 1400 (JEOL (UK) Ltd., Welwyn Garden City, UK).

2.2. Minor Equipment

1. Fine-point stainless steel forceps (e.g., Dumont tweezers 4, T5288, Agar Scientific Ltd., Stansted, UK). 2. Reverse forceps (e.g., Dumont tweezers, NOC, T5071, Agar Scientific Ltd.). 3. Fine artists’ paint brush (Size 00) (e.g., G3444, Agar Scientific Ltd.).

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4. Variable-speed rotary mixer (e.g., R060, TAAB Laboratories Equipment Ltd., Aldermaston, UK) or a variable-speed roller mixer (e.g., 444–1067, VWR International Ltd., Lutterworth, UK). 5. Portable balance; range 0–200 g, with accuracy to two decimal places (e.g., Fisherbrand SG202, Fisher Scientific, Loughborough, UK). 6. Variable-speed magnetic stirrer with stirring bars (e.g., Hanna HI-200M, Fisher Scientific). 7. Embedding oven, chest type (e.g., EO62 or E100, TAAB Laboratories Equipment Ltd.). 8. Embedding capsule press (e.g., C213, TAAB Laboratories Equipment Ltd.); this item is not essential but it is extremely useful. 9. Small bench vice (e.g., “Vacu-Vice,” T576, Agar Scientific Ltd.). 10. Small (6 in.) “Junior” hacksaw. 11. Hotplate (e.g., Stuart SD160, VWR International Ltd.). 2.3. Consumables and Small Items

1. Personal protective equipment, including disposable protective gloves and laboratory coat. 2. Locally approved incineration bins for disposal of solid waste, resin-contaminated material, and sharps. 3. Specimen tubes, 2 mL (T308-2, Simport, Beloeil, Canada). 4. Disposable graduated plastic transfer pipettes, 7 mL (612–1681, VWR International Ltd.). 5. Ordinary glass microscope slides (plain), or white tile. 6. Razor blades, single or double edged (e.g., T585, Agar Scientific Ltd.). 7. Pipettors and tips, to handle volumes ranging from 5  mL to 1 mL (e.g., Finnpipette or Gilson Pipetman). 8. Polyethylene graduated containers with caps (e.g., G332 and G333, Agar Scientific Ltd.). 9. Polyethylene flat-ended embedding capsules (for modified Spurr’s resin or other epoxy resins) (e.g., G3744, Agar Scientific Ltd.). 10. Petri dish, 9-cm diameter, glass (or Conway Dish if available). 11. Plastic sealing film (e.g., Parafilm). 12. Syringe filters, 0.2 mm (e.g., Acrodisc, Pall Corporation, MI, USA). 13. Electron microscope grid storage box (e.g., G276, Agar Scientific Ltd.).

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2.4. Chemistry (see Note 1)

2.4.1. Tissue Fixation

The following is a list of chemicals that are required for the described techniques. We assume that the reader has access to a distilled water supply. 1. Sörensen’s phosphate buffer (SP buffer), comprising disodium hydrogen orthophosphate and sodium dihydrogen orthophosphate (see Subheading 2.5.1). 2. Glutaraldehyde, 25% (w/v) aqueous solution, EM grade. Toxic and harmful to the environment. 3. Paraformaldehyde (prilled), EM grade. Harmful. 4. Osmium tetroxide, 2% (w/v) aqueous solution. Toxic and corrosive. 5. Milk powder. 6. Potassium ferricyanide. Irritant. 7. Uranyl acetate (also used in section staining). Radioactive, very toxic, and may have cumulative effects. 8. Sodium hydroxide, 1 N (1 M), carbonate-free aqueous solution. Corrosive.

2.4.2. Tissue Dehydration and Embedding

1. Ethanol, analytical grade. This is used both pure and diluted (in distilled water) at the following concentrations: 30, 50, 70, and 90% (v/v). Highly flammable. 2. 1,2-Epoxypropane (pseudonym, propylene oxide; PO). Toxic, suspected carcinogen, and highly flammable. 3. Epoxy resin embedding medium. For example, modified Spurr’s resin, comprising: ERL 4221 (irritant); diglycidylether of polypropylene glycol (DER 736, irritant); nonenyl succinic anhydride (NSA, irritant); dimethylaminoethanol (DMAE, flammable and irritant).

2.4.3. Sample Remounting and Section Staining

1. High-strength epoxy resin adhesive (e.g., Araldite, Huntsman Advanced Materials Ltd., Cambridge, UK). Irritant and dangerous to the environment. 2. Reusable pressure adhesive (e.g., Blu-Tack, Bostik Ltd., Leicester, UK). 3. Lead nitrate (to make Reynolds lead citrate). Toxic and dangerous to the environment. 4. Trisodium citrate (to make Reynolds lead citrate).

2.5. Working Buffers and Reagents (see Note 1) 2.5.1. Sörensen’s Phosphate Buffer (0.2 M, pH 7.2) (8)

Sörensen’s phosphate buffer (SP buffer) is a mixture of mono- and dibasic sodium phosphates. Prepare a 0.2 M disodium hydrogen orthophosphate solution (Solution A) as follows: dissolve 35.61 g of Na2HPO4⋅2H2O (or 53.65 g of Na2HPO4·7H2O or 71.64 g of Na2HPO4⋅12H2O, depending on availability) in 1  L of distilled water. Next, prepare a 0.2 M sodium dihydrogen orthophosphate

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solution (Solution B) as follows: dissolve 27.6 g of NaH2PO4·H2O (or 31.21 g of NaH2PO4⋅2H2O) in 1 L of distilled water. To make the 0.2 M SP buffer stock solution, mix 36 mL of Solution A with 14 mL of Solution B; the pH of the final solution should be 7.2, which should be verified. This stock solution is used to make the primary and secondary fixation solutions (see Subheading 2.5.3). A further dilution of the stock is also required: 0.1 M SP buffer is used for washing steps (see Subheading 3.2). 2.5.2. Depolymerised Paraformaldehyde Solution (10% Formaldehyde)

In a fume cupboard, dissolve 10.0 g of prilled paraformaldehyde in 90 mL of distilled water. Warm the mixture to approximately 60°C and stir constantly to ensure that the paraformaldehyde dissolves. If the solution is not completely clear, add, drop-wise, 1 N sodium hydroxide until clarity is achieved. The solution should then be made up to 100.0 mL with distilled water. Ideally, this should be prepared immediately before use (9), though it may be stored frozen for some months. If the solution exhibits any opalescence or traces of flocculate material, it should not be used.

2.5.3. Primary, Secondary, and Tertiary Fixatives

1. Primary fixative. Add 4.0  mL of freshly depolymerised 10% (w/v) paraformaldehyde solution (see Subheading  2.5.2) to 5.0 mL of 0.2 M SP buffer. To this mixture, add 1.0 mL of 25% (w/v) glutaraldehyde solution. The final working solution is 4.0% (w/v) formaldehyde, 2.5% (w/v) glutaraldehyde, in 0.1 M SP buffer (10) (see Note 2). 2. Secondary fixative. To 1.0 mL of 2% (w/v) aqueous osmium tetroxide (see Notes 3 and 4), add 1.0 mL of 0.2 M SP buffer, and to this add 30.0 mg of potassium ferricyanide. The final working solution is 1% (w/v) osmium tetroxide in 0.1 M SP buffer containing 1.5% (w/v) potassium ferricyanide. 3. Tertiary fixative. To 1.0  mL of distilled water, add 20.0  mg of uranyl acetate (see Note 5). The final working solution is 2% (w/v) aqueous uranyl acetate. This solution may also be used as an electron-dense stain for ultrathin sections (see Subheading 3.4.1).

2.5.4. Modified Spurr’s Low-Viscosity Resin (11) (see Notes 6–8)

The following procedure makes approximately 30.0  mL of “standard” hardness resin. Using disposable 7-mL graduated plastic transfer pipettes (see Notes 9 and 10), weigh into a disposable plastic container (see Subheading 2.3, item 8) 12.3 g of ERL 4221, 4.3 g of DER 736 (see Note 11), and 17.7 g of NSA (see Subheading 2.4.2). Add a small magnetic stirring bar and thoroughly mix for 5 min on a magnetic stirrer. Finally, add 0.3 g of DMAE accelerator and mix for a further 5 min. Avoid incorporating air bubbles into the mixture. DMAE and NSA should never be mixed together alone as this may result in a strong exothermic reaction. Test the quality of the resin as follows: fill a polyethylene flatended embedding capsule (see Subheading 2.3, item 9) with the resin mixture and polymerise for at least 8 h at 70°C. Remove the

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polymerised block from the capsule and check the hardness and integrity of the resulting block. This is best judged by using a single-edged razor blade to trim a piece of resin from the block; it should not be soft or brittle. 2.5.5. Reynolds Lead Citrate (12) (see Note 12)

Boil and allow to cool 100 mL of distilled water (see Note 13). Weigh 1.33 g lead nitrate (Pb(NO3)2) and 1.76 g trisodium citrate (Na3(C6H5O7)⋅2H2O) into a 50-mL volumetric flask. Add 30 mL of the boiled distilled water to the flask. Close the flask with a stopper and then shake vigorously for at least 1 min. Leave the mixture to stand, with occasional shaking, to complete the conversion to lead citrate. The mixture will become “milky” in appearance and should contain no solid material. After 30 min, add 8.0 mL of 1 N sodium hydroxide solution (13); the mixture should “clear” quickly. Dilute to 50  mL with additional freshly boiled, distilled water, and mix by inversion. The resulting solution should have a pH of 12 (±0.1) and should be stored at 4°C in a tightly sealed bottle. Before use, allow to reach room temperature and filter through a 0.2-mm syringe filter to remove any particulate matter.

3. Methods The entire procedure, from start to finish, will take approximately four days; therefore, sample collection should ideally commence at the beginning of the week. To process two genotypes (i.e., at least six cotyledons or leaf sections) in 2.0  mL specimen tubes for transmission electron microscopy (TEM) requires approximately 21.0  mL of distilled water, 5.0 mL of 0.2 M SP buffer, 0.8 mL of 10% freshly depolymerised paraformaldehyde (formaldehyde), 0.2 mL of 25% glutaraldehyde, 1.0 mL of 2% osmium tetroxide, 30.0 mg of potassium ferricyanide, 12.0  mL of analytical grade ethanol (including that used for graded dehydration steps), and 8.0 mL of 1,2-epoxypropane (including that used for graded infiltration steps). In practice, larger “stock” volumes of each of the above should be available. Approximately 30.0  mL of freshly mixed modified Spurr’s resin should be made. 3.1. Collection of Samples 3.1.1. Standardisation

When working with young seedlings, one may choose to analyse cotyledons, true leaves, or non-photosynthetic tissues such as the roots. With very young/small specimens, it may prove optimal to excise and process whole cotyledons or even process the entire seedling. It should be normal practice to take samples from seedlings or plants of equivalent age and which have been grown under identical conditions. When analysing true leaves, it is important to select leaves of an equivalent developmental stage.

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When using plate-grown samples, identify and use seedlings whose cotyledons/leaves are not in contact with the growth medium. Cotyledons and small leaves may be excised from the seedling at their base with a razor blade. When possible, it is our practice to also cut away the top 30–50% of the cotyledon or leaf at 90° to the central vein (see Fig.  1) (see Note 14). The retained portion is immediately immersed in the buffered primary fixative solution (see Subheading  3.2.1). When working with true leaves, a slice (maximum 3 × 5 mm) is dissected from the central portion of the leaf lamina, again at 90° to the central vein. When working with roots, ~2–3-mm-long sections approximately 3 mm back from the root apex should be excised for analysis. When working with darkgrown seedlings in order to study etioplast ultrastructure, tissue collection and initial fixation should be conducted under green safe-light conditions. Cotyledons from etiolated plants are collected by harvesting intact apical regions approximately 2–3 mm in length. Particularly small, entire seedlings may be fixed without dissection in primary fixative. These may then be washed and dissected before the secondary fixation stage, under normal lighting. It is normal to retain only one cotyledon, leaf or root specimen per plant, in order that one can be sure that each corresponds to a separate individual. Several (at least six) unique specimens should be collected for processing, per sample (i.e., per genotype or condition), although usually only three will be sectioned, observed, and recorded.

Fig.  1. Collection and dissection of the biological material. Shown here is a typical 10-day-old, plate-grown Arabidopsis seedling. One cotyledon has been removed using a single-edged razor blade. The top 30–40% of the cotyledon has also been removed to aid penetration of the primary fixative solution into which the sample must be immediately transferred (see Subheading 3.1.2).

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3.2. Tissue Processing for TEM (see Note 1)

3.2.1. Day 1

With the exception of the refrigerated steps, all of the following should be conducted in a suitable ducted fume cupboard at room temperature. Materials stored in a refrigerator should be allowed to reach room temperature before opening to avoid the possible formation of condensation. The following procedure assumes the use of 2-mL screw-top specimen tubes (see Subheading 2.3, item 3). Alternative containers (e.g., 7-mL screw-top, flat-bottomed, glass vials) may be substituted with appropriate reagent volume adjustments (see Note 15). A minimum of 1.0  mL of each reagent should be used in all steps. The samples should be regularly agitated; this may be done intermittently by hand, or (preferably) constantly on a rotary or roller mixer. 1. For each sample (i.e., for each plant genotype or condition to be analysed), dispense at least 1.0 mL of freshly prepared primary fixative (see Subheading 2.5.3) into a labelled specimen tube. 2. Transfer the biological material to be sampled to a glass microscope slide (or tile). To avoid physical damage, do not manipulate the area of tissue to be examined. Use a single-edged razor blade to excise the required tissue specimen, as described in Subheading  3.1.2, and immerse it immediately in the fixative solution (see Note 16). Repeat as many times as required with additional plants. 3. Allow the samples to fix for ~3–4 h at room temperature following collection of the last sample (see Note 17). After this time, move the vials to a refrigerator (ensure that the samples are still in contact with the fixative) and leave overnight (see Note 18).

3.2.2. Day 2

4. Remove the samples from the refrigerator and allow them to reach room temperature. 5. Using a disposable plastic transfer pipette, remove the fixative solution and replace it with 0.1 M SP buffer (see Notes 19–21). Wash for 30 min. 6. Repeat the last step two more times. The waste fixative and washes must be disposed of appropriately. 7. Remove the final wash and replace it with 1.0 mL of secondary fixative (see Subheading 2.5.3) (see Note 3). Allow the samples to fix for 1–1.5 h at room temperature. 8. Thoroughly wash the samples (now noticeably dark greenishblack) once for 15 min with 0.1 M SP buffer, and then three times (for 30 min each time) with distilled water (see Note 22). The waste fixative and washes must be disposed of appropriately. 9. Remove the final wash and replace it with 1.0 mL of tertiary fixative (see Subheading  2.5.3) (see Note 5). The samples should now be kept in the dark for 1 h; agitate them regularly.

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10. Wash the samples three times (30 min each time) in distilled water. The waste uranyl acetate and washes must be disposed of appropriately. 11. After removing the final wash, commence dehydration of the samples using a graded ethanol/distilled water series as follows (in order of increasing ethanol concentration): 30, 50, and 70% ethanol, for 30 min at each step. 12. Replace the 70% ethanol with fresh 70% ethanol, transfer the samples to a refrigerator, and leave overnight. 3.2.3. Day 3

13. Remove the samples from the refrigerator and allow them to reach room temperature. 14. Conduct further dehydration steps using (in order) 90% ethanol and 100% analytical grade ethanol for 30  min each (see Note 23). The analytical grade ethanol step should be repeated twice more. 15. During step 14, prepare the required volume (dependent on the number of samples) of modified Spurr’s resin (see Subheading 2.5.4) to be used in the following infiltration and embedding steps. 16. Exchange the ethanol for 1, 2-epoxypropane (PO) and incubate for 10 min, and then repeat. PO is a toxic, potentially carcinogenic, and highly flammable solvent (see Note 24). Do not allow the samples to be exposed to air; leave a minimal residual amount of the previous solution during exchanges. 17. Infiltrate the samples with the modified Spurr’s resin by immersion in progressively increasing concentrations of resin according to the following series (see Note 25): (a) 3 parts PO to 1 part resin, 60–90 min; (b) 1 part PO to 1 part resin, 60–90 min; (c) 1 part PO to 3 parts resin, 60–90 min; (d) 100% resin, 30 min. 18. Replace the resin with fresh 100% resin and leave on the mixer overnight.

3.2.4. Day 4

19. Replace the overnight resin with fresh 100% resin and leave on the mixer at room temperature for a further 3 h. 20. Obtain the required number of polyethylene embedding capsules (see Subheading 2.3, item 9). Each capsule should include a label indicating the identity of the sample (see Note 26). 21. Using a modified 7-mL disposable plastic transfer pipette, collect an individual leaf specimen from the processing container and transfer it (with a small volume of resin) into a separate capsule, i.e., one piece per capsule (see Note 27). Repeat this for each specimen.

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22. Completely fill each capsule with fresh 100% resin and close the lid (see Notes 7 and 28). 23. Leave the loaded capsules for an hour or two and then transfer them into a pre-heated (70°C) embedding oven for at least 8 h to polymerise the resin. 24. Proceed to Subheading 3.3. 3.3. Remounting (see Fig. 2)

For sections to be taken at a specific orientation from the tissue (e.g., perpendicular to the plane of the leaf lamina, and at right angles to the mid-vein), the sample will need to be in a specific position for the process of ultramicrotomy. 1. Once the resin has polymerised, if available, use an embedding capsule press (see Subheading  2.2, item 8) to remove the blocks from the moulds. Alternatively, very carefully cut down two opposing sides of the capsule with a single-edged razor blade and peel the two halves of the capsule apart to release each block. 2. Using a simple dissecting microscope, locate (and mark if necessary) the position of the sample within each block (see Fig. 2a); it may help to draw precise guidelines on the block face for sawing along (see Note 29). 3. Insert the block vertically, specimen up, into a small vice. Position the block to leave 3–4 mm of resin above the jaws of the vice. Close the vice (do not over-tighten it) and use a small hand saw (hacksaw) to remove the identified piece of resin containing the sample from the block (see Fig. 2b and c) (see Note 30). 4. Remove any dust or sawing debris from both the excised piece of resin and the remaining block, hereafter called the base. 5. Remove the base from the vice and position it vertically onto a piece of Blu-Tack pressure adhesive stuck atop a glass microscope slide or similar. 6. Mix, according to the manufacturer’s instructions, approximately 0.5–1.0  mL of Araldite epoxy resin adhesive; place a small quantity of this adhesive onto the upper surface of the base. 7. Using forceps, firmly insert the excised piece of resin containing the sample (longest edge to the base) into the Araldite adhesive. Ensure that the remounted piece is correctly orientated before building up the adhesive around all sides of the piece. 8. Allow the epoxy resin adhesive to harden (usually 24 h), after which time the block is ready for sectioning (see Fig. 2d). 9. Proceed to Subheading 3.4.

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Fig.  2. Re-orientation and mounting of the specimen before ultramicrotomy. Sequential stages in the mounting process are shown (see Subheading 3.3). (a) An embedded cotyledon sample within the polymerised resin block after removal from the embedding capsule. (b) Excess resin has been removed to a depth of 2–3 mm from around sample by careful use of a hacksaw, leaving a trapezium-shaped “island” containing the sample piece. The base is retained for use in step 4, shown in panel (d). (c) Here, the sample piece has been undercut, removed from block, and cleaned to remove any dust generated during the sawing process. (d) Finally, the sample piece, with the longest edge now oriented to the base (such that the cotyledon is vertical), is remounted back onto the original base using Araldite high-strength epoxy adhesive.

3.4. Ultramicrotomy and Staining

This is the process of obtaining ultra-thin sections (slices) of the sample for observation in the TEM. Because of the risk of physical damage to the sample from the initial specimen collection, it is often necessary to obtain and observe many grids of multiple sections so as to ascertain the optimum area of tissue preservation. Ultramicrotomy is a technique that requires much training and specialist equipment and as such must be approached with the help of experienced assistance. Many reference books are available that deal with the intricacies and problems of ultramicrotomy and staining (6, 14). A detailed method is beyond the scope of this chapter, but for general information, we include a brief summary of the technique as follows.

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The block (remounted tissue sample) is first trimmed to remove excess resin from around the tissue. The aim is to produce a block face, approximately 2–3-mm square, from which semi-thick sections of about 0.5 mm can be taken using a glass or special diamond knife. Behind the cutting edge of the knife, there is a trough that is filled with water, upon which the sections float. Three or four of the thick sections are grouped together and transferred on a drop of water (using a wire loop of 3-mm diameter) to the approximate centre of a glass microscope slide. Move the slide to a hotplate (set to 80–90°C) to dry. Stain the sections with a suitable light microscopy stain, e.g., 1% (w/v) toluidine blue in 1% (w/v) sodium borate (14). To do this, cover the sections with a drop of staining solution and leave the slide on the hotplate for ~2  min (do not allow the stain to dry out). Remove the slide from the hotplate and carefully rinse away the stain with distilled water. Dry the slide on the hotplate for ~2 min before observation. The sections are observed using a light microscope in order to determine the exact location within the sample that has been reached, and to locate the area to be used for EM. Once this has been identified, trim the block further to reduce the face to a trapezium shape similar in appearance to Fig. 2c (the longest edge should ideally be only 0.75–1.0  mm). Next, ultra-thin sections (60–90nm thick) are cut for EM purposes. These are collected on the surface of a specimen support (grid) by picking up a clean grid with forceps, submerging it in the water trough, moving it below the sections, and lifting it up through the surface, thus picking up the sections on the grid. It is usual to collect three or four sections on any single grid. Grids have a diameter of 3.05 mm, which is the limiting factor regarding the size of sample that can be observed by TEM. Grids are produced with many different architectures, e.g., square or hexa­ gonal mesh, different mesh sizes (quoted as holes per inch), and metal compositions. The mesh size determines the support given to the sections and, along with the bar thickness, dictates the percentage transmittance of the grid; i.e., smaller mesh sizes (e.g., 300+ holes per inch) give greater section stability and support, but lower transmittance and therefore, potentially, a smaller viewable sample area. For routine TEM applications, a 300 square mesh copper grid may be chosen. Once the sections have been collected on a grid, they may be contrasted using heavy metal stains. For regular TEM observations, we routinely use the following staining protocol. 3.4.1. Primary Staining with Uranyl Acetate (see Notes 1 and 5)

If uranyl acetate has been used in the fixation process, this primary staining may be omitted. In a fume cupboard, for each grid to be stained, pipette a 25–30 mL drop of filtered (0.2 mm) 2% aqueous uranyl acetate stain (see Subheading 2.5.3) onto a freshly exposed Parafilm surface; this is best done contained within a 9-cm diameter glass Petri dish.

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Using suitable forceps (see Subheading  2.2, item 2), submerge the grids in the stain, one grid per droplet. Then, cover the Petri dish by placing a box over it to exclude light and leave it for 15–20 min. Using the forceps, remove each grid from the staining solution and wash by immersion, several times, in a substantial volume (10–20  mL) of distilled water contained in a 25-mL glass beaker. Repeat this process through at least three successive washes to remove all traces of stain. The washes must all be disposed of appropriately. Remove excess water from the grid using a small piece of filter paper and allow the grids to air-dry whilst still held by the forceps. 3.4.2. Secondary Staining with Reynolds Lead Citrate (12) (see Notes 1 and 12)

This process should be carried out in a nitrogen (carbon dioxide free) atmosphere; to this end, we utilise an in-house made Perspex box through which nitrogen gas is allowed to flow at 1–2 L per min (see Note 31). From the filtered (0.2  mm) lead citrate stock (see Subhead­ ing 2.5.5), dispense one 25–30 mL drop of stain for each grid to be stained onto a freshly exposed Parafilm surface contained within a 9-cm glass Petri dish. Submerge the grids in this solution, one grid per droplet, and leave them for 2–3 min. After staining, wash the grids as described in Subheading 3.4.1 by immersion in a substantial volume (10–20 mL) of freshly boiled distilled water (see Note 13).The first wash should contain one or two drops of 1 N sodium hydroxide to reduce the chance of lead carbonate precipitation. The grid should be passed through at least three successive boiled distilled water washes to remove all traces of stain. As with the primary stain, the washes must all be disposed of appropriately. Leave the grids to air-dry as before whilst still held by the forceps. It is advisable to store the stained grids in a labelled grid storage box (see Subheading 2.3, item 13) (see Note 32) prior to observation using the TEM.

3.5. Observation and Recording

Observation of the samples is carried out in the TEM, operated according to the manufacturer’s instructions. We routinely use an accelerating voltage of 80 kV, although this is just one of the parameters that may be adjusted in order to optimise results. We use an Olympus SIS Megaview III digital camera (Olympus Soft Imaging Solutions), which is controlled via dedicated iTEM software to capture and save images as required. Typical TEM images of chloroplasts in wild-type Arabidopsis cotyledons are shown (Fig. 3). In order to enable meaningful comparisons between different genotypes, treatments, or conditions, it is essential to analyse a relatively large number of different organelles from multiple individuals (not least because of the nature of the sectioning process and the influence that it may have on what is observed). We recommend that at least three independent plants per genotype/treatment/condition are analysed, and that at least ten representative whole-organelle

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Fig. 3. Typical transmission electron micrographs of chloroplasts in a wild-type seedling. Cotyledons of 10-day-old, wild-type Arabidopsis seedlings were analysed using the protocol described in this chapter. Images were recorded at three different magnification levels. Scale bars indicate 5 mm (a), 2 mm (b), and 0.5 mm (c).

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images per plant are selected for detailed analysis (the selection of representative images having been made on the basis of a careful overview of the whole section); i.e., a minimum of 30 independent, representative images per sample should be considered. Images from the TEM can be used to derive quantitative data. For example, digital image files can be analysed using Adobe Photoshop or more specialised image analysis software. In Photoshop, the “Measure Tool” function can be used to obtain length and width measurements for each individual organelle. Such values can then be used to provide an indication of chloroplast shape (the length/ width ratio; typically 3 in wild-type Arabidopsis), or to derive an approximate estimation of organellar cross-sectional area (using the following formula, which describes a perfect elliptical shape: p × 0.25 × length × width) (15, 16). The TEM images can also be used to derive other quantitative measures of development, such as the number of granal or stromal lamellae per granal stack.

4. Notes 1. Sample processing for electron microscopy requires the handling of hazardous materials including chemical fixatives, solvents, embedding resins, and heavy metal solutions. Suitable personal protective equipment and the availability of approved containment facilities are essential. It is imperative to both consult and understand each material safety data sheet, and to ensure that appropriate approved codes of practice and local regulations for safe handling and waste disposal are strictly followed. Because of the significant hazards presented, persons using these materials must also be familiar with emergency procedures associated with accidental release or spillage. Ultimate disposal of many of these substances will require an approved chemical waste contractor. 2. The proportions of formaldehyde and glutaraldehyde fixatives may be varied, as may the buffer vehicle (e.g., sodium cacodylate may be used in place of SP buffer). The fixation time, and the dehydration solvent (ethanol or acetone) and/or time may be adjusted and optimised for any particular specimen (17). 3. Osmium tetroxide is very volatile and should always be used in a fume cupboard. It is also advisable to have a plentiful supply of milk powder available (or other suitable absorbent material) that can be used to absorb any spills. Glauert (9) further suggests that waste osmium may be converted to osmium dioxide (black) by mixing with an ethanol/water mixture or with ferrous sulphate. Other suitable methods of inactivation include mixing with vegetable oil or milk; Bozzola (18) recommends

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mixing in the ratio of two or three parts corn oil to one part osmium. All waste material must then be appropriately disposed of according to local regulations. 4. Potassium ferricyanide (usually at 1.5% [w/v]) is an optional addition to the secondary fixative; its presence enhances membrane contrast (9). 5. Uranyl acetate is particularly hazardous, especially in its powder form where inhalation presents a major hazard, as it is radioactive and very toxic, and may have cumulative effects. It must be handled according to local regulations, which should at the very least require containment to eliminate the risk of inhalation and ingestion. Uranyl acetate is also photolabile (9) and should, therefore, be stored in total darkness. It is quite slow to dissolve, so it is best prepared on the day before it is required. 6. The original Spurr’s ultra low-viscosity resin (19) cannot now be purchased due to the withdrawal of one component, vinyl cyclohexene dioxide (VCD, ERL 4206). A modification to the original recipe is the replacement of this component with ERL 4221 (11). 7. Spurr’s resin is particularly susceptible to moisture content. Therefore, to reduce the possibility of atmospheric moisture ingress, always keep component bottles closed when not in use and always cap the mixing vessel whilst stirring. Loaded specimen capsules should also be closed as quickly as possible. Water present in the final mix can lead to blocks of unsatisfactory consistency and prove problematic at the ultramicrotomy stage. 8. Other epoxy resins are obtainable; for plant material, it is always advisable to use the lowest viscosity available. 9. Some of the components used in these methods are very viscous and as such it is neither accurate nor practical to measure them by volume. 10. We prefer plastic transfer pipettes because disposable glass pipettes may have fragments of broken glass that could be introduced into the resin and other solutions. If this occurs, there is a risk that when sectioning, a glass fragment embedded in the resin near the sample (which would be invisible) could damage the ultramicrotomy knife. Indeed, if using a diamond knife, this would destroy that area of the cutting edge. 11. A decrease in the amount of DER 736 used in the mixture will give a harder block, while an increase will give a softer block. This effect may be used to determine the optimum hardness for any given sample. 12. Lead citrate is readily precipitated (as lead carbonate) by the presence of carbon dioxide. It is best to use commercially purchased 1 N sodium hydroxide in the preparation of this stain, to ensure the correct pH of the final solution (13).

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13. To minimise the risk of lead carbonate precipitation, we prefer to use freshly boiled distilled water rather than stored distilled water that may have absorbed atmospheric carbon dioxide. 14. Removing the top 30–50% of the organ is performed to assist penetration of the fixative. Single- or double-edged razors may be used. Double-edged blades are thinner than single-edged blades, and hence there is less potential for tissue damage at the point of cutting. Nonetheless, because of this possible damage, it is advisable to remove 0.5–1.0  mm of the tissue back from the immediate vicinity of the cut during the initial trimming of the block face at the ultramicrotomy stage. 15. Processing containers should ideally be considered disposable, as they will become resin contaminated. Key points to consider when choosing a processing container are (A) resistance of the material to the various chemicals that will be used; (B) efficiency of containment of the chemicals – chemical vapour escape must be avoided; and (C) container volume. Consideration must be given to the efficacy of solution changes at each stage; the replacement solution may be diluted by residual solution from the previous stage. Therefore, to minimise this effect, volumes should not be reduced too much. Small screw-top containers are preferred, as they seal completely and present minimal risk of aerosol formation when opening (which can be a hazard with “snap-top” containers). Test any new container to ensure that it meets all requirements before use. 16. A fine paintbrush is an excellent tool for manipulating the sample pieces; this is far less likely to cause the physical damage that might occur when using forceps. 17. If available, it may help the infiltration of the fixative into the samples to expose them to a slight vacuum for the first 20–30  min of the primary fixation step (in fact, with some sample types this may be essential). This should not to be any more than 0.5 atmospheres (380 mmHg) (9). This should be applied and released intermittently with a frequency of 1–2 min per cycle (20). 18. Because they contain hazardous aldehyde fixative, the sample tubes should ideally be contained within a sealed, labelled container. 19. It is vital at all stages that the samples are not allowed to dry out; for this reason, solution exchanges should be completed one by one and as quickly as possible. 20. The samples must be closely monitored. Leaf specimens have a tendency to “stick” to the sides of any container, and some may float until the 70% ethanol stage. This issue can be a particular problem when removing the previous solution at any exchange step, as the samples may adhere to the transfer pipette or tip. If this happens, a small amount of the next stage

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solution can be used to wash the adherent piece(s) carefully back into the processing tube. 21. To avoid any possibility of crossover between samples (i.e., between different genotypes or conditions), always employ good laboratory practice and never use the same pipette to process different samples. 22. All traces of phosphate must be removed from the samples at this stage. Residual phosphate may lead to uranyl phosphate precipitation, resulting in electron-dense deposits (21). 23. Analytical grade ethanol should be opened immediately before use, and to limit atmospheric water absorption, the stock bottle should not be left open for any length of time. We advise the purchasing of bottles of the minimal volume available (usually 100 mL). 24. 1,2-Epoxypropane (PO) is extremely volatile; this poses two significant problems during pipetting. Firstly, any liquid drawn into a pipette starts to evaporate rapidly, filling the bulb with vapour which rapidly forces the liquid back out of the pipette. To prevent this, fill and drain the pipette two to three times (thus saturating the pipette bulb with vapour) before acquiring the required volume. The second problem can be the formation of condensation on the outside of the transfer pipette; this must never be allowed to enter either the sample tube or the stock bottle. 25. Waste PO/resin mixtures may be left, uncapped, in the fume cupboard to allow the PO to evaporate. All of the waste resin may ultimately be combined and polymerised as per the sample blocks before appropriate disposal. 26. Labels should be a paper band 3–4 mm deep with pencil writing (most inks will be removed by the resin), and this should be positioned around the circumference, half way down the capsule. 27. To transfer each piece of leaf/cotyledon to an embedding capsule, modify a 7-mL transfer pipette by cutting off ~0.5  cm from the tip. This gives a tube of larger diameter that should be sufficiently wide to draw each sample piece, individually, with just a small amount of the resin for transfer to the capsule. 28. If the specimen is located to one side of the capsule, to simplify later remounting, lay the filled/closed embedding capsule on its side. The sample, visible through the base of the capsule, should slowly descend to the centre of the capsule (this should take no more than 2–3 min). When achieved, return the capsule to a vertical orientation. Alternatively, before closing the lid, a toothpick or similar may be used to position the sample very carefully.

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29. If deemed necessary because of size or visibility, a fine-tip marker pen may be used to draw a trapezium on the resin block face enclosing the sample. This may assist subsequent sawing/ removal. Ensure that the longer of the two parallel edges is at 90° to the leaf stem (see Fig. 2). 30. Sawing of resin blocks should be conducted in a fume cupboard to eliminate the risk of inhaling dust. 31. If a nitrogen atmosphere chamber is not available, lead citrate staining may be conducted using a staining (Conway) dish comprising a central wax bed surrounded by an annulus “moat” of sodium hydroxide pellets in distilled water, and covered with a ground glass plate/lid. 32. EM grids can be very susceptible to static charge. For this reason, it is advisable to store them in a dedicated grid storage box rather than in a Petri dish, where they may be readily attracted to the lid of the dish.

Acknowledgments The authors wish to acknowledge Ms. Natalie Allcock of the Core Biotechnology Services Electron Microscopy Laboratory, University of Leicester, for both technical support and comments on the manuscript. References 1. Whatley, J. M. (1978) A suggested cycle of plastid developmental interrelationships. New Phytol. 80, 489–502. 2. López-Juez, E., and Pyke, K. A. (2005) Plastids unleashed: their development and their integration in plant development. Int. J. Dev. Biol. 49, 557–577. 3. Kinsman, E. A., and Pyke, K. A. (1998) Bundle sheath cells and cell-specific plastid development in Arabidopsis leaves. Development 125, 1815–1822. 4. Schelbert, S., Aubry, S., Burla, B., Agne, B., Kessler, F., Krupinska, K., and Hörtensteiner, S. (2009) Pheophytin pheophorbide hydrolase (pheophytinase) is involved in chlorophyll breakdown during leaf senescence in Arabidopsis. Plant Cell 21, 767–785. 5. Kubis, S., Patel, R., Combe, J., Bédard, J., Kovacheva, S., Lilley, K., Biehl, A., Leister, D., Ríos, G., Koncz, C., and Jarvis, P. (2004) Functional specialization amongst the Arabidopsis Toc159 family of chloroplast protein import receptors. Plant Cell 16, 2059–2077.

6. Kuo, J. (2007) Electron Microscopy: Methods and Protocols. Methods in Molecular Biology, Vol. 369. Humana Press, Totowa, NJ, USA. 7. Pfeiffer, S., and Krupinska, K. (2005) Chloroplast ultrastructure in leaves of Urtica dioica L. analyzed after high-pressure freezing and freeze-substitution and compared with conventional fixation followed by room temperature dehydration. Microsc. Res. Tech. 68, 368–376. 8. Hayat, M. A. (1981) Fixation for Electron Microscopy. Academic Press, New York, USA. 9. Glauert, A. M., and Lewis, P. R. (1998) Biological Specimen Preparation for Transmission Electron Microscopy. Practical Methods in Electron Microscopy, Vol. 17. Portland Press Ltd., London, UK. 10. 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. 11. Ellis, E. A. (2006) Solutions to the problem of substitution of ERL 4221 for vinyl cyclohexene

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dioxide in Spurr low viscosity embedding formulations. Microscopy Today 14, 32–33. 12. Reynolds, E. S. (1963) The use of lead citrate at high pH as an electron-opaque stain in electron microscopy. J. Cell Biol. 17, 208–212. 13. Ellis, E. A. (2007) Poststaining grids for transmission electron microscopy: conventional and alternative protocols. In, Electron Microscopy: Methods and Protocols (Kuo, J., ed.) Humana Press, Totowa, NJ, USA, pp. 97–106. 14. Hunter, E. (1993) Practical Electron Microscopy: A Beginner’s Illustrated Guide. Cambridge University Press, Cambridge, UK. 15. Kovacheva, S., Bédard, J., Patel, R., Dudley, P., Twell, D., Ríos, G., Koncz, C., and Jarvis, P. (2005) In vivo studies on the roles of Tic110, Tic40 and Hsp93 during chloroplast protein import. Plant J. 41, 412–428. 16. Aronsson, H., Boij, P., Patel, R., Wardle, A., Töpel, M., and Jarvis, P. (2007) Toc64/OEP64 is not essential for the efficient import of proteins

into chloroplasts in Arabidopsis thaliana. Plant J. 52, 53–68. 17. Hall, J. L., and Hawes, C. (1991) Electron Microscopy of Plant Cells. Academic Press, London, UK. 18. Bozzola, J. J. (2007) Conventional specimen preparation techniques for transmission electron microscopy of cultured cells. In, Electron Microscopy: Methods and Protocols (Kuo, J., ed.) Humana Press, Totowa, NJ, USA, pp. 1–18. 19. Spurr, A. R. (1969) A low-viscosity epoxy resin embedding medium for electron microscopy. J. Ultrastruct. Res. 26, 31–43. 20. Kuo, J. (2007) Processing plant tissues for ultrastructural study. In, Electron Microscopy: Methods and Protocols (Kuo, J., ed.) Humana Press, Totowa, NJ, USA, pp. 35–45. 21. Louw, J., Williams, K., Harper, I. S., and WalfeCoote, S. A. (1990) Electron dense artefactual deposits in tissue sections: the role of ethanol, uranyl acetate and phosphate buffer. Stain Technol. 65, 243–250.

Chapter 9 Transplastomics in Arabidopsis: Progress Toward Developing an Efficient Method Kerry Ann Lutz, Arun Azhagiri, and Pal Maliga Abstract Protocols developed for plastome engineering in Nicotiana tabacum rely on biolistic delivery of the transforming DNA to chloroplasts in intact leaf tissue; integration of the foreign DNA into the plastid genome by homologous recombination via flanking plastid DNA (ptDNA) targeting regions; and gradual dilution of non-transformed ptDNA during cultivation in vitro. Plastid transformation in Arabidopsis was obtained by combining the tobacco leaf transformation protocol with Arabidopsis-specific tissue culture and plant regeneration protocols. Because the leaf cells in Arabidopsis are polyploid, this protocol yielded sterile plants. Meristematic cells in a shoot apex or cells of a developing embryo are diploid. Therefore, we developed a regulated embryogenic root culture system that will generate diploid tissue for plastid transformation. This embryogenic culture system is created by steroid-inducible expression of the BABY BOOM transcription factor. Plastid transformation in Arabidopsis will enable the probing of plastid gene function, and the characterization of posttranscriptional mechanisms of gene regulation and the regulatory interactions of plastid and nuclear genes. Key words: Arabidopsis thaliana, BABY BOOM, Dexamethasone, Plastid transformation, Plant regeneration, Steroid-inducible gene expression, Tissue culture

1. Introduction The plastid genome (plastome or ptDNA) of Arabidopsis thaliana is highly polyploid. A leaf cell contains ~120 chloroplasts and a total of 1,000–1,700 ptDNA copies per cell (1, 2). The ptDNA is ~145.5 kb in size (3), with significant sequence variation in intergenic regions that enables distinction of the ptDNA of different ecotypes (4–6). In contrast, genes encoded in the ~367-kb mitochondrial genome (mtDNA) (7) are present in 40–280 copies per leaf cell shared by 300–450 mitochondria, suggesting that in leaf cells,

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the number of mtDNA copies may be lower than the number of organelles and that the genes may be present on non-stoichiometric subgenomic molecules rather than on master chromosomes (8). The ~100 genes encoded in the organellar genomes complement the estimated 29,454 nuclear genes encoded in the 125-Mb nuclear genome, of which ~10% targets proteins to plastids and another 10% to mitochondria (9, 10). The plant’s small size, rapid life cycle, and the available genomic resources make Arabidopsis thaliana the most advanced model species in flowering plants. Transformation of the nuclear genome of Arabidopsis is routine using tissue cultureindependent flower dip protocols (11). Transformation of the Arabidopsis plastid genome has been accomplished (12), but the protocol does not reproducibly yield fertile plants. Transformation of the mitochondrial genome in Arabidopsis, and in all flowering plant species, remains elusive, although a routine protocol has been developed in Chlamydomonas reinhardtii, a unicellular alga (13). Below, we give an overview of the current state of plastid transformation followed by a protocol that is most likely to overcome the limitations of the published Arabidopsis plastid transformation protocol. Protocols developed in our laboratory for plastome engineering in Nicotiana tabacum rely on biolistic delivery of the transforming DNA to chloroplasts in intact leaf tissue; integration of the foreign DNA into the plastid genome by homologous recombination via flanking ptDNA targeting regions; and gradual dilution of non-transformed ptDNA with transformed copies during protracted (3–12 weeks) cultivation in vitro (14–16). Because the plastid genome is polyploid, selection for marker genes is essential to obtain genetically uniform plants. Marker genes that are suitable for selective enrichment of plastid genomes confer resistance to spectinomycin and streptomycin (17), kanamycin (18–20), or the amino acid analogs 4-methylindole (4MI) and 7-methyl-DL-tryptophan (7MT) (21). Genetically stable, homoplastomic plants are obtained through a gradual process of ptDNA replication and sorting, and preferential maintenance of transgenic ptDNA copies on antibiotic-containing tissue culture medium. Because selective enrichment of transformed ptDNA is carried out in tissue culture, a suitable protocol for plant regeneration is essential to obtain transformed plants. Fig. 1 shows a schematic representation of the Arabidopsis plastid transformation vector pKAL1, the targeted region of the plastid genome (At-wt), and the resulting transplastome after integration of the aadA gene encoding streptomycin/ spectinomycin resistance. In Fig. 1, the aadA gene is flanked by triangles symbolizing loxP target sites recognized by the CRE sitespecific recombinase. Marker genes flanked by loxP target sites are stable in the plastid genome in the absence of the recombinase. However, post-transformation excision of the marker gene is readily obtained when a uniform population of transgenic ptDNA

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Fig. 1. Plastid transformation in Arabidopsis thaliana with vector pKAL1. Maps of the plastid targeting region of vector pKAL1, the cognate region in the wild-type Arabidopsis plastid genome (At-Wt), and transplastomic DNA (At-pKAL1) after integration of the plasmid DNA by homologous recombination are shown. rps12/7, trnV, and rrn16 are plastid genes. The aadA coding region is represented by a red box; a c-myc tag is represented as a black box at the 3¢-end of the aadA coding region. The black box upstream of the rrn16 gene is the native Prrn promoter.

is obtained. Excision of the loxP-flanked plastid marker genes is achieved by a plastid-targeted CRE recombinase expressed from a nuclear Cre gene. The Cre gene is introduced by Agrobacteriummediated transformation of the nucleus, or by crossing (14). Thus, after marker excision, only one loxP site and the gene of interest remain in the transplastome. The Cre gene can then be segregated away in the seed progeny. Plastid transformation in Arabidopsis was obtained by combining the tobacco leaf transformation protocol with Arabidopsisspecific tissue culture and plant regeneration protocols (12). Because the leaf cells in Arabidopsis are polyploid (1, 22, 23), we ended up with sterile plants. Meristematic cells in a shoot apex or cells of a developing embryo are diploid. We, therefore, developed an embryogenic culture system for plastid transformation in Arabidopsis. Constitutive expression of BABY BOOM (BBM), a member of the AP2/ERF family of transcription factors, was reported to sustain spontaneous production of somatic embryos

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but did not yield fertile plants (24–26). Therefore, we decided to regulate BBM function by fusing BBM with the glucocorticoid receptor steroid-binding domain. Direct fusion of plant transcription factors with a steroid receptor has been successfully employed to regulate expression, for example, of the maize transcriptional regulator R (27), and recently BBM (28). Although the fusion proteins are constitutively expressed, transcription of the downstream target genes is dependent on the supply of steroid hormones in the culture medium. In the absence of the steroid hormone, the fusion protein remains in the cytoplasm. Association of steroid with the hormone-binding domain of the fusion protein allows for its translocation into the nucleus where it can activate transcription (29). The protocol we describe here for plastid transformation utilizes Arabidopsis lines carrying the pKO216 steroidinducible BBM construct in the RLD and Landsberg erecta (Ler) backgrounds (to be described elsewhere). Ongoing experiments aim at refining plastid transformation in Arabidopsis thaliana in steroid-inducible BBM root cultures. We already obtained fertile spectinomycin-resistant Arabidopsis mutants using the protocol reported here. Although no plastid transformants have been obtained to date, we believe that the improved tissue culture system will yield fertile plants. Fertile transplastomic plants have been reported in the related Brassica napus (30, 31), Brassica oleracea (cabbage) (32), Brassica oleracea var. botrytis (cauliflower) (33), and Lesquerella fenderli (34). The opportunity to express multiple genes in operons, exclusive uniparental-maternal inheritance, and readily obtainable high protein levels are the driving force behind engineering the plastid genome for applications in agriculture, metabolic engineering, and molecular pharming (35, 36). In the model plant Arabidopsis, plastid transformation will enable the probing of plastid gene function, translational regulation, and the regulatory interactions of plastid and nuclear genes.

2. Materials 2.1. Propagation of Arabidopsis Roots in Culture

1. Transgenic Arabidopsis thaliana lines stably transformed with Agrobacterium binary plasmid pKO216 containing a steroidinducible BBM construct. The lines RLD-pKO216-22-2, LerpKO216-T4-10, and Ler-pKO216-2-31 are designated by the ecotype, the plasmid name, and a serial number, and are nonsegregating, gentamycin-resistant lines. 2. Agar, plant tissue culture tested (Sigma, St. Louis, MO, USA). 3. Screening medium (SCM), modified from Zuo et al. (37). Per liter add 100  mL of 10× macronutrient solution, 10  mL of

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100× micronutrient solution, 2% (w/v) glucose, 0.8% (w/v) agar, and 5  mL of 1% (w/v) Fe-ethylenediaminetetraacetic acid (Fe-EDTA); adjust pH to 5.7 using 1 M KOH. Prepare and filter-sterilize vitamin stock solution (10 mg/mL vitamin B1, 1 mg/mL vitamin B6, 1 mg/mL nicotinic acid, and 1 mg/mL glycine). After the medium has cooled to 45°C following autoclaving, add 1 mL of vitamin stock solution, 1.5 mL of 1 mg/mL indoleacetic acid (IAA) stock, 0.3 mL of 1 mg/mL isopentenyladenine (IPA) stock, and 5  mL of 100  mg/mL 2-(N-morpholino)-ethanesulfonic acid (MES) stock. 4. Arabidopsis regeneration medium B (ARM-B) (38) plant tissue culture medium, based on Murashige and Skoog (MS) salts medium (39). Per liter add 100  mL of 10× macronutrient solution, 10  mL of 100× micronutrient solution, 3% (w/v) sucrose, 5 mL of 1% (w/v) Fe-EDTA, and 200 mg of myoinositol; adjust pH to 5.8 using 1 M KOH. Prepare and filtersterilize a stock solution of biotin (0.1  mg/mL). After the medium has cooled to 45°C following autoclaving, add 1 mL of biotin stock solution, 1 mL of vitamin stock solution (see point 3 above), 3 mg of IAA, 0.6 mg of benzyladenine (BA), and 0.3 mg of IPA. For solid ARM-B media, add 0.8% (w/v) agar before autoclaving. 5. All stock solutions of plant hormones were made as described below, filter-sterilized, and added to media cooled to 45°C after autoclaving. To prepare a 1  mg/mL stock solution of IAA, dissolve 0.1 g of IAA powder into 0.5 mL of 1 N KOH. Once dissolved, slowly add warm water up to 100 mL. To prepare a 1 mg/mL stock solution of BA, dissolve 0.1 g of BA powder in 0.5  mL of 1  N HCl. Once dissolved, slowly add warm water up to 100 mL. To prepare a 1 mg/mL stock solution of IPA, dissolve 0.1 g of IPA powder into 0.5 mL of 1 N HCl. Once dissolved, slowly add warm water up to 100 mL. 6. 10× Macronutrient solution. Per liter add 19 g KNO3, 3.7 g MgSO4·7H2O, 4.4 g CaCl2·2H2O, 1.7 g KH2PO4, and 16.5 g NH4NO3. Autoclave for long-term storage. 7. 100× Micronutrient solution. Per liter add 1,560  mg MnSO4·2H2O, 620 mg H3BO3, 860 mg ZnSO4·7H2O, 83 mg KI, 2.5  mL of Na2MoO4·2H2O (10  mg/mL), 2.5  mL of CuSO4·5H2O (1  mg/mL), and 0.25  mL of CoCl2·6H2O (1 mg/mL). Autoclave for long-term storage. 8. Culture room, with temperature set to 22°C and cool white fluorescent bulbs providing illumination for 16  h per day at 2000 lux (~40 mmol photons/m2/s). 9. Desiccator, heavy glass (Fisher Scientific, Pittsburgh, PA, USA). 10. Dexamethasone (DEX; Sigma). Prepare a 10 mM stock solution by dissolving 4 mg DEX in 1 mL of dimethyl sulfoxide (DMSO).

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Filter-sterilize with a 0.2-mm, 25-mm DMSO-Safe Acrodisc® syringe filter (Pall Corporation, Port Washington, NY, USA). Aliquot into 1  mL volumes and place at 4°C; note that the solution will freeze at 4°C. DEX should be used at 5 mM final concentration in the culture medium. 11. DMSO-Safe Acrodisc® syringe filter, 0.2  mm, 25  mm, sterile (Pall Corporation; catalog no. 4433). 12. 70% (v/v) ethanol. 13. ClingWrap (The Glad Products Co., Oakland, CA, USA), cut into 2.5-cm strips. 14. Gyratory shaker. 15. 125-mL glass flasks with cotton stoppers. 16. Tissue culture plates, 100 × 20  mm (Sarstedt, Newton, NC, USA). 2.2. Coating Gold Particles with Plasmid DNA

1. Spermidine-free base, 0.1 M stock (Sigma). 2. Gold microcarrier (Bio-Rad Laboratories, Hercules, CA, USA; catalog no. 165–2262). 3. 70% (v/v) ethanol. 4. 50% (w/v) glycerol. 5. 5 mg plasmid DNA (such as pKAL1; see Fig. 1). 6. 2.5 M CaCl2. 7. 100% Ethanol. 8. Fisher Vortex Genie 2 (catalog no. 12–812), or equivalent.

2.3. Introduction of Plasmid DNA into Arabidopsis Chloroplasts

1. Helium, 99.999% pure, moisture free. 2. ClingWrap (The Glad Products Co.), cut into 2.5-cm strips. 3. ARM5 (38), based on MS salts medium (39). Per liter add 100 mL of 10× macronutrient solution, 10 mL of 100× micronutrient solution, 5% (w/v) sucrose, 0.8% (w/v) agar, 5 mL of 1% (w/v) Fe- EDTA, and 200 mg myo-inositol; adjust pH to 5.8 using 1 M KOH. After the medium has cooled to 45°C following autoclaving, add 1  mL biotin stock solution and 1 mL of vitamin stock solution (see Subheading 2.1). 4. Particle gun macrocarriers (Bio-Rad Laboratories; catalog no. 165–2335). 5. Particle gun PDS-1000/He (Bio-Rad Laboratories; catalog no. 165–2257). A suitable vacuum pump for use with the gun is the Thermo Savant VLP285 (Thermo Savant, Holbrook, NY, USA). 6. Particle gun rupture disks, 1,100 psi (Bio-Rad Laboratories; catalog no. 165–2329). 7. Particle gun stopping screens (Bio-Rad Laboratories; catalog no. 165–2336).

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1. Materials listed in Subheading 2.1. 2. ARM. This plant tissue culture medium is identical with the ARM5 medium (see Subheading 2.3) except that it contains only 3% (w/v) sucrose. 3. Magenta GA7 vessels (Sigma; catalog no. V8505-100EA) (see Note 1). 4. Secador 4.0 auto-desiccator cabinet (ISC BioExpress, Kaysville, UT, USA).

3. Methods 3.1. Propagation of Arabidopsis Roots in Culture

This protocol describes propagation of Arabidopsis roots in sterile culture that represent suitable target tissue for plastid transformation and readily regenerate fertile plants. The protocol was developed using information provided in refs. 37 and 38. 1. To vapor-sterilize seeds, place a 1.5-mL Eppendorf tube with ~50 seeds along with a flask containing 100 mL of bleach into a glass desiccator located in a fume hood (see Note 2). Then add 3 mL of concentrated (37.8%) HCl directly to the bleach already in the flask in the desiccator. Close the desiccator lid. 2. Let vapor-sterilization proceed for between 3 and 16 h. The time needed will vary based the extent to which seeds are contaminated. Longer vapor-sterilization times can result in killing of some of the seeds, especially if the seed is not fully mature and dry. 3. When sterilization is complete, open the desiccator in a fume hood and close the Eppendorf tube containing the seed. Open the tube in sterile laminar flow hood and sprinkle the seed onto SCM plates containing 5 mM DEX (see Note 3). 4. Place the plates at 4°C for three days to break seed dormancy and then transfer them to the culture room (16-h light cycle; 22°C). 5. Three weeks after germination, excise hypocotyls with roots from ~25 seedlings and transfer them into a 125-mL flask with 25 mL liquid ARM-B medium containing 5 mM DEX (i.e., 25 individuals per flask). Incubate the flasks on a gyratory shaker (90 rpm) at 22°C under low-intensity illumination in the culture room. Subculture the tissue every 2 weeks into new 125-mL flasks containing fresh ARM-B + DEX medium. Excise and discard all green tissue from each root culture before transferring into a new flask. If necessary, split the tissue into two flasks to ensure availability of nutrients. 6. Two months after germination, there should be enough tissue for bombardment (see below).

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3.2. Coating Gold Particles with Plasmid DNA

With one DNA construct, 20–30 tissue samples are bombarded. Each tube (30 mg) of gold is sufficient for 50 bombardments (two DNA constructs). You may use freshly prepared gold particles, or stored gold. If using stored gold, vortex the tube for 5 min before coating with DNA. This protocol was modified from Bio-Rad Bulletin 9075. 1. Weigh 30  mg of 0.6  mm gold microcarrier in a 1.5-mL Eppendorf tube and add 1 mL of ice-cold 70% ethanol. 2. Place the tube in a vortex microtube holder and vortex vigorously for 5 min. Let the particles settle at room temperature (20–25°C) for 15 min. 3. Spin in a microcentrifuge at 3,000 rpm (600 × g) for 1 min to compact the gold. 4. Remove the ethanol with a pipette and add 1 mL of ice-cold, sterile distilled water. 5. Vortex the tube to suspended the particles. Allow the particles to settle at room temperature for 10 min. 6. Sediment the gold by spinning in a microcentrifuge at 3,000 rpm (600 × g) for 1 min. 7. Remove the water with a pipette and add 1  mL of ice-cold, sterile distilled water. 8. Wash the gold one more time with water by repeating steps 5–7. 9. Resuspended the particles by vortexing and store the tube at room temperature for 10 min to let the particles settle. 10. Microcentrifuge at 5,000  rpm (1,700 × g) for 15  s and then remove the water completely. 11. Add 500 mL of 50% glycerol and vortex for 1 min to resuspend the particles. The gold concentration will be 60 mg/mL. Clean gold can be stored for 2 weeks at room temperature. If the gold has been stored, place the Eppendorf tube containing gold in a vortex microtube holder and shake at setting 3 (moderate speed) for 5 min. 12. While the tube is shaking, remove 50 mL aliquots of gold and pipette them into ten 1.5-mL Eppendorf tubes in a rack. 13. Place the tubes containing the gold aliquots into a vortex microtube holder and shake at setting 3. While the tubes are shaking, add 5 mL of DNA (1 mg/mL), 50 mL of 2.5 M CaCl2, and 20  mL of 0.1  M spermidine-free base. Make sure to add these components in the order described, and that the contents are thoroughly mixed before adding the next component. 14. Shake the tubes on a vortex at setting 3 for 5 min.

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15. Sediment the gold by spinning in a microcentrifuge at 3,000 rpm (600 × g) for 1 min. 16. Discard the supernatant and add 140  mL of 70% ethanol to each tube. 17. Tap the tube lightly until the pellet just becomes loose, to make sure that the pellet is not tightly packed. If the gold does not go into solution by gently tapping the tube, break the pellet by pipetting up and down. 18. Sediment the gold by spinning in a microcentrifuge at 3,000 rpm (600 × g) for 1 min. 19. Remove the supernatant and add 140  mL of ice-cold 100% ethanol to each tube. 20. Lightly tap the tube until the pellet just starts to come into solution. 21. Sediment the gold by spinning in a microcentrifuge at 5,000 rpm (1,700 × g) for 15 s. 22. Resuspend the coated gold pellet in 50 mL of 100% ethanol by gently tapping the tube. The pellet should easily enter the solution. Shake the tubes on a vortex at setting 3 while waiting to use them for bombardment. If the tubes are left sitting for a long period of time before bombardment, replace the ethanol in the tube with fresh 100% ethanol. 3.3. Introduction of Plasmid DNA into Arabidopsis Chloroplasts

The protocol we describe here is a modified version of the protocol routinely used for transformation of tobacco leaves using the Bio-Rad PDS1000/He biolistic gun. With this gun, 20–30 plates containing Arabidopsis tissue are bombarded per DNA construct. 1. Grow BBM root culture for bombardment as described above (see Subheading 3.1). 2. Remove the root tissue for biolistic transformation from the flask and dab it dry on a stack of sterile Whatman No. 4 filter papers. Place the tissue on top of two sterile Whatman No. 4 filter papers on a Petri plate (10 cm) containing 20 mL of solid ARM5 + DEX medium. If bombardment is to be done the next day, place a plastic bag over the plates and incubate them in the culture room overnight. 3. Bombardment with the gun should be carried out in a sterile laminar flow hood. Before bombardment, sterilize the gun main chamber, rupture disk retaining cap, microcarrier launch assembly, and target shelf by wiping off with a cloth soaked in 70% ethanol. 4. Sterilize the rupture disks (1,100 psi), macrocarriers, macrocarrier holders, and stopping screens by soaking in 100% ethanol (for 5  min), and then air-dry them in the tissue culture hood in an open container.

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5. Turn on the helium tank and set the delivery pressure in the regulator (distal to the tank) to 1,300  psi (i.e., 200–300 psi above the rupture disk value). 6. Turn on the vacuum pump and gene gun. Set the vacuum rate on the gene gun to 7 and the vent rate to 2. 7. Prepare DNA-coated gold particles as described above (see Subheading  3.2). Pipette 10  mL of DNA-coated gold onto one flying disk (placed in holder) and allow it to air-dry for 5 min. Five samples may be made up at one time. 8. Place a rupture disk into the retaining cap and screw in tightly. 9. Put a stopping screen and a flying disk (face down) into the microcarrier launch assembly and place it into the chamber just below the rupture disk. For a more detailed description, see Bio-Rad Bulletin 9075. 10. Place the plate with the tissue into the chamber 9 cm below the microcarrier launch assembly (fourth shelf from the top) and close the door. 11. Press the vacuum button to open the valve. When the vacuum reaches 28  in. Hg, hold down the fire button until the pop from the gas breaking the rupture disk is heard. 12. Immediately release the vacuum and remove the sample (see Note 4). 13. Repeat steps 7–12 until all samples are bombarded. When finished, turn off the helium tank and release the pressure by holding down the fire button while the vacuum is on. Turn off the vacuum pump. 14. Place a plastic bag over plates containing the bombarded samples and incubate them in the culture room (conditions as in Subheading 3.1) for up to 40 h. This incubation allows time for marker gene expression to commence before selection is started. 3.4. Selection and Identification of Transplastomic Clones by Streptomycin/ Spectinomycin Resistance

We describe below a protocol for the selection of plastid transformants using the Arabidopsis BBM root cultures. This protocol is modified from one that routinely yields plastid transformants in bombarded tobacco leaf cultures. Transplastomic tobacco clones are identified as regenerating shoots on a bleached, callusing leaf. In the BBM root system, we expect recovery of transplastomic Arabidopsis clones by formation of green embryogenic callus. 1. Two days after bombardment, split the bombarded tissue into two to three 125-mL flasks each holding 25  mL of ARM-B medium containing DEX and spectinomycin (25 mg/L). 2. Incubate the flasks in the culture room on a gyratory shaker. The tissue should be transferred to new flasks with fresh medium

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Fig.  2. Plastid transformation using Arabidopsis root tissue. A spectinomycin-resistant, transplastomic line identified as green embryogenic callus (white arrow ) in root culture is shown.

every 2 weeks. If necessary, continue splitting the root culture into two to three flasks to ensure availability of nutrients. 3. Green, spectinomycin-resistant tissue may form in the root cultures. Figure  2 shows such green, spectinomycin-resistant embryogenic callus in an Ler-216-2 BBM root culture. In bombarded tobacco leaf cultures, spectinomycin-resistant clones appear within 4–12 weeks after bombardment. When the Arabidopsis tissue is big enough (about the size of a quarter; ~25 mm across), cut the embryogenic callus into smaller pieces (1 mm2) and transfer them onto solid ARM-B medium containing DEX and 100 mg/L spectinomycin. Each sector at a distinct location derives from an independent transformation event and, therefore, is an independently derived clone. We identify putative plastid transformants by the plasmid name and a unique serial number. 4. Spectinomycin resistance may be due to expression of the aadA gene incorporated from the vector, or to a spontaneous mutation in the plastid small ribosomal RNA (rrn16) gene (17). The aadA gene confers resistance to spectinomycin and streptomycin, whereas spontaneous spectinomycin-resistant mutants are resistant only to spectinomycin (17). Therefore, transgenic clones can be distinguished from mutants by testing a sample of each clone for resistance to streptomycin and spectinomycin. For testing, place a piece of the spectinomycin-resistant

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tissue on ARM5 medium containing 25 mg/L spectinomycin and 25 mg/L streptomycin. Since streptomycin delays shoot regeneration, continue subculturing the remaining spectinomycin-resistant tissue on solid ARM-B medium containing 100  mg/L spectinomycin to obtain homoplastomic tissue. The aadA gene rarely inserts and expresses in the nuclear genome. Transplastomic clones should be positively identified by confirming incorporation of aadA in the plastid genome by DNA gel blot analysis. 5. Clones that are resistant to streptomycin and spectinomycin should be regenerated into plants using the tissue on the plates containing spectinomycin only. Plants regenerated from the same clone (initial transformant) are considered subclones and we distinguish them by adding letters to the clone number. Southern blot analysis will show that some of these plants are homoplastomic (all ptDNA copies transformed), that some are heteroplastomic (containing transformed and wild-type ptDNA copies), and that some contain only wild-type ptDNA copies. 6. Embryogenic callus confirmed to have transformed plastids should be transferred to Magenta boxes containing ARM medium without DEX. After about 2 months, some callus will form shoots, flower, and produce seed. Shoots that flower and produce seed will not develop roots. Seed can be collected when the tissue has completely dried out (approximately 1 month after seed formation) (see Note 5). 7. Seed collected should be placed in a 1.5-mL Eppendorf tube in a Secador desiccator for 1 week to dry the seed completely. 8. Germinate sterilized seed on ARM medium containing spectinomycin (25 mg/L). Homoplastomic seedlings will be dark green, whereas sensitive seedlings will be white. Observing 100% green seedlings confirms the homoplasmic state of the parental plant; no segregation for spectinomycin resistance should be seen.

4. Notes 1. Sigma is currently not selling Magenta boxes with vented lids. Vented lids can be made by cutting a round hole (18 mm) on a Magenta lid and closing it with an autoclavable Sun cap closure that is available from Sigma (catalog no. S5939). 2. Chlorine gas is poisonous to humans; therefore, work with proper ventilation. 3. Be sure to open and vent the tube after vapor-sterilization; otherwise the seed rapidly loses viability upon storage.

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4. If the gun is fired at lower pressure, DNA-coated particles will lack sufficient momentum to penetrate cells and no transplastomic lines will be obtained. If you have no experience with biolistic transformation, we recommend that you first test the particle coating and DNA delivery procedures by using transient expression of a nuclear uidA gene, which encodes the b-glucuronidase enzyme, the activity of which can be readily detected by histochemical staining (40, 41). 5. Be careful when handling boxes to prevent seed from falling out of the seed pods.

Acknowledgments This research was supported by Grant MCB-039958 from the NSF Eukaryotic Genetics Program. K.A.L. was the recipient of a Charles and Johanna Busch Predoctoral Fellowship. References 1. Zoschke, R., Liere, K., and Börner, T. (2007) From seedling to mature plant: Arabidopsis plastidial genome copy number, RNA accumulation and transcription are differentially regulated during leaf development. Plant J. 50, 710–722. 2. Pyke, K. A., and Leech, R. M. (1994) A genetic analysis of chloroplast division and expansion in Arabidopsis thaliana. Plant Physiol. 104, 201–207. 3. Sato, S., Nakamura, Y., Kaneko, T., Asamizu, E., and Tabata, S. (1999) Complete structure of the chloroplast genome of Arabidopsis thaliana. DNA Res. 6, 283–290. 4. Provan, J., and Campanella, J. J. (2003) Patterns of cytoplasmic variation in Arabidopsis thaliana (Brassicaceae) revealed by polymorphic chloroplast microsatellites. Syst. Bot. 28, 578–583. 5. Sall, T., Jakobsson, M., Lind-Hallden, C., and Hallden, C. (2003) Chloroplast DNA indicates a single origin of the allotetraploid Arabidopsis suecica. J. Evol. Biol. 16, 1019–1029. 6. Azhagiri, A., and Maliga, P. (2007) DNA markers define plastid haplotypes in Arabidopsis thaliana. Curr. Genet. 51, 269–275. 7. Unseld, M., Marienfeld, J. R., Brandt, P., and Brennicke, A. (1997) The mitochondrial genome of Arabidopsis thaliana contains 57 genes in 366,924 nucleotides. Nat. Genet. 15, 57–61.

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biolistic transformation of Chlamydomonas mitochondria can be used to insert mutations in complex I genes. Proc. Natl. Acad. Sci. USA 103, 4771–4776. 1 4. Lutz, K. A., Svab, Z., and Maliga, P. (2006) Construction of marker-free transplastomic tobacco using the Cre-loxP site-specific recombination system. Nat. Protocols 1, 900–910. 15. Lutz, K. A., and Maliga, P. (2007) Transformation of the plastid genome to study RNA editing. Methods Enzymol. 424, 501–518. 16. Maliga, P., and Svab, Z. (2010) Engineering the plastid genome of Nicotiana sylvestris, a diploid model species for plastid genetics. In, Plant Chromosome Engineering: Methods and Protocols (Birchler, J. J., ed.), Humana Press, Totowa, NJ, USA, in press. 17. Svab, Z., and Maliga, P. (1993) High-frequency plastid transformation in tobacco by selection for a chimeric aadA gene. Proc. Natl. Acad. Sci. USA 90, 913–917. 18. Carrer, H., Hockenberry, T. N., Svab, Z., and Maliga, P. (1993) Kanamycin resistance as a selectable marker for plastid transformation in tobacco. Mol. Gen. Genet. 241, 49–56. 19. Huang, F. C., Klaus, S. M. J., Herz, S., Zuo, Z., Koop, H. U., and Golds, T. J. (2002) Efficient plastid transformation in tobacco using the aphA-6 gene and kanamycin selection. Mol. Genet. Genomics 268, 19–27. 20. Lutz, K., Corneille, S., Azhagiri, A. K., Svab, Z., and Maliga, P. (2004) A novel approach to plastid transformation utilizes the phiC31 phage integrase. Plant J. 37, 906–913. 21. Barone, P., Zhang, X. H., and Widholm, J. M. (2009) Tobacco plastid transformation using the feedback-insensitive anthranilate synthase [a]-subunit of tobacco (ASA2) as a new selectable marker. J. Exp. Bot. 60, 3195–3202. 22. Galbraight, D. W., Harkins, K. R., and Knapp, S. (1991) Systemic endopolyploidy in Arabidopsis thaliana. Plant Physiol. 96, 985–989. 23. Melaragno, J. E., Mehrotra, B., and Coleman, A. W. (1993) Relationship between endopolyploidy and cell size in epidermal tissue of Arabidopsis. Plant Cell 5, 1661–1668. 24. Nole-Wilson, S., Tranby, T. L., and Krizek, B. A. (2005) AINTEGUMENTA-like (AIL) genes are expressed in young tissues and may specify meristematic or division-competent states. Plant Mol. Biol. 57, 613–628. 25. Riechmann, J. L., Heard, J., Martin, G., Reuber, L., Jiang, C.-Z., Keddie, J., Adam, L., Pineda, O., Ratcliffe, O. J., Samaha, R. R., Creelman, R., Pilgrim, M., Broun, P., Zhang, J.

Z., Ghandehari, D., Sherman, B. K., and Yu, G. L. (2000) Arabidopsis transcription factors: genome-wide comparative analysis among eukaryotes. Science 290, 2105–2110. 26. Boutilier, K., Offringa, R., Sharma, V. K., Kieft, H., Ouellet, T., Zhang, L., Hattori, J., Liu, C. M., van Lammeren, A. A. M., Miki, B. L. A., Custers, J. B. M., and van Lookeren Campagne, M. M. (2002) Ectopic expression of BABY BOOM triggers a conversion of vegetative to embryonic growth. Plant Cell 14, 1737–1749. 27. Lloyd, A. M., Schena, M., Walbot, V., and Davis, R. W. (1994) Epidermal cell fate determination in Arabidopsis: patterns defined by a steroid-inducible regulator. Science 266, 436–439. 28. Srinivasan, C., Liu, Z., Heidmann, I., Supena, E. D., Fukuoka, H., Joosen, R., Lambalk, J., Angenent, G., Scorza, R., Custers, J. B., and Boutilier, K. (2007) Heterologous expression of the BABY BOOM AP2/ERF transcription factor enhances the regeneration capacity of tobacco (Nicotiana tabacum L.). Planta 225, 341–351. 29. Zuo, J., and Chua, N. H. (2000) Chemicalinducible systems for regulated expression of plant genes. Curr. Opin. Biotechnol. 11, 146–151. 30. Cheng, L., Li, H. P., Qu, B., Huang, T., Tu, J. X., Fu, T. D., and Liao, Y. C. (2010) Chloroplast transformation of rapeseed (Brassica napus) by particle bombardment of cotyledons. Plant Cell Rep. 29, 371–381. 31. Hou, B. K., Zhou, Y. H., Wan, L. H., Zhang, Z. L., Shen, G. F., Chen, Z. H., and Hu, Z. M. (2003) Chloroplast transformation in oilseed rape. Transgenic Res. 12, 111–114. 32. Liu, C. W., Lin, C. C., Chen, J. J., and Tseng, M. J. (2007) Stable chloroplast transformation in cabbage (Brassica oleracea L. var. capitata L.) by particle bombardment. Plant Cell Rep. 26, 1733–1744. 33. Nugent, G. D., Coyne, S., Nguyen, T. T., Kavanagh, T. A., and Dix, P. J. (2006) Nuclear and plastid transformation of Brassica oleracea var. botrytis (cauliflower) using PEG-mediated uptake into protoplasts. Plant Sci. 170, 135–142. 34. Skarjinskaia, M., Svab, Z., and Maliga, P. (2003) Plastid transformation in Lesquerella fendleri, an oilseed Brassicacea. Transgenic Res. 12, 115–122. 35. Bock, R. (2007) Plastid biotechnology: prospects for herbicide and insect resistance, metabolic engineering and molecular farming. Curr. Opin. Biotechnol. 18, 100–106.

9  Transplastomics in Arabidopsis: Progress Toward Developing an Efficient Method 36. Daniell, H., Kumar, S., and Dufourmantel, N. (2005) Breakthrough in chloroplast genetic engineering of agronomically important crops. Trends Biotechnol. 23, 238–245. 37. Zuo, J., Niu, Q. W., and Chua, N. H. (2002) The WUSCHEL gene promotes vegetative-toembryonic transition in Arabidopsis. Plant J. 30, 349–359. 38. Marton, L., and Browse, J. (1991) Facile transformation of Arabidopsis. Plant Cell. Rep. 10, 235–239.

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39. Murashige, T., and Skoog, F. (1962) A revised medium for the growth and bioassay with tobacco tissue culture. Physiol. Plant 15, 473–497. 40. Jefferson, R. A., Kavanagh, T. A., and Bevan, M. W. (1987) GUS fusions: beta-glucuronidase as a sensitive and versatile gene fusion marker in higher plants. EMBO J. 6, 3901–3907. 41. Gallagher, S. R. (1992) GUS Protocols: Using the GUS Gene as a Reporter of Gene Expression. Academic Press, San Diego, CA, USA.

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Part II Gene Expression and Protein Accumulation

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Chapter 10 Isolation, Quantification, and Analysis of Chloroplast DNA Beth A. Rowan and Arnold J. Bendich Abstract Many areas of chloroplast research require methods that can assess the quality and quantity of chloroplast DNA (cpDNA). The study of chloroplast functions that depend on the proper maintenance and expression of the chloroplast genome, understanding cpDNA replication and repair, and the development of technologies for chloroplast transformation are just some of the disciplines that require the isolation of high-quality cpDNA. Arabidopsis thaliana offers several advantages for studying these processes because of the sizeable collection of mutants and natural varieties (accessions) available from stock centers and a broad community of researchers that has developed many other genetic resources. Several approaches for the isolation and quantification of cpDNA have been developed, but little consideration has been given to the strengths and weaknesses and the type of information obtained by each method, especially with respect to A. thaliana. Here, we provide protocols for obtaining high-quality cpDNA for PCR and other applications, and we evaluate several different isolation and analytical methods in order to build a robust framework for the study of cpDNA with this model organism. Key words: Chloroplast DNA, DNA fluorophore, Fluorescence microscopy, Flow cytometry, Quantitative PCR

1. Introduction The metabolic processes performed by chloroplasts and other plastid types are essential to plants and are achieved through the interactions between the prokaryotic chloroplast genome and the eukaryotic nuclear genome (1, 2). The copy number of the chloroplast genome varies substantially among individual chloroplasts and between populations of chloroplasts during leaf development (3–6). Our understanding of the mechanism responsible for generating such variation depends on the ability to assess accurately the quantity and quality of cpDNA. Additionally, many other fields of

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_10, © Springer Science+Business Media, LLC 2011

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chloroplast research depend on reliable methods for isolating highquality DNA. Although several methods have been developed for quantifying and analyzing cpDNA, there has been little discussion in the literature of either the relative advantages of each method or whether each method provides the same type of information (7). In this chapter, we describe several methods for quantifying and analyzing cpDNA at three levels of organization: individual chloroplasts, populations of chloroplasts, and the fraction of cellular DNA represented by cpDNA. We discuss the type of information obtained and the relative strengths or weaknesses of each approach in order to provide a consistent strategy for researchers interested in determining the quantity and quality of cpDNA. We focus our attention on A. thaliana because it has many genomic and genetic resources. While it might be expected that changes in the quantity of cpDNA among individual chloroplasts or populations of chloroplasts would reflect the amount of cpDNA as a fraction of cellular DNA, this expectation may not be realized, as explained below. We also describe two methods to evaluate the structure and integrity of cpDNA molecules that may be useful for chloroplast transformation (8).

2. Materials 2.1. The Analysis of DNA from Individual Chloroplasts 2.1.1. Chloroplast Isolation

1. Sarkosyl, dissolved in water at 0.5% (w/v). 2. High salt buffer (HSB): 1.25 M NaCl, 40 mM 4-(2-hydroxyethyl)1-piperazineethanesulfonic acid (HEPES)-KOH, pH 7.6, 2  mM ethylenediaminetetraacetic acid (EDTA), 0.1% (w/v) bovine serum albumin (BSA), and 0.1% (v/v) b-mercaptoethanol (bME). Add bME to HSB immediately before use. 3. Miracloth (Calbiochem, San Diego, CA, USA); Percoll (GE Healthcare, Piscataway, NJ, USA). 4. Sorbitol dilution buffer (SDB): 0.33  M sorbitol, 20  mM HEPES-KOH, pH 7.6, 1 mM MgCl2, 2 mM EDTA, and 0.1% (w/v) BSA. 5. 5× SDB for Percoll gradient solutions: 1.65  M sorbitol, 100 mM HEPES-KOH, pH 7.6, 5 mM MgCl2, 10 mM EDTA, and 0.5% (w/v) BSA. 6. Tabletop microcentrifuge (e.g., Eppendorf 5415 D with standard rotor) that can be placed at 4°C. 7. Standard laboratory centrifuge with temperature control (e.g., Beckman J21M with fixed-angle rotor JA-21 [BeckmanCoulter, Brea, CA, USA]; for large preps). 8. Blender that can be cooled to 4°C (e.g., Waring Pro stainless steel blender).

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9. 1.5–50-mL plastic centrifuge tubes. 10. Mortars and pestles (Coors, Golden, CO, USA). 11. Antifoam A emulsion (Sigma–Aldrich, St. Louis, MO, USA). 12. Fire-polished glass Pasteur pipettes. 13. Glutaraldehyde, grade II, 25% (v/v) aqueous solution (Sigma–Aldrich). 2.1.2. Staining Chloroplasts with DNA Fluorophores and Analyzing the DNA Content of Individual Chloroplasts Using Fluorescence Microscopy

1. SDB (see Subheading 2.1.1). 2. Microscope slides (75 × 25 mm; 1-mm thick) with square coverslips (22 × 22 mm; thickness #1). 3. Fast-drying colorless nail polish. 4. 4¢,6-Diamidino-2-phenylindole (DAPI) fluorescent nucleic acid stain: 5 mg/mL stock and 10 mg/mL staining solutions of DAPI in deionized water. 5. Other fluorescent nucleic acid stains: dilute the SYBR Green (Invitrogen, Carlsbad, CA, USA) stock solution to 1,000× (1,000 mg/mL) in dimethyl sulfoxide (DMSO), or the SYTO 42 or SYTO 45 (Invitrogen) to 5 mM in DMSO, before preparing staining solutions. 6. Fluorescence microscope (such as Nikon EpiPhot-FX, Tokyo, Japan) with excitation and emission filter sets for DAPI (ex 350/50 nm, em 460/50 nm), SYBR Green (ex 470/20 nm, em 515 nm), or SYTO 42 (ex 436/20 nm, em 480/30 nm). For optimal observation of chloroplasts, use a 60× oil immersion objective. 7. Digital camera and image processing software, such as NIH ImageJ (http://rsbweb.nih.gov/ij/) or OpenLab™ (Perkin Elmer, Waltham, MA, USA) (optional). 8. Razor blades. 9. Dessicator jar with vacuum attachment. 10. Deoxyribonuclease I, type IV (DNase I, Sigma–Aldrich). 11. Vaccinia virus particles (ATCC, Manassas, VA, USA). 12. Fume hood.

2.1.3. Analyzing the DNA Contents of Individual Chloroplasts Using Flow Cytometry with SYBR Green or SYTO 42

1. Eosinophil counting slide (Speirs-Levy, Bluebell, PA, USA) with 22 × 50 mm coverslips (or similar cell-counting device). 2. Fluorescent nucleic acid stains: stock solution of 1,000× (1,000 mg/mL) SYBR Green (Invitrogen) prepared in DMSO, or of 5 mM SYTO 42 (Invitrogen) prepared in DMSO. 3. SDB (see Subheading 2.1.1). 4. Flow cytometer (such as a Becton Dickinson LSRII, Franklin Lake, NJ, USA) equipped with a 405-nm (for SYTO 42) or 488-nm laser (for SYBR Green).

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5. Analysis software for flow cytometry data, such as CellQuest (Becton Dickinson) or FlowJo (Treestar, Ashland, OR, USA). 6. 5-mL polystyrene tubes. 2.1.4. In-Gel Preparation of Ethidium-Stained cpDNA for the Analysis of Individual Molecules

1. SDB (see Subheading 2.1.1). 2. Lysis solution I: 40  mM EDTA, 1% (w/v) sarkosyl, and 200 mg/mL proteinase K. 3. Lysis solution II: 1 M NaCl, 5 mM EDTA, 1% (w/v) sarkosyl, and 200 mg/mL proteinase K (this solution is optional). 4. Tris–EDTA solution (TE): 10 mM 2-amino-2-hydroxymethylpropane-1,3-diol (Tris)–HCl and 1 mM EDTA, pH 8. 5. Stock solution of 3% (w/v) low-melting-point agarose prepared in TE. 6. bME/ethidium bromide (EtBr) solution (BET): 3% (v/v) bME, 0.1 mg/mL EtBr, 90 mM Tris–HCl, 90 mM boric acid, and 2 mM EDTA, pH 8. 7. BET with agarose (ABET): 1% (w/v) low-melting-point agarose prepared in BET. 8. Disposable plastic plug mold with ten 80-mL wells (Bio-Rad, Hercules, CA, USA). 9. Fluorescence microscope (see Subheading 2.1.2, item 6) with filter set for EtBr (ex 546/5, em 590 nm) with 60× oil immersion objective. 10. Digital camera and software for recording and measuring microscopic images (such as NIH ImageJ or OpenLab™). 11. 1.5-mL plastic centrifuge tubes. 12. 1× Tris–Boric acid EDTA (TBE) solution: 90 mM Tris–HCl, pH 8, 90 mM boric acid, and 2 mM EDTA (this solution is optional). 13. Water bath set at 37°C. 14. Aluminum block heated to 62°C.

2.2. The Analysis of DNA from Pools of Chloroplasts 2.2.1. Extracting DNA from Isolated Chloroplasts

1. Eosinophil counting slide (Speirs-Levy) with 22 × 50  mm coverslips. 2. Lysis solution III: SDB containing 1% sodium dodecyl sulfate (SDS), 2.5 mM EDTA, and 200 mg/mL proteinase K. 3. Stock solution of 100  mM phenylmethylsulfonyl fluoride (PMSF), prepared in isopropanol; 50–100 mL aliquots should be stored at −20°C. 4. Potassium acetate, 20 mM stock solution. 5. Tabletop microcentrifuge (e.g., Eppendorf 5415 D with standard rotor). 6. 1.5-mL plastic centrifuge tubes.

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1. SYBR Green: prepare a 1,000× (1,000 mg/mL) stock of SYBR Green (Invitrogen) in DMSO and then a 100× stock in deionized water. 2. iQ™ SYBR Green Supermix (Bio-Rad) (optional). 3. DNA isolation kit. For example, the Nucleon PhytoPure™ kit (GE Healthcare) (optional). 4. PCR reagents: Taq DNA polymerase supplied with buffer and MgCl2 (e.g., GenScript, Piscataway, NJ, USA), and deoxyribonucleotides (dNTPs) (e.g., Fermentas, Glen Burnie, MD, USA). 5. Real-time PCR detection system, such as Chromo4 (Bio-Rad). 6. 96-Well skirted PCR plates with optically clear strip caps (BioRad).

2.2.3. Analysis of cpDNA by Pulsed-Field Gel Electrophoresis

1. SDB (see Subheading 2.1.1). 2. 0.5× TBE: see Subheading  2.1.4, item 12, for preparing 1× TBE, and then dilute 1:1 in water. 3. 1.5% (w/v) agarose in 0.5× TBE. 4. EtBr: prepare a 1 mg/mL stock solution in deionized water. 5. EtBr staining solution: 0.15 mg/mL EtBr in deionized water or 0.5× TBE. 6. Destaining solution: deionized water. 7. Digital camera and software for recording EtBr-stained gel images (optional). 8. Rotating platform pulsed-field gel electrophoresis system (manufactured in-house), or CHEF pulsed-field gel electrophoresis system (Bio-Rad).

2.2.4. Blot Hybridization Using Alkaline Phosphatase Direct Labeling

1. Depurination solution: 0.25 N HCl. 2. Alkaline transfer solution: 0.4 M NaOH and 0.6 M NaCl. 3. Neutralization solution (2× SSPE): 0.15  M NaCl, 1  mM EDTA, and 10  mM NaH2PO4 (adjust pH to 7.4 with NaOH). 4. Alkaline phosphatase direct or indirect labeling kit, such as the AlkPhos direct labeling and CDP-Star chemiluminescent detection systems (GE Healthcare). 5. Positively charged nylon membrane (Roche Applied Sciences, Indianapolis, IN, USA). 6. Filter paper (3M, St. Paul, MN, USA). 7. Paper towels. 8. Plastic wrap, such as Saran Wrap™ (SC Johnson Brands, Racine, WI, USA).

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9. UV source for cross-linking, such as a Stratalinker 2400 (Stratagene, La Jolla, CA, USA). 10. X-ray film (Kodak, Rochester, NY, USA). 2.3. The Analysis of Chloroplast DNA as a Fraction of Total Tissue DNA 2.3.1. CTAB Method for Isolating Total Tissue DNA from Plants

1. CTAB solution: 2% (w/v) cetyltrimethylammonium bromide (CTAB), 1.4 M NaCl, 8 mM EDTA, and 20 mM Tris–HCl, pH 8. 2. TE (see Subheading 2.1.4, item 4). 3. Chloroform:isoamyl alcohol (24:1) for aqueous extraction of DNA. 4. Isopropanol or 100% ethanol and 70% ethanol for DNA precipitation. 5. Small plastic pestle for 1.5-mL microcentrifuge tube, such as those available from RayLabs (Auckland, New Zealand). 6. 1.5-mL plastic centrifuge tubes. 7. Tabletop microcentrifuge (e.g., Eppendorf 5415 D with standard rotor) that can be placed at 4°C.

2.3.2. Quantifying cpDNA by Blot Hybridization of Restriction-Digested Genomic DNA

1. Commercially available restriction enzyme of choice. HindIII, BamHI, and SpeI are suitable for use with Arabidopsis. 2. Probes designed for hybridization to chloroplast- and nuclearspecific genes. A nuclear-specific probe for SpeI-digested total cellular Arabidopsis DNA can be generated by PCR using the following primer sequences: forward primer 5¢-CTGA­ TATTTCTCTCCGGGGAG-3¢ and reverse primer 5¢-TACC­ TGT­CCGTTGCCAGTAAC-3¢. A chloroplast-specific probe for SpeI-digested total cellular Arabidopsis DNA can be generated by PCR using the following primer sequences: forward primer 5¢-TCGCTCCATATCTGTCTCACTC-3¢ and reverse primer 5¢-GTGCCAAAACAACCGATCCT-3¢. 3. 1.5-mL plastic centrifuge tubes. 4. DNA isolation kit. For example, the Nucleon PhytoPure™ kit (GE Healthcare) (optional). 5. Densitometer (X-Rite, Neu-Isenberg, Germany).

2.3.3. Measuring the Ratio of cpDNA to Nuclear DNA Using Real-Time Quantitative PCR

1. SYBR Green: prepare a 1,000× (1,000 mg/mL) stock of SYBR Green (Invitrogen), then dilute to 100× in water, and use this solution to prepare the PCR. 2. iQ™ SYBR Green Supermix (Bio-Rad) (optional). 3. Primers designed to amplify chloroplast- and nuclear-specific DNA. A suitable primer set for amplifying Arabidopsis nuclear DNA is forward primer 5¢-CCCCTACTTAACCGGTGGTC-3¢ and reverse primer 5¢-GAAGCGGCGAATATCTCACA-3¢.

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A suitable primer set for amplifying Arabidopsis chloroplast DNA is forward primer 5¢-AGAGACGCGAAAGCGAAAG-3¢ and reverse primer 5¢-CTGGAGGAGCAGCAATGAA-3¢.

3. Methods 3.1. The Analysis of DNA from Individual Chloroplasts

The methods described in this section provide information about the quantity and quality of DNA for individual chloroplasts within the population of chloroplasts in a tissue. These methods require only small amounts of tissue and provide information on the variation in DNA content among chloroplasts from, for example, an individual plant, an individual leaf, or a pool of individual leaves. The methods described in Subheadings 3.1.2 and 3.1.4, however, are time consuming and may not be suitable for large numbers of samples. The method described in Subheading  3.1.3 can be used to measure relative cpDNA amounts for individual chloroplasts quickly, but the initial experiments to determine the proper machine settings are time consuming. All four of these methods require prior chloroplast isolation (see Subheading  3.1.1). The method in Subheading  3.1.4 provides information about the structure of individual cpDNA molecules. We provide a comparison among the methods described in this section and the two following sections in Table 1.

3.1.1. Chloroplast Isolation

1. To reduce microbial contamination (for plants that have not been grown under aseptic conditions), wash plant tissue for 3–5 min with 0.5% sarkosyl. Rinse four times with tap water, followed by four rinses with distilled water. 2. Keep all solutions and plant tissue/chloroplasts on ice at all times. Homogenize tissue in 2–5 mL of HSB per gram of tissue on ice with a precooled mortar and pestle. Alternatively, the tissue can be mixed with HSB and 0.5–1 mL of antifoam in a chilled blender and pulsed three times for 1–2  s to homogenize. 3. Filter the homogenate through 1–3 layers of Miracloth. The material retained by the Miracloth can be re-homogenized and filtered again to increase yield. 4. Centrifuge the filtered liquid for 20 s at 12,000–16,000 × g or for 5 min at 3,000 × g for larger volumes. Discard the supernatant and resuspend the pellet in sorbitol dilution buffer (SDB) by pipetting or with a small paintbrush for larger pellets. Resuspend in 250 mL–4 mL of SDB, according to the size of the chloroplast pellet. 5. Prepare a step gradient of 30/70% Percoll (v/v) in SDB according to the volume of chloroplast suspension obtained in

Method

Staining of chloroplasts with DNA fluorophores

Flow cytometry

In-gel ethidiumstaining of cpDNA

Real-time qPCR of cpDNA from pooled chloroplasts

PFGE/blot hybridization of cpDNA

Blot hybridization of restriction-digested genomic DNA

Real-time qPCR of genomic DNA

Subheading

3.1.2

3.1.3

3.1.4

3.2.2

3.2.3/3.2.4

3.3.2

3.3.3

Low

High

High

Medium

Low-medium

Medium

Low

Amount of tissue required

Low

Medium

Medium

Medium

High

Low

High

Relative time investment

Yes

No

No

No

No

Yes

No

Suitable for analyzing a large number of samples?

Table 1 A comparison of methods for the isolation, quantification, and analysis of cpDNA

Quantitative determination of the proportion of cellular DNA represented by cpDNA

Quantitative determination of the proportion of cellular DNA represented by cpDNA

Quantitative measurement of the average DNA amount for a chloroplast population and qualitative assessment of structural forms

Quantitative measurement of the average DNA amount for a chloroplast population

Quantitative measurement of DNA amount per chloroplast and qualitative assessment of structural forms

Quantitative measurement of DNA amount per chloroplast

Qualitative or quantitative measurement of DNA amount per chloroplast

Type of information obtained

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step 4. For chloroplast suspensions of 250 mL, prepare the gradient by adding 350 mL 30% Percoll to a 1.5-mL plastic centrifuge tube. Slowly pipette 700 mL 70% Percoll underneath the 30% Percoll to form the step gradient. Gently layer the chloroplast suspension from step 4 onto this gradient and centrifuge for 10  min at 12,000–16,000 × g. For chloroplast suspensions greater than 250  mL but less than 2  mL, prepare a step gradient as above with 3  mL 30% Percoll and 3 mL 70% Percoll in a 15-mL plastic centrifuge tube and centrifuge for 30  min at 1,500 × g at 4°C. For chloroplast suspensions of 2–4 mL, prepare a step gradient as above with 15  mL 30% Percoll and 15  mL 70% Percoll and centrifuge for 30 min at 1,500 × g at 4°C. Alternatively, 70% Percoll in SDB (not a step gradient) can be used to obtain a higher yield (see Note 1). 6. Remove the purified chloroplasts from the band at the interface between the 30 and 70% Percoll layers using a fire-polished glass Pasteur pipette. If the alternative procedure is used, remove the chloroplasts from the band at the top of the 70% Percoll. 7. Wash with 10 volumes of SDB. Centrifuge at 12,000–16,000 × g for 20 s or at for 5 min at 3,000 × g. Repeat the wash twice with half the amount of SDB. 8. Pellet the chloroplasts by centrifugation at 12,000–16,000 × g. 9. Resuspend in a small volume of SDB to complete the procedure. A ratio of 1.72 mL SDB per 1 mL of packed chloroplasts is recommended for pulsed-field gel electrophoresis or other subsequent procedures. Chloroplasts can be stored for several hours in SDB on ice before proceeding with subsequent procedures or for longer periods of time in 25% glycerol at −20°C; alternatively, they can be fixed in 0.8–1.6% (v/v) glutaraldehyde and stored at 4°C (see Note 2 for safety information regarding glutaraldehyde). However, fixed chloroplasts should not be used for the procedures described in Subheadings 3.1.4 and 3.2 because glutaraldehyde can cause cross-links between proteins and DNA that could impair visualization of cpDNA molecules and inhibit PCRs. 3.1.2. Staining Chloroplasts with DNA Fluorophores and Analyzing the DNA Content of Individual Chloroplasts Using Fluorescence Microscopy

1. This method can be used for isolated chloroplasts (see Subheading 3.1.1) or for hand sections (see step 2 below). For fluorescence microscopy, chloroplasts should be fixed in 0.8– 1.6% glutaraldehyde before staining (see Note 2 for safety information regarding glutaraldehyde). For staining isolated chloroplasts with DAPI, a concentration of 1–2 mg/mL with 1% bME in SDB is recommended. To stain chloroplasts with SYBR Green, SYTO 42, or SYTO 45, use a concentration of

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3.5 mg/mL (3.5×) for SYBR Green and 20–25 mM for SYTO 42 or 45 with 1% bME in SDB. 2. To prepare hand sections for staining with DNA fluorophores, place a single leaf in a Petri dish in a fume hood. Add enough 0.8% glutaraldehyde to immerse the leaf completely. While wearing gloves, hold the leaf with its thinnest edge flush against the Petri dish. Use a scalpel or razor blade to slice the leaf tissue carefully into 0.5–1.0-mm sections. Incubate the sections in a microcentrifuge tube containing 1–2  mg/mL DAPI, and 1% bME in SDB for 30 min in the dark. Optionally, leaf sections may be placed under a vacuum for 30 min to 2 h to facilitate infiltration of the solutions into the tissue. 3. To examine isolated chloroplasts, spread 10 mL of the stained chloroplasts on a microscope slide and gently place a coverslip on top of the liquid. Seal the edges of the coverslip with quickdry colorless nail polish. Allow the nail polish to dry before proceeding to the next step. For hand sections, place a section (with the cut surface facing up) on a microscope slide along with 10 mL of the staining solution. Apply a coverslip and seal with quick-dry clear nail polish. 4. Stained chloroplasts and hand sections can be examined using a fluorescence microscope. The intensity of the fluorescence per chloroplast can be evaluated qualitatively by subjective scoring (i.e., strong, weak, and none) or measured quantitatively using digital image processing software. Fig.  1 shows

Fig. 1. Changes in DAPI-DNA fluorescence during chloroplast development. Proplastid (a, d), immature chloroplast (b, e), and mature chloroplast (c, f) isolated from Arabidopsis plants. (a–c) Bright-field images. (d–f) DAPI-DNA fluorescence images. Scale bar is 10 mm.

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examples of isolated chloroplasts after DAPI staining and examination by fluorescence microscopy. 5. When measuring DNA fluorophore-stained chloroplasts quantitatively using digital imaging, it is important to consider how the images are acquired and analyzed. The exposure time should be adjusted so that none of the images are overexposed. Intensity of the fluorescence of the DNA fluorophore can be measured using grayscale images, where each pixel is assigned a brightness value. Software programs (such as NIH ImageJ or OpenLab™) can be used to capture and analyze images. These programs allow the user to select the region of the photograph that represents a single chloroplast and measure the average pixel intensity and the area. These two values should be multiplied in order to obtain a value that is proportional to the total amount of DNA present. It is also important to consider a number of controls in order to adjust for field-to-field variation in background fluorescence (fluorescence that arises from the staining solution) and the autofluorescence arising from chlorophyll or from the glutaraldehyde fixative. A slide preparation that does not include the DNA fluorophore, or a preparation of DNase-I-treated, stained chloroplasts can be used to measure fluorescence arising from chlorophyll or the fixative. These values can then be used to adjust those of chloroplasts from the experimental sample. To adjust for field-to-field variation of the background fluorescence, measure the fluorescence intensity of a region of the image that does not contain chloroplasts or debris and record the average intensity. The relative fluorescence intensity (Rfl) values obtained using this method can be used to assess the relative DNA amounts of individual chloroplasts. A comparison of the chloroplast Rfl values to those of particles with a known DNA amount and base composition (e.g., Vaccinia virus particles) can be used to calculate the number of genome equivalents per chloroplast. See Note 3 for more details. 3.1.3. Analyzing the DNA Contents of Individual Chloroplasts Using Flow Cytometry with SYBR Green or SYTO 42

1. Most flow cytometers consume 50–100 mL per run. We recommend at least three replicates, so that a sample volume of 500 mL should be prepared. However, the chloroplasts can be concentrated if a lower sample volume is desired. 2. Isolate chloroplasts according to the protocol described in Subheading  3.1.1. Adjusting the chloroplasts to a specific concentration is not necessary. However, if the chloroplast concentration is very low (and you cannot see any green color in the chloroplast solution after step 3 below), then it will take a longer time to analyze the sample on the flow cytometer. Prepare the staining solution by diluting SYTO 42 to a concentration of 10  mM (or SYBR Green to a concentration of 3.5× [3.5 mg/mL]) in SDB. Add 5 mL of each chloroplast sample

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to a 1.5-mL microcentrifuge tube. The chloroplasts are permeable to SYTO 42 and SYBR Green with or without prior fixation. It is important to note that certain fixatives can generate an autofluorescence. Thus, fixed samples should not be compared to samples that have not been fixed. 3. Add 495 mL (or the amount needed to obtain the desired sample volume) of staining solution to the chloroplasts. 4. Analyze the stained chloroplasts on the flow cytometer. Collect data from forward-scatter, side-scatter, and the appropriate emission detectors for the DNA fluorophore (460  nm for SYTO 42 and 520 nm for SYBR Green) and chlorophyll (640– 680 nm). Data should be collected in log scale for the DNA fluorophores and chlorophyll. A control sample of chloroplasts that have been treated with DNase (fixed chloroplasts are permeable to DNase) should be used to distinguish chloroplasts from debris and determine the appropriate voltage settings that will maximize the separation between chloroplasts that contain DNA and those that do not. This will improve the accuracy and sensitivity of the analysis. 5. We recommend collecting 5,000–10,000 events per replicate and performing three replicates per sample. 3.1.4. In-Gel Preparation of Ethidium-Stained cpDNA for the Analysis of Individual Molecules

1. Isolate the chloroplasts according to the protocol described in Subheading 3.1.1. After resuspending the chloroplast pellet in SDB, prepare 1:100, 1:500, and 1:1,000 dilutions of this stock suspension in SDB. 2. Add 3% molten low-melting-point agarose to adjust the chloroplast solution to a final concentration of 0.7% low-meltingpoint agarose and add 2 M sorbitol to maintain a concentration of 0.33 M. Before adding the agarose, briefly warm the chloroplasts (previously kept on ice) in a 37°C waterbath. Using a wide-bore pipette tip, gently mix and pipette the agaroseembedded chloroplasts into a reusable plug mold and incubate at 4°C for 10–20 min to solidify the agarose. 3. Remove the agarose plugs from the mold and lyse overnight at 48°C in lysis solution I (see Subheading 2.1.4). Alternatively, lysis solution II can be used to remove more of the chlorophyll from the agarose plug. However, the high ionic strength in the latter solution may lead to the formation of G-quartet structures at the ends of DNA molecules. It is not necessary to inactivate the proteinase K before proceeding, but the proteinase K can be inactivated using PMSF if desired (see Subheading 3.2.1, step 2). Wash the samples at least five times in TE, 20–30 min per wash. 4. Cut a piece from the plug (about one-quarter the size of the 80-mL agarose plug) and stain in 50–100 mL of BET for 30 min at room temperature.

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5. Remove the staining solution and repeat the staining procedure described in step 4. 6. Cut the stained plug in half and place it on a glass microscope slide along with 10–20 mL of ABET. 7. Place a coverslip on top of the agarose and ABET, and heat the slide until the agarose plug melts. We use an aluminum block kept at 62°C. Move the slide to room temperature and allow the agarose to resolidify. Use clear nail polish to seal the top and bottom but not the sides of the coverslip. This will allow for an electric field to be imposed across the gel underneath the coverslip (see step 8 below). 8. The agarose-embedded cpDNA can now be examined using fluorescence microscopy, as described in the previous section. If desired, an electric field can be imposed across the slide using 1× TBE and a power supply connected to a cathode and an anode placed on either side of the coverslip. As DNA is negatively charged, the fibers of cpDNA will stretch toward the anode, which will provide better visualization of the structure of cpDNA molecules and allow measurement of the fiber lengths. 3.2. The Analysis of DNA from Pools of Chloroplasts

3.2.1. Extracting DNA from Isolated Chloroplasts

The methods described in this section provide information about the quality and structure of DNA from a population of chloroplasts. All of these methods require prior isolation of chloroplasts (see Subheading 3.1.1). The method described in Subheading 3.2.2 can quickly and accurately measure the amount of DNA for an average chloroplast within the population. Information about both the structural forms and the amount of cpDNA can be obtained by employing the methods described in Subheadings 3.2.3 and 3.2.4. However, these methods require a large amount of tissue. 1. Isolate chloroplasts according to the protocol described in Subheading  3.1.1. Determine the concentration of chloroplasts using an eosinophil counter or a similar method. Lyse a known number of chloroplasts in lysis solution III, vortex briefly, and incubate for at least 1 h at 37°C. 2. Inactivate the proteinase K by adding 0.1 mM PMSF and incubate for 1 h at room temperature. 3. Add 20 mM potassium acetate and incubate for 10 min on ice to precipitate the SDS and denatured proteins. 4. Centrifuge at 12,000 × g for 10 min at 4°C in a microcentrifuge. 5. Transfer the supernatant to a fresh microcentrifuge tube. Record the volume and store at 4°C. The lysate is ready for qPCR analysis (see Subheading 3.2.2). Ensure that no SDS is present in the lysate, otherwise downstream applications such as PCR will be inhibited. If necessary, repeat steps 3 and 4

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if you see bubbles in the lysate, or repeat step 4 only if any white precipitate remains. 3.2.2. Quantifying DNA from Isolated Chloroplasts Using Real-Time Quantitative PCR

1. Prepare a set of cpDNA standards. The cpDNA can be prepared by isolating chloroplasts using the protocol described in Subheading 3.1.1, followed by the CTAB protocol described in Subheading  3.3.1 starting from step 2 (or the method described above in Subheading  3.2.1). Alternatively, a commercially available DNA preparation kit can be used (see Subheading  2.2.2). Determine the DNA concentration and prepare five tenfold serial dilutions ranging from 5 fg/mL to 50 pg/mL in the same solution as used for the lysates. This will ensure that the amplification of the standards and unknowns will have a similar efficiency. 2. Amplify 1 mL template DNA in a 25-mL reaction mixture 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 using primers designed to amplify a 100–200-bp fragment of cpDNA. Alternatively, a commercially available mastermix containing SYBR Green can be used. Monitor the increase in SYBR Green fluorescence using a real-time PCR detection system. Follow the manufacturer’s instructions to set the threshold and determine the cycle at which the amplification curves cross the threshold Ct. Then, calculate the concentration of the sample chloroplast lysates using the standard curve. See Note 4 for additional recommendations regarding replicates. See Note 5 for determining the efficiency of amplification. 3. Divide the concentration of cpDNA in the lysate by the concentration of chloroplasts originally present in the solution. This will give the average amount of DNA per chloroplast. We find that the data obtained using this method are consistent with the average DNA amount for individual chloroplasts from the same sample as measured by DAPI staining using Vaccinia virus particles as a standard.

3.2.3. Analysis of cpDNA by Pulsed-Field Gel Electrophoresis

1. Isolate the chloroplasts according to Subheading 3.1.1. After resuspension in SDB, perform steps 2 and 3 of Subheading 3.1.4. 2. Use one-quarter to one-half of the cpDNA-agarose plug for PFGE in a 1.5% agarose gel in 0.5× TBE at 5 V/cm overnight at 6°C or other suitable conditions. Adjust the pulse time and direction of the electric field for the best resolution. For resolution of genomic monomers, dimers, and trimers, we recommend using a pulse time of 50 s and rotating the gel (or the electric field) at 105°. 3. Stain the gel for 1–1.5 h in EtBr staining solution and destain the gel as needed (a 1-h destain is sufficient but a 2-h destain with a change of solution is better, giving a reduced background).

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Fig. 2. Images of cpDNA from pooled Arabidopsis chloroplasts after PFGE. The cpDNA was visualized by (a) staining with ethidium bromide and (b) blot hybridization. The cpDNA from both wild-type (wt) and drt100-1 mutant plants shows a wild-type pattern of separation by PFGE, where bands of cpDNA are detected at the position of the monomer, dimer, and trimer. There is also a prominent cpDNA signal from the well and the compression zone (cz). This figure was reproduced with permission from Oxford Journals (Rowan et al. 2010).

4. Software programs (such as NIH ImageJ) can be used to quantify the relative intensity of the EtBr fluorescence among samples. It should be noted that this technique can be used to determine both the amount and the structural forms of cpDNA. Blot hybridization (see Subheading 3.2.4) can also be used to analyze the cpDNA. Fig.  2 shows PFGE-separated cpDNA after staining with EtBr and the same cpDNA samples after blot hybridization. 3.2.4. Blot Hybridization Using Alkaline Phosphatase Direct Labeling

1. Incubate the gel in depurination solution (0.25  N HCl) for 10 min with gentle agitation. 2. Transfer the gel to a solution containing alkaline transfer solution and incubate for 20 min with agitation. 3. Transfer the DNA from the gel to a nylon membrane overnight. Arrange two long sheets of filter paper with both ends

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in alkaline transfer solution. Place the gel upside down on one piece of filter paper cut to the size of the gel, lay the membrane on top, and then two pieces of filter paper. Place Saran Wrap or a plastic barrier along the edges of the gel/membrane sandwich to direct the flow of transfer solution through the gel. Place many paper towels on top, weigh down the paper towels (a half-full 500-mL bottle works well), and then allow the transfer to proceed overnight. 4. Neutralize the nylon membrane by incubating in 2× SSPE (neutralization solution) for 10 min. Test the pH of the membrane by touching a piece of pH paper to a corner of the wet membrane. If the pH is >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

11  Measurement of Transcription Rates in Arabidopsis Chloroplasts plastid RNA polymerase and in vitro reconstitution of the holoenzyme. Plant J. 40, 164–172. 5. Ogrzewalla, K., Piotrowski, M., Reinbothe, S., and Link, G. (2002) The plastid transcription kinase from mustard (Sinapis alba L.). A nuclear-encoded CK2-type chloroplast enzyme with redox-sensitive function. Eur. J. Biochem. 269, 3329–3337. 6. Pfannschmidt, T., Ogrzewalla, K., Baginsky, S., Sickmann, A., Meyer, H. E., and Link, G. (2000) The multisubunit chloroplast RNA polymerase A from mustard (Sinapis alba L.): integration of a prokaryotic core into a larger complex with organelle-specific functions. Eur. J. Biochem. 267, 253–261. 7. Pfalz, J., Liere, K., Kandlbinder, A., Dietz, K.-J., and Oelmüller, R. (2006) pTAC2, -6 and −12 are components of the transcriptionally active plastid chromosome that are required for plastid gene expression. Plant Cell 18, 176–197. 8. Allison, L. A., Simon, L. D., and Maliga, P. (1996) Deletion of rpoB reveals a second distinct transcription system in plastids of higher plants. EMBO J. 15, 2802–2809. 9. Krause, K., Maier, R. M., Kofer, W., Krupinska, K., and Herrmann, R. G. (2000) Disruption of plastid-encoded RNA polymerase genes in tobacco: expression of only a distinct set of genes is not based on selective transcription of the plastid chromosome. Mol. Gen. Genet. 263, 1022–1030. 10. Legen, J., Kemp, S., Krause, K., Profanter, B., Herrmann, R. G., and Maier, R. M. (2002) Comparative analysis of plastid transcription profiles of entire plastid chromosomes from tobacco attributed to wild-type and PEPdeficient transcription machineries. Plant J. 31, 171–188. 11. Hess, W. R., and Börner, T. (1999) Organellar RNA polymerases of higher plants. Int. Rev. Cytol. 190, 1–59. 12. Liere, K., and Börner, T. (2007) Transcription and transcriptional regulation in plastids. In, Topics in Current Genetics: Cell and Molecular Biology of Plastids (Bock, R., ed.) Springer, Berlin/Heidelberg, Germany, pp. 121–174. 13. Lerbs-Mache, S. (1993) The 110-kDa polypeptide of spinach plastid DNA-dependent RNA polymerase: single-subunit enzyme or catalytic core of multimeric enzyme complexes? Proc. Natl. Acad. Sci. USA 90, 5509–5513. 14. Hedtke, B., Börner, T., and Weihe, A. (1997) Mitochondrial and chloroplast phage-type RNA polymerases in Arabidopsis. Science 277, 809–811.

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15. Chang, C.-C., Sheen, J., Bligny, M., Niwa, Y., Lerbs-Mache, S., and Stern, D. B. (1999) Functional analysis of two maize cDNAs encoding T7-like RNA polymerases. Plant Cell 11, 911–926. 16. Liere, K., Kaden, D., Maliga, P., and Börner, T. (2004) Overexpression of phage-type RNA polymerase RpoTp in tobacco demonstrates its role in chloroplast transcription by recognizing a distinct promoter type. Nucleic Acids Res. 32, 1159–1165. 17. Hricová, A., Quesada, V., and Micol, J. L. (2006) The SCABRA3 nuclear gene encodes the plastid RpoTp RNA polymerase, which is required for chloroplast biogenesis and mesophyll cell proliferation in Arabidopsis. Plant Physiol. 141, 942–956. 18. Baba, K., Schmidt, J., Espinosa-Ruiz, A., Villarejo, A., Shiina, T., Gardestrom, P., Sane, A. P., and Bhalerao, R. P. (2004) Organellar gene transcription and early seedling development are affected in the RpoT;2 mutant of Arabidopsis. Plant J. 38, 38–48. 19. Swiatecka-Hagenbruch, M., Emanuel, C., Hedtke, B., Liere, K., and Börner, T. (2008) Impaired function of the phage-type RNA polymerase RpoTp in transcription of chloroplast genes is compensated by a second phagetype RNA polymerase. Nucleic Acids Res. 36, 785–792. 20. Courtois, F., Merendino, L., Demarsy, E., Mache, R., and Lerbs-Mache, S. (2007) Phagetype RNA polymerase RPOTmp transcribes the rrn operon from the PC promoter at early developmental stages in Arabidopsis. Plant Physiol. 145, 712–721. 21. Hedtke, B., Legen, J., Weihe, A., Herrmann, R. G., and Börner, T. (2002) Six active phagetype RNA polymerase genes in Nicotiana tabacum. Plant J. 30, 625–637. 22. Weihe, A., Hedtke, B., and Börner, T. (1997) Cloning and characterization of a cDNA encoding a bacteriophage-type RNA polymerase from the higher plant Chenopodium album. Nucleic Acids Res. 25, 2319–2325. 23. Young, D. A., Allen, R. L., Harvey, A. J., and Lonsdale, D. M. (1998) Characterization of a gene encoding a single-subunit bacteriophagetype RNA polymerase from maize which is alternatively spliced. Mol. Gen. Genet. 260, 30–37. 24. Ikeda, T. M., and Gray, M. W. (1999) Identification and characterization of T7/T3 bacteriophage-like RNA polymerase sequences in wheat. Plant Mol. Biol. 40, 567–578. 25. Kobayashi, Y., Dokiya, Y., and Sugita, M. (2001) Dual targeting of phage-type RNA

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polymerase to both mitochondria and plastids is due to alternative translation initiation in single transcripts. Biochem. Biophys. Res. Commun. 289, 1106–1113. 26. Emanuel, C., Weihe, A., Graner, A., Hess, W. R., and Börner, T. (2004) Chloroplast development affects expression of phage-type RNA polymerases in barley leaves. Plant J. 38, 460–472. 27. Kusumi, K., Yara, A., Mitsui, N., Tozawa, Y., and Iba, K. (2004) Characterization of a rice nuclear-encoded plastid RNA polymerase gene OsRpoTp. Plant Cell Physiol. 45, 1194–1201. 28. Hedtke, B., Börner, T., and Weihe, A. (2000) One RNA polymerase serving two genomes. EMBO Rep. 1, 435–440. 29. Bensing, B. A., Meyer, B. J., and Dunny, G. M. (1996) Sensitive detection of bacterial transcription initiation sites and differentiation from RNA processing sites in the pheromoneinduced plasmid transfer system of Enterococcus faecalis. Proc. Natl. Acad. Sci. USA 93, 7794–7799. 30. Sambrook, J., and Russel, D. W. (2001) Molecular Cloning: A Laboratory Manual, 3rd edn. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, NY, USA. 31. Fromont-Racine, M., Bertrand, E., Pictet, R., and Grange, T. (1993) A highly sensitive

method for mapping the 5¢ termini of mRNAs. Nucleic Acids Res. 21, 1683–1684. 32. Gruissem, W., Greenberg, B. M., Zurawski, G., and Hallick, R. B. (1986) Chloroplast gene expression and promoter identification in chloroplast extracts. Methods Enzymol. 118, 253–270. 33. Orozco, E. M., Jr., Mullet, J. E., HanleyBowdoin, L., and Chua, N. H. (1986) In vitro transcription of chloroplast protein genes. Methods Enzymol. 118, 232–253. 34. Sugiura, M. (1995) The chloroplast genome. Essays Biochem. 30, 49–57. 35. Georg, J., Honsel, A., Voß, B., Rennenberg, H., and Hess, W. R. (2010) A long antisense RNA in plant chloroplasts. New Phytol. 186, 615–622. 36. Mullet, J. E., and Klein, R. R. (1987) Transcription and RNA stability are important determinants of higher plant chloroplast RNA levels. EMBO J. 6, 1571–1579. 37. Church, G. M., and Gilbert, W. (1984) Genomic sequencing. Proc. Natl. Acad. Sci. USA 81, 1991–1995. 38. Deng, X. W., Stern, D. B., Tonkyn, J. C., and Gruissem, W. (1987) Plastid run-on transcription. Application to determine the transcriptional regulation of spinach plastid genes. J. Biol. Chem. 262, 9641–9648.

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 (<15  mL and <5  mg, respectively), and to use fresh reagents. Northern blotting provides considerably more information than quantitative reverse transcription – polymerase chain reaction (qRT-PCR) or microarray hybridization, which report solely the abundance of a specific RNA segment. By employing short oligonucleotides as probes, Northern blots can even be used to map the termini of specific transcripts to within ~7 nucleotides (6). Primer extension is a particularly simple method for the precise mapping of 5¢ termini. A 5¢-end-labeled oligonucleotide complementary to sequences downstream of the terminus of interest is hybridized in solution to total RNA and used to prime reverse transcription. The radiolabeled products are then resolved on a denaturing polyacrylamide gel. To obtain high-resolution data, the primer should be positioned ~30–100 nucleotides from the terminus under investigation. Parallel analysis of a sequencing ladder obtained with the same primer allows the precise determination of the nucleotides at which reverse transcription terminates. Stable structures in the RNA template can cause reverse transcriptase to pause, generating artifactual bands in primer extension assays.

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This problem can be minimized by performing the reverse transcription reactions at the highest temperature that is compatible with enzymatic activity (~48°C, but the precise temperature depends upon the source of the reverse transcriptase). To confirm that bands detected reflect true 5¢ termini, parallel reactions can be performed with an in vitro RNA transcript generated from a cloned chloroplast DNA template that spans the region of interest (for an example, see ref. 6). The protocol provided here for poisonedprimer extension can be modified to serve as a standard primer extension assay simply by including all four nucleotides as dNTPs and excluding ddNTPs from the reverse transcription reactions. 1.1.3. Poisoned-Primer Extension

Poisoned-primer extension is a modified primer-extension procedure that includes a single dideoxynucleotide in the reverse transcription reaction; reverse transcription terminates at the first nucleotide in the RNA template that is complementary to the dideoxynucleotide in the reaction mix. A primer and dideoxynucleotide combination is designed such that reverse transcription stops after a different number of nucleotides on differentially processed RNA templates (e.g., spliced versus unspliced RNA). Poisoned-primer extension is a simple method for quantifying the ratio of edited versus unedited RNA at a particular site, or the ratio of spliced versus unspliced RNA from a particular gene. The method reports on the RNA sequence over a very short region, such that answers can be obtained even with RNA samples that have suffered some degradation. A detailed protocol for monitoring chloroplast RNA splicing by poisoned-primer extension is provided below.

1.1.4. Circular RT-PCR

Circular RT-PCR (cRT-PCR) can be used to map the positions of 5¢ and 3¢ termini of specific RNAs to high resolution (7). Total RNA is circularized by ligation with T4 RNA ligase. Regions spanning the 5¢/3¢ junctions of specified RNAs are then amplified by RT-PCR and the products are cloned and sequenced. cRT-PCR is the method of choice for mapping 3¢ termini. The 3¢ termini of chloroplast RNAs are typically heterogeneous (for an example, see ref. 6), so it is important to sequence at least ten different cRT-PCR clones to obtain an accurate representation of their distribution. A detailed protocol for cRT-PCR has been published previously (8).

1.1.5. Ribonuclease Protection

Ribonuclease protection assays can be used to approximate the positions of 5¢ and 3¢ RNA termini, and to quantify the relative abundance of alternative processed RNA isoforms. A body-labeled radioactive in vitro RNA transcript that is complementary to the RNA terminus or splice junction of interest is hybridized in solution to total RNA; the unhybridized RNA is then degraded with single-strand specific ribonucleases. The sizes of the surviving probe fragments reflect the positions of discontinuity between the probe and its complement in the RNA sample. Ribonuclease protection

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assays are more sensitive than primer-extension and Northern blot assays, but they are considerably more labor intensive. In addition, this method cannot map RNA termini to high resolution because the probe is body labeled, and the ribonucleases do not trim the probe to their precise point of discontinuity with the complementary RNA. 5¢ Termini are more precisely mapped by primer extension, 3¢ termini are more precisely mapped by cRT-PCR, and different splice isoforms are more easily quantified by poisoned-primer extension and/or by Northern blotting. 1.1.6. Recommended Methods for Assaying Specific Aspects of RNA Structure and Processing

To map  5¢ termini, primer-extension assays are typically the best choice because they are simple and precise. Primer extension is also an excellent method for comparing the relative use of different 5¢ termini in mutant and wild-type strains, or in plants grown under different growth conditions. If the use of radioactive isotopes is problematic, 5¢-rapid amplification of cDNA ends (RACE) and cRT-PCR provide alternatives, but these methods are more labor intensive and are less suited to quantitative analyses. The precise mapping of 3¢ termini is more challenging and requires the use of a ligation-mediated method such as cRT-PCR. However, ribonuclease-protection assays are ideal for comparing the relative abundance of different 3¢ RNA isoforms in plants of different genotype or grown under different conditions. RNA splicing can be monitored in several different ways, each with distinct advantages and disadvantages. Northern blotting is an effective means to quantify spliced and unspliced isoforms from “simple” transcription units such as those encoding spliced tRNAs. However, Northern blots are less suitable for analyzing the splicing of RNAs derived from complex transcription units (i.e., all introncontaining protein-coding genes in chloroplasts) because the complex banding patterns complicate the quantification of spliced and unspliced isoforms. Poisoned-primer extension assays are recommended in such cases because they generate just one product representing all unspliced RNAs and one representing all spliced RNAs (see Fig. 1). RT-PCR is a poor choice for quantifying the ratio of spliced to unspliced chloroplast RNAs because of its nonquantitative nature, which is exacerbated by the fact that chloroplast introns are long and highly structured. Nonetheless, a qRT-PCR assay has been used effectively for detecting chloroplast splicing defects in mutant lines (9), and is worth developing if the splicing of numerous chloroplast introns will be assayed repeatedly. The ratio of spliced to unspliced RNA from a particular gene is related to splicing efficiency and is often equated to this. However, this ratio is also influenced by the stability of the spliced product. Therefore, it is important to monitor both products of splicing – the spliced RNA and the excised intron – to draw strong conclusions about differences in splicing efficiency between samples. Excised introns can be detected by probing Northern blots with intron-specific probes.

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Fig. 1. Representative Results of a Poisoned-Primer Extension Splicing Assay. RNA was purified from the leaves of Arabidopsis seedlings of the indicated genotypes, and analyzed with a primer that binds immediately downstream of the ndhB 3¢ splice junction (5¢-AAGCAACGACTGGAGTGGGAGA-3¢). The reaction included the chain terminator ddATP, such that reverse transcription terminated after the addition of 4 or 9 nucleotides on unspliced and spliced RNA templates, respectively. These results show that the cfm3-4 mutant fails to splice the ndhB intron.

1.2. Methods for Identifying the Binding Sites of Chloroplast RNA-Binding Proteins 1.2.1. Approaches for Identifying the Native RNA Ligands of Chloroplast RNA-Binding Proteins

Genetic screens have revealed numerous RNA-binding proteins (RBPs) that influence chloroplast gene expression, with hundreds more predicted from in silico evidence. A major challenge is to define the RNA sequences with which each chloroplast RBP interacts, and to determine how these interactions influence RNA structure and function. This problem is typically approached from one of two directions: genetic analysis, in which the function of an RBP is inferred through the analysis of chloroplast gene expression defects in mutant lines, or biochemical analysis, in which RBPs are “fished” from chloroplast extract by affinity chromatography with specific RNA sequences. An advantage of genetic analysis as the starting point is that it avoids false leads arising from artifactual interactions that can occur in vitro. However, the inferences made from genetic data are limited because direct and indirect effects of a mutation can be difficult to distinguish. This is especially problematic when the mutation compromises the chloroplast translation machinery, as the loss of plastid ribosomes causes stereotypical downstream effects on chloroplast transcription and RNA processing (see, for example, refs. 10, 11). The identification of RNAs that coimmunoprecipitate with an RBP from chloroplast extract provides a powerful complement to genetic approaches. RNA coimmunoprecipitation can confirm that a protein interacts with particular RNAs in  vivo, and can

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clarify the direct function of an RBP even when its mutant phenotype is pleiotropic (see, for example, refs. 12–14). In a “RIPchip” assay, coimmunoprecitated RNAs are detected in an unbiased fashion by hybridization to a chloroplast genome tiling microarray (15). Alternatively, the coimmunoprecipitated RNAs can be probed for specific candidate RNAs by slot-blot hybridization. The latter method in conjunction with short oligonucleotide probes can pinpoint the native binding sites of chloroplast RBPs to a resolution of <50 nucleotides (6, 15). Detailed protocols for chloroplast RNA coimmunoprecipitation assays involving either microarray or slot-blot hybridization were published previously (16). 1.2.2. Assaying the RNA-Binding Properties of Chloroplast RBPs In Vitro

The combination of genetic and RNA coimmunoprecipitation data can provide a strong foundation for understanding the RNA ligands and biological functions of chloroplast RBPs. However, in  vitro assays with purified RBPs are necessary to achieve a detailed mechanistic understanding. There is no “one-size-fits-all” approach to assay the RNA-binding activities of RBPs: the choice of binding conditions and binding assay must be tailored to each protein. The various choices to be made and the parameters to consider are discussed below. Reference is made to publications that illustrate these points and that describe the methods themselves.

1.2.3. Protein Expression and Purification

RBPs for in  vitro assays are generally produced as recombinant proteins in Escherichia coli. This is often more complicated than it sounds because many RBPs are prone to aggregation when expressed in E. coli. It is common that multiple expression vectors and purification strategies must be explored to obtain soluble protein of adequate purity (see, for example, ref. 17). Expression of RBPs as fusions to certain affinity tags (e.g., maltose-binding protein, glutathione S-transferase, or thioredoxin) can enhance solubility and aid purification, but oftentimes the passenger protein precipitates upon removal of the tag. Furthermore, many recombinant proteins that appear to be “soluble” (i.e., material does not pellet after extended microcentrifugation) are actually in microaggregates whose detection requires a method such as gel filtration chromatography or dynamic light scattering. Proteins found in microaggregates cannot be used reliably to monitor the protein’s native biochemical activities. Therefore, recombinant RBPs for in vitro assays should be purified by at least two successive purification steps: an initial affinity purification step that exploits an affinity tag, followed by a gel filtration step to remove contaminants and to ensure that the RBP is monodisperse. Ideally, proteins used for biochemical assays are separated from the affinity tag by proteolytic cleavage and subsequent purification. However, if removal of the tag causes protein insolubility (which it often does), reliable results can sometimes be obtained with the uncleaved (but purified and monodisperse) fusion protein.

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1.2.4. Methods for Detecting RNA-Binding Activity

In vitro RNA-binding assays with purified RBPs are used to establish the specificity of the RBP for different RNA sequences or for different nucleic acid forms (e.g., single-stranded versus doublestranded versus structured RNA). The most commonly employed methods are gel mobility shift (GMS) assays (also called EMSAs; see Chapter 13, Vol. 1), filter-binding assays, and UV cross-linking assays. These assays are typically performed with labeled RNA (usually 32P) and unlabeled protein. GMS assays use native gel electrophoresis to resolve “free” RNA from RNA bound to protein. Filter-binding assays exploit the protein-binding capacity of nitrocellulose membranes to separate free and bound RNA (18). UV crosslinking assays identify proteins that are found in close proximity to labeled RNA by label transfer after UV-induced cross-linking. Of these methods, GMS assays provide the most information and UV cross-linking assays provide the least. UV cross-linking assays are the least quantitative of these methods because they capture transient interactions, and the signal is influenced by the proximity of the labeled nucleotide to those amino acid side chains that are amenable to cross-linking. Both filter-binding and GMS assays can be used to generate binding curves that are suitable for the estimation of equilibrium Kd, whereas this is not possible with UV cross-linking. Additional advantages of GMS assays are their ability to resolve RNA/protein complexes with different stoichio­ metries and to display the integrity of the labeled RNA in the output data; neither filter-binding nor UV cross-linking assays have these features. Although UV cross-linking is a poor choice for discerning the sequence specificity of purified RBPs, it is useful for identifying proteins in a crude mixture that can bind a particular labeled RNA substrate. Because GMS assays provide the most information, we recommend their use when possible. An informative discussion of the parameters to consider when designing GMS assays and an example of a GMS protocol can be found in ref. (19). Unfortunately, some RBP/RNA complexes do not resolve well during native gel electrophoresis. This is true, for example, of proteins bound to large RNAs such as group II introns. In such instances, filter-binding assays provide a good alternative (20). Once an interaction is localized to a general region of an RNA by a GMS or filter-binding assay, the details of the interaction can be explored with high-resolution methods such as hydroxyl radical footprinting (20).

1.2.5. Assessing “Sequence Specificity”

“Sequence specificity” is, in essence, the affinity of a protein for a “specific” RNA sequence in comparison to its affinity for a “nonspecific” RNA sequence. Two methods are commonly used to explore sequence specificity: (1) competition assays, which score the ability of unlabeled RNAs to compete with a labeled RNA for interaction with the RBP, and (2) comparison of the affinity of the RBP for different RNAs through the estimation of dissociation

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constants (Kd). We feel that the latter approach is less prone to artifacts for the following reasons. The interpretation of competition assays relies on the intactness and accurate quantification of the competitor RNA; these are parameters that cannot be “seen” in the actual experiment. This is especially problematic when in vitro transcripts (as opposed to synthetic RNA oligonucleotides) are used as the competitors because it can be difficult to quantify them accurately and to keep them intact throughout their purification. In contrast, when binding curves are generated with a constant, low concentration of labeled RNA (i.e., well below the Kd) and increasing concentrations of protein, the precise quantification of the RNA is unnecessary because complex formation is driven solely by protein concentration. If (and only if) the RNA is present at much lower concentration than the Kd, the Kd can be estimated as the protein concentration at which 50% of the RNA is bound (19). Sequence specificity can then be assessed by comparing the Kd of the RBP for RNAs of similar length but different sequence. 1.2.6. Optimizing RNA-Binding Reaction Conditions

Most RNA-binding proteins will bind in vitro to any RNA they are presented with unless conditions are optimized to reduce nonspecific interactions (see, for example, refs. 17, 20). It is difficult to evaluate which conditions are “optimal” without prior genetic and/or RNA coimmunoprecipitation data that provide evidence for the native RNA ligand(s). For example, the fact that the chloroplast RBPs CRS1 and PPR5 were already known to interact with specific group II introns in vivo guided the development of the in  vitro assay conditions that revealed their precise, direct binding sites (17, 20). The key variables to consider while exploring conditions to reveal sequence-specific interactions in vitro are (1) the length and concentration of the RNA; (2) the concentrations of monovalent and divalent salts; (3) the conditions used to promote or restrict the “folding” of the RNA; and (4) the concentration of nonspecific competitors such as tRNA and heparin. Each of these parameters is discussed below. 1. Experiments should compare an RNA anticipated to harbor a specific binding site to several other RNAs of similar length. The longer the RNA, the more opportunities arise for nonspecific interactions with the RBP. Sequence specificity can be difficult to detect when a binding site is embedded in a long RNA molecule, because nonspecific interactions can overwhelm the “specific” signal and because internal RNA:RNA interactions can mask the binding site. For example, the specificity of PPR5 for its group II intron ligand could not be detected when the intact intron (~900 nucleotides) was used in binding assays. Specificity began to emerge upon analysis of intron fragments of ~200 nucleotides, and was ultimately clarified by the analysis

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of 50-mers (17). Labeled RNAs greater than ~50 nucleotides are generally obtained by in vitro transcription of PCR products that incorporate a promoter for T7 RNA polymerase; a radiolabeled nucleotide can be incorporated during in vitro transcription. Shorter RNAs can be purchased as synthetic oligoribonucleotides, and 5¢ end labeled by polynucleotide kinase. In either case, the labeled RNA should be purified on a denaturing polyacrylamide gel prior to its use in binding reactions. Except in the case of exceptionally high-affinity interactions, an RNA concentration of 50  pM is sufficiently low that the Kd can be estimated as the protein concentration at which half-maximal binding is observed. 2. Salts have a strong influence on the affinity and specificity of RNA– protein interactions. Because high concentrations of monovalent salts inhibit nonspecific electrostatic interactions, sequence specificity is often apparent only under “high” salt conditions (e.g., >300  mM) (see, for example, ref. 20). Optimization experiments should explore a range of Na+ or K+ concentrations, starting at roughly physiological concentrations (~125  mM) and increasing to ~350 mM. Mg++ can strongly influence the binding of RBPs by stabilizing RNA tertiary structures (21). Proteins that recognize a particular RNA structure may require Mg++ for a specific interaction (see, for example, ref. 20), whereas the presence of Mg++ can hinder interactions for RBPs that bind unstructured RNA. Optimization experiments should test the effects of varying Mg++ concentrations between 0 and ~10 mM. 3. RBPs that recognize single-stranded RNA bind optimally when the RNA is treated to minimize secondary structure formation. This can be achieved by heating the RNA in the absence of salts to ~80°C, followed by snap-cooling on ice. Conversely, RNAs whose recognition involves a particular three-dimensional structure should be treated to encourage “folding,” e.g., heating the RNA followed by slow-cooling in the presence of monovalent salts (at least 100 mM) and Mg++ (~2–10 mM). Optimization experiments should explore both “folding” and “unfolding” conditions, unless it has already been established that the RBP binds preferentially to either single-stranded or structured RNA. When working with an RBP that binds single-stranded RNA, it is important to consider the inhibitory effects of RNA structure. Otherwise, preferential binding to RNAs with minimal structure can be misinterpreted as sequence-specific RNA recognition. 4. The detection of specific RBP/RNA interactions usually requires the use of nonspecific competitors such as tRNA and/ or heparin (an anionic polymer that mimics a nucleic acid backbone). Optimization experiments should seek a competitor

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concentration that reduces nonspecific interactions without disrupting the specific interaction. It is difficult to predict which competitor will be effective for any particular RBP, so both types should be explored. Typical concentration ranges are between 0.1 and 1  mg/mL heparin, and between 1 and 50 mg/mL tRNA. Heparin at 1 mg/mL proved to be an excellent nonspecific competitor in binding reactions involving the pentatricopeptide repeat proteins PPR5 and PPR10 (6, 17), whereas tRNA (50 mg/mL) or high salt (330  mM Na+) was more effective for the CRM domain protein CRS1 (20).

2. Materials for Poisoned Primer Extension Assay 2.1. RNA Extraction and Storage

2.2. Experimental Design

Total RNA can be extracted from plant tissue with commercial organic extraction reagents such as Trizol (Invitrogen). Extraction of RNA from isolated chloroplasts is not necessary and is less likely to yield intact RNA. Tissue to be used for RNA extraction should be stored at −80°C. Tissue should be ground to a fine powder in liquid nitrogen using a mortar and pestle. After the addition of extraction buffer, the material should be ground further until it is a homogenous slurry. Care should be taken to avoid any thawing of the tissue prior to its immersion in the extraction buffer, as this will result in RNA degradation. All aqueous solutions used for RNA extraction, storage, and analysis should be treated to remove contaminating ribonucleases. Filtration of autoclaved solutions through 0.2-m nitrocellulose disposable syringe filters provides a simple and effective means to remove trace protein contaminants such as ribonucleases. “RNAse-free” solutions should be stored in small aliquots in disposable plastic tubes. RNA samples should be stored at −80°C. A custom primer for reverse transcription is paired with a specific dideoxynucleotide such that they yield products of different size from spliced and unspliced RNA templates. Each splice junction is assayed with a different primer/dideoxynucleotide combination. The primer is a ~20-mer oligodeoxynucleotide that is complementary to sequences several nucleotides downstream of the splice junction to be assayed. One dideoxynucleotide is included in the reaction, which terminates reverse transcription at the first occurrence of its complement in the RNA template. The distance from the primer to the first nucleotide complementary to the chain terminator must differ between spliced and unspliced RNA templates. The greater the size difference between the free primer and the two primer extension products, the easier it is to resolve them cleanly on gels. For unknown reasons, ddATP, ddTTP, and ddCTP give better results than ddGTP. Primer/dideoxynucleotide

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combinations that can be used to monitor the splicing of some chloroplast introns in Arabidopsis were described previously (22). 2.3. Stock Solutions

1. 10× Polynucleotide kinase (PNK) buffer: 700 mM Tris–HCl, pH 7.6, 100  mM MgCl2, and 50  mM dithiothreitol (DTT). 2. 5× Annealing buffer: 50 mM Tris–HCl, pH 8.5, and 300 mM KCl. 3. 5× RT buffer: 250 mM Tris–HCl, pH 8.5, 40 mM MgCl2, and 50 mM DTT. 4. Formamide dye mix: 90% deionized formamide (v/v), 20 mM Tris–HCl, pH 8, 20  mM ethylenediaminetetraacetic acid (EDTA), 0.02% (w/v) bromophenol blue, and 0.02% (w/v) xylene cyanol. 5. 10× TBE: 890 mM Tris base, 890 mM boric acid, and 20 mM EDTA. 6. 40% (w/v) acrylamide/2% (w/v) bis-acrylamide. 7. 10% (w/v) ammonium persulfate (APS). 8. 7.5 M ammonium acetate (NH4OAc). 9. dNTP/ddNTP mixes: 10 mM of the relevant dideoxynucleoside triphosphate (ddNTP) and 10 mM of each of the other three deoxynucleoside triphosphates (dNTPs). For example, the mix for reactions to be terminated with ddCTP should consist of 10 mM ddCTP, 10 mM dATP, 10 mM dTTP, and 10 mM dGTP. Store at −20°C. 10. Reverse transcription master mix (per reaction): 2 mL of H2O, 2 mL of 5× RT buffer, 0.5 mL of dNTP/ddNTP mix, 0.25 mL of RNAsin, and 0.25 mL of transcriptor reverse transcriptase. Immediately before the experiment, prepare a sufficient volume of master mix for one more than the total number of reactions to be performed. Store on ice.

2.4. Reagents and Equipment

1. “RNAse-free” water: double-distilled water that has been autoclaved and then filtered through a disposable 0.2-m nitrocellulose filter to remove trace protein contaminants. 2. 100% ethanol. 3. (g32P]ATP, 6,000 Ci/mmole. 4. Glycoblue (Ambion). 5. N,N,N ¢,N ¢-tetramethylethylenediamine (TEMED). 6. Urea. 7. Transcriptor reverse transcriptase (Roche) (see Note 1). 8. T4 polynucleotide kinase (T4 PNK). 9. RNAsin ribonuclease inhibitor (Promega).

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10. Total leaf RNA suspended in 10  mM Tris–HCl, pH 8, and 1 mM EDTA, at a concentration of at least 1 mg/mL. 11. Custom synthetic oligodeoxynucleotide primers (~20-mers, HPLC purified), suspended in water to 10  pmol/mL. These should be designed to monitor the splicing of specific introns as described above. 12. Standard equipment for polyacrylamide gel electrophoresis, microcentrifugation, sample drying (e.g., SpeedVac), imaging of radioactive gels (e.g., a phosphorimager), and controlled temperature incubations. A scintillation counter, together with vials and fluid, is also required.

3. Methods All steps should be performed at 4°C unless otherwise indicated. For each primer/ddNTP pair, a control reaction should be performed that includes RNA suspension buffer instead of RNA. 3.1. Radiolabeling the Primer

1. In a 500-mL microfuge tube, combine 2  mL of primer (10 pmol/mL), 1.5 mL of 10× PNK buffer, 1 mL of T4 PNK (10 U/mL), 1 mL of (g32P]-ATP, and 9.5 mL of H2O. Incubate at 37°C for 30 min. 2. Add 35 mL of water, 13 mL of 7.5 M NH4OAc, 125 mL of icecold ethanol, and 1 mL of glycoblue. Mix and store at −20°C for at least 1 h. Centrifuge for 20 min at 12,000 ´ g in a microcentrifuge. Carefully pipette off the supernatant and discard in radioactive waste. 3. Rinse the pellet with ~200 mL of 70% ethanol and microcentrifuge for 10 min at ~15,000 rpm. Pipette off the ethanol and dry the pellet by centrifugation under vacuum in a SpeedVac. Avoid overdrying the pellet, as this can make it difficult to resuspend. 4. Resuspend the pellet in 50 mL of 10 mM Tris–HCl, pH 8.3. Estimate the incorporation of radiolabel by counting 1 mL in a scintillation counter. Dilute with 10 mM Tris–HCl, pH 8.3, to 75,000 cpm/mL. Store at −20°C for up to 1 week.

3.2. Preparing the Denaturing Polyacrylamide Gel

The gel should be poured at least several hours and up to 1 day prior to running the reactions. Gels that are 0.4-mm thick and ~40-cm long provide sufficient resolution. Dissolve 24 g of urea in 9 mL of water, 5 mL of 10× TBE, and 18.75 mL of 40% acrylamide/2% bis-acrylamide. Add 300 mL of 10% APS and 30 mL of TEMED to initiate polymerization. (This is sufficient to pour one gel using a 20  cm × 40  cm × 0.4  mm gel mold). Immediately pour into the gel mold and insert the comb. Let it sit at room temperature for at least 2 h and up to 1 day.

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3.3. Annealing the Labeled Primer to the RNAs

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1. Combine 1 mL of radiolabeled primer, 3 mg of leaf RNA, 1 mL of 5× annealing buffer, and H2O to bring the total volume to 5 mL. (It is best to combine sufficient primer, buffer, and water for all reactions, and to then distribute this mix to tubes containing the different RNA samples.) Place in a beaker containing ~200 mL of water at ~95°C and allow to cool on the bench to ~48°C. 2. During this incubation, assemble the reverse transcription master mix.

3.4. Primer Extension by Reverse Transcription

1. Microcentrifuge the annealing reactions briefly to pellet the condensate. Add 5 mL of the reverse transcription master mix to each tube. Incubate at 48˚–55°C (depending on the melting temperature (Tm) of the primer; see Note 2) for 30 min. 2. Stop the reaction by adding 15 mL of formamide dye mix and heating for 3 min at 95°C.

3.5. Denaturing Polyacrylamide Gel Electrophoresis

1. Pre-run the gel in 1× TBE for ~30  min at 25  mA (constant power, room temperature). Immediately prior to loading the samples, turn off the power and clean the wells with reservoir buffer expelled through a syringe. 2. Load 10 mL of each sample. Run the gel at 25 mA (constant power, room temperature) until the bromophenol blue (the more rapidly migrating dye) is ~4 cm from the bottom of the gel (see Note 3). This should take several hours. 3. Disassemble the gel apparatus. Transfer the gel to a used piece of X-ray film by laying the film on the gel and gently peeling the film back to coax the gel from the plate. 4. Wrap the gel/film in plastic wrap and expose to a phosphor screen for ~6–12 h (see Note 4). An example of a successful experiment is shown in Fig. 1.

4. Notes 1. AMV reverse transcriptase can also yield suitable results. However, signals are considerably stronger with transcriptor reverse transcriptase. If AMV reverse transcriptase is used, the 5× RT and 5× annealing buffers should be considered to be “4×” stocks, to account for a small difference in the salt optima of the two enzymes. 2. Higher temperatures reduce the formation of RNA structures that could otherwise inhibit reverse transcription. However, the reaction temperature must be below the Tm of the RNA/ primer duplex. Transcriptor reverse transcriptase is active up to

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at least 55°C; incubation at 50°C is suitable for primers that are ~40% GC (the average GC content of land plant chloroplast DNA). Reactions performed with AMV reverse transcriptase should be incubated at 48°C. 3. In this type of gel, the bromophenol blue comigrates with 12-mer deoxyribonucleotides, and the xylene cyanol comigrates with 35-mers. The products of poisoned-primer extension are typically between 25 and 30 nucleotides. 4. Products of poisoned-primer extension are small (typically 25–30 nucleotides) and can diffuse noticeably after exposures of ~12 h. If a gel drying apparatus is available that can dry a 15% polyacrylamide gel to completion, we recommend the use of 0.2-mm-thick gels, and drying the gel before exposing it to the phosphor screen.

Acknowledgments I thank Yukari Asakura and Kenny Watkins for their input into the development of the protocol for poisoned-primer extension, and Yukari Asakura for permission to present the data in Fig. 1. The guidelines for RNA-binding assays evolved from discussions with and data obtained by Kenny Watkins, Rosalind Williams-Carrier, Oren Ostersetzer, and Margarita Rojas. References 1. Stern, D.B., Goldschmidt-Clermont, M., and Hanson, M.R. (2010) Chloroplast RNA metabolism. Annu. Rev. Plant. Biol. 61, 125–155. 2. Barkan, A., Klipcan, L., Ostersetzer, O., Kawamura, T., Asakura, Y., and Watkins, K. (2007) The CRM domain: an RNA binding module derived from an ancient ribosomeassociated protein. RNA 13, 55–64. 3. Kroeger, T., Watkins, K., Friso, G., Wijk, K.v., and Barkan, A. (2009) A plant-specific RNA binding domain revealed through analysis of chloroplast group II intron splicing. Proc. Natl. Acad. Sci. USA 106, 4537–4542. 4. Schmitz-Linneweber, C., and Small, I. (2008) Pentatricopeptide repeat proteins: a socket set for organelle gene expression. Trends Plant Sci. 13, 663–670. 5. Barkan, A. (1998) Approaches to investigating nuclear genes that function in chloroplast biogenesis in land plants. Methods Enzymol. 297, 38–57.

6. Pfalz, J., Bayraktar, O., Prikryl, J., and Barkan, A. (2009) Site-specific binding of a PPR ­protein defines and stabilizes 5’ and 3’ mRNA termini in chloroplasts. EMBO J. 28, 2042–2052. 7. Kuhn, J., and Binder, S. (2002) RT-PCR analysis of 5’ to 3’-end-ligated mRNAs identifies the extremities of cox2 transcripts in pea mitochondria. Nucleic Acids Res. 30, 439–446. 8. Slomovic, S., Portnoy, V., and Schuster, G. (2008) Detection and characterization of ­polyadenylated RNA in Eukarya, Bacteria, Archaea, and organelles. Methods Enzymol. 447, 501–520. 9. Falcon de Longevialle, A., Hendrickson, L., Taylor, N., Delannoy, E., Lurin, C., Badger, M., Millar, A.H., and Small, I. (2008) The pentatricopeptide repeat gene OTP51 with two LAGLIDADG motifs is required for the cis-splicing of plastid ycf3 intron 2 in Arabidopsis thaliana. Plant J. 56, 157–168.

12  Studying the Structure and Processing of Chloroplast Transcripts 10. Jenkins, B., Kulhanek, D., and Barkan, A. (1997) Nuclear mutations that block group II RNA splicing in maize chloroplasts reveal several intron classes with distinct requirements for splicing factors. Plant Cell 9, 283–296. 11. Watkins, K., Kroeger, T., Cooke, A., WilliamsCarrier, R., Friso, G., Belcher, S., Wijk, K.v., and Barkan, A. (2007) A ribonuclease III domain protein functions in group II intron splicing in maize chloroplasts. Plant Cell 19, 2606–2623. 12. Beick, S., Schmitz-Linneweber, C., WilliamsCarrier, R., Jensen, B., and Barkan, A. (2008) The pentatricopeptide repeat protein PPR5 stabilizes a specific tRNA precursor in maize chloroplasts. Mol. Cell Biol. 28, 5337–5347. 13. Schmitz-Linneweber, C., Williams-Carrier, R.E., Williams-Voelker, P.M., Kroeger, T.S., Vichas, A., and Barkan, A. (2006) A pentatricopeptide repeat protein facilitates the transsplicing of the maize chloroplast rps12 pre-mRNA. Plant Cell 18, 2650–2663. 14. Ostheimer, G., Williams-Carrier, R., Belcher, S., Osborne, E., Gierke, J., and Barkan, A. (2003) Group II intron splicing factors derived by diversification of an ancient RNA binding module. EMBO J. 22, 3919–3929. 15. Schmitz-Linneweber, C., Williams-Carrier, R., and Barkan, A. (2005) RNA immunoprecipitation and microarray analysis show a chloroplast pentatricopeptide repeat protein to be associated with the 5'-region of

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mRNAs whose translation it activates. Plant Cell 17, 2791–2804. 16. Barkan, A. (2008) Genome-wide analysis of RNA-protein interactions in plants. In, Plant Systems Biology (Belostotsky, D., ed.) Humana Press, Totowa, NJ, USA, pp. 13–37. 17. Williams-Carrier, R., Kroeger, T., and Barkan, A. (2008) Sequence-specific binding of a chloroplast pentatricopeptide repeat protein to its native group II intron ligand. RNA 14, 1930–1941. 18. Wong, I., and Lohman, T. (1993) A doublefilter method for nitrocellulose-filter binding: application to protein-nucleic acid interactions. Proc. Natl. Acad. Sci. USA 90, 5428–5432. 19. Setzer, D.R. (1999) Measuring equilibrium and kinetic constants using gel retardation assays. Methods Mol. Biol. 118, 115–128. 20. Ostersetzer, O., Watkins, K., Cooke, A., and Barkan, A. (2005) CRS1, a chloroplast group II intron splicing factor, promotes intron folding through specific interactions with two intron domains. Plant Cell 17, 241–255. 21. Draper, D.E. (2008) RNA folding: thermodynamic and molecular descriptions of the roles of ions. Biophys. J. 95, 5489–5495. 22. Asakura, Y., and Barkan, A. (2006) Arabidopsis orthologs of maize chloroplast splicing factors promote splicing of orthologous and speciesspecific group II introns. Plant Physiol. 142, 1656–1663.

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Chapter 13 In Vitro RNA-Binding Assay for Studying Trans-Factors for RNA Editing in Chloroplasts Toshiharu Shikanai and Kenji Okuda Abstract In plant organelles, specific C residues are modified to U by RNA editing. Short RNA sequences surrounding the target site (i.e., cis-elements) are recognized by trans-factors, which were recently shown to be pentatricopeptide repeat (PPR) proteins. PPR proteins consist of tandem arrays of a highly degenerate unit of 35 (pentatrico) amino acids, and PPR motifs are believed to recognize specific RNA sequences. In Arabidopsis thaliana, more than 450 sites are edited in mitochondria and plastids, and a similar number of PPR proteins are encoded in the nuclear genome. To study how the tandem array of a PPR motif facilitates the recognition of RNA sequences, an efficient biochemical strategy is an in vitro binding assay of recombinant PPR proteins with target RNA. This analysis is especially powerful with a combination of in vivo analyses based on the phenotypes of mutants and transgenic plants. In this chapter, we describe methods for the expression of recombinant PPR proteins in Escherichia coli, preparation of probe RNAs, and RNA gel shift assays. These methods can also be utilized for other RNA-binding proteins. Key words: PPR protein, RNA-binding protein, RNA editing, RNA electrophoresis mobility shift assay, Trans-factor

1. Introduction RNA editing is a posttranscriptional process that alters the genetic information on RNA molecules (1). In land plants, this process involves the conversion of specific C residues to U, although the reverse reaction from U to C often occurs in mosses, hornworts, and ferns (2). In Arabidopsis thaliana, 34 and approximately 450 sites are edited in plastids and mitochondria, respectively (3, 4). Two experimental technologies have contributed to clarifying the mechanism by which specific C residues are recognized by the RNA editing machinery in tobacco. Using plastid transformation,

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sequences encoding RNA editing sites can be introduced into plastids, where the sites are precisely edited (5, 6) (see Note 1). By introducing RNA editing sites with different lengths of surrounding sequences, the RNA sequences required for site recognition for RNA editing were determined. In general, 20–30 nucleotides surrounding the editing site (fewer than 20 upstream and 10 downstream nucleotides) are sufficient to be recognized by the RNA editing machinery and are called cis-elements. This discovery was followed by a complementary study using an in vitro RNA editing system in which a competitor for the site-specific factor (transfactor) was added quantitatively (7). Trans-factors recognizing cis-elements were first discovered in the study of photosynthetic electron transport. Eleven plastid ndh genes (ndhA-ndhK) encode subunits of chloroplast NADH dehydrogenase-like complex (NDH), which is involved in cyclic electron transport around photosystem I (8). Arabidopsis crr4 (chlororespiratory reduction 4) mutants are specifically defective in the RNA editing event that generates the translational start codon of ndhD (9). The CRR4 gene encodes a member of the pentatricopeptide repeat (PPR) family of proteins that contain a tandem array of a PPR motif, a highly degenerate unit of 35 amino acids. According to a recent model, PPR proteins act as site-recognition factors (trans-factors) that recognize cis-elements and recruit still unknown RNA editing enzymes to catalyze the conversion of C residues to U at target sites (Fig. 1). A surprising feature of the PPR family is its huge size, especially in flowering plants: the Arabidopsis genome encodes approximately 450 members, the majority of which are predicted to target to plastids or mitochondria (10). The PPR family is specific to eukaryotes, and its size increased during the evolution of land plants. The size of the PPR family is well correlated with the number

Fig. 1. A model of the RNA editing machinery in plant organelles. PPR proteins recognize and bind to cis-elements surrounding the target C residues. The tandem array of the PPR motif is involved in RNA sequence recognition. PPR proteins may recruit still unknown editing enzymes that catalyze the C to U conversion.

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of RNA editing sites, and this observation is consistent with the idea that each PPR protein recognizes a single or several RNA editing sites. Accumulating genetic information supports this idea in both plastids and mitochondria (11). In addition to RNA editing, PPR proteins are involved in many steps of gene expression in organelles, including RNA stabilization, RNA splicing, RNA processing, and translation. In these processes, PPR proteins are thought to recognize target RNAs, as in RNA editing, and somehow recruit the machinery for RNA maturation (12). One of the hot topics in this research area is clarifying how a tandem array of PPR motifs is able to recognize different target sequences. In such studies, powerful tools include in vitro RNAbinding assays using a recombinant PPR protein and in vivo analysis by transforming PPR protein genes. In this chapter, we describe a method for evaluating the specific binding of PPR proteins to their target RNA molecules. This technique depends on the expression of recombinant proteins in Escherichia coli and analysis of their binding with a probe RNA using an electrophoresis mobility shift assay (EMSA).

2. Materials 2.1. Vector Construction

1. Expression vector (e.g., pGEX-6P and pBAD/Thio-TOPO).

2.2. Purification of Recombinant Protein

1. E. coli (e.g., Rosetta 2 [DE3] and TOP10), growth media, and incubators. 2. Isopropyl thio-b-galactoside (IPTG). Prepare a 0.1 M solution in water. Sterilize by filtration and store at −20°C. 3. Buffer I: 50  mM Tris–HCl, pH 8.0, 500  mM NaCl, 1  mM phenylmethylsulfonyl fluoride (PMSF), and 7  mM b-mercaptoethanol. 4. Sonicator for disrupting bacterial cells. 5. GSTrap FF column (GE Healthcare, Fairfield, CT, USA). This column is for pGEX-6P; alternative columns may be necessary if a different expression vector is to be used. 6. Binding buffer: 140 mM NaCl, 2.7 mM KCl, 10 mM Na2HPO4, and 1.8 mM KH2PO4, pH 7.3. 7. Dialysis cassette (Pierce, Rockford, IL, USA). 8. Centrifugal concentrator (e.g., Centricon, Millipore, Billerica, MA, USA). 9. Coomassie (Bradford) protein assay kit (Takara, Kyoto, Japan).

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2.3. Preparation of Probe or Competitor RNA

1. SP6/T7 transcription kit (Roche Applied Science, Penzberg, Germany). 2. TE buffer: 10 mM Tris–HCl, pH 8.0, and 1 mM ethylenediaminetetraacetic acid (EDTA). 3. 80% (v/v) glycerol. 4. 5¢ EndTagTM nucleic acid labeling system (Vector Laboratories, CA, USA). 5. Fluorescein maleimide (Vector Laboratories). 6. 30% (w/v) acrylamide/bis-acrylamide solution Caution: this is a neurotoxin when unpolymerized.

(29:1).

7. 10× TBE buffer: 890  mM Tris, 890  mM boric acid, and 20 mM EDTA. 8. 10 M urea. 9. 10% (w/v) ammonium persulfate (APS). Store at 4°C and use within 1 month. Caution: this is harmful. 10. N,N,N ¢,N ¢-tetramethylethylenediamine (TEMED). Caution: this is corrosive. 11. Electrophoresis system and power supply (e.g., AE-6200 and AE-8155, ATTO, Tokyo, Japan). 12. Glass plates (110 × 100  mm), 1  mm spacers, and 14-well combs. 13. Ethidium bromide (EtBr) solution (1 mg/mL). Caution: this is a mutagen; handle with plastic gloves at all times. 14. UV illuminator for visualizing EtBr-stained gels. 15. RNA extraction buffer: for 200 mL, mix 180 mL of TE, 20 mL of 3  M sodium acetate, and 2  mL of 10% (w/v) sodium dodecyl sulfate (SDS). 2.4. RNA EMSA

1. 20× Buffer IVT: 75 mM MgCl2, 40 mM dithiothreitol (DTT), and 2700 mM KCl. 2. Buffer E: 20 mM 4-(2-hydroxyethyl)piperazine-1-ethanesulfonic acid (HEPES)-KOH, pH 7.9, 60 mM KCl, 12.5 mM MgCl2, 0.1 mM EDTA, 17% (v/v) glycerol, and 2 mM DTT. 3. 6% Polyacrylamide gel without urea prepared in the electrophoresis system (e.g., AE-6200, ATTO). 4. 5× TGE: 30.3 g Tris, 3.9 g EDTA, and 142.7 g glycine (for 1 L). 5. 10× TBE buffer: 890  mM Tris, 890  mM boric acid, and 20 mM EDTA. 6. Loading buffer: 80% (v/v) formamide, 10  mM EDTA, pH 8.0, and 0.025% (w/v) bromophenol blue. Store at −20°C. 7. Hybond N+ nylon membrane (GE Healthcare). 8. Whatman 3MM paper.

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9. UV cross-linker (e.g., Stratalinker, Agilent Technologies, Santa Clara, CA, USA). 10. Gene Image CDP-Star detection kit (GE Healthcare). 11. Chemiluminescence analyzer (e.g., LAS1000, Fuji Film, Tokyo, Japan). 12. A cold room or a chromato-chamber for electrophoresis (set at 4°C).

3. Methods 3.1. Vector Construction

In order to express PPR proteins in E. coli, DNA sequences encoding mature PPR proteins are cloned into an expression vector. For the expression of PPR proteins in an active form, the choice of vector is critical. CRR4 was successfully expressed as a fusion protein with glutathione S-transferase (GST) using the pGEX-6P-1 vector (GE Healthcare) (13).

3.2. Expression and Purification of Recombinant Protein

Another key step is optimization of the protein expression conditions. The choice of host cells, growth temperature, and induction conditions such as the concentration of IPTG should be carefully investigated. To assay RNA-binding activity, the recombinant proteins should be expressed in an active form, and it is necessary to optimize the conditions for soluble expression. The following steps are optimized for the expression of GST-CRR4. 1. Culture at least 2–3 L of E. coli (e.g., Rosetta 2 [DE3]) carrying the expression vector at 37°C to OD600 = 0.4. 2. Transfer the E. coli to 22°C and continue culturing for an additional 30 min. 3. Add 0.4 mM IPTG and continue culturing at 22°C for 6 h to induce expression of the recombinant protein. The induction conditions should be optimized for each recombinant protein. 4. Centrifuge the cells at 5,000 × g for 30 min at 4°C. 5. Suspend the cells in precooled buffer I and disrupt them by sonication. 6. Centrifuge the lysate at 15,000 × g for 30 min at 4°C. 7. Pre-filter the supernatant through a 0.45-mm pore filter (Millipore). 8. Load the filtrate onto a GSTrap FF column (GE Healthcare) pre-equilibrated with binding buffer. 9. Elute the recombinant protein with the same buffer containing 10 mM reduced glutathione. 10. Pool the eluted protein and concentrate it with a centrifugal concentrator.

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11. Dialyze the proteins overnight at 4°C in buffer E (see Subheading 2.4). 12. Determine the protein concentration using the Bradford method. Bovine serum albumin can be used as a standard. 13. Analyze a sample of the purified protein by SDS-PAGE using a mini-gel system followed by Coomassie blue staining, to assess the purity and integrity of the protein. 3.3. Preparation of Probe or Competitor RNA 3.3.1. Template Construction

3.3.2. In Vitro Transcription

DNA templates for probes or competitor RNAs are prepared and placed under the control of T7 RNA polymerase in a suitable vector. The T7 promoter sequence should be attached to the 5¢ end of the forward primer for PCR (see Note 2). Alternatively, the template DNA can be cloned into a vector that already carries the T7 promoter. The probe RNA is designed to include a putative target sequence for the PPR protein being studied. For cold competitor RNA, the same RNA sequence is used without labeling. As a negative control competitor, an RNA sequence unrelated to the target site is used. As an example, Fig. 2 summarizes the positions of the probe and competitor RNAs used in the analysis of CRR4 (13). An SP6/T7 transcription kit is used for in vitro transcription. To obtain enough RNA, we usually increase the reaction volume to 80 mL (×4). 1. Add the following to a fresh 1.5-mL microfuge tube: Template DNA (0.2 mg of PCR product or 1 mg of plasmid DNA), 2 mL each of 10 mM ATP, 10 mM GTP, 10 mM CTP and 10 mM UTP, 2 mL of 10× buffer (1× final), 1 mL of RNase inhibitor (20  U/mL), and 2  mL of T7 RNA polymerase (10  U/mL). Make up the volume to 20 mL with RNase-free H2O. 2. Incubate the reaction for 2 h at 37°C. 3. Add 2 mL of DNase I (10 U/mL) and incubate for 15 min at 37°C.

Fig. 2. Design of the probe and competitor RNAs. CRR4 is involved in an RNA editing event that converts ACG to AUG to create the translational initiation codon of ndhD. The minimum cis-element was determined to be from −25 to +10 where the first A of ndhD is +1 (12). RB1 was used for both the probe and competitor, while RB2, which does not include the cis-element, was used as negative competitor.

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4. Add 20 mL of reaction mixture to 76 mL of TE, 10 mL of 3 M sodium acetate, and 250 mL of ethanol, and then incubate for 20 min at −20°C. 5. Precipitate the RNA by centrifugation at 15,000 × g for 20 min at 4°C. 6. Dissolve the pellet in 10 mL of RNase-free H2O. 3.3.3. Gel Purification of RNA

1. Denature the RNA at 90°C for 30  s and place it on ice immediately. 2. Add 2 mL of 80% glycerol to the 20 mL sample. 3. Prepare a 7M urea-6% polyacrylamide gel (see Note 3). 4. Pre-run the gel using 1× TBE for 10 min at a constant voltage of 200 V. 5. Load the samples onto the gel. 6. Run the gel for 80 min (for 100–200-base RNAs) at a constant voltage of 200 V. 7. Stain the gel in EtBr solution. Under UV illumination, use a clean scalpel to excise a gel slice containing the RNA and place it into a 1.5-mL tube. 8. Add 200  mL of RNA extraction buffer and incubate at 4°C overnight to elute the RNA from the gel. 9. Add 500 mL of ethanol to the 200-mL sample and incubate for 20 min at −20°C. 10. Precipitate the RNA by centrifugation at 15,000 × g for 20 min at 4°C. 11. Dissolve the pellet in 10 mL of RNase-free H2O (see Note 4).

3.3.4. Labeling the RNA Probe

3.4. RNA EMSA Assay 3.4.1. Incubation of RNA and Protein

The 5¢ end of the RNA is labeled with fluorescein maleimide as described in the instruction manual for the 5¢ EndTagTM nucleic acid labeling system. After labeling, the probe RNA is gel purified again (see Subheading 3.3.3). 1. Add the following to fresh 1.5-mL microfuge tubes: 1 mL of 20× buffer IVT (1× final), P mL of protein (1 fmol to 1 pmol) dialyzed in buffer E, B mL (P + B = 5 mL) of buffer E, and Up to 13 mL of RNase-free H2O (final reaction volume after adding RNA should be 20 mL). 2. Mix gently and incubate for 10 min at room temperature. 3. Separately, incubate the RNA (probe and competitor) at 70°C for 30 s and then place the tubes on ice immediately. 4. For the competition test, add cold RNA to the reaction, mix carefully, and then incubate at 25°C for 5 min. 5. Add 1 mL of the labeled RNA to each reaction, mix carefully, and then incubate at 25°C for 15 min.

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6. Add 2 mL of 80% glycerol to each 20-mL sample and mix carefully (see Note 5). 7. Apply 10–15 mL of sample immediately to a pre-run polyacrylamide gel (see Subheading 3.4.2). 3.4.2. Polyacrylamide Gel Electrophoresis

The appropriate acrylamide concentration depends on the size of the RNA probe or complexes being studied. In most cases, a 6–8% native polyacrylamide gel in 1× TBE or 1× TGE buffer is optimal. When the RNA probe is less than 100 nucleotides, an 8% polyacrylamide gel is suitable. 1. Prepare a native polyacrylamide gel with 6% or 8% acrylamide in 1× TBE or 1× TGE buffer. 2. Pre-run the gel at a constant current of 30 mA for 30 min in a cold room. 3. Load the samples (see Subheading 3.4.1) onto the gel. 4. Run the gel at a constant current of 30 mA in the cold room until the dye migrates to the bottom of the gel (see Note 6).

3.4.3. Blotting and Cross-Linking

1. Carefully remove one glass plate from the gel. 2. Place a dry Hybond N+ nylon membrane on the gel. 3. Place three layers of dry Whatman 3MM paper on the membrane. 4. Put a glass plate and a 1,500–2,000-g weight on the top of the stack. 5. Allow the RNA transfer to proceed for more than 60 min at room temperature. 6. After blotting, place the membrane on a fresh piece of Whatman 3MM paper and cross-link at 120 mJ with a Stratalinker. 7. The membrane can be stored at room temperature between two sheets of Whatman 3MM paper.

3.4.4. Chemiluminescent Detection

1. Detect the signal from the labeled RNA using a Gene Image CDP-Star detection kit. 2. Quantify the signal using a chemiluminescence analyzer. A representative pattern of EMSA is shown in Fig. 3.

4. Notes 1. If the trans-factor is conserved among plants, heterologous sequences (e.g., Arabidopsis editing sites) can also be analyzed by this system. 2. T7 promoter TATAGGG-3¢.

sequence:

5¢-ATGTAATACGACTCAC

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Fig. 3. EMSA with the recombinant CRR4 and its target RNA. This figure is modified from ref. 13. We also added 1.5 nM yeast tRNA (optional), which was absent in the lanes indicated by asterisks. pRB1 is a probe RNA including the cis-element. cRB1 is a cold RNA with the same sequence, while cRB2 is a cold RNA whose sequence is not related to the CRR4 target site. See the positions of probe and competitor RNAs in Fig. 2. The positions of the CRR4–RNA complex and free RNA are indicated by C and F, respectively.

3. 7M urea-6% polyacrylamide gel: Mix 2 mL of 30% acrylamide solution, 1  mL of 10× TBE, and 7  mL of 10  M urea. Add 90  mL of 10% APS and 10  mL of TEMED, pour into glass plates, and wait for 30 min until the gel is solidified. 4. The concentration and integrity of RNA can be confirmed by agarose gel electrophoresis with a dilution series of tRNA standards. For this purpose, load 1 or 2  mL of the sample. For labeled RNA, go to Subheading 3.3.4. For cold RNA, directly use the extracted RNA. 5. Do not use a loading buffer with any dye such as bromophenol blue. 6. Load the dye to the empty lanes. Make sure that the gel does not heat up during the run. The shift pattern depends on the reaction and electrophoresis conditions. If necessary, these conditions should be optimized (e.g., by changing protein concentration or buffer system). References 1. Shikanai, T. (2006) RNA editing in plant organelles: machinery, physiological function and evolution. Cell Mol. Life Sci. 63, 698–708. 2. Yoshinaga, K., Iinuma, H., Masuzawa T., and Uedal, K. (1996) Extensive RNA editing of U to C in addition to C to U substitution in the rbcL transcripts of hornwort chloroplasts and the origin of RNA editing in green plants. Nucleic Acids Res. 24, 1008–1014. 3. Tsudzuki, T., Wakasugi, T., and Sugiura, M. (2001) Comparative analysis of RNA editing sites in higher plant chloroplasts. J. Mol. Evol. 53, 327–332.

4. Giege, P., and Brennicke, A. (1999) RNA editing in Arabidopsis mitochondria effects 441 C to U changes in ORFs. Proc. Natl. Acad. Sci. USA 96, 15324–15329. 5. Chaudhuri, S., Carrer, H., and Maliga, P. (1995) Site-specific factor involved in the editing of the psbL mRNA in tobacco plastids. EMBO J. 14, 2951–2957. 6. Bock, R., Hermann, M., and Kössel, H. (1996) In vivo dissection of cis-acting determinants for plastid RNA editing. EMBO J. 15, 5052–5059. 7. Hirose, T., and Sugiura, M. (2001) Involvement of a site-specific trans-acting factor and a

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c­ ommon RNA-binding protein in the editing of chloroplast mRNAs: development of a chloroplast in vitro RNA editing system. EMBO J. 20, 1144–1152. 8. Shikanai, T. (2007) Cyclic electron transport around photosystem I; genetic approaches. Annu. Rev. Plant Biol. 58, 199–217. 9. Kotera, E., Tasaka, M., and Shikanai, T. (2005) A pentatricopetide repeat protein is essential for RNA editing in chloroplasts. Nature 433, 326–330. 10. Lurin, C., Andrés. C., Aubourg, S., Bellaoui, M., Bitton, F., Bruyère, C., Caboche, M., Debast, C., Gualberto, J., Hoffmann, B., Lecharny, A., Le Ret, M., Martin-Magniette, M. L., Mireau, H., Peeters, N., Renou, J. P., Szurek, B., Taconnat, L., and Small, I. (2004) Genome-wide analysis of Arabidopsis penta­

tricopeptide repeat proteins reveals their essential role in organelle biogenesis. Plant Cell 16, 2089–2103. 11. Schmitz-Linneweber, C., and Small, I. (2008) Pentatricopeptide repeat proteins: a socket set for organelle gene expression. Trends Plant Sci. 13, 663–670. 12. Okuda, K., Myouga, F., Motohashi, R., Shinozaki, K., and Shikanai, T. (2007) Conserved domain structure of pentatricopeptide repeat proteins involved in chloroplast RNA editing. Proc. Natl. Acad. Sci. USA 104, 8178–8183. 13. Okuda, K., Nakamura, T., Sugita, M., Shimizu, T., and Shikanai, T. (2006) A pentatricopeptide repeat protein is a site-recognition factor in andchloroplast RNA editing. J. Biol. Chem. 281, 37661–37667.

Chapter 14 Studying Translation in Arabidopsis Chloroplasts Paolo Pesaresi Abstract Chloroplasts as descendents of a cyanobacterial endosymbiont have retained, during evolution, their own genome together with the gene expression machinery, including the translation apparatus. Therefore, chloroplast protein synthesis is not only a key process in organello biogenesis and maintenance, but it also represents the major regulatory step in chloroplast gene expression. In fact, several independent evidences have shown that the accumulation of template messengers is not limiting in the expression of chloroplast genes. On the contrary, translation regulatory processes based on selection of translatable mRNA by either nucleus-encoded activation factors or sensors of the assembly status of chloroplast multiprotein complexes have been reported. Additionally, we have shown that organelle translation rate triggers an organelleto-nucleus signaling cascade aimed to modulate nuclear gene expression according to the organelle’s needs. Therefore, the study of chloroplast translation appears to be essential for the comprehension of several aspects of chloroplast activity. Here, we describe the in vivo pulse-chase and the polysome isolation approaches. Taken together, the two methods allow one to assess rates of protein synthesis and degradation as well as defects during the initial steps of protein synthesis. Key words: Chloroplast, In vivo labeling, Protein translation rate, Polysome assembly

1. Introduction The chloroplasts present in plant and algal cells are descendents of an original cyanobacterial endosymbiont. Reflecting their origin, chloroplasts still resemble bacteria: they carry their own DNA and gene expression machinery including bacteria-like ribosomes. However, molecular genetic studies of plastid genomes have found that most of cyanobacteria-derived genes are no longer inside the chloroplasts, but they have been transferred to the cell nucleus during evolution (1). Hence, chloroplast multi-protein complexes such as ribosomes or photosystems are mosaics of nucleus-encoded and plastid-encoded subunits, implying the existence of mechanisms that coordinate gene expression in the different compartments. 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_14, © Springer Science+Business Media, LLC 2011

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In particular, several studies have demonstrated that chloroplasts developed their major regulatory steps in gene expression at the translational and posttranslational levels (2, 3). Therefore, comparison of chloroplast protein synthesis in Arabidopsis wild-type and mutant plants appears to be an indispensable and straightforward approach to gain insight on the molecular basis of chloroplast biogenesis and maintenance. Here, two methods for the investigation of protein synthesis in the chloroplasts are described: (1) in  vivo radiolabeling of intact seedlings to assess rates of protein synthesis and degradation and (2) polysome isolation to investigate the presence of defects during the initiation or early elongation phases of protein synthesis.

2. Materials 2.1. Plant Propagation

1. Plastic trays with peat moss-based soil mix consisting of one part of perlite, one part of fine vermiculite, and one part of sphagnum moss. Perform fertilization with “Osmocote Plus” (15% N, 11% P2O5, 13% K2O, and 2% MgO; Scotts Deutschland GmbH, Nordhorn, Germany) according to the manufacturer’s instructions (see Note 1). 2. Plants in soil are grown in a growth chamber with long-day conditions: 16 h of 100 mmol/m2/s light with 50% humidity and 22°C; 8 h dark with 65% humidity and 18°C. 3. MS medium: dissolve 1× Murashige and Skoog (MS) mineral salts in the appropriate amount of water. Adjust to pH 5.8 with KOH and add 0.8% (w/v) agar. Sterilize by autoclaving. After autoclaving, let the liquid medium to cool down to 60°C and pour approximately 25 mL of it into each 9-cm Petri dish in a laminar flow hood. Store at room temperature. 4. Seed surface sterilization reagents: 95% (v/v) ethanol; 10% (v/v) Clorox bleach together with 0.2% (w/v) sodium dodecyl sulfate (SDS); sterile water. 5. Plants on MS medium are grown in a growth chamber with long-day conditions: 16 h of 50 mmol/m2/s light at 22°C; 8 h dark at 18°C. 6. Pasteur glass pipettes; these will be employed to add the seeds to the MS medium containing Petri dishes. 7. Laminar flow hood; this will be necessary for operating in a sterile environment.

2.2. SDSPolyacrylamide Gel Electrophoresis

1. Peqlab minigel system (Peqlab Biotechnologie GmbH, Erlangen, Germany), or any other electrophoresis apparatus. 2. Separating polyacrylamide gel (2×): 30% acrylamide/bis-acrylamide solution (37.5/1). Store at 4°C.

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3. Stacking polyacrylamide gel (6×): 30% acrylamide/bis-acrylamide solution (37.5/1). Store at 4°C. 4. Separating buffer (4×): 1.5  M Tris–HCl, pH 8.8, and 0.4% (w/v) SDS (4). Store at room temperature. 5. Stacking buffer (4×): 0.5 M Tris–HCl, pH 6.8, and 0.4% (w/v) SDS. Store at room temperature. 6. Gel formation initiators: ammonium persulfate (APS), 10% (w/v) aqueous solution (divide into aliquots of 500  mL and store at −20°C), and N,N,N ¢,N ¢-tetramethylethylenediamine (TEMED). 7. Running buffer (10×): 1.9 M glycine and 0.25 M Tris, pH 8.3. Store at room temperature (see Note 2). 8. Isopropanol; this will be used to produce a smooth, completely level surface on top of the separating polyacrylamide gel. 9. Sample buffer (4×): 10% (v/v) glycerol, 0.06 M Tris–HCl, pH 6.8, 2% (w/v) SDS, 5% (v/v) b-mercaptoethanol, and 0.01% (w/v) bromophenol blue. Divide into aliquots of 500 mL and store at −20°C. 10. A plastic syringe of 5 cm3; this will be employed to rinse the wells of the SDS-PAGE before sample loading. 11. Pre-stained molecular weight markers. Store at −20°C. 12. A Hamilton syringe of 50 mL capacity; this will be used to load the protein samples onto the SDS-PAGE. 13. Heating block (Thermomixer Comfort, Eppendorf AG, Hamburg, Germany); this will be employed for protein denaturation. 14. Gel-dryer (Quick-Gel-Dryer, International PBI, Milan, Italy). 2.3. In Vivo Labeling and Radiolabel Detection

1. Pulse-labeling mix: 0.1 mCi/mL of [35S]-l-methionine, 1 mM K2HPO4–KH2PO4, pH 6.3, and 0.1% (w/v) Tween-20 (5, 6). Prepare fresh prior to use (see Note 3). 2. Chase mix: 10  mM l-methionine, 1  mM K2HPO4–KH2PO4, pH 6.3, and 0.1% (w/v) Tween-20. Prepare fresh prior to use. 3. A cork drill of 0.5-cm diameter or, as an alternative, a plastic drinking straw of the same diameter; this will be employed to obtain leaf disks from young Arabidopsis leaves. 4. Coarse sand paper; this will be employed to facilitate leaf infiltration. 5. A transparent plastic syringe of 50 cm3 with a 2-way stopcock replacing the needle. The stopcock will be essential to regulate air flow into the syringe. Air vacuum in the syringe will be generated by pulling the plunger while the stopcock is fully closed. 6. Petri dishes (9-cm diameter); these will be employed to incubate leaf disks in either the pulse-labeling mix or the chase mix for the desired period, under the desired light intensity.

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7. Liquid nitrogen; this will be employed to stop the reactions at the desired time and to store leaf disks. 8. Assay of [35S]-l-methionine incorporation (7): Whatman DE 81 filter disks; 0.5 M Na2HPO4; 95% (v/v) ethanol; scintillation cocktail; and distilled water. 9. Geiger counter; this will be employed to detect the radioactive signal intensity on the dried gel, thus allowing to estimate the length of SDS-PAGE exposure to X-ray films. 10. Autoradiography films (Hyperfilm MP, GE Healthcare, Piscataway, NJ, USA) and cassettes (Amersham Hypercassette, GE Healthcare, Piscataway, NJ, USA) will be employed to detect radiolabeled proteins after SDS-PAGE fractionation. Alternatively, a phosphorimager (Typhoon Trio, GE Healthcare, Piscataway, NJ, USA) and the required storage phosphor screens and cassettes can be used. 2.4. Protein Extraction and Quantification

1. Homogenization buffer (8): 0.4  M sucrose, 10  mM NaCl, 5 mM MgCl2, 10 mM Tricine-KOH, pH 7.5, 100 mM ascorbate, 5  mg/mL fraction V bovine serum albumin (BSA), 0.2 mM phenylmethylsulfonyl fluoride (100 mM stock solution dissolved in isopropanol. Store at 4°C), 1 mM benzamidine (100 mM stock solution dissolved in 20 mM HEPES-KOH, pH 7.5. Store at 4°C), and 5 mM e-aminocaproic acid (500 mM stock solution dissolved in water. Store at 4°C). Prepare fresh prior to use (see Note 4). 2. Hypotonic buffer: 5  mM Tricine-KOH, pH 7.9. Autoclave and store at 4°C. 3. Resuspension buffer I: 0.4  M sucrose, 10  mM NaCl, 5  mM MgCl2, 10 mM Tricine-KOH, pH 7.5, and 20% (w/v) glycerol. Store at 4°C. 4. Handled homogenizer (RW 16 basic, IKA Werke, Staufen, Germany); this will be employed to disaggregate leaf tissue and cell wall. 5. Refrigerated benchtop centrifuge (Z 216 MK, Hermle Labortechnik, Wehingen, Germany). 6. Plastic syringe of 3 cm3 equipped with 20-mm pore-size nylon filter (Millipore, Billerica, MA, USA). 7. Bradford assay kit (Quick Start™ Bradford protein assay, Biorad, Hercules, CA, USA) together with BSA. 8. Spectrophotometer (Biophotometer, Eppendorf AG, Hamburg, Germany); this will be employed for protein quantification together with the Bradford assay kit.

2.5. Polysome Isolation

All buffers and solutions (unless stated otherwise) are made with diethyl pyrocarbonate-treated water (DEPC water) in order to inactivate RNAase enzymes (9, 10) (see Note 5).

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1. Polysome extraction buffer: 0.2 M Tris–HCl, pH 9.0, 0.2 M KCl, 35 mM MgCl2, 25 mM EGTA, 0.2 M sucrose, 1% (w/v) Triton X-100, and 2% (v/v) polyoxyethylene-10-tridecyl ether. Filter the buffer through 0.2-mm-pore sterile filtration unit and store in aliquots at −20°C. Immediately prior to use, add 0.5  mg/mL heparin (stock solution 100  mg/mL. Store at −20°C), 100  mM b-mercaptoethanol, 100  mg/mL chloramphenicol (stock solution 50 mg/mL dissolved in ethanol. Store at −20°C), and 25  mg/mL cycloheximide (stock solution 10 mg/mL. Store at −20°C). 2. Polysome gradient buffer (10×): 0.4 M Tris–HCl, pH 8.0 (see Note 5), 0.2 M KCl, and 0.1 M MgCl2. Store at 4°C. 3. Polysome sucrose step gradient solutions: dilute 70% (w/v) sucrose stock solution to make 15, 30, 40, and 55% sucrose solutions with 3 mL of polysome gradient buffer (10×), 150 mL of heparin, 60 mL of chloramphenicol, 75 mL of cycloheximide, and DEPC water to reach 30 mL of final volume. 4. Step sucrose gradients: add 1.1 mL of 55% sucrose solution to 5-mL ultracentrifuge tubes and freeze at −80°C. Pipette 1.1 mL of 40% solution on top of the 55% frozen solution and freeze again at −80°C. Repeat this with the 30 and 15% sucrose solutions. Step gradients can be stored at −80°C for long periods and thawed overnight at 4°C just prior to use. 5. Solubilization solution: 10% (w/v) sodium deoxycholate. Store in 500 mL aliquots at −20°C. 6. Denaturing solution: 5% (w/v) SDS and 0.2  M EDTA, pH 8.0. Store at room temperature. 7. Phenol/chloroform/isoamyl alcohol (25:24:1) is used to isolate RNA from ribosomal proteins. Keep the solution in a bottle wrapped in aluminium foil at 4°C. 8. 95% (v/v) ethanol; this will be employed for RNA precipitation. 9. Resuspension buffer II (TE): 10  mM Tris–HCl, pH 8, and 1 mM EDTA. Store at room temperature. 10. Mortar, pestle, and liquid nitrogen; these will be employed to grind leaf tissue. 11. Plastic syringe of 3 cm3 equipped with 20-mm-pore-size nylon filter (Millipore, Billerica, MA, USA). 12. Refrigerated benchtop centrifuge (Z 216 MK, Hermle Labortechnik, Wehingen, Germany). 13. Beckman L7-55 ultracentrifuge equipped with a Kontron TST 60.4 rotor. 14. Spectrophotometer (Biophotometer, Eppendorf AG, Hamburg, Germany); this will be employed for RNA quantification.

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2.6. Northern Blot Analysis

1. Horizontal midi gel (Peqlab Biotechnologie GmbH, Erlangen, Germany), or any other horizontal electrophoresis apparatus. 2. 2% (w/v) agarose dissolved in 200 mL of water. 3. MEN buffer (10×): 200  mM MOPS, 10  mM EDTA, and 50 mM sodium acetate. Adjust to pH 7 using either acetic acid or 10 N NaOH. Do not autoclave; store at 4°C. 4. Formaldehyde (37%); this is used to denature RNA. 5. Denaturing agarose gel (11). Boil 2% (w/v) agarose solution in a microwave oven in order to dissolve the agarose completely in water. Let the agarose solution cool down to about 60°C at room temperature or in a water-bath, and add 43 mL of formaldehyde (37%) and 27 mL of MEN buffer (10×). After gently stirring, pour the solution into the gel tray and insert the comb at one side of the gel. The gel will become solid in around 20 min, and after that the comb can be removed. Transfer the solidified agarose gel into the buffer chamber and add 1× MEN buffer (diluted in water), making sure to submerge the agarose gel fully (see Note 6). 6. RNA denaturing buffer (2×): 3  mL of formamide (³99.5%, deionized), 1 mL of formaldehyde (37%), and 1 mL of MEN buffer (10×). Prepare fresh prior to use. 7. RNA loading buffer (10×): 15% (w/v) Ficoll, 0.25% (w/v) xylene cyanol, and 0.25% (w/v) bromophenol blue. Store in 500 mL aliquots at −20°C. 8. SSC buffer (20×): 3  M NaCl and 300  mM sodium citrate. Adjust to pH 7 with HCl. Sterilize by autoclaving. 9. A buffer reservoir, positively charged nylon membrane (Roche Diagnostics, Penzberg, Germany), 3-MM blotting paper, dry paper towels, two glass plates, and a 200–500-g weight are all needed for the agarose gel-to-membrane RNA transfer. 10. UV cross-linker (UV Stratalinker 1800, Stratagene, CA, USA); this is needed to cross-link RNA to the membrane, after blotting. 11. Methylene blue RNA staining solution: 0.03% (w/v) methylene blue is dissolved in 0.3 M sodium acetate–acetic acid, pH 5.5. Store at room temperature. 12. Hybridization buffer: 7% (w/v) SDS and 0.25  M Na2HPO4NaH2PO4, pH 7. Store at room temperature. Immediately prior to use, pre-warm 20 mL of buffer at 65°C and add 160 mL of herring sperm DNA previously denatured at 95°C for 5 min. 13. OLB buffer (5×): 250 mM Tris–HCl, pH 8.0; 25 mM MgCl2; 0.35% (v/v) b-mercaptoethanol; 100 mM each dGTP, dATP, and dTTP; 1 M HEPES-KOH, pH 6.0; and 0.54 mg/mL pdN6 random hexamers. Store in 20 mL aliquots at −20°C.

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14. Klenow DNA polymerase and [32P]-dCTP; these are needed for probe labeling. 15. PCR product purification kit (QIAquick PCR purification kit, Qiagen, Hilden, Germany); this will be employed for labeledprobe purification. 16. SSPE buffer (20×): 3 M NaCl, 0.2 M NaH2PO4, and 20 mM EDTA. Adjust to pH 7.4 with NaOH. Store at room temperature. 17. Washing solution I: 0.1% (w/v) SDS is added to 2× SSPE and the solution is used for washing the nylon membrane at 65°C. Store at room temperature and pre-warm at 65°C in a waterbath just prior to use. 18. Washing solution II (10×): 50 mM Na2HPO4–NaH2PO4, pH 7, 10 mM EDTA, and 2% (w/v) SDS. The solution is used for washing the nylon membrane at room temperature. Store at room temperature. 19. Microwave oven; this will be employed to dissolve agarose. 20. Heating block (Thermomixer Comfort, Eppendorf AG, Hamburg, Germany); this will be employed for DNA and RNA denaturation. 21. Hybridization oven (HB-100 Hybridization Oven 115 VAC, Hoefer, Holliston, MA, USA) and corresponding hybridization bottles; these will be employed for filter hybridization with the [32P]-labeled probe. 22. Water-bath; this will be employed to warm hybridization buffer and washing solution I. 23. Geiger counter; this will be employed to detect the radioactive signal intensity on the filter, thus allowing to estimate the quality of hybridization and the length of filter exposure to X-ray films. 24. Detection of filter radioactivity can be performed as described above (see Subheading 2.3, item 10).

3. Methods In vivo pulse-labeling and chase experiments on leaf disks represent a very effective approach to assess rates of protein synthesis and to detect proteolytic processing activities. The method is very reliable and, due to its high sensitivity, only few leaf disks are necessary to detect radiolabeled proteins. For the proteins showing a reduced translation rate in pulse-labeling experiments, the association of ribosomes with the corresponding mRNAs should also be addressed to gain insight into the translation step that is disrupted.

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Indeed, the reduced association of ribosomes with a particular mRNA represents a confirmation of a defect in protein translation during the initiation or early elongation steps. Both pulse-chase and polysome isolation methods are generally used to compare chloroplast translation in Arabidopsis wild-type and mutant plants (2, 12). However, for most mutations affecting plastid translation, mutant plants are notably smaller and paler than their wild-type seedlings and show a delay in development. Moreover, chloroplast translation defects are particularly evident in young developing leaves with highly active chloroplasts, whereas older mutant leaves resemble the wild-type phenotype in many cases. Therefore, for optimal estimation of plastid translation, it is important to analyze pools of young leaves isolated from wild-type and mutant plants at the same developmental stage (identical number of rosette leaves), thus excluding differences in protein translation/degradation rates due to different leaf ages. 3.1. Plant Growth

1. Place Arabidopsis seeds on a wet paper and incubate them for 3 days at 2–5°C, in the dark. This treatment, also called stratification, is very important to break dormancy, improve germination rate, and synchronize germination. 2. Transfer the seeds onto the soil surface with the help of a tooth-stick. Cover the tray with a transparent plastic lid and transfer the tray to the growth chamber. Remove the plastic lid immediately after germination. 3. In the case of growth on sterile MS medium, Arabidopsis seeds must be surface sterilized by soaking, twice, the desired amount of seeds in 1 mL of 95% ethanol for 5 min. After decanting the ethanol, add 1 mL of 10% Clorox bleach together with 0.2% (w/v) SDS and soak the seeds for 5 min, exactly. Afterward, rinse the seeds three times with 1 mL sterile water. Carry out the entire procedure in a laminar flow hood (see Note 7). 4. Add sterile seeds to the MS medium containing Petri dishes with the help of a Pasteur pipette. Incubate them for 3 days at 2–5°C, in the dark, to break dormancy. Afterward, transfer the Petri dishes to the growth chamber.

3.2. Preparation of SDS-PAGE

The instructions assume the use of Peqlab minigel system, but they are adaptable to any electrophoresis apparatus. 1. Assemble glass plates using 1-mm thick spacers. Prepare 10 mL of separating gel solution using 5 mL of separating polyacrylamide gel (2×), 2.5 mL of separating buffer (4×), 100 mL of APS (10%), and 10  mL of TEMED. Pour the gel, leaving enough space for the stacking gel and overlay with isopropanol. 2. After polymerization of separating gel, remove isopropanol and rinse the top of the gel abundantly with water. Prepare 5 mL of

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stacking gel solution using 0.83 mL of stacking polyacrylamide gel (6×), 1.25  mL of stacking buffer (4×), 50  mL of APS (10%), and 10 mL of TEMED. Pour the stacking gel immediately on top of the polymerized separating gel and insert the comb (see Note 8). 3. Prepare 1× running buffer by diluting 50 mL of 10× running buffer with 450 mL of water and add 0.1% (w/v) SDS. 4. Once the stacking gel has polymerized, remove the comb gently and rinse the wells with 1× running buffer using a 5-cm3 syringe. 5. Add 1× running buffer to both lower and upper chambers of the electrophoresis unit before loading the gel with samples and the pre-stained molecular weight marker. 3.3. In Vivo Labeling of Protein Leaves

1. Prepare leaf disks of 0.5-cm diameter from the three youngest leaves of plants at the six-leaf rosette stage, grown in either soil or MS medium, by using a cork drill of the appropriate size or, as an alternative a plastic drinking straw. To facilitate leaf infiltration with pulse-labeling mix, press leaf disks extremely gently against a coarse sand paper (5, 6). Three leaf disks are used for each pulse-chase time point. For Arabidopsis leaves, 15-min pulse appears to be the minimum length to get sufficient protein labeling. For most of the chloroplast proteins, optimal translation kinetics are obtained after pulse labeling for 0, 15, 30, and 60 min (see Fig. 1a). The chase-point length strictly depends on the stability of the protein of interest. In the example reported in Fig.  1a, 8 chase points, each 1  h in length, optimally described the degradation kinetics of photosystem II-D1 (PSII-D1) subunit (see Note 9). 2. Add 10 mL of pulse-labeling mix, which is sufficient to infiltrate 40 leaf disks, into a 50-cm3 transparent plastic syringe devoid of plunger and with a 2-way stopcock replacing the needle. At this stage, the stopcock is closed and the syringe is pointing downward. Add leaf disks into the pulse-labeling mix in the syringe and replace the plunger. Afterward, point the syringe upward, open the stopcock, and push out all of the air in the syringe. Then, close the stopcock and gently pull the plunger down. Repeat this step until all the leaf disks sink, thus indicating that all the air into the leaf disks has been replaced with the pulse-labeling mix. 3. After vacuum infiltration (above), freeze three leaf disks in liquid nitrogen (t0 of pulse-labeling kinetic). Transfer the rest of the leaves together with the pulse-labeling mix to one or more 9-cm Petri dishes and expose them to the desired light conditions for the desired labeling period. Always freeze three leaf disks in liquid nitrogen at the end of each pulse-labeling time point.

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Fig. 1. Thylakoid protein turnover and polysome accumulation in wild-type chloroplasts. (a) Autoradiogram of wild-type thylakoid membrane proteins resolved by SDS-PAGE after pulse-labeling with [35S]-l-methionine for 0, 15, 30, and 60 min (lanes 1–4) and subsequent chase in unlabeled medium for 60, 120, 180, 240, 300, 360, 420, and 480 min (lanes 5–12). An increased or decreased accumulation of radiolabeled proteins in Arabidopsis mutants during the pulse labeling (lanes 1–4) indicates a higher or lower translation rate, respectively. Alternatively, a rapid or slower decrease of radiolabel proteins during the chase period implies a higher or lower protein degradation rate, respectively (see Note 9). The result shown indicates high rates of synthesis and turnover for D1 protein, in contrast with LHCII which is considerably less dynamic; adapted from Bonardi et  al. (6). (b) Association of chloroplast mRNA with ribosomes monitored by polysome fractionation on sucrose gradients. Ten fractions of equal volume were collected from the top to the bottom of the sucrose gradients. An equal proportion of the RNA purified from each fraction was analyzed by Northern blot hybridization with a RbcL-specific probe. If the distribution of the RbcL mRNA is shifted toward the top of the gradient in an Arabidopsis mutant relative to the wild type, this provides evidence for a defect in the translation of RbcL mRNA; adapted from Pesaresi et al. (12). On the contrary, a shift of the RbcL mRNA toward the bottom of the gradient might indicate either enhanced translation or a defect in translation termination. D1, Photosystem II-D1 subunit; LHCII, Light harvesting complex of photosystem II; RbcL, Ribulose bisphosphate carboxylase large subunit.

4. At the end of the pulse-labeling period, collect the remaining leaf disks and wash them abundantly in 9-cm Petri dishes containing the chase mix solution. Afterward, carry out the vacuum infiltration, as described above (see step 2), with the chase mix. Subsequently, transfer the Petri dishes containing leaf disks immersed in the chase mix solution again to the desired light conditions for the desired chase period. Freeze three leaf disks in liquid nitrogen at the end of each chase time point. 3.4. Extraction of Stroma- and Thylakoid-Enriched Protein Fractions

1. Grind three frozen leaf disks in 0.3 mL of ice-cold homogenization buffer with a handheld homogenizer until a homogenous green suspension is obtained (6, 8).

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2. Transfer the green slurry to a 3-cm3 syringe equipped with 20-mm-pore-size nylon filter and collect the filtrate in a 1.5mL microfuge tube. 3. Spin down the solution at 5,000 × g for 10  min at 4°C in a benchtop centrifuge. 4. Resuspend the pellet, enriched in chloroplasts, in 0.1 mL icecold hypotonic buffer and leave in ice for 15 min in darkness. 5. Centrifuge the solution at 11,200 × g for 10 min at 4°C in a benchtop centrifuge. The supernatant, enriched in stromal proteins, is transferred to a new microfuge tube, whereas 50 mL of resuspension buffer I is added to the green pellet, enriched in thylakoid membranes. 6. Assay the incorporation of radiolabeled [35S]-l-methionine by spotting 10 mL of each sample on Whatman DE 81 filter disks (7). Wash the filters five times (2  min each) with 0.5  M Na2HPO4, twice with distilled water (3 min each), and twice with 95% ethanol (30 s each). Add the filters to a scintillation cocktail and measure radiolabeled incorporation in a scintillation counter. Scintillation values higher than 5,000  cpm/mL are optimal for estimating protein translation and degradation rates. 7. Quantify protein amount using the Bradford assay and BSA as standard, following the manufacturer’s instructions. 8. Load 10  mg of protein per sample onto an SDS-PAGE gel; alternatively, store the samples at −80°C in 10  mg protein aliquots to avoid repeated freeze–thaw cycles. 3.5. SDS-PAGE and Radiolabeled Protein Detection

1. Denature 10 mg protein samples in the appropriate volume of sample buffer (4×) for 10 min at 70°C. 2. Load the protein samples and the pre-stained molecular weight marker onto 15% SDS-PAGE, prepared as previously described (see Subheading  3.2), with the help of a 50-mL Hamilton syringe. 3. Run the gel at constant voltage, 50 V, until the samples reach the separating gel (about 30 min). Afterward, turn up the voltage to 100 V until the bromophenol blue reaches the bottom of the gel (about 2 h). 4. After electrophoresis, place the gel on a piece of absorbent paper slightly larger than the gel itself, avoiding formation of air bubbles. Then, place them, gel uppermost, on top of the porous polyethylene sheet of a gel-dryer and cover with some of the nonporous cling film. All of this construction is then covered over with the gel-dryer silicon rubber sheet. 5. Apply vacuum, which should draw all layers tightly together. After checking that there is no air leak, apply heat (~60°C) until

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the gel is dry. At this point, the silicon rubber sheet over the gel should assume a completely flat appearance (see Note 10). 6. When the gel is completely dry and bound onto the absorbent paper, it can be removed from the gel-dryer and the cling film over it peeled off and discarded. The gel is ready for autoradiography or, as an alternative, for phosphorimaging detection (Fig. 1a). 3.6. Polysome Isolation

1. Grind 0.2 g of leaf tissue to a fine powder in liquid nitrogen using a mortar and pestle (9). 2. Transfer the frozen leaf powder to a 1.5-mL microfuge tube and add 1 mL of polysome extraction buffer. Gently shake the tube until a green slurry is obtained. 3. Remove debris by passing the solution through a 3-cm3 syringe equipped with 20-mm-pore-size nylon filter and collect the filtrate into a 1.5 mL-microfuge tube on ice. 4. Keep the solution on ice for 10 min to solubilize membranes. Afterward, centrifuge the sample at 157,000 × g in a benchtop centrifuge for 15  min at 4°C to remove unsolubilized material. 5. Transfer the supernatant to a new 1.5-mL tube and incubate with 1/20 volume of 10% sodium deoxycholate (solubilization solution). Solubilization is performed on ice for 5 min and the unsolubilized material is then spun down in a benchtop centrifuge at 157,000 × g for 15 min at 4°C. 6. Layer 0.5 mL of solubilized material onto 15–55% step sucrose gradients, prepared as described previously (see Subheading 2.5). Balance carefully the tubes by the addition of polysome extraction buffer and centrifuge in a Beckman L7-55 ultracentrifuge with a Kontron TST 60.4 rotor at 210,000 × g for 65 min at 4°C. 7. During centrifugation, label ten 1.5-mL microfuge tubes for each gradient tube and add to each of them 50 mL of denaturing solution. At the end of the centrifugation, fractionate the sucrose gradient tubes by collecting 0.45  mL fractions from the top of the gradients, add them to the prepared denaturing solution-containing tubes, and store on ice (see Note 11). 8. Add to each fraction 0.4 mL of phenol/chloroform/isoamyl alcohol, vortex vigorously, and centrifuge in a benchtop centrifuge at 157,000 × g for 10 min at 4°C. Transfer the aqueous phases to new tubes. 9. Add 1 mL of 95% ethanol to each fraction and invert the tubes several times to mix the solution thoroughly. Centrifuge in a benchtop centrifuge at 157,000 × g for 15 min at room temperature to precipitate RNA.

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10. Remove the supernatants and drain the pellets until almost dry. Add to each tube 30 mL of resuspension buffer II (TE), quantify the RNA by using a UV spectrophotometer, and store the samples at −80°C. 3.7. Polysome Analysis by Northern Blot Hybridization

1. Add 5 mL of RNA denaturing buffer (2×) to 5 mL of polysome RNA and incubate at 65°C for 15 min. Transfer the samples on ice for 5  min and then add 1  mL of RNA loading buffer (10×) to each of them. 2. Load the samples onto denaturing agarose gel, prepared as previously described (see Subheading 2.6), and run the gel at a constant voltage, 60 V, until the bromophenol blue reaches the end of the gel. 3. Place the agarose gel on top of a glass plate together with a piece of 3-MM blotting paper, acting as buffer bridge, and two pieces of 3-MM blotting paper of the same size of the gel. Submerge the two ends of the 3-MM buffer bridge into a reservoir containing 20× SSC buffer. Afterward, place the nylon membrane directly on top of the agarose gel, followed by two pieces of 3-MM blotting paper and a stack of dry paper towels. A glass plate and a 200–500-g weight on top of the stack of paper towels will favor the capillary flow. The RNA will be transferred from the gel onto the nylon membrane by capillarity together with the 20× SSC buffer that is driven by the towel papers from the reservoir through the 3-MM blotting paper bridge, then through the gel, the nylon membrane, and into the 3-MM blotting paper and the stack of paper towels. RNA transfer will take place overnight (see Note 12). 4. The day after, cross-link the RNA to the filter by employing a UV cross-linker and stain the filter with methylene blue to verify equal loading and transfer efficiency (see Note 13). 5. The same day, transfer the filter to a glass hybridization bottle and add 20 mL of hybridization buffer pre-warmed at 65°C, as previously described. Pre-hybridize overnight at 65°C using a hybridization oven. 6. The same day, bring about 100  ng of probe (cDNA/CDS probe of 350–450 bp) to 12 mL with water and denature for 5 min at 95°C. Place on ice for 5 min to cool it down and add 4 mL OLB buffer (5×), 1 mL of Klenow DNA polymerase, and 3 mL of radioactive [32P]-dCTP. Mix by pipetting the solution carefully up and down and let it stand at room temperature till the next day (see Note 3). 7. At the end of the labeling incubation, add 80 mL of water to the probe and purify it using any spin column-based PCR purification kit, following the manufacturer’s instructions.

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8. Dilute the labeled probe to 300 mL with water, add 40 mL of herring sperm DNA, and denature for 5  min at 95°C. Place the probe on ice for 5 min to cool it down and then add it to the hybridization bottle. Hybridize overnight at 65°C. 9. At the end of the hybridization period, throw away the probe from the hybridization bottle and wash the filter with 10 mL of pre-warmed washing solution I for 30 min at 65°C. Wash the filter again with the same buffer for an additional 15 min at 65°C. 10. Take the filter out from hybridization bottle and place it into a reservoir containing 1× washing solution II. Wash the filter at room temperature for 1 h under gentle shaking. 11. At the end of the three washing steps, drain the solution from the membrane and seal it in plastic wrap. The filter is ready for autoradiography (Fig. 1b) or, as an alternative, for phosphorimaging detection (see Note 14).

4. Notes 1. Mix the soil with the fertilizer according to the manufacturer’s instructions. Add the soil to the plastic tray and pour water until the soil is completely wet. Prepare prior to use. 2. Do not adjust the pH of the buffer. The Tris is titrated with glycinate ion to get the correct pH. 3. When working with radioactivity, take appropriate precautions to avoid contamination of the experimenter and the surroundings. Carry out the experiment and dispose of wastes in an appropriately designated area, following the guidelines provided by the local radiation safety officer. 4. Protease inhibitors such as phenylmethylsulfonyl fluoride, benzamidine, and e-aminocaproic acid are harmful if inhaled or absorbed through the skin. Thus, wear gloves and a dust mask during protease inhibitor stock preparation. Keep the protease inhibitor stock solutions at 4°C for not longer than 1 month. 5. Add 0.1% (v/v) of DEPC to water and incubate at room temperature overnight. Autoclave for 20 min to remove residual DEPC. DEPC cannot be used with Tris buffer since it inactivates DEPC by reacting with it. Therefore, nuclease-free Tris buffer is obtained through sterile filtration. 6. The entire Northern blot procedure must be carried out in a fume hood and gloves have to be worn. Formaldehyde is a denaturing agent that protects RNA from degradation; thus it is toxic if inhaled or absorbed through the skin.

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7. Carry out the entire procedure in 1.5-mL microfuge tubes and shake the tubes vigorously at each step. The amount of seeds put into each 1.5-mL tube should not exceed the 100 mL volume. Larger amounts of seeds will hamper the sterilization procedure. Additionally, do not expose the seeds to Clorox bleach for more than 5 min; longer exposure diminishes the germination rate drastically. After the last washing step, leave a few water drops in the tube. That will help with placing the seeds on the surface of MS medium using a Pasteur pipette. 8. Before casting the gel, clean the plates accurately. Lay them out on paper towels and spray with water and then 95% ethanol, scrubbing with a Kimwipe each time. Before assembling the plates, check the bottom for chips. If there is a chip that runs all the way across, discard the plate because these will not form a seal. In addition, it has to be considered that acrylamide is a toxic substance; therefore, it is recommended to use care and wear gloves while handling solutions that contain it. 9. Note that in the case of low abundant proteins with a high turnover rate, a 15-min labeling period might be rather long and the possibility that the protein is very rapidly degraded during the labeling period cannot be excluded. Therefore, under these circumstances, a further assay, such as an in organello pulse-labeling experiment, is recommended (9). 10. Before starting the gel-drying procedure, soak the SDS-PAGE gel in 2% glycerol for 15 min. This pre-treatment will help to prevent gel cracking. The time taken to dry a gel is dependent on various factors, including size and composition of the gel, but a gel of 0.5–1-mm thickness will take 1–2 h to dry. 11. Collection of gradient fractions, each of them of 0.45  mL, allows the isolation of ten different polysome populations. However, in case a higher resolution is needed, collection of 0.35-mL fractions, thus allowing the isolation of 14 polysome populations, is recommended. 12. To save solution and materials, blot only the portion of agarose gel containing fractionated RNA, cutting off all the empty lanes on the sides. Cut the membrane slightly larger than the gel and pre-wet membrane and 3-MM blotting paper in 10× SSC buffer for 5 min at room temperature. During the assembly of the blot “sandwich,” avoid formation of air bubbles by rolling a sterile pipette over the sandwich. 13. Stain the membrane with 0.03% (w/v) methylene blue until the lanes become visible (~15 min). Delineate the RNA marker bands on the side of the membrane with a pencil and cut one corner of the membrane away to establish the orientation of the lanes. Destain the membrane with 2× SSC for about 10 min.

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14. Check the radioactivity on the membrane with a Geiger counter to define the exposure length. The radioactive signal should be detected in a very limited region of the membrane, whereas almost no background signal should be present elsewhere. For [32P] detection with autoradiography films, the cassette must be incubated at −80°C for a variable time length, depending on the intensity of the radioactive signal. References 1. Pesaresi, P., Schneider, A., Kleine, T., and Leister, D. (2007) Interorganellar communication. Curr. Opin. Plant Biol. 10, 600–606. 2. Pesaresi, P., Masiero, S., Eubel, H., Braun, H. P., Bhushan, S., Glaser, E., Salamini, F., and Leister, D. (2006) Nuclear photosynthetic gene expression is synergistically modulated by rates of protein synthesis in chloroplasts and mitochondria. Plant Cell 18, 970–991. 3. Choquet, Y., and Wollman, F. A. (2002) Translational regulations as specific traits of chloroplast gene expression. FEBS Lett. 529, 39–42. 4. Laemmli, U. K. (1970) Cleavage of structural proteins during the assembly of the head of bacteriophage T4. Nature 227, 680–685. 5. Aro, E. M., McCaffery, S., and Anderson, J. M. (1993) Photoinhibition and D1 protein degradation in peas acclimated to different growth irradiances. Plant Physiol. 103, 835–843. 6. Bonardi, V., Pesaresi, P., Becker, T., Schleiff, E., Wagner, R., Pfannschmidt, T., Jahns, P., and Leister, D. (2005) Photosystem II core phosphorylation and photosynthetic acclimation require two different protein kinases. Nature 437, 1179–1182. 7. Mans, R. J., Novelli, G. D. (1961) Measurements of the incorporation of radioactive amino acids

into protein by a filter-paper disk method. Arch. Biochem. Biophys. 94, 48–53. 8. Haldrup, A., Naver, H., and Scheller, H. V. (1999) The interaction between plastocyanin and photosystem I is inefficient in transgenic Arabidopsis plants lacking the PSI-N subunit of photosystem I. Plant J. 17, 689–698. 9. Barkan, A. (1998) Approaches to investigating nuclear genes that function in chloroplast biogenesis in land plants. In, Photosynthesis: Molecular Biology of Energy Capture (McIntosh, L., ed.) Academic Press, New York, USA pp. 38–57. 10. Pesaresi, P., Gardner, N. A., Masiero, S., Dietzmann, A., Eichacker, L., Wickner, R., Salamini, F., and Leister, D. (2003) Cytoplasmic N-terminal protein acetylation is required for efficient photosynthesis in Arabidopsis. Plant Cell 15, 1817–1832. 11. Sambrook, J., Fritsch, E. F., Maniatis, T. (1989) Molecular Cloning: A Laboratory Manual, 2nd edn. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, NY, USA. 12. Pesaresi, P., Varotto, C., Meurer, J., Jahns, P., Salamini, F., and Leister, D. (2001) Knock-out of the plastid ribosomal protein L11 in Arabidopsis: effects on mRNA translation and photosynthesis. Plant J. 27, 179–189.

Chapter 15 Studying Proteases and Protein Turnover in Arabidopsis Chloroplasts Lars L.E. Sjögren and Adrian K. Clarke Abstract Proteolysis is a key process for maintaining homeostasis in all living cells. The ability to degrade specific metabolic enzymes and regulatory proteins is essential for both cellular integrity and function. Equally important is the efficient removal of damaged or otherwise inactive polypeptides, especially during periods of developmental change or stress adaptation. Being one of the most metabolically active plant organelles, chloroplasts require various proteases to control overall protein quality. Much has been revealed about these chloroplast proteases over the last decade, and yet the identity of their native protein substrates remains elusive. In this chapter, we describe a variation upon a classic genetic approach to identify protease substrates based on the comparative protein degradation rates in wild-type and transgenic lines with impaired proteolytic activity. We have successfully used this approach with an in organello assay to identify numerous substrates for the stromal Clp protease from Arabidopsis thaliana, using both gene knockout mutants and antisense repression lines. In principle, the technique can be readily adapted for the study of other chloroplast proteases, and in other plant and algal species as the necessary genetic resources become available. Key words: Arabidopsis thaliana, Chloroplasts, Mutants, Proteases, Protein degradation

1. Introduction Chloroplasts perform a range of metabolic processes within plant cells and contain anywhere from 2,000 to 3,000 proteins functioning at any given time or developmental stage. Various proteases help maintain this dynamic protein environment, by removing short-living enzymes and regulatory proteins as well as those that are misfolded and otherwise damaged. This protein quality control within chloroplasts is performed by proteases of bacterial ancestry, including Clp, FtsH, Deg, and Lon (see Note 1). Each protease is localized within distinct sub-chloroplastic compartments, and most are essential for chloroplast functionality and plant viability (1). 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_15, © Springer Science+Business Media, LLC 2011

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Much effort has recently been focused on the characterization of chloroplast proteases in the model plant species Arabidopsis thaliana. Tremendous progress has been made on elucidating the subunit composition of these proteases, their regulation during the plant lifecycle, and their overall importance for chloroplast function (1). This is exemplified by the stromal ATP-dependent Clp protease, a two-component enzyme composed of a central proteolytic core complex flanked on one or both sides by an Hsp100 molecular chaperone partner. The chaperone partner targets the protein substrates and translocates them in an unfolded state into the inner chamber of the proteolytic core, within which they are degraded (2). The chloroplast Clp protease in Arabidopsis is relatively intricate compared to all other orthologs. The proteolytic core complex consists of eleven distinct subunits (ClpP1, ClpP3-6, ClpR1-4, and ClpT1-2) (3), with three potential chaperone partners (ClpC1, ClpC2, and ClpD). By analogy to bacterial counterparts, the core complex presumably consists of two heptameric rings, one of which contains ClpP3-6 and the other ClpP1 plus ClpR1-4 (4). Little redundancy exists within this subunit configuration (5), and loss of any one subunit typically disrupts the assembly of the entire protease. Several genetic studies have now demonstrated the overall importance of the Clp protease (4, 6–9), whose role in chloroplasts is primarily as an essential constitutive housekeeping enzyme. Despite the many advances in characterizing chloroplast proteases in higher plants, identifying their native protein substrates remains an ongoing challenge. To this end, we have recently developed an in organello assay to identify protein substrates for the Clp protease in Arabidopsis (4). This genetic approach compares the degradation profiles of stromal proteins in wild-type and transgenic lines in which the amount of the Clp protease has been reduced to 5–20% of the wild-type levels. The assay as described in detail below has revealed to date 25 potential protein substrates for the Clp protease (5), a number that will almost certainly increase as better protein separation and detection systems are adopted.

2. Materials 2.1. Mechanical Isolation of Intact Chloroplasts and In Organello Degradation Assay

1. Resuspension buffer: 20  mM 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid (HEPES)-NaOH, pH 8, 0.33  M ­sorbitol, 5 mM MgCl2, and 10 mM NaHCO3. 2. Homogenization buffer: 20 mM HEPES-KOH, pH 8, 0.33 M sorbitol, 10  mM ethylenediaminetetraacetic acid (EDTA), 5 mM MgCl2, and 10 mM, NaHCO3.

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3. Mechanical grinder: Ultra-Turrax T25 basic (IKA-Werke, Germany). 4. Sigma 4K15C centrifuge and rotors 11150 (swing-out) and 12169 (fixed angle) (Sigma Labozentrifugen GmbH, Germany). 5. Soft-bristled paintbrush for resuspending intact chloroplasts. 6. Gradient buffer (4×): 80 mM HEPES-NaOH, pH 8, 1.32 M sorbitol, and 20 mM MgCl2. 7. Rupture buffer: 20 mM HEPES-NaOH, pH 7.6, and 10 mM MgCl2. 8. Olympus BX50 phase-contrast microscope (Olympus, Japan) and standard improved Neubauer hemocytometer (Assistant, Germany). 9. Eppendorf centrifuge 5417C and rotor F45-30-11. 2.2. One-Dimensional Denaturing PAGE

1. XCell SureLock® Mini-cell gel electrophoresis system using precast NuPAGE® Novex 3–8% Tris–acetate gels (Invitrogen, USA). 2. Protein concentration is determined using the Coomassie Plus (Bradford) assay (Thermo Scientific, Pierce Biotechnology, USA). 3. NuPAGE LDS sample buffer: 250 mM Tris–HCl, pH 8.5, 2% (w/v) lithium dodecyl sulfate (LDS), 10% (v/v) glycerol, 0.5 mM EDTA, 0.02% (w/v) bromophenol blue, and 50 mM dithiothreitol (DTT). 4. Running buffer: 50 mM Tricine, 50 mM Tris-base, and 0.1% (w/v) sodium dodecyl sulfate (SDS), pH 8.24 (unadjusted). 5. Novex® sharp unstained protein standard (Invitrogen).

2.3. Protein Staining with Colloidal Coomassie Blue G-250

2.4. Western Blotting

1. Fixation solution: 2% (v/v) phosphoric acid and 50% (v/v) ethanol. 2. GFL 3017 orbital shaker (GFL, Germany). 3. Staining solution: 34% (v/v) methanol, 17% (w/v) ammonium sulfate, and 3% (v/v) phosphoric acid. 1. XCell II™ blot module CE mark (Invitrogen). 2. Trans-Blot® supported nitrocellulose membrane (0.2  mm) (Bio-Rad). 3. Transfer buffer: 25  mM Bicine, 25  mM Bis-Tris (free base), 1  mM EDTA, and 0.05  mM chlorobutanol (see Note 2), pH 7.2 (unadjusted). 4. Block buffer: 1× TBS (20 mM Tris–HCl, pH 7.5, and 150 mM NaCl), 5% (w/v) low-fat dried milk powder, and 0.05% (v/v) Tween-20 (see Note 3).

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5. Wash buffer: 1× TBS and 0.05% (v/v) Tween-20. 6. Antibody buffer: 1× TBS, 2% (w/v) milk powder, and 0.25% (v/v) Triton X-100. 7. Secondary antibody: ECL rabbit IgG, horseradish peroxidaselinked whole antibody (from donkey) (GE Healthcare). In  general, the secondary antibody must match the primary antibody used. 8. Enhanced chemiluminescent (ECL) reagents from GE Healthcare (e.g., ECL Advance Western blotting detection kit, GE Healthcare, USA). 9. Plastic film (Glad Cling wrap, The Glad Products Company, USA). 10. ChemiGenius2 imaging system (Syngene, UK). 11. Hyperfilm ECL (GE Healthcare). 2.5. Two-Dimensional Denaturing PAGE

1. ReadyPrep 2D cleanup kit (Bio-Rad, USA). 2. Isoelectric focusing (IEF) buffer: 8  M urea, 2  M thiourea, 50  mM DTT, 4% (w/v) CHAPS, 0.2% (v/v) Bio-Lyte pH 3–10 ampholytes, and 0.0002% (w/v) bromophenol blue. 3. IEF ReadyStrips (24 cm, IPG nonlinear, pH 4–7; Bio-Rad). 4. Ettan™ IPGphor™ 3 IEF system (GE Healthcare). 5. DTT buffer: 6 M urea, 50 mM Tris–HCl, pH 8.8, 2.5% (w/v) SDS, 20% (v/v) glycerol, 2% (w/v) DTT, and 0.002% (w/v) bromophenol blue. 6. Iodoacetamide buffer: 6  M urea, 50  mM Tris–HCl, pH 8.8, 2.5% (w/v) SDS, 20% (v/v) glycerol, 2.5% (w/v) iodoacetamide, and 0.002% (w/v) bromophenol blue. 7. Ettan™ DALTsix large vertical electrophoresis system using precast DALT 12.5% gels (GE Healthcare). 8. Flamingo fluorescent gel stain (Bio-Rad); detection sensitivity of £0.5 ng protein. 9. Ettan™ DIGE imager and ImageMaster™ 2D Platinum 7.0 software (GE Healthcare).

3. Methods A classic genetic approach that has been successful in identifying protein substrates for bacterial proteases has been the use of selected gene mutants. In theory, mutants lacking a certain proteolytic activity will exhibit significantly reduced degradation of substrates native to the affected protease compared to that in the wild type. Protein degradation itself is typically monitored using the “pulse-chase” method.

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This involves cells being first grown for a brief time in the presence of a radiolabeled amino acid (e.g., 35S-methionine) to label or “pulse” all nascent polypeptides. This pulse step is then followed by the “chase,” in which the radiolabeled amino acid is replaced with the non-labeled form. Protein degradation can then be followed over time as the radiolabeled proteins are replaced with nonlabeled ones during normal protein turnover. In order to identify protein substrates for the chloroplast Clp protease in higher plants, we developed a modified form of the above genetic strategy; one that in principle can be adapted for other organellar proteases. Arabidopsis was chosen mainly because of the vast genetic resources currently available for this plant species. Not only has the complete genome been sequenced, but a huge number of T-DNA gene insertion mutants have also been mapped and are freely available. The relative ease of transforming Arabidopsis also enables the targeted repression of selected genes by techniques such as antisense inhibition and RNAi, a particular advantage when the desired knockout mutants are either unavailable or prove embryo lethal. As for the degradation assay itself, we have adopted an alternative approach to that previously used. Although theoretically applicable to study protein turnover in chloroplasts, the pulse-chase method has several drawbacks when applied to plants such as Arabidopsis, especially the need for relatively large quantities of radiolabeled amino acids. Instead, we have developed an in organello technique based on the fact that most chloroplast-localized proteins originate from nuclear genes and are posttranslationally imported from the cytosol. By using isolated intact chloroplasts, the degradation of proteins can readily be followed over time by simply quantifying the amounts of each protein. The method can also be easily adapted for chloroplast-encoded proteins by including in the assay an antibiotic that blocks translation by 70S ribosomes (e.g., lincomycin). Moreover, the method can be readily used to study other plant species once proteasedeficient transgenic lines become available. A schematic overview of the protease assay is shown in Fig. 1. 3.1. Plant Growth Conditions

Due to the comparative nature of the protease assay, all wild-type and transgenic plants should be grown under identical conditions, preferably at the same time and in the same chamber/cabinet. In our experiments, seeds of Arabidopsis thaliana wild-type (ecotype Columbia-0) and transgenic lines are sown in a perlite/soil mix (1:5) following vernalization at 4°C for at least 48 h to break dormancy. All plants are grown as lawns under the following standard conditions: 8 h photoperiod with white light (ca. 150 mmol photons/m2/s), 23/18°C day/night temperatures, and 65% relative air humidity. The two transgenic lines we have so far used in these studies are a clpR1 ethyl methanesulfonate-induced knockout mutant (5) and clpP6 antisense lines (4) (see Note 4).

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Fig. 1. Schematic overview of the in organello procedure to identify protein substrates for chloroplast proteases. The method was originally designed for studying the stromal ATP-dependent Clp protease in Arabidopsis, but in principle it is also suitable for other chloroplast proteases and in other plant and algal species. Intact chloroplasts are isolated from wild type (WT) and a specific protease-deficient mutant or transgenic line (mutant) grown under identical conditions. Equal numbers of intact chloroplasts are then incubated for 3 h in light with ATP, with samples taken every hour. Chloroplast samples are fractionated and stromal proteins separated by denaturing 1D- or 2D-PAGE (proteins >60 and £60  kDa, respectively). Proteins are detected by either staining with various colored/fluorescent dyes or Western blotting using a specific antibody. Proteins are then imaged and their degradation profiles quantified. Unknown proteins whose degradation in the wild type is significantly reduced in the mutant/transgenic lines are identified by various mass spectrometry techniques.

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1. Before starting the isolation, ensure that all buffers and glassware are pre-chilled and that all necessary equipment is setup and ready to operate (see Notes 5 and 6). 2. Prepare four two-step Percoll gradients in 50-mL centrifuge tubes with the top layer having 20 mL of 40% Percoll in 1× gradient buffer and the lower layer 5  mL of 75% Percoll in 1× gradient buffer. 3. For each chloroplast isolation, soak three layers of Miracloth of appropriate size in cold distilled water and then rinse in homogenization buffer. Place the wet cloth in a large funnel sitting in a 500-mL Erlenmeyer flask on ice in a cold room (set at 4°C). 4. Cut 30–45 g of leaves from 3–4-week-old Arabidopsis lawns using sharp scissors (see Note 7). Leaves are typically separated into three batches of 10–15 g. Each batch is placed in a 100-mL beaker and ground in 50 mL of cold homogenization buffer for three 5  s pulses using a suitable mechanical grinder. Repeat if necessary until all material is homogenized (see Note 8). 5. Filter the homogenate from all three batches through the Miracloth into the 500-mL beaker at 4°C. Apply gentle hand pressure to squeeze the excess liquid out of the Miracloth. 6. When performing a second chloroplast isolation, keep the filtrate from the first on ice in darkness while steps 4 and 5 are repeated. When ready, continue with both filtrates simultaneously for the remainder of the isolation procedure, ensuring that each is clearly labeled throughout. 7. Distribute the filtrates into suitably sized tubes/buckets and pellet the chloroplasts by centrifugation at 1,200 × g for 5 min in a fixed-angle rotor. Carefully decant the supernatants, holding the tubes upside-down and wiping the inside with a tissue to remove residual solution. Resuspend the crude chloroplast pellets in a small volume (2–6  mL) of ice-cold resuspension buffer using a soft-bristled paintbrush. 8. Use a Pasteur pipette to remove the crude chloroplast suspensions gently and overlay them on top of the pre-chilled Percoll gradients (two per chloroplast preparation). Run the suspensions slowly down the side of each tube being careful not to disturb the gradient. Centrifuge the gradients at 1,500 × g for 10 min in a swing-out rotor with the brake off. During centrifugation, the intact chloroplasts will sediment at the interface of the 40/75% Percoll layers. 9. After centrifugation, carefully aspirate the top Percoll layer down to the gradient interface and then gently collect the intact chloroplasts using a glass pipette. For each preparation, pool the extracted chloroplasts into a single 50-mL Falcon tube.

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10. Wash the intact chloroplasts by slowly adding five volumes of resuspension buffer and mixing by gentle inversion. Re-pellet the chloroplasts by centrifuging at 1,000 × g for 5 min in a fixedangle rotor. Decant the supernatant and resuspend the chloroplast pellet in ca. 500 mL of resuspension buffer (see Note 9). 11. Use a hemocytometer to count the number of purified intact chloroplasts by phase-contrast microscopy. Use resuspension buffer to equalize the concentration of intact chloroplasts in both preparations (i.e., wild type and transgenic/mutant). Keep the chloroplast preparations on ice in darkness until ready to begin the degradation assay. 3.3. In Organello Degradation Assay

1. Dilute chloroplasts in resuspension buffer to 1.5 × ­­06  chloroplasts/mL. Included in the buffer are 5  mM ATP, 2.5 mM phosphocreatine, and 50 mg/mL creatine phosphokinase (final concentration) (see Note 10). 2. For each preparation, aliquot an equal volume (³20  mL) of chloroplasts into transparent 1.5-mL Eppendorf tubes, one tube for each time point. 3. Incubate chloroplasts for 0–3 h at 25°C in ca. 60 mmol photons/ m2/s light, gently mixing the suspension every 20  min (see Note 11). Samples are taken each hour (see Note 12) by removing a tube from the assay and adding five volumes of rupture buffer. The tube is left on ice in darkness for 10 min to ensure complete breakage of all chloroplasts and then frozen in liquid nitrogen to await further analysis. 4. Once all samples are ready, they are thawed and then centrifuged at 20,000 × g for 10 min in a fixed-angle rotor to fractionate stromal and thylakoid membrane proteins (see Note 13). Supernatants containing stromal proteins are transferred to fresh tubes and stored to await separation by polyacrylamide gel electrophoresis (PAGE).

3.4. Detection of Protein Substrates

Various methods can be used to analyze the isolated chloroplast proteins to identify potential substrates for the protease of interest. A global search for new substrates can be done by separating the  protein samples using different denaturing PAGE systems. In  our studies, we have used one-dimensional (1D)-PAGE for large  molecular mass proteins (>60  kDa) and two-dimensional (2D)-PAGE for smaller ones (£60  kDa). Separated proteins can then be visualized using a range of colored stains and fluorescent dyes depending on the desired sensitivity and suitability for mass spectrometry. Selected proteins are later identified by different mass spectrometry techniques, such as matrix-assisted laser desorption/ ionization – time of flight (MALDI-TOF) or high-­performance ­liquid chromatography – mass spectrometry/mass spectrometry (HPLC-MS/MS). Degradation of a specific protein can also be tested by Western blotting if an appropriate antibody is available.

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1. The following instructions are based on the XCell SureLock® Mini-Cell gel electrophoresis system using precast gels, but other commercial PAGE systems using precast or self-poured gels work equally well. 2. Determine stromal protein content using an appropriate assay. 3. Mix an equal volume with 2 × NuPAGE LDS sample buffer to each sample and then heat at 75°C for 5 min. Cool down the samples on ice and then briefly centrifuge to spin down any condensation. Leave on ice to await gel loading. 4. Prepare an ice bucket for the Mini-Cell system and remember to put a small magnetic stir bar into the outer buffer chamber (in the round stir bar holder). 5. Remove the precast gel from the plastic pouch and rinse in deionized water. Peel off the tape from the bottom of the gel cassette. 6. Gently remove the comb from the cassette and rinse the sample wells with 1× running buffer. Shake out the buffer and repeat twice more. Load the wells once more with buffer to avoid later formation of air bubbles. 7. Place two gels (or one gel and one plastic dam) in the Mini-Cell so that the well side of the cassette faces inward toward the buffer core. Position the gel on the bottom of the Mini-Cell and lock into place. 8. Fill the inner buffer chamber with running buffer until its reaches the level of the wells (ca. 200 mL) (see Note 14). 9. Fill the outer buffer chamber with running buffer until it reaches the same level as the inner chamber (ca. 600 mL). 10. Load the stromal protein samples according to the time course of the degradation assay (see Note 15). The 0-h control samples are loaded on the basis of 15 mg protein, with the remaining samples then loaded on the same volume as the controls. A  suitable protein standard is also included on each gel for accurate size determination (see Note 16). 11. Electrophorese the gel at 150 V (equivalent to 18.75 V/cm) for 1–2 h. 12. Once the run is complete, turn off the power and remove the cassettes from the Mini-Cell. Rinse the cassettes briefly with deionized water. 13. Remove the gels from the cassettes. The separated proteins in each gel are then either stained with Coomassie blue G-250 (see Subheading 3.4.2) or transferred to a nitrocellulose membrane for Western blotting (see Subheading 3.4.3). An example of a stained gel to detect large protein substrates for the chloroplast Clp protease in Arabidopsis is shown in Fig. 2.

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Fig. 2. Identification of large molecular mass protein substrates for the chloroplast Clp protease in Arabidopsis. (a) Equal numbers of intact chloroplasts from wild-type (WT) and clpP6 antisense lines (clpP6 ) were incubated for 3 h in the presence of light and ATP. At 1-h intervals, aliquots were taken and the chloroplasts were ruptured. Following fractionation, stromal proteins were separated by denaturing 1D-PAGE and visualized by colloidal Coomassie blue G-250 staining. The proteins showing significant degradation over the 3-h time course in the wild-type but not in the clpP6 antisense lines were identified by MALDI-TOF mass spectrometry. Shown is a representative degradation assay, with three protein substrates – Elongation factor-Ts (EF-Ts), Hsp90, and RNA helicase – indicated on the right. (b) Degradation profiles for the three substrate proteins from wild-type and clpP6 antisense lines. Values shown are averages ± SE (n = 3) plotted as a percentage of the amount of each protein at time 0, which was set to 100%. Reproduced from ref. 4 with permission from the American Society of Plant Biologists.

3.4.2. Protein Staining with Colloidal Coomassie Blue G-250

1. Place each gel into a suitable plastic container. Add sufficient volume of fixation solution to cover the entire gel surface (ca. 50  mL) and gently shake for at least 3  h (can be left overnight). 2. Wash each gel in distilled water for 3 × 10 min with continuous shaking. 3. Add 50 mL of staining solution to each gel and gently shake for 1 h. During this time, dissolve 33 mg of Coomassie blue G-250 in 2  mL of methanol (per gel) and then add to the staining solution at the end of the 1-h period. 4. Shake the gels in this solution for 6–48  h depending on the sensitivity required.

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5. Pour off the staining solution (see Note 17) and destain the gels by gently shaking in distilled water. 6. Stained proteins on each gel are then imaged and quantified using suitable software (see Note 18). Proteins whose degradation, as seen in the wild type, is significantly reduced in the mutant/transgenic line are then excised from the gel and identified by either MALDI-TOF or HPLC-MS/MS. 3.4.3. Western Blotting for Specific Protein Substrates

1. Protein samples that have been separated by 1D-PAGE are transferred to supported nitrocellulose membranes electrophoretically. These instructions assume the use of a commercial minigel transfer tank system (XCell II™ Blot Module CE Mark, Invitrogen), but other similar commercial transfer systems work equally well (when using another system, follow the manufacturer’s instructions). 2. Soak five blotting (Scotch Brite) pads in transfer buffer. 3. Carefully open the gel cassette and remove the top plate, allowing the gel to remain on the bottom plate. Cut away the wells. 4. Place a piece of presoaked filter paper (see Note 19) on top of the gel. Remove any air bubbles by gently rolling a glass pipette over the filter paper’s surface. 5. Turn the cassette over, so the gel and filter paper are facing downward over a gloved hand or a clean flat surface. Gently push the foot of the gel out of the slot in the cassette; the gel will then easily fall off the cassette plate. Remove the protruding gel foot so that the gel surface is flat. 6. Wet the gel surface with transfer buffer and position the presoaked nitrocellulose membrane on the gel, ensuring that all air bubbles have been removed. Place the presoaked anode filter paper on top of the membrane and again remove any trapped air bubbles. 7. Place two soaked blotting pads into the cathode core of the blot module. Carefully place the gel membrane assembly onto the pad, so that the gel is closest to the cathode plate. 8. Add enough presoaked blotting pads so they reach ca. 0.5 cm over the rim of the cathode core, and then place the anode core on top of the pads. Hold the gel/membrane assembly together firmly and slide it into the guide rails on the lower buffer chamber. Place the gel tension wedge against the blot module and lock by pulling the lever forward. 9. Fill the blot module with transfer buffer until the gel/membrane assembly is covered, but do not fill all the way to the top as this will only generate extra conductivity and heat. 10. Fill the outer buffer chamber with ca. 650 mL of cold deionized water or the remaining transfer buffer. The water level should reach ca. 2 cm from the top of the lower buffer chamber.

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11. Place the lid on the top of the unit and blot at 30 V (~170 mA) for 1 h. 12. When the run is complete, disassemble the module carefully and remove the membrane. If not used directly for immunodetection, the membrane can be wrapped in plastic film and stored at 4°C. 13. Immunodetection begins by first washing the membrane in block buffer (25 mL per membrane) for 1 h at room temperature (RT) (see Note 20). This step will minimize nonspecific binding of antibodies to the membrane. 14. Wash the membrane in ca. 40  mL of wash buffer for 5  min at RT. 15. Add the primary antibody to 20 mL of antibody buffer (dilution range 1:1,000–20,000) (see Note 21) and then add to the membrane. Incubate for 1 h at RT with constant mixing. 16. Briefly rinse the membrane twice with wash buffer and then wash it in ca. 40 mL of wash buffer for 2 × 5 min and 1 × 15 min at RT. 17. Add the secondary antibody (see Note 22) to 20 mL of antibody buffer at a dilution of 1:75,000 (depending on the manufacturer’s instructions) and then add to the membrane. Incubate for 1 h at RT with constant mixing. 18. Wash the membrane for 4 × 5 min in ca. 40 mL of wash buffer. Rinse the membrane in distilled H2O and continue with the ECL detection. 19. During the final wash, 1 mL aliquots of solutions A and B of the ECL reagents are warmed to RT and then mixed into a suitable sized plastic tray at the end of the wash step. 20. Place the membrane in the tray containing the ECL reagents and shake for 5  min, ensuring that solution constantly flows over the membrane surface. 21. Remove the membrane from the ECL solution, blot the excess liquid on a tissue, and then wrap the membrane in plastic film, removing all air bubbles from the surface. 22. Chemiluminescent signals are detected and quantified using a suitable imaging system and associated software (see Note 18). If an imaging system is not available, conventional x-ray film and cassettes can also be used. 3.4.4. Two-Dimensional Denaturing PAGE

1. These instructions are based on available commercial systems with only minor variations as indicated. 2. Stromal protein fractions are purified further using the ReadyPrep 2D cleanup kit (Bio-Rad). The kit precipitates and concentrates the protein sample while removing most substances

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that interfere with the IEF step, such as ionic detergents, salts, lipids, nucleic acids, and phenolics. If the kit is unavailable, the stromal protein sample can be used directly without any further purification. 3. Purified protein (450  mg) is resuspended in IEF buffer and loaded on 24-cm IEF ReadyStrips (IPG nonlinear pH 4–7). 4. The strips are rehydrated for 12  h at 20°C in 24-cm strip holders, followed by IEF on an Ettan IPGphore using the standard separation program. 5. After focusing, the strips are immediately equilibrated for SDSPAGE in DTT buffer for 15 min followed by a 10-min incubation in iodoacetamide buffer at RT with slow constant mixing on an orbital shaker. 6. IEF strips are then placed on precast 12.5% polyacrylamide SDS-PAGE gels and run overnight at 20°C in an Ettan™ DALTsix large vertical electrophoresis system (see Note 23). 7. The gels are stained with a high-sensitivity stain compatible with mass spectrometry and then scanned using a suitable imaging system, and the protein spots are analyzed using associated software. Spot volumes are normalized to the total spot volume on each gel. 8. Spots are selected for mass spectrometric identification based on significant differences in the three replicates between sample pairs; identification is done with MALDI-TOF MS or HPLCMS/MS.

4. Notes 1. The abbreviation “Clp” derives from the name Caseinolytic protease. The names of the other proteases listed come from mutant phenotypes in Escherichia coli (i.e., FtsH, temperaturesensitive strain locus H; Deg, degradation-deficient strain; Lon, long form radiation-sensitive strain). 2. Chlorobutanol is used as a preservative and can be excluded without affecting the efficient transfer of proteins. 3. Milk powder may be replaced with either bovine serum albumin (BSA) or gelatin if blocking is insufficient. 4. Plants should be used at similar developmental stages, particular when the growth rate of the transgenic lines used is significantly different from that of the wild type. 5. Thorough and careful preparation is crucial for the reliable preparation of a high-yield of intact chloroplasts, so as to minimize the time from leaf disruption to chloroplast purification.

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Smaller items needed include scissors, paintbrushes, ice bucket, tubes, racks, pipettes, tips, and Miracloth. 6. Ideally, chloroplasts from both the wild-type and the transgenic/mutant line should be purified simultaneously in order to minimize experimental variation in the subsequent degradation assay performed later that day. 7. Leaves are typically taken from plants around 1 h before the day cycle commences. This minimizes leaf starch content and improves the yield of intact chloroplasts during the isolation. 8. Since mechanical isolation involves disrupting the leaf but not the chloroplasts, the optimal grinding conditions need to be determined by trial and error depending on the plant species, amount of leaf material used, buffer volume, type of grinder, grinding speed, and duration. 9. The final volume of buffer in which to resuspend the purified intact chloroplasts should be determined by trial and error, based on the amount of chloroplasts needed for the degradation assay. 10. ATP is included in the resuspension buffer for the activity of the ATP-dependent Hsp100 molecular chaperone. Phospho­ creatine and creatine phosphokinase are also added as an ATP regeneration system in order to maintain a constant ATP concentration throughout the assay and to minimize the amount of ADP formed (ADP is a potent inhibitor of large AAA [ATPase associated with various cellular activities] proteins such as those in the Hsp100/Clp family). 11. In the first trials of this method, it is important to check that the numbers of intact chloroplasts from wild-type and mutant/ transgenic lines remain similar throughout the assay. Since ­broken chloroplasts have little or no observable proteolytic activity, it is necessary to ensure that apparent changes in the degradation of certain proteins are not due to differences in the proportion of intact chloroplasts after 3  h between the wild-type and mutant/transgenic preparations. In our experiments, although ca. 50% of chloroplasts remained intact by 3 h (which is why the assay was not prolonged beyond this time point), there was no significant difference in the survivability of chloroplasts isolated from the wild-type and clp mutant/ antisense lines. 12. Additional time points between 0 and 3  h can be included depending on the amount of intact chloroplasts available from each preparation. 13. Although originally designed to fractionate stromal proteins, the above method can easily be adapted for analyzing thylakoid membrane proteins. After the first centrifugation, the pellet is resuspended in 1 mL of rupture buffer and then centrifuged

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again at 20,000 × g for 10  min. The supernatant is removed and the thylakoid membrane pellet is resuspended in 20–100 mL of resuspension buffer. 14. The tightness of the seal between the cassette and the buffer core can first be checked by filling the inner buffer chamber with a small amount of running buffer. If a leak is detected from the inner chamber, recollect the buffer, reseal the chamber, and refill. 15. Samples for both the wild-type and mutant/transgenic lines should be loaded on the same gel whenever possible for better comparison of the protein degradation profiles. 16. For the gels destined for protein staining, we use the BenchMarkTM ladder (Invitrogen) as the unstained protein standard, whereas for those destined for Western blotting, we use the pre-stained MagicMarkTM XP Western protein standard (Invitrogen). 17. The staining solution can be reused several times until little or no colloidal Coomassie blue G-250 remains. 18. In our experiments, gel imaging and protein quantification were done using the ChemiGenius2 imaging system (Syngene) and associated software. 19. All filter paper and nitrocellulose membrane should be cut to the same size as the gel (i.e., 7 × 8.5 cm). 20. All washing steps are done by shaking the membrane (preferably on an orbital shaker) so that the solution continuously covers the surface of the membrane. 21. The optimal concentration of the primary antibody needs to be determined for each individual antibody used. 22. The type of secondary antibody used depends on the origin of the primary antibody (from which animal species it came) and the detection system being used for the Western blotting. In  our laboratory, since most of our antibodies are made in rabbits, we typically use anti-rabbit sera from donkey, which is also conjugated to horseradish peroxidase for the ECL detection system. 23. Gels were electrophoresed in simultaneous sets of four to reduce variation, with triplicate gels run for each treatment.

Acknowledgments This work is supported by grants given to A.K.C. from the Swedish Research Council for Environment, Agricultural Science and Spatial Planning.

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References 1. Sakamoto, W. (2006) Protein degradation machineries in plastids. Annu. Rev. Plant Biol. 57, 599–621. 2. Kress, W., Maglica, Z., and Weber-Ban, E. (2009) Clp chaperone-proteases: structure and function. Res. Microbiol. 160, 618–628. 3. Peltier, J. B., Ripoll, D. R., Friso, G., Rudella, A., Cai, Y., Ytterberg, J., et al. (2004) Clp protease complexes from photosynthetic and nonphotosynthetic plastids and mitochondria of plants, their predicted three-dimensional structures, and functional implications. J. Biol. Chem. 279, 4768–4781. 4. Sjögren, L. L. E., Stanne, T. M., Zheng, B., Sutinen, S., and Clarke, A. K. (2006) Structural and functional insights into the chloroplast ATP-dependent Clp protease in Arabidopsis. Plant Cell 18, 2635–2649. 5. Stanne, T. M., Sjögren, L. L. E., Koussevitzky, S., and Clarke, A. K. (2009) Identification of new protein substrates for the chloroplast ATPdependent Clp protease supports its constitutive role in Arabidopsis. Biochem. J. 417, 257–268.

6. Shikanai, T., Shimizu, K., Ueda, K., Nishimura, Y., Kuroiwa, T., and Hashimoto, T. (2001) The chloroplast clpP gene, encoding a proteolytic subunit of ATP-dependent protease, is indispensable for chloroplast development in tobacco. Plant Cell Physiol. 42, 264–273. 7. Rudella, A., Friso, G., Alonso, J. M., Ecker, J. R., and van Wijk, K. J. (2006) Downregulation of ClpR2 leads to reduced accumulation of the ClpPRS protease complex and defects in chloroplast biogenesis in Arabidopsis. Plant Cell 18, 1704–1721. 8. Zheng, B., MacDonald, T. M., Sutinen, S., Hurry, V., and Clarke, A. K. (2006) A nuclearencoded ClpP subunit of the chloroplast ATPdependent Clp protease is essential for early development in Arabidopsis thaliana. Planta 224, 1103–1115. 9. Koussevitzky, S., Stanne, T. M., Peto, C. A., Giap, T., Sjögren, L. L. E., Zhao, Y. et al. (2007) An Arabidopsis thaliana virescent mutant reveals a role for ClpR1 in plastid development. Plant Mol. Biol. 63, 85–96.

Part III Protein Transport, Localization, and Topology

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Chapter 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 Abstract Numerous experimental and in silico approaches have been developed for attempting to identify the ­subcellular localisation of proteins. Approximately 2,000–4,000 proteins are thought to be targeted to plastids in plants, but a complete and unambiguous catalogue has yet to be drawn up. This article reviews the various prediction methods that identify plastid targeting sequences, and those that can help estimate location and topology within the plastid or plastid membranes. The most successful approaches are described in detail, with detailed notes to help avoid common pitfalls and advice on interpreting conflicting or ambiguous results. In most cases, it is best to try multiple approaches, and we also cover the powerful new integrated databases that provide a selected blend of experimental data and predictions. Key words: Chloroplast proteins, Transit peptide, Protein import, Targeting predictions, Protein topology, Transmembrane helices

1. Introduction Plant cells are partitioned into membrane-bound compartments such as the nucleus, the endomembrane system, mitochondria, chloroplasts, and peroxisomes, as well as other cellular and molecular features. In each of these subcellular compartments, specific sets of proteins carry out characteristic biochemical processes. In endosymbiont-derived organelles with autonomous genomes, such as chloroplasts and mitochondria, some of these proteins are encoded by the organellar genome. However, the majority of proteins in these organelles are encoded by the host nuclear genome (1), translated into polypeptides in the cytosol, subsequently targeted to the 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_16, © Springer Science+Business Media, LLC 2011

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organelle, and imported via specialised protein import apparatuses. For additional details on chloroplast protein import, we recommend several excellent reviews (2–6). Defining the intracellular location of a protein provides useful information about its metabolic or biochemical role. However, experimental localisations of proteins such as immunolocalisation (see Chapters 3 and 4, Vol. 1), import studies (see Chapters 17–21, Vol. 1), tagging of proteins by fluorescent markers (see Chapters 4 and 5, Vol. 1), and subcellular proteomics (see Chapters 10–13, Vol. 2) cannot keep pace with the rapid accumulation of highthroughput genomic data, such as genome sequences of entire organisms. For these reasons, computational prediction methods have become increasingly important for annotating newly sequenced genes and for providing testable hypotheses regarding protein localisation and function that may be followed up by experimental studies. In this chapter, we will focus on current computational approaches designed to identify the organellar and suborganellar location of Arabidopsis chloroplast proteins, including membraneassociated proteins. 1.1. Import into the Chloroplast Stroma

Chloroplasts are the organelles that carry out the vital photosynthetic function of plants. They originated from an endosymbiotic event, in which an ancestral photosynthetic cyanobacterium was taken up by a host cell that already contained mitochondria (7). The subsequent substantial transfer of genetic information from the endosymbiont to the host nucleus (8) was only successful with the development and establishment of a protein-import machinery to import chloroplast-localised polypeptides that are now synthesised in the cytosol (9). Chloroplasts are highly structured and contain three distinct membrane systems: the outer and inner envelope membranes and the thylakoid membrane network. These form the borders of three separate soluble compartments: the inter-envelope space, the stroma, and the thylakoid lumen. Most chloroplast proteins, encoded by nuclear genes, have N-terminal targeting sequences, called chloroplast transit peptides (cTPs), which aid in targeting the polypeptides to the correct chloroplast membrane or compartment (10, 11). After translation in the cytosol, the cTPs of precursor proteins (preproteins) are recognised by receptors of the outer envelope translocon, the TOC complex (12, 13). They cross the outer envelope through an aqueous pore and are then transported across the inner envelope membrane by the TIC complex (14). Once imported, the stromal processing peptidase cleaves the cTP to produce the mature form of the protein, which can fold into its native form (15).

1.2. Targeting Signals for Chloroplast Import

The cTPs, directing nucleus-encoded proteins into the chloroplast stroma, are specific and sufficient for protein targeting. However, what precisely comprises a cTP is still poorly understood. The length

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of a cTP varies considerably from 13 to 146 amino acid residues (16). There is little conservation in the primary sequence of cTPs, apart from an alanine as the second residue (in 57% of the 831 Arabidopsis sequences examined), a lack of acidic residues across the entire cTP (except for the second position), and, in Arabidopsis, an enrichment of serines (10, 17). A relatively weak motif, VRA↓AA, surrounds the cTP cleavage site (↓) (17). Approximately 50 Arabidopsis proteins have been reported to be dual targeted to both chloroplasts and mitochondria by an ambiguous dual targeting peptide (dTP) (18–25). The overall properties of the dTPs resemble the characteristics of both cTPs and mitochondrial targeting peptides (mTPs), in that they are enriched in positively charged residues and significantly deficient in acidic residues and in glycine (22, 24, 26). Therefore, the receptors responsible for binding the dTPs in the two organelles probably recognise common features. This highlights one of the major problems faced by prediction software – how to distinguish cTPs from other TPs, particularly mTPs. Further complications come from the fact that a few proteins have been reported to be imported into chloroplasts via the secretory pathway (27–29). These proteins contain an endoplasmic reticulum (ER)-targeting signal peptide at their N-terminus and are initially transported into the ER. They are then translocated to plastids or are dual targeted to both the extracellular space and plastids (27). 1.3. Insertion into the Chloroplast Envelope

Protein sorting in chloroplasts is complex as nucleus-encoded proteins must not only be targeted to the organelle, but must also be sorted between at least six different suborganellar compartments. The mechanisms of targeting nucleus-encoded proteins to the three envelope compartments are not well understood. Most outer envelope proteins are synthesised without a cleavable transit sequence and their targeting information seems to reside within hydrophobic transmembrane (TM) domains (30). It has been suggested that these sequences are sufficient to mediate spontaneous insertion into the membrane bilayer, without assistance from an import apparatus (31). However, more recent data indicate that these proteins use the same translocation channel (Toc75) as those used by transit peptides (32). Targeting to the inter-envelope space has been obtained from studies on Tic22 and monogalactosyldiacylglycerol synthase (MGD1) (33, 34). Both proteins contain a cleavable N-terminal targeting signal and use the TOC complex to cross the outer envelope. MGD1 also engages the TIC complex because it is processed by the stromal processing peptidase, whereas Tic22 is not (34). It is unknown how they are diverted into the inter-envelope space. Proteins localised to the inner envelope membrane contain a cTP and utilise the TOC/TIC system for import. The intrinsic

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TM domains of mature inner envelope proteins may function as stop-transfer domains and proteins diffuse laterally into the inner membrane (35). Alternatively, inner envelope proteins may undergo complete translocation into the stroma prior to membrane integration (36–38). 1.4. Transport into or Across the Thylakoid Membranes

Many fundamental chloroplast proteins (such as photosynthesisrelated proteins) are located in the membrane and the lumen of the thylakoid. Owing to the endosymbiotic origin of chloroplasts, the process of translocating a protein from the stroma to the thylakoid lumen is topologically equivalent to (and evolutionarily reminiscent of) prokaryotic protein secretion (39, 40). In particular, the corresponding N-terminal signals are similar to the secretory signal peptides (SPs). The signals are arranged in a bipartite pre-sequence with the cTP (containing the stromal targeting information) at the N-terminus, immediately followed by the SP-like signal (41). Lumenal proteins are imported into the stroma by the default TOC/TIC pathway. In most cases, the cTP is removed by the stromal processing peptidase, exposing the SP-like signal that directs subsequent translocation across the thylakoid membrane (41). For proteins targeted to the thylakoid lumen, this signal, termed the luminal transit peptide (lTP), shares significant features with the SP of secreted proteins in bacteria, i.e., the three-domain structure and a strongly conserved -3, -1 motif (42–44). The translocation of proteins across the thylakoid membrane to the lumen can occur by one of two very different pathways (41, 42). The Sec-dependent pathway resembles the well-characterised Sec export mechanism in bacterial inner membranes (45, 46). The core elements are SecA, an ATP-driven translocation motor, and the integral membrane proteins SecY and SecE, forming the translocation channel (47). The alternative import pathway, the twin-arginine translocation (Tat) pathway, is a DpH-dependent mechanism (48, 49) characterised by two conserved sequential arginines in the lTP (50). This pathway uses a secretion pore that is separate from the standard translocon built by Sec proteins. Remarkably, proteins targeted to the lumen via the Tat pathway are translocated in a folded state (51–54), whereas proteins targeted via the Sec pathway are generally translocated in an unfolded state (52, 55). Proteins imported into the lumen by either pathway are processed by the same thylakoid processing peptidase to remove the lTP (56). Two additional translocation pathways are responsible for proteins inserted into the thylakoid membrane. The signal recognition particle (SRP) pathway is used by light-harvesting chlorophyllbinding proteins, such as Lhcb1. These proteins are synthesised with a cTP that directs import into the chloroplast. After import, the cTP is removed and the mature protein, containing the targeting information, interacts with the SRP and its partner protein,

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FtsY (41, 57–59). They direct the targeting of the protein to the membrane and insertion occurs in a poorly understood process that depends on the integral membrane protein Alb3 (60). Other thylakoid-bound proteins, including the CFoII subunit of the ATP synthase, and the photosystem I and II subunits PsaG, PsaK, PsbW, PsbX, and PsbY, do not require any of the known protein transport machinery, or any form of free energy (61–66). Therefore, it has been suggested that these proteins insert spontaneously in the thylakoid membrane. This “simple” Sec/SRP-independent pathway for membrane protein insertion appears to be unique to chloroplasts (67). 1.5. Computational Methods for Predicting Chloroplast Localisation

In silico methods designed for predicting chloroplast localisation of proteins fall into several broad categories: (1) methods that rely on comparisons with well-characterised chloroplast proteins to detect homologues or characteristic protein domains, (2) methods that predict N-terminal signal/targeting peptides, (3) methods based on global properties of the protein, and (4) approaches that utilise a mixture of the above strategies. All of these require the amino acid sequence of the protein as input. Other methods exist that use additional input data, such as Gene Ontology terms, expression levels, and phylogenetic profiles. The accuracy is likely to be higher where additional information is provided compared to that in sequence-based methods. However, the applicability is more limited as this additional information is not always available. Annotation by homology to proteins with known subcellular location can yield very good predictive outcomes, but obviously can fail when very similar paralogues are present in different compartments, when protein localisation differs between species, or when previous annotations are in error. Methods that predict the presence of N-terminal signal/targeting peptides are the most widely used, with the majority of available programs exclusively relying on the presence of such a signal. Prediction methods based on global properties use the full protein sequence and obtain successful results through patterns in surface residues that correlate with the biochemical conditions specific to different compartments (e.g., pH). These methods can be useful for genomic and EST sequences where the N-terminal sequence of the protein is unknown. Methods differ not only in the chloroplast-specific features they detect, but also in the computational classification methods they are based on. Several machine-learning methods have been used: hidden Markov models (HMMs), neural networks (NNs), and support vector machines (SVMs). These methods are data driven and adjust their parameters by repeatedly being trained on a dataset of proteins of known localisation and attempting to construct general rules from these. The details of machine-learning algorithms are outside the scope of this chapter.

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1.5.1. Methods for Predicting Protein Localisation to the Chloroplast

An early method to predict cTPs and their cleavage sites in plant proteins was the NN-based ChloroP 1.1 (68). A successor of ChloroP 1.1 is TargetP 1.1 (69), which assigns proteins to four different locations: the secretory pathway, chloroplasts, mitochondria, and “all other compartments”. TargetP, like ChloroP, uses a combination of NNs to calculate a transit or signal peptide score, and a weight matrix to locate the transit peptide cleavage sites. The location is determined based on the predicted presence of one of the N-terminal pre-sequences: cTP, mTP, or SP. For studying plant proteins of unknown localisation, it is recommended to use TargetP first. Once a protein is predicted to be localised in chloroplasts, the cleavage site can be further predicted with ChloroP (70). Detailed explanations of how to use these two predictors are given in Subheadings 3.1 and 3.2. Another tool to predict chloroplast proteins via their characteristic N-terminal targeting sequences is the NN-based Predotar 1.03 (Prediction of Organelle Targeting sequences) program (71). This program was specifically designed to screen whole genomes/ proteomes systematically and has a very low rate of false positives compared with similar programs, including TargetP, though this comes at a price of lowered sensitivity (71, 72). A description of how to use Predotar 1.03 is given in Subheading 3.3. Many other software tools have been developed subsequently to perform the same or similar tasks (Table 1), but none have yet proved to be convincingly superior on real-world data (72–76).

1.5.2. Methods for Predicting Protein Localisation to the Lumen

Because lTPs are quite similar to the signal peptides that target proteins for secretion in bacteria, one way to identify lTPs is to wsearch first for cTPs, followed by a search for signal peptides using the SignalP 3.0 predictor (77). This predictor can be used because the two targeting pathways to the lumen (Sec-dependent and Tat pathways) have analogous systems in Gram-negative bacterial secretory pathways (78). LumenP (79), a neural network-based predictor, was developed for the identification of proteins targeted to the thylakoid lumen of plant chloroplasts and prediction of their cleavage sites. When coupled with TargetP, LumenP achieves significantly better results than previous efforts in predicting thylakoid lumen location, mostly due to a lower false-positive rate (79). Using the combination of TargetP and LumenP predicts that around 1.5–3% of proteins encoded in the Arabidopsis thaliana genome are located in the lumen of the thylakoid. A more recent method, PredSL (73), performs classification of eukaryotic proteins to chloroplast, thylakoid, mitochondrion, secretory pathway, and “other” based on the presence of an N-terminal targeting sequence. When PredSL is compared to TargetP, iPSORT (80), and Predotar, it offers comparable reliability for the prediction to the chloroplast, the mitochondrion, and

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Table 1 Software for predicting the subcellular localisation of plant proteins, particularly chloroplast proteins Program

Description

URL (http://…)

Ref.

AAIndexLoc

Predicts the subcellular location by using amino acid composition and the physico-chemical properties of amino acids. This system has been trained using the dataset from MultiLoc on animal, fungal and plant sequences. Ten locations are predicted in the plant version.

aaindexloc. bii.a-star.edu.sg/

(98)

AdaBoost Learner

Subcellular localisation predictions are based on the amino acid composition. Predicts 12 localisations (ch, cs, cy, er, ex, go, ly, mi, nu, pe, pm, and va) in animals and plants.

chemdata.shu.edu. cn/sl12/

(99)

ATP

Ambiguous Targeting Predictor. Specifically useful for predicting ambiguous dual targeting to chloroplasts and mitochondria. This SVM tool makes use of 12 different amino acid features and tests whether the first 70 amino acids of a plant protein sequence are likely to encode an ambiguous dTP.

www.cosmoss.org/ bm/ATP

(100)

BaCelLo

The Balanced subCellular Localisation predictor discriminates five different localisations in plants (ch, cy, mi, nu, secretory pathway). TM proteins are not considered. Similar to LOCtree, it analyses the protein sequence and its evolutionary profile considering the whole sequence and its N- and C-terminal regions separately.

gpcr.biocomp. unibo.it/bacello

(101)

ChloroP 1.1

Predictor of cTPs in plants including cleavage sites. Should only be used to confirm ch prediction results obtained from TargetP. ChloroP gives a more detailed report of the cTP scores and thereby aids in a higher ­confidence level of both the cTP and cleavage site prediction results.

www.cbs.dtu.dk/ services/ ChloroP/

(68)

EpiLoc

This predictor is a text-based system for ­predicting animal, plant, and fungal protein subcellular locations. Ten locations are ­predicted in the plant version.

www.epiloc.cs. queensu.ca/

(102)

iPSORT

Predictor for N-terminal sorting signals (cTP, mTP, or SP). Consists of an extensive search for simple rules and various attributes, which are partially guided by human intuition.

www.ipsort.hgc.jp/

(80)

(continued )

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Table 1 (continued) Program

Description

URL (http://…)

Ref.

LOCtree

This program utilises a variety of sequence and predicted structural features, with evolutionary information contributing to the predictions’ reliability. Plant proteins are classified into six localisations: secreted, organelles of secretory pathway, nu, cy, mi, and ch. Localisation of membrane proteins is not possible.

www.cubic.bioc. columbia.edu/ cgi/var/nair/ loctree/query

(74, 103)

LumenP

Predictor of putative lTPs to identify proteins targeted to the thylakoid lumen and to predict their cleavage sites. Best used in combination with TargetP.

available from: gunnar@dbb. su.se

(79)

MultiLoc 2

Predictor of animal, plant, and fungal subcellular localisations. Phylogeny and gene ontology terms are integrated in this updated version. A low-resolution version, specialised for globular proteins and predicting five locations, and a high-resolution version, predicting ten locations, are available for plants.

www-bs.informatik. uni-tuebingen. de/Services/ MultiLoc2

(104)

PCLR 0.9

Method to predict chloroplast localisation of proteins in plant cells using a principal component logistic regression (PCLR). Unlike ChloroP, this NN-based method uses the amino acid distribution in the N-terminus with no positional information. Cleavage site is not predicted.

www.andrewschein. com/pclr/

(105)

PlantmPLoc 2.0

Plant-mPLoc discriminates plant proteins among 12 subcellular locations: ch, cy, cw, er, ex, go, mi, nu, pe, pl, pm, and va. It includes predictions of plant proteins with multiple location sites.

www.csbio.sjtu. edu.cn/bioinf/ plant-multi/

(106–108)

Predotar 1.03

www.urgi.versailles. Predictor that assigns a location to mi, pl, er, inra.fr/predotar/ or “elsewhere” based on the identification of a predotar.html putative N-terminal sorting signal. Designed for high-throughput studies and can handle large numbers of input sequences with relatively high accuracy but at a loss of sensitivity.

PredSL

Predictor based on the N-terminal targeting sequence and classifies proteins into five groups: ch, thylakoid, mi, secretory pathway, and “other”. If a sequence is predicted as a chloroplast protein, a HMM is used to ­ determine the existence of an lTP.

www.bioinformatics. (73) biol.uoa.gr/ PredSL/

ProLoc-GO

Sequence-based prediction method that uses gene ontology terms to predict subcellular location of proteins.

www.iclab.life.nctu. (109) edu.tw/prolocgo/ index.php

(71)

(continued )

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Table 1 (continued) Program

Description

URL (http://…)

Ref.

Protein Prowler 1.2

Predictor uses a multilayer classifier system to predict eukaryotic targeting signals as secretory, mi, ch, or “other”.

www.pprowler.imb. uq.edu.au/

(110, 111)

PSORT

PSORT combines a database of N-terminal sorting signals with predictions based on composition and can distinguish 11 subcellular locations. The predictive accuracy is not as good as many more recent tools.

www.psort. ims.u-tokyo. ac.jp/form.html

(112, 113)

SignalP 3.0

Predictor of N-terminal secretory SPs in eukaryotes and bacteria. Results include cleavage site predictions as well as discrimination between SP-containing sequences and non-secretory sequences (signal anchor probability).

www.cbs.dtu.dk/ (77) services/SignalP/

SherLoc2

Hybrid method that combines sequence-based and text-based features for prediction of subcellular localisation of animal, fungal, and plant proteins. The plant version covers ten subcellular locations (ch, cy, er, ex, go, mi, nu, pe, pm, and va). In addition, phylogenetic profiles and Gene Ontology (GO) terms are incorporated to improve the prediction performance considerably. Another feature to improve prediction is DiaLoc, which allows users to provide background knowledge by describing a protein in a short abstract.

www-bs.informatik. uni-tuebingen. de/Services/ SherLoc2

(114)

SLP-Local

Subcellular Location Predictor based on local features of amino acid sequence. Users are able to select from three organism categories plant (eukaryote), non-plant (eukaryote), and prokaryote. Predicted localisations are ch, mi, secretory pathway, and “other” (nu or cy).

www.sunflower. kuicr.kyoto-u.ac. jp/~smatsuda/ slplocal.html

(115)

SLPFA

Predicts localisation by combining methods for sequence alignment and feature vectors based on amino acid composition. Subcellular locations include ch, mi, secretory pathway, and other locations (nu or cy).

www.sunflower. kuicr.kyoto-u.ac. jp/~tamura/ slpfa.html

(116)

SLPS

Subcellular Localisation Predicting System, predicts localisation using a Nearest Neighbor algorithm and incorporating a protein ­functional domain profile.

www.pcal.biosino. (117) org/sub_loc.html

(continued )

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Table 1 (continued) Program

Description

URL (http://…)

Ref.

SubCellPredict Uses the amino acid composition as input into AdaBoost, an algorithm that tries to improve the prediction results by combining several basic and weak predictors. Proteins are ­classified into 12 subcellular locations: ch, cs, cy, er, ex, go, ly, mi, nu, pe, pm, and va.

www.chemdata.shu. edu.cn/sl12/

(99)

SubCellProt

The method uses two machine-learning approaches (k Nearest Neighbor and Probabilistic Neural Network) to classify proteins into one of 11 subcellular locations (ch, cs, cy, er, ex, go, ly, mi, nu, pe, pm, and va).

www.databases. niper.ac.in/ SubCellProt

(118)

TargetP 1.1

Predictor of putative N-terminal cTPs, mTPs, and secretory SPs to identify proteins targeted to ch, mi, secretory pathway, and “other”. Most widely used prediction program for chloroplast proteins.

www.cbs.dtu.dk/ services/ TargetP/

(69)

TatP 1.0

Identifies putative proteins that are imported across the thylakoid membranes via the Tat pathway by predicting the twin-arginine signal peptide. The program also reports a potential cleavage site of the predicted signal peptide.

www.cbs.dtu.dk/ services/TatP/

(81)

WoLF PSORT Successor to PSORTII for eukaryotes. Predictions are based on sorting signals, amino acid content, functional motifs, and sequence length. It classifies proteins into more than ten localisations (ch, cy, er, ex, go, mi, nu, pe, pm, and va), including dual localisation.

www.wolfpsort.org/ (119)

YLoc

www.multiloc.org/ YLoc

Predicts localisation signals or motifs relevant to protein sorting in animals and plants. In addition to the predicted location, it explains why this prediction was made. YLoc+ is able to predict multiple locations per protein.

(120)

Abbreviations for subcellular localisations: ch chloroplast, cp cell plate, cs cytoskeleton, cw cell wall, cy cytosol, en endosome, er endoplasmic reticulum, ex extracellular space, go Golgi apparatus, ly lysosome, mi mitochondrion, nu nucleus, pe peroxisome, pl plastid, pm plasma membrane, va vacuole. Other abbreviations: cTP chloroplast transit peptide, dTP dual targeting peptide, HMM hidden Markov model, lTP luminal transit peptide, NN neural network, SP signal peptide, SVM support vector machine, TM transmembrane.

the secretory pathway (73). In addition, PredSL has the advantage of offering an integrated prediction for the lTP, making it simpler to use for this purpose than the combination of TargetP and LumenP described above. The results of lTP prediction compared to those of LumenP are claimed to be significantly better (73).

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cTP-containing proteins imported across the thylakoid ­ embranes via the Tat pathway can be predicted using the TatP m 1.0 program (81), which is based on an NN in combination with a simple pattern matching of the twin-arginine motif. 1.5.3. Integrated Methods for Predicting Subcellular Localisation

Obviously, it is preferable to use experimental data on protein location where available, rather than relying solely on bioinformatics predictions. Unfortunately, these data are scattered in the literature and it is not always easy to compare evidence obtained using different approaches. Several databases have been developed to integrate predicted and experimental data (Table 2).

Table 2 Integrated databases for identifying the subcellular localisation of plant proteins, particularly chloroplast proteins Program

Description

URL (http://…)

Ref.

ARAMEMNON 6.2

Database of membrane proteins in Arabidopsis thaliana and ~300 other seed plants. Up to 18 predictors contribute to build a consensus for a-helix TM segments. Information from various predictors is shown for N-terminal TM anchors and TM b-barrels. It also classifies proteins into three subcellular locations (ch, mi, and secretory pathway) based on prediction and experimental data.

www.aramemnon. uni-koeln.de

(93)

AT_CHLORO

This accurate mass and time tags (AMT) database is dedicated to the chloroplast proteome from Arabidopsis. It stores information for proteins that have been identified in different chloroplast sub-fractions, including subplastidial localisations.

www.grenoble. prabi.fr/ at_chloro/

(95)

PPDB

Plant Proteome DataBase of subcellular locations in Arabidopsis, maize, and rice: 11 locations and 10 suborganellar locations for chloroplasts based on prediction and experimental proteomic datasets.

www.ppdb.tc. cornell.edu/

(92)

SUBA II

Database of Subcellular localisations for Arabidopsis proteins that unifies unrelated datasets and provides a Web accessible interface for the construction of user-based queries. Assignment of 13 locations (cp, cs, cy, en, er, ex, go, mi, nu, pe, pl, pm, and va) that are based on prediction and experimental datasets (GFP tagging and MS/MS).

www.suba. plantenergy. uwa.edu.au/

(1, 82)

Abbreviations for subcellular localisations: ch chloroplast, cp cell plate, cs cytoskeleton, cw cell wall, cy cytosol, en endosome, er endoplasmic reticulum, ex extracellular space, go Golgi apparatus, ly lysosome, mi mitochondrion, nu nucleus, pe peroxisome, pl plastid, pm plasma membrane, va vacuole. Other abbreviations: GFP green fluorescent protein, MS mass spectrometry.

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The SUBcellular localisation database for Arabidopsis proteins (SUBA) (1, 82) represents such an integrator by bringing together protein localisation information provided by ten different prediction algorithms (iPSORT, LOCtree, MITOPRED, MitoProt 2, MultiLoc, Predotar 1.03, PeroxiP, SubLoc 1.0, TargetP 1.1, and WoLF-PSORT) as well as experimental data from proteomic studies (mass spectrometry), chimeric fusion studies (green fluorescent protein (GFP) tagging), and annotations from Uni-ProtKB/SwissProt (83, 84), AmiGO (85), and The Arabidopsis Information Resource (TAIR) (86). SUBA II provides a web-accessible interface that allows advanced combinatorial queries on the data, resulting in a one-stop shop for protein localisation in Arabidopsis. Subcellular locations in SUBA are represented as 13 locations (cell plate, cytoskeleton, cytosol, endosome, endoplasmic reticulum, extracellular, Golgi, mitochondrion, nucleus, peroxisome, plasma membrane, plastid, and vacuole). At the time of writing, SUBA II contains 195,475 pieces of assembled data on a set of 33,518 Arabidopsis proteins, including plastid- and mitochondrion-encoded proteins. Of these data, 126,185 are calls by predictors and 12,719 are calls from experimental evidence (GFP tagging and mass spectrometry data). The experimental data include data on 6316 distinct proteins localised by GFP tagging and/or mass spectrometry. The use of SUBA II is described in detail in Subheading 3.4. 1.5.4. Methods for Predicting Membrane Protein Localisation

To be able to distinguish whether a protein is located in the stroma/ lumen or is inserted in the envelope (outer envelope membrane, inter-envelope space, or inner envelope membrane) or the thylakoid membrane, in silico predictors have been developed (Tables 3 and 4) that predict TM spanning regions (a-helix and b-barrel). The HMM method of SignalP 3.0 (SignalP-HMM) (77), in the eukaryotic version, is able to distinguish between three types of sequences: SP, signal anchor, and “other”. A signal anchor, just like an SP, contains a region of hydrophobic residues close to the N-terminus, but is not cleaved. Instead, it remains as a TM a-helix in the membrane and anchors the protein to the membrane in an N-in/C-out orientation (87). There are also inverted signal anchors that insert in the membrane with the N-terminus first and anchor the protein in an N-out/C-in orientation (87). SignalP-HMM is not trained to recognise inverted anchor sequences, but TMHMM and other membrane protein topology predictors recognise both signal anchor and inverted signal anchor sequences. To check whether the protein is an integral membrane protein, a TM a-helix predictor can be used (Table  3). For example, TMHMM 2.0 (88) is widely used and has been found to perform well consistently (89, 90). Instead of a TM a-helix, some proteins contain a TM b-barrel that forms a pore with a polar interior and a nonpolar surface (91). These proteins are found in the outer

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Table 3 Software for predicting transmembrane a-helices and membrane topology Program

Description

URL (http://…)

ALOM2

Investigates local properties of amino acid sequences Available via the PSORT to decide which sub-sequences are likely to span server at the membrane. It correctly predicts N-terminal psort.nibb.ac. targeting sequences (i.e., not as TM domains) and jp/ performs well in confirming TM regions with a very low number of false positives (89). This strongly under-predicting tool might serve to increase the degree of confidence in individual TM regions.

(121)

DAS-TMfilter

www.enzim. This tool is a modification of the dense alignment hu/DAS/ surface (DAS) method and authors claim a DAS.html ­substantial decrease in the false-positive error rate. Sequences that include possible TM regions are compared with a sequence library that contains TM segments of documented TM proteins.

(122)

HMM-TM

This approach predicts the TM regions of a-helical membrane proteins and was trained on crystallographically solved data. It incorporates prior topological information in HMMs.

(123)

HMMTOP 2.0

www.enzim. Predicts the localisation of helical TM segments hu/hmmtop/ and the topology of TM proteins. This global approach is based on the difference in amino acid distributions in various structural parts of the protein rather than on the specific amino acid compositions of these parts.

(124, 125)

LIPS

LIPS (LIPid-facing Surface) can be used for predic- gila.bioengr. uic.edu/lab/ tion of TM helix orientation. This method is larisa/ based on a canonical model of the a-helix and lips.html identifies helix–lipid interfaces of TM helices from sequence information alone.

(126)

MEMSAT3

Predicts signal peptide, TM segments, and the TM protein topology. It employs a neural network to determine which residues are on the cytoplasmic side of the membrane and which residues are within TM helices.

www.psipred. net

(127, 128)

MINNOU

Predicts a-helical and b-sheet TM domains. Relies on structural profiles that consist of predicted relative solvent accessibility and secondary structure of each amino acid residue.

minnou. cchmc.org/

(129)

Philius 3.0

Philius is an updated version of HMM Phobius and combines a signal peptide submodel with a TM submodel. Philius uses a two-stage dynamic Bayesian network decoder.

www.yeastrc. org/philius

(130)

bioinformatics. biol.uoa.gr/ HMM-TM

Ref.

(continued )

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Table 3 (continued) Program

Description

URL (http://…)

Ref.

Phobius

This HMM method predicts signal peptides and TM a-helices. The output provides a list of the locations of the predicted TM helices, the predicted location of the intervening loop regions (cy, non-cy), and whether a signal peptide is present.

phobius.sbc. su.se/

(131, 132)

SCAMPI

SCAMPI allows for requests of up to 100,000 protein sequences, and is appropriate for large benchmark sets and full proteome scans.

scampi.cbr. su.se/

(133)

SOSUI 1.11

bp.nuap. SOSUI predicts inner membrane proteins from nagoya-u. amino acid sequences. The parameters used for ac.jp/sosui/ the prediction are based on the physico-chemical properties of the amino acid residues. Output includes details of each TM region, a hydropathy plot, and a helical wheel diagram of the predicted segments.

(134–136)

SPOCTOPUS

A method for combined prediction of signal peptides octopus.cbr. su.se/ and membrane protein topology, suitable for genome-scale studies of TM proteins. It consists of an NN and a dynamic programming algorithm based on a HMM.

(137)

TMHMM 2.0

A global approach that implements circular HMMs. www.cbs.dtu.dk/ (88) services/ Output shows number of predicted TM helices TMHMM/ and length of each TM segment. Orientation of protein in membrane is shown in posterior ­probability plot.

TMPred

www.ch.embnet. (138) Predictor of TM regions and their orientation. org/software/ TMPred investigates local properties of amino TMPRED_ acid sequences and uses a sliding window form.html approach. The algorithm is based on the statistical analysis of TMbase (database of naturally occurring TM proteins).

HMM hidden Markov model, NN neural network, TM transmembrane.

membranes of bacteria, mitochondria, and chloroplasts. TM b-barrels are more difficult to predict compared to a-helices. A selection of programs is listed in Table 4. 1.5.5. Integrated Methods for Predicting Suborganellar Localisation

The Plant Proteome DataBase (PPDB) (92) is a useful database that contains data for subcellular locations of Arabidopsis thaliana and Zea mays proteins. Initially, PPDB was dedicated to plant plastids, but has now expanded to the whole plant proteome. The PPDB stores experimental data from in-house proteome and

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Table 4 Software for predicting transmembrane b-barrels and membrane topology Program

Description

URL (http://…)

Ref.

B2TMR

This predictor locates TM spanning segments believed to be inserted into the outer membrane. There are two different predictors: the HMM predictor should be used when ­discrimination between b-barrels and other proteins is needed. The HMM model can predict the topology of b-barrel proteins using, as input, evolutionary information.

www.gpcr.biocomp. unibo.it/ predictors/

(139, 140)

BOMP

www.services.cbu. This program predicts whether or not uib.no/tools/ a polypeptide sequence from a Grambomp negative bacterium is an integral b-barrel outer membrane protein. It is based on recognising a C-terminal pattern typical of many integral b-barrel proteins and calculates an integral b-barrel score of the sequence.

Freeman–Wimley b-Barrel Analyzer

Predictor based on the physico-chemical properties of experimentally characterised TM b-barrel structures. The method reduces the large rate of false-positive predictions, and authors claim that the Freeman–Wimley algorithm has an accuracy of 99% when using the most efficient ­prediction criteria.

TMB-Hunt

www.bmbpcu36. This program uses a modified k-nearest leeds.ac.uk/~andy/ neighbour algorithm to predict protein betaBarrel/ sequences as TM b-barrel or non-TM AACompPred/ b-barrel on the basis of whole sequence aaTMB_Hunt.cgi amino acid composition. Composition profiles can either be calculated using single sequences or with evolutionary information, where composition is calculated using the query sequence and its close homologues.

(143)

TMBETAGENOME

A database for annotated b-barrel TM proteins in genomic sequences using different discrimination algorithms. Statistical algorithms based on amino acid composition, reside pair preference, and motifs are used, as well as machine-learning approaches based on amino acid and dipeptide composition.

(144)

www.tulane. edu/~biochem/ WW/apps.html

www.tmbeta-genome. cbrc.jp/annotation/

(141)

(142)

(continued )

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Table 4 (continued) Program

Description

URL (http://…)

Ref.

transFold

This web server predicts secondary structure and TM b-barrel topology and additionally predicts the sidechain orientation of TM b-strand residues, inter-strand residue contacts, and TM b-strand inclination with respect to the membrane. Instead of using machine-learning methods, such as HMM or NN, transFold employs multi-tape S-attribute grammars to predict the supersecondary structure of TM b-barrels.

www.bioinformatics. bc.edu/clotelab/ transFold/

(145)

HMM hidden Markov model, NN neural network, TM transmembrane.

mass spectrometry analysis, as well as curated information about protein function, protein properties, and subcellular localisation. Importantly, proteins are particularly curated for suborganellar plastid location and function, and this involves thylakoid membrane and envelope membrane proteins along with the proteins’ orientation in the membrane. ARAMEMNON 6.2 (93) is a database that was designed to aid in the interpretation of sequence data by integrating features that are presently only available from individual sources. This database has specialised on identifying putative integral membrane proteins of Arabidopsis thaliana, rice (Oryza sativa), grape (Vitis vinifera), maize (Zea mays), and about 4,000 putative membrane proteins of about 300 other seed plants. The database uses up to 24 individual prediction programs that contribute to the identification of putative TM a-helices and b-barrels. In addition, up to 17 individual prediction programs and experimental data contribute to the prediction of the subcellular location to four compartments (chloroplast, mitochondrion, secretory pathway, and “other”). A detailed description of how to use ARAMEMNON 6.2 is given in Subheading 3.5.

2. Materials 2.1. Equipment

A computer is required with access to the Internet and a Web browser.

2.2. Data Preparation

The input protein sequences should be written in the standard one-letter code with the following legitimate characters: ABCDEFGHIKLMNPQRSTUVWY and X for unknown. Spaces

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and line breaks will be ignored by all programs and will not interfere with the prediction. On the prediction servers described here, the sequences can be entered into an input window either by copy/ paste commands or by uploading a plain text file containing the sequence(s). Most prediction programs take, as input, sequences in FASTA format, which features a preceding definition line beginning with the “>” symbol. 2.3. Programs

Prediction programs and databases described in this chapter and their corresponding Web addresses are listed in Tables 1–4.

3. Methods A wide variety of programs predicting the subcellular location of proteins have proven to be useful, and some, such as TargetP and Predotar, are both widely known and used by the research community. However, all of the prediction programs have their individual strengths and weaknesses. High accuracy methods such as those based on homology and sequence motifs have low coverage and can only provide annotations for less than one-third of known sequences (94). However, these methods will become increasingly important in the future, as more high-quality protein localisation data become available and annotations improve. Prediction methods using the N-terminal targeting sequence have been shown to be quite successful in identifying subcellular localisation, particularly for chloroplast proteins. They have the advantage of being applicable to any protein sequence, whatever its similarity to previously studied proteins. When the protein sequence to be screened is only partial and might not include the N-terminal targeting region, global properties-based methods are the only alternative. It is recommended to use a mixture of different tools as this provides the most thorough prediction possible, but to take carefully into account the strengths and weaknesses of each. To determine correctly which predictor is best to use for a particular location and to draw reliable conclusions from a prediction, it is essential to be able to estimate the accuracy of the method properly. Ideally, comparative performance tests would be available, but in practice, it is impossible to do this in an unbiased way, because of the difficulty in avoiding contamination of the test data by data used to train the machine-learning algorithms used in most of the predictors. Equally, users should be wary of the claimed success rates of the various programs, which are almost always overestimates when compared with experimental localisation data (72). Generally, the performance of the predictors is assessed using the false-negative rate (also called “sensitivity”) and the false-positive rate (also called “specificity”). Users should have a clear idea of

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which measure is most important to them for the project that they want the predictions for, and which prediction programs perform best assessed by this criterion. These choices may be clearer with a couple of examples. Imagine a list of putative chloroplast proteins generated by subcellular fractionation and proteomics. It is likely that a small fraction of the proteins are contaminants, but most are probably truly chloroplast proteins. If one wanted to verify the list with a prediction program, one would probably choose one with a high sensitivity (such as ChloroP 1.1 (68)) to avoid erroneously throwing out many correctly localised proteins. On the contrary, if one were starting with a whole genome sequence and looking to generate a list of probable chloroplast-targeted proteins for functional genomics studies, then one would choose a predictor with a very low false-positive rate (such as Predotar 1.03 (71)) to avoid including many incorrect targets. To some extent, false-negative or false-positive rates can be minimised by adjusting the prediction cut-offs (the score at which a prediction is considered significant). This is discussed in the various program protocols below. Falsepositive rates can also be minimised by combining results from several different predictors. Such a combination of various predictors is precisely what integrators, such as SUBA II (82), PPDB (92), AT_CHLORO (95), and ARAMEMNON (93), are providing. They were designed to overcome the individual weaknesses of various types of predictors by uniting their predictions and including experimental data as location evidence. Hence, they integrate features from many individual sources. Integrators are superseding the use of individual predictors and are the tools we recommend when investigating the subcellular location of a protein. In this chapter, we describe two widely used predictors, TargetP (69) (in combination with ChloroP (68)) and Predotar (71), and the two integrators SUBA II (82) and ARAMEMNON (93) in detail. 3.1. Protocol for TargetP 1.1

TargetP (69) assigns proteins to four different locations: chloroplasts, mitochondria, the secretory pathway, and “all other compartments”. If a protein is predicted to be targeted to the chloroplast, TargetP provides a prediction of the cTP. 1. Go to TargetP 1.1 (http//www.cbs.dtu.dk/services/TargetP/). TargetP will predict the subcellular location of eukaryotic proteins. This location assignment is based on the predicted presence of an N-terminal sorting signal: cTP, mTP, SP, or other (no sorting signal). 2. Choose a sequence. Paste a single amino acid sequence or ­several sequences in FASTA format into the field provided (see Subheading 2.2). It is also possible to upload a sequence file. Make sure the sequence(s) contain the 130 N-terminal

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amino acid residues (see Note 1), as shorter sequences make the prediction less reliable. 3. Customise TargetP. Under “Organism group” select “Plant”. To obtain a prediction of the cleavage sites of the predicted pre-sequences, select “Perform cleavage site predictions” under “Prediction scope” (see Note 2). Cut-offs for the prediction can also be chosen. The default (“no cut-offs”) is recommended, where the highest score determines the prediction without this score having to be above a particular value. Predefined settings can be chosen that correspond to a specificity of 0.90 and 0.95, or cut-offs can be defined for a particular pre-sequence. Keep in mind that the predicted score is required to be both the highest and above the defined cut-off in order for a prediction to be made. 4. Run TargetP. Select “Submit” to initiate TargetP prediction. 5. Interpret TargetP results. For every submitted protein, TargetP reports the length of the submitted sequence (under “Len”) and a prediction score for each of the four possible outcomes (cTP, mTP, SP, and other). The predicted pre-sequence appears in the “Loc” column: “C” for chloroplast, “M” for mitochondrion, “S” for secretory signal peptide, or “_” for other. If a cut-off was defined for a particular pre-sequence and the highest score is below this cut-off, the program will not assign any targeting sequence and the output will be “*” for “not known”. TargetP gives a reliability coefficient (“RC”), which is based on the scores and assesses how confident TargetP is in each prediction. The RC ranges from 1 (very reliable prediction) to 5 (not reliable). If cleavage site prediction was selected, the predicted length of the cTP is indicated in the “TPlen” column. It is highly recommended to complement a sequence-based prediction of subcellular localisation by a database search (see Note 3). 3.2. Protocol for ChloroP 1.1

When TargetP predicts a cTP, a more detailed report of the cTP scores can be obtained from ChloroP (68). This is recommended, as it will aid in a higher confidence level of both the cTP and cleavage site prediction results. 1. Go to ChloroP 1.1 (http://www.cbs.dtu.dk/services/ ChloroP/). ChloroP will provide a detailed report of the scores of a predicted cTP. Only use ChloroP with sequences from plant or algal proteins. Submit a sequence by copy/paste or upload a sequence file as described in Subheading 2.2. Select “Detailed output” for a detailed score report and “Submit” to initiate the search.

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2. Interpret ChloroP results. As in TargetP, the length of the ­submitted sequence is reported. Next to it a “Score”, ranging from 0.4 to 0.6. The cut-off for the prediction of a cTP is 0.5. Values above this cut-off will show a “Y” in the cTP column, values below 0.5 will show an “N”. Any score above 0.55 indicates a fairly strong prediction. The “CS-score” is the maximum score of a weight matrix used to predict the cleavage site, and is used to decide the length of the cTP (“cTP-length”). See Note 4 for a description of the detailed output. To examine the suborganellar location of a protein and to investigate whether a protein contains a thylakoid lTP in addition to the predicted cTP, one of the following methods can be used: SignalP 3.0 (77) (see Note 5), LumenP (79), PredSL (73), or TatP 1.0 (81) (Table 1). The latter can only be used if the cTP containing proteins are imported across the thylakoid membranes via the Tat pathway. Alternatively, the PPDB (92) or AT_CHLORO (95) can be searched to identify a suborganellar location (Table 2). 3.3. Protocol for Predotar 1.03

For high-throughput analysis, Predotar (71) can be used to screen whole genomes/proteomes systematically. It has a very low rate of false positives compared with similar programs (including TargetP (69)), but with a concomitant loss of sensitivity at default settings (i.e., a higher number of false negatives) (71, 72). 1. Go to Predotar 1.03 (http://urgi.versailles.inra.fr/predotar/ predotar.html). Predotar will assign a location (mitochondrion, plastid, ER, or “elsewhere”) based on the identification of a putative N-terminal sorting signal. 2. Choose a sequence. A name for the sequence query can be entered in the box provided. Paste the sequence(s) in FASTA format into the field provided (see Subheading 2.2). Make sure the sequence(s) contain the N-terminal methionine followed by at least 60 amino acid residues (see Note 6). If required, selecting ”Reset” will clear any input data and a fresh search can be started. 3. Run Predotar. Select “plant sequences”, which includes the possibility of plastid targeting, and “Submit Query” to initiate the prediction by Predotar. 4. Interpret Predotar results. For every protein sequence, Predotar provides probability scores for each of the four locations (mitochondrion, plastid, ER, or “elsewhere”). These output scores are values between 0 (low probability) and 1 (high probability) to indicate the likelihood of a protein location to a particular compartment. The predicted location of the protein is reported in the column “Prediction”. A “plastid” score of over 0.5 is likely to result in a firm prediction of plastid targeting (unless other columns also have higher or similar scores); a plastid

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score of 0.25 or greater is likely to result in a prediction of “probably plastid” (again, unless other columns also have higher or similar scores). For whole-proteome screens, the 0.5 cut-off is recommended to keep the false-positive rate low. The 0.25 cut-off gives approximately similar sensitivity/specificity to the default settings for TargetP. 3.4. Protocol for SUBA II

SUBA II (1, 82) represents an integrator for subcellular localisation by uniting protein localisation information from ten different predictors as well as experimental data from proteomic studies (mass spectrometry), chimaeric fusion studies (GFP tagging), and annotations from Uni-ProtKB/Swiss-Prot (83, 84), AmiGO (85), and TAIR (86). SUBA II assigns 13 subcellular locations, including plastid. 1. Go to SUBA II (http://suba.plantenergy.uwa.edu.au/). Select the “SEARCH” tab from the four tabs (SEARCH, RESULT, ABOUT, and HELP) available at the top of the page. The phrase “Search for Arabidopsis proteins where…” displayed at the top, represents the beginning of the Boolean sentence that will form the query. Each row of pull-down menus works in combination to construct a query and each combination can be added to the main query pane by selecting the “Add” button at the end of the row (Fig. 1). 2. Search SUBA: particular protein. The subcellular location of proteins can be investigated through the “Arabidopsis Gene Identifier (is in list/is not in list)” menu row (last row of pulldown menus; Fig. 1). To define the subcellular location of particular protein(s), keep the pull-down menu as “is in list” and enter the Arabidopsis Gene Identifier (AGI) number(s) in the box below (see Note 7). Select the “Add” button at the end of the row. The built query sentence will appear in the top window under “Search for Arabidopsis proteins where…” with the statement “Arabidopsis Gene Identifier is in list (AGI numbers entered)” (Fig. 1). 3. Search SUBA: simple query. The first row of pull-down menus provides access to the majority of experimental localisation data. Using the three pull-down menus, a query can be built to investigate for example plastid-localised proteins obtained by mass spectrometry. To access such a set of proteins, keep the first pull-down menu (Experimentally observed location) as “is”. In the second menu (inferred by), select the option “MS/ MS assay”. The third pull-down menu (to be in) contains the subcellular location tags, select “plastid”. Then select the “Add” button at the end of the row. The built query sentence in the top window will state: “Protein location is inferred by MS/MS assay to be in plastid”.

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Fig. 1. A screenshot of the SUBA II interface, Search page. Arabidopsis Gene Identifier (AGI) numbers were used to search SUBA II. After selecting the “Add” button at the end of the row, the Boolean sentence (on top) reads: Search for Arabidopsis proteins where Arabidopsis Gene Identifier is in list (AT1G15510.1,AT1G47580.1,…). If not specified, the software will choose the first gene model by default and add “.1” to the AGI number (as shown).

4. Search SUBA: more complex query. Select the Boolean linkers “AND” or “OR” command buttons to build more complex queries. For example to search for plastid proteins identified through mass spectrometry and GFP analysis requires the extension of the simple query sentence from above by selecting the Boolean linker “AND”. The “Add” buttons at the end of each row become activated again. Keep the first pull-down menu (Experimentally observed location) as “is”. In the second menu (inferred by), select the option “GFP assay”. In the third pull-down menu (to be in), select “plastid”. The built query sentence in the top window will now state “Protein location is inferred by MS/MS assay to be in plastid AND Protein location is inferred by GFP assay to be in plastid”. The linker buttons “AND” or “OR” can also be used to include the following: (a) Pre-computed predictions of subcellular localisation (see Note 8) by accessing the data through the “Predicted

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location (is/is not) predicted by (predictor) to be in (location)” menu row; (b) Subcellular claims by different research papers using the “Literature referenced location (is/is not) described in (paper)” menu row; (c) Text-based searches of descriptor fields through the “Protein description (contains/does not contain) keyword (e.g., protein kinase)” menu row; and (d) Physico-chemical parameters such as molecular weight, expressed sequence tag (EST) number, chromosomal location using the “Physical properties” and “Underlying gene model (is on/is not on) chromosome (number)” menu rows. 5. Run SUBA. Select the “Submit” button at the bottom of the page (Fig. 1) to initiate the search. If required, use the “Undo” button to remove any mistakes from the forming query. This removes the last entered query. Use the “Clear” button to remove the entire constructed query in the top window for a fresh search. 6. View SUBA results. Once a query has been submitted, the contents of the “RESULT” tab will automatically be displayed (Fig. 2). By default, eight columns will be displayed: AGI number, TAIR description, location summaries of all predictors, location by mass spectrometry, location by GFP tagging, location by annotation (TAIR), location by AmiGO, and location by Uni-Prot/Swiss-Prot. Results can be sorted ascending or descending using the function menu, which is activated by tracking the mouse over the column header and then selecting the emerging arrow. In addition, new columns can be added to the RESULT window by tracking the mouse over “Columns” in the function menu and selecting the required columns. Columns can then be organised using the mouse drag and drop functionality. Fifty rows of data are displayed, with further rows available using the “Next Page” or “Last Page” buttons at the bottom left of the RESULT window (see Note 9; Fig. 2). The codes or unique identifiers (uid) beside each location are links to the primary data source for each entry at PubMed or ISI Web of Knowledge. Selecting such a link will open another window in the browser and display the abstract in PubMed or ISI Web of Knowledge. Each AGI number provides a link to a summary page for the entry. This SUBA flatfile contains a detailed breakdown of subcellular localisation information (experimental and predicted), description and sequence information (TAIR9), physico-chemical characteristics, a hydropathy plot, and several links to the same entry in other Arabidopsis databases.

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Fig. 2. A screenshot of the SUBA II interface, Result page. The default result page shows eight columns. Each AGI number represents a link to the flatfile of a particular locus giving a detailed description of subcellular localisation information. The hyperlinks beside each location represent unique identifiers (UID) and are links to the abstract in PubMed or ISI Web of Knowledge. The “Download All Results” button at the top left of the RESULT window will download the results (including the columns not displayed) as a tab-delimited file and can be opened in Microsoft Excel.

7. Download SUBA results. All results (including the columns not displayed) can be downloaded as a tab-delimited file using the “Download All Results” button at the top left of the RESULT window (Fig.  2). By default, this file opens in Microsoft Excel. 3.5. Protocol for ARAMEMNON 6.2

ARAMEMNON 6.2 (93) is specialised for identifying putative integral membrane proteins in plants, including Arabidopsis thaliana. To examine whether a chloroplast protein is membrane bound, the database can be searched for the presence of putative a-helical TM segments, N-terminal TM anchors, and b-barrel TM segments. Respectively, up to 18 and up to six individual prediction programs contribute to the identification of TM a-helices and b-barrels. In addition, three programs predict the presence of putative N-terminal TM anchors. 1. Go to ARAMEMNON 6.2 (http://aramemnon.botanik. uni-koeln.de/ or http://aramemnon.uni-koeln.de/). Select the “Search” tab to query the database. A search for membrane proteins in nuclear or organellar genomes of Arabidopsis thaliana (and other seed plants) can be carried out in three different

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Fig. 3. A screenshot of the ARAMEMNON interface, Search page. Search by sequence (main window ) or keyword (top right ). Databases of “all” plants will be searched when submitting a protein sequence (displayed here). Begin the search by selecting the loupe symbol (bottom right ). One or more keywords can be entered into the search box at the top right corner (AGI number entered here). Use the pull-down tab to the left to select all seed plants or one of four plants (Arabidopsis thaliana selected here). To start the search, select the “go” button to the right of the search box.

ways: submitting a sequence (Step 2), entering keywords (Step 3), or selecting properties (Step 4) (Fig. 3). 2. Search by sequence. To search for nucleus- and organellarencoded putative membrane proteins, a DNA or protein sequence can be entered in the provided “Sequence” box using the copy/paste command (Fig.  3). When submitting a protein sequence, databases of “all” plants will be searched (see Note 10). When submitting a nucleotide sequence, the search is restricted to one of four plants, including Arabidopsis thaliana (see Note 10). The search is based on sequence alignments performed by Basic Local Alignment Search Tool (BLAST) 2.2. A BLAST option “low complexity filter on” is selected by default. Initiate the search by selecting the loupe symbol at the bottom right of the search window (Fig. 3). The related sequences will be displayed together with a link to the BLAST alignment details. 3. Search by keyword. One or more keywords can be entered into the search box at the top right corner (Fig. 3). The pulldown tab provides access to information on all seed plants in the database or on one of four plants (Arabidopsis thaliana, Oryza sativa, Vitis vinifera, or Zea mays). Single keywords (such as AGI number, functional description, etc.) or multiple keywords can be used (Fig.  3). The latter can combine

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descriptions (i.e., transporter AND malate), use one of the keywords (i.e., transporter OR malate), or exclude one description from another (i.e., transporter NOT malate) (see Note 11). To initiate the search, select the “go” button next to the search box (Fig. 3). 4. Search by selecting properties. Under “Search by” click on “Properties”. This will allow searching by a combination of different parameters: plant species, type of membrane association (TM a-helices, TM b-barrel, or membrane anchored), and predicted subcellular location (chloroplast, mitochondrion, or secretory pathways). See Note 12 for further details. The search is initiated by selecting the loupe symbol at the bottom right of the search window. 5. ARAMEMNON results – External links. The first column (“External links”) on the result page lists the gene identifier number (i.e., AGI number) and the plant species (i.e., Arabidopsis thaliana), respectively linking to the same entry at TAIR or the NCBI Taxonomy Browser (Fig. 4). 6. ARAMEMNON results – Expression. For most Arabidopsis thaliana genes, coexpression/expression data are available and links to the same entry at the Arabidopsis eFP browser, Genevestigator, and ATTED-II coexpression networks exist in the “Expression” column (Fig. 4). 7. ARAMEMNON results – Description/Gene Ontologies. A description that also represents a link to bibliographic references (if available), a transporter classification that links to the family in the Transporter Classification Database (if applicable),

Fig. 4. A screenshot of the ARAMEMNON interface, Result page. The result page following a keyword search using an AGI number (as shown here), displays various links to external databases containing coexpression/expression data, bibliographic references, transporter classifications, and links to gene ontology, sequence, topology, and cluster data. Most relevant for the identification of membrane proteins is the topology link providing data on transmembrane spans, subcellular locations, and lipid modifications.

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and a link to gene ontology data of protein function (see Note 13) are displayed under “Description/Gene Ontologies (GO)” (Fig. 4). 8. ARAMEMNON results – Sequence. Selecting the link in the “Sequence” column will access protein, cDNA, and genomic sequence data with exon/intron predictions (Fig.  4). The details of this section are beyond the scope of this chapter and do not aid in identifying membrane-bound proteins. 9. ARAMEMNON results – Topology. The link in the “Topology” column (Fig. 4) provides data on transmembrane spans (Fig. 5), subcellular locations, and lipid modifications. The “overview” section displays the length of the protein sequence, its molecular weight, and the consensus of both the subcellular location and the TM a-helical segments. Up to 18 individual programs contribute to the prediction of TM a-helices (Fig. 5; Table 3).

Fig. 5. A screenshot of the ARAMEMNON interface, Result page, link Topology, section transmembrane spans. The diagram shows the position of the predicted TM a-helical segments on the protein and indicates average hydrophobicities. The numbers of TM segments are indicated on the right of the diagrams with the N-terminus orientation of the protein next to it. A maximum of three consensus diagrams (“TmConsens”, “ConPred v2”, “TmMultiCon”) are displayed below the individual predictor diagrams and a consensus score is reported below each TM segment. The prediction results for an N-terminal TM anchor and for b-barrel TM segments are also displayed. The confidence level of each prediction is indicated by different colour intensities.

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A diagram in the section “transmembrane spans” displays the position of the predicted TM a-helical segments on the protein and indicates average hydrophobicities by different colour intensities (Fig. 5). A detailed output of the TM a-helices predicted by each program can be viewed by selecting an individual protein prediction line from the diagram (see Note 14). The individual predictions for each protein are combined to a built-in consensus prediction (“TmConsens”) using a Bayes method. The consensus diagram is displayed below the individual predictor diagrams and a consensus score is reported below each TM segment (Fig. 5). TM segments are shown if the score is above 0.10 and counted if this score is equal to or above 0.42 (see Note 15). An additional consensus prediction (performed by ConPred 2) is available for many proteins. Notice that ConPred 2 uses a different set of individual predictions. A built-in extended consensus prediction is performed by combining consensus predictions of several homologous proteins, resulting in the consensus “TmMultiCon”. The consensus diagram for “TmMultiCon” shows TM segments with a score above 0.10 and counts all with a score equal to or above 0.38. The counts of TM segments are indicated on the right of the diagrams (Fig. 5). The orientation of the protein N-terminus is reported next to the counts of the TM segments (Fig.  5). For the orientation of plasma membrane proteins, “N-in” indicates a cytosolic location of the N-terminus, and “N-out” indicates a non-cytosolic location. For the orientation of membrane proteins in chloroplasts, the definition uses the stroma as reference point instead of the cytosol. Thus, the topology of chloroplast membrane proteins is as follows. For outer envelope proteins, “N-in” indicates a location of the N-terminus in the cytosol, and “N-out” indicates intermembrane space. For inner envelope proteins, “N-in” indicates a location of the N-terminus in the stroma, and “N-out” indicates intermembrane space. For thylakoid membrane proteins, “N-in” indicates a location of the N-terminus in the stroma, and “N-out” indicates thylakoid lumen. Such topology predictions made for chloroplast membrane proteins are not as reliable as those made for plasma membrane proteins. If the cleavage site of the cTP is known, we recommend using the protein sequence without its cTP, as this is likely to yield a more reliable prediction result. Below the diagrams, presence of an N-terminal TM anchor is predicted (Fig.  5). Three prediction programs (Phobius, HMM-based SignalP 3.0, and SosuiSignal; Table 3) indicate whether an N-terminal TM segment (position <50 amino acids) could be responsible for the  membrane association of the protein (orange colour) or not (white colour). Up to six individual programs (BOMP, MCMBB, TMB-hunt, TMB-KNN, TmBetaDisc, and transFold) contribute to the

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prediction of TM b-barrels present in outer membrane proteins (Table  4), and the confidence level of the prediction is indicated by different colour intensities (Fig. 5). 10. ARAMEMNON results – Topology (subcellular location). In the “subcellular location” section (scroll down from what is seen in Fig. 5), the protein is predicted to be located in the chloroplast, mitochondrion, or secretory pathways. Up to 17 individual prediction programs contribute to the subcellular location prediction (Table 1). However, not all programs perform predictions for all three locations (indicated by a dash instead of a circle). The confidence levels of each prediction are indicated by different colour intensities. Similar to the “TmConsens”, the individual predictions for each protein are combined to a builtin consensus prediction using a Bayes method (see Note 16). If available, experimental evidence for the subcellular location is listed and a link to the publication is provided. 11. ARAMEMNON results – Topology (lipid modification). Several proteins are attached to a membrane by hydrophobic anchors, which are added by a post-translational lipid modification, specified in the “lipid modification” section (again, scroll down from what is seen in Fig. 5). However, such modifications are not relevant for membrane-bound chloroplast proteins and are thus not further discussed here. 12. ARAMEMNON results – Cluster. Selecting the link in the “Cluster” column (Fig. 4) will display an orthologue cluster of related plant protein sequences. The cluster view includes links to an ARAMEMNON result sheet (showing all the loci from this cluster from one particular species), a cladogram that includes all cluster member proteins, a list of isospecic homologues (paralogues), and a cladogram that includes paralogous protein sequences only.

4. Notes 4.1. TargetP

1. TargetP predictions are based on the 130 N-terminal residues of each input sequence. Therefore, it is strongly recommended to include the N-terminus for all sequences; missing N-terminal residues make the prediction more difficult and less reliable. Submitting more than 130 residues does not influence the ­prediction in any way (apart from making it slower). 2. ChloroP is automatically used to predict the cleavage sites of cTPs and the results are incorporated into the TargetP output. 3. A database search to complement a sequence-based prediction of subcellular localisation can be performed with BLAST (96)

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at NCBI (http://blast.ncbi.nlm.nih.gov/Blast.cgi). Choose “Arabidopsis thaliana” as the species genome to search and select “protein blast” (blastp) to search a protein database using a protein query. Enter the query sequence and choose a database, for example “Swissprot protein sequences (swiss-prot)”. If no annotated close homologues are found, search the entire protein sequence universe by choosing “Non-redundant protein sequences (nr)”. Then use the best hits and run their sequences through the predictors. If most of them yield the same predicted localisation as the query sequence, one can have more confidence in the prediction. 4.2. ChloroP

4. In the detailed output, three scores for each amino acid residue of the submitted sequence are presented: the “Raw” NN-score for each amino acid residue, a derivative of that score (“Deriv.”), which is used to define the region that the weight matrix is applied to and then search for the cleavage site, and the cleavage site score for each residue (“CS-score”). The cleavage site is defined to be directly N-terminal of the amino acid with the highest CS-score.

4.3. SignalP

5. SignalP can be used because the targeting pathways to the lumen have analogous systems in Gram-negative bacterial secretory pathways (78). Thus, the Gram-negative version of SignalP should be used. Under “Truncation”, make sure to adjust the “Truncate each sequence to max. 130 residues”, as 96% of the combined cTP and lTP signals lie within 130 amino acid residues (79). Before submitting a protein sequence to SignalP in order to identify an lTP, the cTP part of the sequence could be removed. However, the cTP cleavage site prediction is relatively unreliable, and instead of just removing the predicted cTP, the submission of several truncated versions of the protein is recommended (i.e., remove the N-terminal residues in steps of five). This progressive truncation method could also be useful when submitting protein sequences to LumenP and TatP. PredSL, however, offers an integrated prediction for the lTP.

4.4. Predotar

6. Predotar recognises the N-terminal targeting sequences of classically targeted precursor proteins. It can neither recognise targeted proteins from internal sequences, nor can it work with short N-terminal segments. Thus, it will not give a result if the submitted sequence lacks an N-terminal methionine or if the sequence is shorter than 60 amino acids.

4.5. SUBA II

7. If the gene model is not specified (i.e., At1g23450.2), by default the SUBA software will chose the first gene model and add “.1” at the end of the AGI number (i.e., At1g23450.1). 8. For the pre-computed predictions of subcellular localisations, the protein sequences from TAIR9 of all 33,518 Arabidopsis proteins were run through the ten individual prediction programs that are reported in SUBA II.

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9. If no scroll bars are visible on the right side of the RESULT window and only a part of the results can be viewed, the page needs to be reloaded. Simply select the “Next Page” button at the bottom left of the window. If less than 50 rows of data are reported, highlight the page number “1” and replace it with any other number (1 or 2 or 3, it does not matter). Press “Enter” on your keyboard, the page will reload and a scroll bar should appear on the right side of the window. 4.6. ARAMEMNON

10. When submitting a protein sequence to ARAMEMNON, “all” databases need to be selected; otherwise related sequence(s) will not be displayed. When submitting a nucleotide sequence, one of the organism databases (Arabidopsis thaliana, Oryza sativa, Vitis vinifera, or Zea mays) needs to be selected; otherwise related sequence(s) will not be displayed. 11. The search by multiple keywords is performed by a search engine where the index created for that search engine contains data about protein description terms, ontology terms, PFAM domain description terms, and plant common and scientific names as well as plant taxonomic order and family information. 12. Searching by “Properties” allows selecting for proteins that belong to a certain plant species, have a specific membrane association, and/or a predicted subcellular location. “All seed plants” or one of eleven plant species (Arabidopsis thaliana, Oryza sativa, Vitis vinifera, Zea mays, Triticum aestivum, Nicotiana tabacum, Medicago truncatula, Solanum lycopersicon, Solanum tuberosum, Hordeum vulgare, or Glycine max) or “other seed plants” (except for the eleven listed) can be selected using the pull-down menu. The membrane association can either be “not specified” or specified by choosing TM spans (TM a-helices and TM b-barrels) or post-translational lipid modifications (“glycosyl phosphatidylinositol attachment”, “prenylation”, “myristoylation”, or “palmitoylation sites”). Similarly, the predicted subcellular location can either be “not specified” or limited to “chloroplast”, “mitochondrion” or “secretory pathways”. A “strict” or “tolerant prediction score” can be selected for each location. 13. When clicking on the “GO”: button under “Description/ Gene Ontologies (GO)”, three ontology terms according to the Gene Ontology Consortium are displayed: (1) biological process to which the protein contributes, (2) molecular function that the protein performs, and (3) cellular component where the protein is active. 14. The details of the predicted TM a-helices include positions on protein, mean hydrophobicities, maximal amphiphilicities, and hydropathy profile. The position data are taken from the output of the TM prediction programs. They are highlighted in the protein sequence and in the hydropathy profile. The beginning

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and end positions of each TM a-helix are listed next to their mean hydrophobicities and maximal amphiphilicities. 15. The consensus scores are based on the ratio of the posterior probabilities of both hypotheses, i.e., that a certain sequence is a TM region given that the program predicts positively, and that a certain sequence is not a TM region although the program predicts positively. The corresponding likelihoods of the hypotheses were estimated using a set of training proteins with known TM segments. The consensus score value is normalised to a maximal value of 1. (Note that a score value of 1 does not reflect a probability of 1.) The centre and edge positions for each consensus TM segment are calculated by the average positions. Extra procedures are added to avoid overlapping or unreasonably short consensus segments. 16. The consensus score is based on the ratio of the posterior probabilities of both hypotheses, i.e., that a protein is targeted and that it is not targeted given positive prediction results. The corresponding likelihoods of the hypotheses were estimated using a set of training plant proteins with known subcellular location (97). References 1. Heazlewood, J. L., Tonti-Filippini, J., Verboom, R. E., and Millar, A. H. (2005) Combining experimental and predicted datasets for determination of the subcellular location of proteins in Arabidopsis. Plant Physiol. 139, 598–609. 2. Soll, J., and Schleiff, E. (2004) Protein import into chloroplasts. Nat. Rev. Mol. Cell Biol. 5, 198–208. 3. Hörmann, F., Soll, J., and Bölter, B. (2007) The chloroplast protein import machinery: a review. Methods Mol. Biol. 390, 179–193. 4. Inaba, T., and Schnell, D. J. (2008) Protein trafficking to plastids: one theme, many variations. Biochem. J. 413, 15–28. 5. Jarvis, P. (2008) Targeting of nucleus-encoded proteins to chloroplasts in plants. New Phytol. 179, 257–285. 6. Li, H. M., and Chiu, C. C. (2010) Protein transport into chloroplasts. Annu. Rev. Plant Biol. 61, 157–180. 7. Gould, S. B., Waller, R. F., and McFadden, G. I. (2008) Plastid evolution. Annu. Rev. Plant Biol. 59, 491–517. 8. Martin, W., Rujan, T., Richly, E., Hansen, A., Cornelsen, S., Lins, T., Leister, D., Stoebe, B., Hasegawa, M., and Penny, D. (2002) Evolutionary analysis of Arabidopsis, cyanobacterial, and chloroplast genomes reveals plastid

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39. Bonen, L., and Doolittle, W. F. (1975) On the prokaryotic nature of red algal chloroplasts. Proc. Natl. Acad. Sci. USA 72, 2310–2314. 40. Moreira, D., Le Guyader, H., and Philippe, H. (2000) The origin of red algae and the evolution of chloroplasts. Nature 405, 69–72. 41. Aldridge, C., Cain, P., and Robinson, C. (2009) Protein transport in organelles: Protein transport into and across the thylakoid membrane. FEBS J. 276, 1177–1186. 42. Robinson, C., Thompson, S. J., and Woolhead, C. (2001) Multiple pathways used for the targeting of thylakoid proteins in chloroplasts. Traffic 2, 245–251. 43. Shackleton, J. B., and Robinson, C. (1991) Transport of proteins into chloroplasts. The thylakoidal processing peptidase is a signaltype peptidase with stringent substrate requirements at the −3 and −1 positions. J. Biol. Chem. 266, 12152–12156. 44. von Heijne, G. (1990) The signal peptide. J. Membr. Biol. 115, 195–201. 45. Yuan, J., Henry, R., McCaffery, M., and Cline, K. (1994) SecA homolog in protein transport within chloroplasts: evidence for endosymbiont-derived sorting. Science 266, 796–798. 46. Schuenemann, D., Amin, P., Hartmann, E., and Hoffman, N. E. (1999) Chloroplast SecY is complexed to SecE and involved in the translocation of the 33-kDa but not the 23-kDa subunit of the oxygen-evolving complex. J. Biol. Chem. 274, 12177–12182. 47. Dalbey, R. E., and Chen, M. (2004) Sectranslocase mediated membrane protein biogenesis. Biochim. Biophys. Acta 1694, 37–53. 48. Mould, R. M., and Robinson, C. (1991) A proton gradient is required for the transport of two lumenal oxygen-evolving proteins across the thylakoid membrane. J. Biol. Chem. 266, 12189–12193. 49. Cline, K., Ettinger, W. F., and Theg, S. M. (1992) Protein-specific energy requirements for protein transport across or into thylakoid membranes. Two lumenal proteins are transported in the absence of ATP. J. Biol. Chem. 267, 2688–2696. 50. Chaddock, A. M., Mant, A., Karnauchov, I., Brink, S., Herrmann, R. G., Klosgen, R. B., and Robinson, C. (1995) A new type of signal peptide: central role of a twin-arginine motif in transfer signals for the delta pH-dependent thylakoidal protein translocase. EMBO J. 14, 2715–2722. 51. Creighton, A. M., Hulford, A., Mant, A., Robinson, D., and Robinson, C. (1995) A monomeric, tightly folded stromal intermediate on the delta pH-dependent thylakoidal

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99. Jin, Y. H., Niu, B., Feng, K. Y., Lu, W. C., Cai, Y. D., and Li, G. Z. (2008) Predicting subcellular localization with AdaBoost Learner. Protein Pept. Lett. 15, 286–289. 100. Mitschke, J., Fuss, J., Blum, T., Hoglund, A., Reski, R., Kohlbacher, O., and Rensing, S. A. (2009) Prediction of dual protein targeting to plant organelles. New Phytol. 183, 224–235. 101. Pierleoni, A., Martelli, P. L., Fariselli, P., and Casadio, R. (2006) BaCelLo: a balanced subcellular localization predictor. Bioinformatics 22, e408-416. 102. Brady, S., and Shatkay, H. (2008) EpiLoc: a (working) text-based system for predicting protein subcellular location. Pac. Symp. Biocomput. 13, 604–615. 103. Nair, R., and Rost, B. (2005) Mimicking cellular sorting improves prediction of subcellular localization. J. Mol. Biol. 348, 85–100. 104. Blum, T., Briesemeister, S., and Kohlbacher, O. (2009) MultiLoc2: integrating phylogeny and Gene Ontology terms improves subcellular protein localization prediction. BMC Bioinformatics 10, 274. 105. Schein, A. I., Kissinger, J. C., and Ungar, L. H. (2001) Chloroplast transit peptide prediction: a peek inside the black box. Nucleic Acids Res. 29, E82. 106. Chou, K. C., and Shen, H. B. (2007) Largescale plant protein subcellular location prediction. J. Cell. Biochem. 100, 665–678. 107. Chou, K. C., and Shen, H. B. (2008) CellPLoc: a package of web servers for predicting subcellular localization of proteins in various organisms. Nat. Protoc. 3, 153–162. 108. Chou, K. C., and Shen, H. B. (2010) PlantmPLoc: a top-down strategy to augment the power for predicting plant protein subcellular localization. PLoS One 5, e11335. 109. Huang, W. L., Tung, C. W., Ho, S. W., Hwang, S. F., and Ho, S. Y. (2008) ProLoc-GO: utilizing informative Gene Ontology terms for sequence-based prediction of protein subcellular localization. BMC Bioinformatics 9, 80. 110. Boden, M., and Hawkins, J. (2005) Prediction of subcellular localization using sequencebiased recurrent networks. Bioinformatics 21, 2279–2286. 111. Hawkins, J., and Boden, M. (2006) Detecting and sorting targeting peptides with neural networks and support vector machines. J. Bioinform. Comput. Biol. 4, 1–18. 112. Nakai, K., and Horton, P. (1999) PSORT: a program for detecting sorting signals in proteins and predicting their subcellular localization. Trends Biochem. Sci. 24, 34–36.

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Chapter 17 Rapid Isolation of Arabidopsis Chloroplasts and Their Use for In Vitro Protein Import Assays Henrik Aronsson and R. Paul Jarvis Abstract In vitro chloroplast protein import assays have been performed since the late 1970s, initially with plant species (e.g., pea and spinach) that readily provide an abundant source of starting material and also, subsequently, a good yield of chloroplasts for import assays. However, the sequencing of the Arabidopsis genome paved the way for an additional model system that is more amenable to genetic analysis, as a complement to the more biochemically orientated models such as pea and spinach. A prerequisite for this change was an efficient and reliable protocol for the isolation of chloroplasts for use in protein import assays, enabling biochemical approaches to be combined with the genetic potential of the plant. The method described here was developed as a rapid and low-cost procedure that can be accessed by everyone due to its simplicity. Despite its rapidity and simplicity, the method yields highly pure chloroplasts, and in addition works well with mutant plants that exhibit pale or chlorotic phenotypes. The protocol is also optimized for work with material from young plants (10–14 days old), when protein import is believed to be at its peak, and so plant growth can be conducted in vitro on Murashige and Skoog medium. The isolation method has been used not only for protein import assays, but also for proteomic analysis and further subfractionation studies. Key words: Arabidopsis, Chloroplast, Chloroplast isolation, In vitro translation, Plastid, Preprotein, Protein import, Protein targeting

1. Introduction In the early 1970s, Blobel and Sabatini proposed that proteins ­destined to the endoplasmic reticulum (ER) each carry an address tag, within the amino acid sequence, that directs the protein to the ER (1); this was known as the signal hypothesis. Soon afterward, the hypothesis was confirmed (2, 3), opening the way for speculation that similar targeting mechanisms exist for other cellular ­compartments. In plants, the chloroplast compartment is highly

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_17, © Springer Science+Business Media, LLC 2011

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interesting because these organelles are vital for photosynthesis, which provides the energy-rich molecules and oxygen that are essential for mankind. Chloroplasts also play many other roles, including N and S metabolism, and the production of amino acids and fatty acids (4). Today we know that of the many different proteins present in chloroplasts, roughly 95% (~3,000 proteins) are nucleus encoded (5) and must therefore be targeted from the cytosol, where they are made, to the chloroplast. While some nucleus-encoded chloroplast proteins do not have a defined cleavable targeting signal (6–8), and others reach the organelle via the ER and Golgi (9, 10), the majority have a cleavable N-terminal transit peptide that mediates posttranslational targeting directly into the organelle from the cytosol (11). Because of its posttranslational and direct nature, it is possible to reconstitute protein import in a test tube using a minimal system comprising isolated chloroplasts and precursor proteins (12, 13). The in vitro import into chloroplasts of proteins with a defined transit peptide (so-called preproteins) can provide much important information concerning chloroplast biogenesis. By manipulating the concentration of nucleotides in an import reaction, or by using chemical cross-linkers, one can obtain information on the energy requirements for import, or on the proteins that interact with preproteins during import (i.e., import apparatus components) (see Chapter 18, Vol. 1). By conducting post-import fractionation experiments, or by treating the chloroplasts with specific proteases, it is possible to probe the final destination and topology of a particular protein of interest (see Chapters 20 and 21, Vol. 1). Such in vitro assays can also be used to estimate protein import rates in mutant plants that lack putative components of the import machinery, thereby testing the importance of the missing component in the overall process (14, 15). When chloroplast protein import assays were first conducted in the late 1970s and 1980s, pea and spinach plants were mainly used. This was due to the ease with which one can get a lot of starting material and, subsequently, a good yield of intact, physiologically active chloroplasts for import assays with these species. However, in the modern era of plant molecular biology, completion of the genome sequencing of Arabidopsis thaliana has led to the adoption of Arabidopsis as an alternative and complementary model plant for import studies (16). In contrast to pea and spinach, however, Arabidopsis plants do not so readily provide a lot of leaf material in a short period of time, due to their diminutive stature. This issue has been overcome by taking advantage of the remarkable fecundity of Arabidopsis: since it is easy to obtain large quantities of seeds, it is possible to grow very large numbers of plants to generate the material needed. Protocols have been described for the isolation of Arabidopsis chloroplasts from protoplasts (17), and subsequently developed

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for use in protein import studies (18). However, the requirement for a protoplastation step means that such methods require a lot of time and also costly enzymes for cell wall degradation (18). On the other hand, the direct isolation of chloroplasts from leaves by homogenization is both fast and cheap, and Arabidopsis chloroplasts isolated in this way have been shown to be useful for protein import studies and other purposes (14). Our protocol typically employs plants grown on Murashige and Skoog (MS) medium plates, harvested after 10–14 days of growth. The short growth period is advantageous because chloroplasts in young leaves have a greater capacity to import proteins, due to the fact that the chloroplast interior is undergoing rapid development at this stage (19). This aspect is also useful when analyzing import mutants, which tend to have a more pronounced mutant phenotype during early development (16). The use of MS medium enables the provision of a carbon source (i.e., sucrose) which facilitates the growth of mutants with particularly severe phenotypes, which otherwise would be very difficult to grow (e.g., on soil) (20, 21). The chloroplast isolation method provided in this chapter (14) has been widely used, not only in import studies (15, 22–26) but also for proteomic work (22, 23, 27), subfractionation experiments (28), and a range of other analyses (29–32). An overview of the procedure is presented in Fig. 1.

2. Materials 2.1. Growth of Arabidopsis Plants

1. Arabidopsis thaliana seeds stored in 1.5-mL microfuge tubes at room temperature (18–23°C). 2. Sterilization solution 1: 70% (v/v) ethanol and 0.05% (v/v) Triton X-100. For 50 mL, mix 35 mL of 100% ethanol, 15 mL of deionized water, and 100 mL of Triton X-100. Store at room temperature. 3. Sterilization solution 2: 96–100% (v/v) ethanol. Store at room temperature. 4. Laminar flow hood. 5. Circular filter papers, 9 cm in diameter. 6. 70% (v/v) ethanol in a glass bottle. 7. Orbital platform shaker. 8. Plastic Petri plates, 9 cm in diameter. 9. Murashige and Skoog (MS) medium. For 1 L, use 4.41 g MS salt and vitamin mixture (Duchefa, Haarlem, the Netherlands), 0.5  g 2-(N-morpholino) ethane sulfonic acid (MES), 5  g sucrose (0.5% [w/v]), and 8 g agar (0.8% [w/v]). Adjust pH to 5.7 using KOH. Sterilize the medium in an autoclave

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Fig. 1. Flow chart for the chloroplast isolation and protein import procedures. The procedures described in this chapter are summarized schematically at the right of the figure. Important steps in the procedure are illustrated with photographs at the left side of the figure. The photos show, from top to bottom (1) a filter paper folded in half to create a crease, facilitating even seed sowing on MS plates. (2) Petri plates containing seeds spread inside a growth cabinet, making efficient use of the available space; the number of plates shown (39, with ~100–150 seeds per plate) provides sufficient material after 14 days growth for two chloroplast isolations (~20 plates each), each typically yielding 100–200 million chloroplasts which is enough for 10–20 import reactions (i.e., 20–40 reactions in total). (3) Homogenization of the harvested plant material using a 50-mL beaker, with plant material almost filling up the whole volume. (4) The equipment setup used for performing import reactions. The light source to the left powers photosynthesis in the organelles, the water-filled heat trap in the middle prevents overheating of the samples by the lamp, while the water bath to the right keeps the import reactions at an optimal temperature; import is performed using microfuge tubes inserted into a homemade metal holder, preventing the samples from getting flooded by water so that the tube lids may be left open during the procedure (which simplifies aliquot removal during time-course experiments).

(30 min, 121°C), cool to 50°C in a water bath, and pour ~20– 25 mL per Petri plate in a laminar flow hood; let the plates dry for 45–60 min in the hood before adding the lids. If not used directly, store the plates upside down and sealed in a clean plastic bag at 4°C (see Note 1). 10. Surgical tape (Micropore, 3M, St. Paul, MN, USA). 11. Cold room or refrigerator (4°C). 12. Plant growth chamber, with the following settings: 20°C, 120–180  mmol/m2/s white light, 16-h light/8-h dark (see Note 2).

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1. Template cDNA coding for a preprotein of interest. The ­precursor of the small subunit of Rubisco (pSS) is a commonly used substrate for import studies, and we routinely use RbcS ats1A from Arabidopsis (At1g67090, EST 188D4T7). The cDNA is cloned in a high-copy plasmid downstream of a T7 polymerase promoter, and the plasmid can be prepared using a standard plasmid preparation mini kit. 2. Universal M13 forward (5¢-TGT AAA ACG ACG GCC AGT3¢) and reverse (5¢-CAG GAA ACA GCT ATG ACC-3¢) primers for amplification of the template. Store at −20°C. 3. Freezers (−20°C and −80°C) for the storage of Escherichia coli stocks, cDNA templates, primers, and translation reagents and products. 4. Microfuge. 5. Polymerase chain reaction (PCR) reagents. Taq polymerase buffer (10×), deionized sterile water, 10 mM dNTP mixture (2.5 mM each), 10 mM of each universal M13 primer (or other appropriate primers), 1–100  ng cDNA template per 20  mL reaction, and 1 U/20 mL Taq polymerase. Store at −20°C. 6. Thermocycler. 7. TAE buffer. For 1 L, use 4.84 g Tris base, 1.09 g glacial acetic acid, and 0.292 g ethylenediaminetetraacetic acid (EDTA; free acid); the pH should be in the range of 8.18–8.29. 8. Agarose gel electrophoresis system. Gels should contain of 1% (w/v) agarose in TAE buffer. A microwave is useful for dissolving the agarose in TAE, and a UV light box is required for DNA detection following staining with ethidium bromide or similar. 9. Radiolabeled [35S] methionine, 11  mCi/mL (Perkin-Elmer, Norwalk, CT, USA). Store at −80°C. 10. Transcription and translation (TNT) kit. We frequently use TNT T7 Quick for PCR DNA (Promega, Madison, WI, USA), a transcription and translation coupled reticulocyte lysate system using the T7 polymerase promoter. Other wheat germ extract systems and reticulocyte lysate systems (Promega) are also useful (see Note 3). Store at −80°C. 11. Water bath, set at 30°C. 12. SDS-polyacrylamide gel electrophoresis (SDS-PAGE) system (e.g., Novex NuPAGE, Invitrogen, Carlsbad, CA, USA) (see Subheading 2.6).

2.3. Chloroplast Isolation

1. For one standard isolation procedure, 20–40 Petri plates of 10–14-day-old plants are required, each plate containing ~100– 150 seedlings (see Note 4). Additional material is required for

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more complex studies or for the analysis of multiple samples (e.g., different genotypes). 2. Two ice buckets with ice. 3. Two 1-L glass beakers, one 50-mL glass beaker, one 100-mL measuring cylinder, one funnel, and a sieve. 4. A plant tissue homogenizer. For example, an Ultra-Turrax T25 Basic (IKA, Staufen, Germany) with an 18-mm diameter rotor, and speed set to 3 (19,000 rpm) on a scale of 7 (see Note 5). 5. Cold room (4°C). 6. Miracloth (Calbiochem, Merck, Darmstadt, Germany); a double layer of about 20 × 20 cm is required. 7. One 250-mL and two 30-mL centrifugation tubes with screw caps (Nalgene, Rochester, NY, USA). 8. Isolation buffer: 0.3  M sorbitol, 5  mM MgCl2, 5  mM ethylene glycol-bis(2-aminoethylether)-N,N,N¢,N¢-tetraacetic acid (EGTA), 5 mM EDTA, 10 mM NaHCO3, and 20 mM 4-(2-hydroxyethyl)piperazine-1-ethanesulfonic acid (HEPES)KOH, pH 8.0. This is prepared as a twofold strong (2×) stock. Store at −20°C for long-term storage, and at 4°C for shortterm storage (see Note 6). 9. A high-speed centrifuge. For example, a Beckman J2-MC High-Speed Centrifuge with rotors JA-14, JA-20, and JS-13 (Beckman Coulter, Brea, CA, USA) (see Note 7). 10. Percoll medium; an opened bottle can be stored at 4°C for several months. To prolong its life, avoid inserting pipettes into the bottle but pour it out instead. 11. Linear Percoll gradient. Prior to use, mix 13  mL of Percoll medium with 13 mL of 2× isolation buffer, and 5 mg glutathione (roughly equals a small spatula tip of the powder; see Note 6). Mix all components in a 30-mL centrifugation tube until the glutathione is dissolved. Pre-spin before use in a fixed angle rotor at 43,000× gmax for 30  min (brake off) at 4°C; this is equivalent to 19,000 rpm in a JA-20 rotor in a Beckman centrifuge, deceleration set to 1. If the function is available on the centrifuge, select the slowest deceleration setting (see Note 8). 12. 1× HEPES–MgSO4–sorbitol (HMS) buffer: 50 mM HEPESNaOH, pH 8.0, 3 mM MgSO4, and 0.3 M sorbitol. Store at −20°C for long-term storage and at 4°C for short-term storage (see Note 9). 13. Aspirator device. 2.4. Determining Yield and Intactness of Chloroplasts

1. Hemocytometer with a 0.1-mm deep counting chamber and a ruling pattern of 0.0025 mm2. 2. Cover glass.

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3. Phase-contrast microscope. 4. 1× HMS buffer (see Subheading 2.3). 5. Tissue paper. 2.5. Chloroplast Protein Import

1. 10× HMS buffer: 500 mM HEPES-NaOH, pH 8.0, 30 mM MgSO4, and 3.0  M sorbitol. Store at −20°C for long-term storage and at 4°C for short-term storage (see Note 9). 2. 1 M gluconic acid (potassium salt). Store at −20°C. 3. 1 M NaHCO3. Store at −20°C. 4. 20% (w/v) bovine serum albumin (BSA). Store at −20°C. 5. 100 mM Mg-ATP. Store at −20°C. 6. 200 mM methionine. Store at −20°C. 7. Radiolabeled preprotein (see Subheading 3.2). 8. Freshly isolated chloroplasts (see Subheading  3.3). Typically, 10 × 106 organelles are required per import reaction. 9. Water bath set at 25°C with white light source (100  mmol/ m2/s) for illumination of the sample. 10. Import stop solution. Ice-cold 50 mM EDTA dissolved in 1× HMS buffer (see Subheading 2.3). 11. Microfuge for spinning down chloroplasts after import. 12. 4× Stacking gel buffer. Combine 6.06 g Tris base and 4 mL of 10% (w/v) SDS, and then add sterile deionized water to a final volume of 10 mL. Adjust the pH to 6.8 using HCl. Store at room temperature. 13. Denaturation buffer. Combine 2.0 mL of glycerol, 2.0 mL of 10% (w/v) SDS, 0.25  g bromophenol blue, 2.5  mL of 4× stacking gel buffer, and 0.5  mL of b-mercaptoethanol, and then add sterile deionized water to a final volume of 10 mL. Ideally, the b-mercaptoethanol should be added to the buffer just before use. Store at room temperature. 14. SDS-PAGE system (e.g., Novex NuPAGE, Invitrogen) (see Subheading 2.6).

2.6. SDS-PAGE for Analysis of Results

1. Heating block, set to 95°C for sample denaturing using denaturation buffer (see Subheading 2.5). 2. NuPAGE electrophoresis system using Novex high-performance precast gels with size 8 × 8 cm (Invitrogen) (see Note 10). 3. SDS running buffer. One may either use 3-(N-morpholino) propane sulfonic acid (MOPS) or MES in this buffer. The choice of MOPS or MES buffer affects the migration pattern of the loaded proteins. To make 500  mL of 20× MOPS/MES buffer, use 104.6 g MOPS or 97.6 g MES (1 M), 60.6 g Tris base (1  M), 10.0  g SDS (2% [w/v]), and 3.0  g EDTA

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(20.5 mM); make up to volume with sterile deionized water. Dilute to 1× concentration just before use. The final 1× buffer should be of pH 7.7; do not use acid or base to adjust the pH. 4. Coomassie stain. To make 100 mL, add 0.1 g of Coomassie Brilliant Blue R-250 to 10 mL of glacial acetic acid, 45 mL of methanol, and 45 mL of sterile water (see Note 11). 5. Destaining solution. To make 100 mL, use 10 mL of glacial acetic acid, 10  mL of methanol, and 80  mL of deionized water. 6. Orbital platform shaker. 7. Fluorescent signal amplifier, NAMP100 (GE Healthcare, Chalfont St. Giles, UK). 8. Whatman 3MM filter paper and plastic wrap (cling film). 9. Gel dryer (Bio-Rad, Hercules, CA, USA), typically set to dry at 80°C for 90 min. 10. Film exposure cassette, and Hyperfilm MP X-ray film (GE Healthcare) (see Note 12). 11. Film developing solutions. These are developer (Kodak, Rochester, NY, USA) in a 1-L glass bottle, stop solution (3% [v/v] acetic acid) in a 1-L glass bottle, and fix solution (Kodak) in a 1-L glass bottle. All steps should be performed in a dark room with red safety light. Alternatively, one may use a developing machine. 12. Scanner and ImageQuant software (GE Healthcare) (see Note 13).

3. Methods 3.1. Growth of Arabidopsis Plants

Although the method described below uses plants grown on MS medium, the isolation procedure after the harvesting step is essentially the same if the plants are grown on soil; chloroplasts isolated from soil-grown plants using this method are also import competent (15). When working with soil-grown material, the plants are preferably slightly older (to aid harvesting), and one should be very careful when harvesting to avoid soil contamination in the homogenization step, which may reduce the yield of intact chloroplasts due to mechanical breakage (H. Aronsson, personal observation). 1. Transfer a sufficient quantity of seeds (e.g., for 20 Petri plates each carrying ~100–150 plants, roughly 200 mL of seeds will be required) into a sterile 1.5-mL microfuge tube, and add 1 mL of sterilization solution 1 (see Note 14).

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2. Suspend all of the seeds in the solution (by shaking by hand) before placing the tube, orientated horizontally, onto an orbital shaker. Shake at 200–250 rpm for 5 min. 3. Let the seeds settle to the bottom of the tube and then remove the supernatant. Add 1  mL of sterilization solution 2, shake the tube by hand, and then place the tube on the orbital shaker at 200–250 rpm for 10 min. 4. Sterilize the laminar flow hood with 70% ethanol. Take out one filter paper per microfuge tube filled with seeds, and fold the filter papers in half to create a crease, which facilitates seed sowing (see Fig. 1). Soak the filter papers in 70% ethanol in the hood and let them dry. 5. Transfer the seeds onto the filter paper, using a cut 1-mL pipette tip (cut ~5 mm from the fine end, to increase the aperture size), and then leave them to dry for ~15 min. 6. Sow ~100–150 seeds per Petri plate containing MS medium (see Notes 1 and 14). 7. Seal each Petri plate with surgical tape. 8. Leave the plates upside down (to avoid condensation on the surface of the agar) at 4°C for at least two days to break seed dormancy and synchronize germination. 9. Grow the plants for 10–14 days in a plant growth chamber (see Fig. 1) (see Note 15). 3.2. Preparation of Radiolabeled Preproteins

Use a TNT-coupled wheat-germ extract kit or a TNT-coupled reticulocyte lysate system to obtain radiolabeled preproteins. Described below is the usage of a kit that employs PCR product as template: TNT T7 Quick for PCR DNA. Other kits use plasmids (linearized or not) as template. We generally use PCR templates to obtain radiolabeled preproteins using the aforementioned kit, but this is not always optimal (see Note 3). To use the TNT T7 Quick for PCR DNA kit, one has to obtain a cDNA plasmid clone encoding the preprotein of interest, run a PCR using that clone as template, and then check that the amplified product is of the correct size prior to starting the TNT reaction. However, the TNT kit does not require any purification of the PCR product prior to use (although it may be advisable to test this if problems are encountered). To obtain the plasmid DNA template for PCR amplification, one has to perform standard overnight growth of E. coli to amplify the plasmid of interest, and then purify the plasmid using a standard plasmid purification kit. 1. Purify your plasmid according to the instructions of the plasmid purification mini kit used. Measure the DNA concentration using a spectrophotometer using the absorbance at 260 nm.

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2. Dilute the purified plasmid 10–20-fold depending on the ­concentration of the cDNA template to a final concentration not exceeding 100 ng/mL. From this point onward, this protocol assumes use of pSS as the template/preprotein. 3. Run a 20 mL PCR (for 35 cycles) using the pSS cDNA as template. The reaction may be prepared as follows: 2 mL

10× polymerase buffer

0.4 mL

10 mM dNTPs

1 mL

10 mM M13 forward primer

1 mL

10 mM M13 reverse primer

1 mL

Diluted (1:10–20) pSS cDNA template not exceeding 100 ng

1 U

Taq polymerase

X mL

Deionized sterile water to take the total reaction volume to 20 mL (added prior to the addition of the polymerase)

4. Run 5 mL of the PCR product on a 1% agarose gel (using TAE buffer) to verify correct amplification of the cDNA and to quantify the relevant band (alternatively, the DNA concentration can be determined photometrically). Store the rest of the product at −20°C for further applications. 5. Prepare a 50  mL TNT T7 Quick for PCR DNA reaction as follows: 40 mL

TNT reticulocyte lysate from the kit (thaw on ice prior to use)

2.5 mL Radiolabeled [35S] methionine, 11 mCi/mL (bring out from −80°C storage just prior to use) 2.5 mL Deionized sterile water (optional: use nuclease-free water) 5 mL

pSS PCR product (100–800 ng)

6. Incubate for 90 min in a 30°C water bath. Stop the reaction by placing the sample on ice. 7. Take out 5 mL as a test sample for verification on SDS-PAGE. Add 10 mL of denaturation buffer to the test sample, boil at 95°C for 90 s, electrophorese on a precast gel, and then stain and dry the gel (see Subheading 3.6). Expose to film in a cassette overnight (~14–18 h). A good result is indicated by observing a distinct and strong band corresponding to the molecular weight of pSS (see Fig. 2) (see Note 16). 8. Store the translation product at −80°C for later applications.

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Fig. 2. Analysis of radiolabeled translation products. Multiple translation products of pSS were prepared using a reticulocyte kit and a PCR product corresponding to pSS cDNA as template; each transcription and translation reaction was performed as described in Subheading 3.2. Equivalent volumes (5 mL) of the translation products were resolved by SDS-PAGE. The gel was incubated with NAMP before drying and was then exposed to an X-ray film for 16 h. Labels 1–4 indicate different pSS translation products. For a single import reaction (total volume 200 mL), it would be appropriate to add 5 mL from sample number 1, 10 mL from sample number 2, and 20 ml from sample number 3 or 4. The position of the precursor form of pSS (which is 20 kDa in size) is indicated (p).

3.3. Chloroplast Isolation

Start the isolation early in the morning to avoid starch ­accumulation inside chloroplasts (which can hamper the isolation of intact organelles). Doing so also leaves more time left over for further ­applications downstream of the isolation procedure (e.g., protein import). Once the plant material has been harvested, it is important to work as quickly as possible, without compromising accuracy, in order to avoid unnecessary protein degradation due to release of proteases during mechanical breakage. The biological material and apparatus should be kept at 4°C during the whole procedure (either in a cold room or on ice). The homogenization step should be carried out at 4°C in a cold room. 1. The day before the isolation, place the following items in a cold room or refrigerator: 250  mL of 2× isolation buffer, 50 mL of 1× HMS buffer, 200 mL of deionized water, a double layer of Miracloth, a funnel, a 100-mL measuring cylinder, two 1-L glass beakers, one 50-mL glass beaker, 250- and 30-mL centrifugation tubes, and Beckman JA-14, JA-20, and JS-13.1 rotors. Thus, all solutions will be thawed and all equipment precooled prior to use. 2. On the day of the isolation, prepare isolation buffer by mixing well 200  mL of deionized H2O with 200  mL of 2× isolation buffer; keep on ice. Put 100 mL of the isolation buffer into a 1-L beaker and keep on ice. Place a second 1-L beaker on ice, with a funnel on top containing a double layer of Miracloth. Place the 250- and 30-mL centrifugation tubes on ice prior to use. 3. Prepare a continuous Percoll gradient as described in Subheading  2.3 (item 11), and keep the tubes on ice after

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­ re-centrifugation (see Note 17). It may be optimal to ­perform p harvesting and homogenization (steps 4–10 below) while the Percoll gradient is being formed, to save time. 4. Fill the 100-mL measuring cylinder with 100 mL of isolation buffer (1×). During homogenization, a total of 100 mL of isolation buffer is used per sample; this is used in five consecutive rounds of homogenization, each one using 20 mL of isolation buffer. 5. Bring the Petri plates from the growth chamber and remove the sealing tape. Remove the plants from the medium by gently scraping them off with a gloved hand, and place them in the 100 mL of isolation buffer in the 1-L glass beaker sitting on ice (see Note 18). 6. Place 20 mL of isolation buffer from the measuring cylinder into the 50-mL glass beaker, and then transfer the harvested seedlings into the beaker. When transferring the plant material, it is useful to pull up gently a handful of plants and let the excess buffer run through your fingers before adding the plants into the beaker. It may be optimal to fill the beaker almost full with plants, and then continue with a new beaker with any remaining plant material. Usually, the harvest from 20 Petri plates of plants fits into one 50-mL beaker. 7. Place the 50-mL glass beaker containing the plant material under the rotor of the Ultra Turrax (or Polytron) and homogenize for 3–4 s (see Fig. 1). The optimal conditions for the homogenization have to be established empirically (see Note 19). 8. Filter the homogenate through a double layer of Miracloth (mounted on the funnel) into the 1-L beaker on ice. Gently squeeze the Miracloth around the plant material to speed up the gravity flow. 9. Place a second 20 mL aliquot of isolation buffer from the measuring cylinder into the 50-mL glass beaker, and return the plant material remaining after filtration (debris) from the Miracloth to the beaker. Check the debris between each homogenization round to avoid an unwanted low final yield of intact chloroplasts (see Note 20). 10. Repeat steps 7–9 above until all 100 mL of isolation buffer (in 5 × 20 mL aliquots; see step 4) has been used; i.e., perform five rounds of homogenization and filtration in total. The plant material will gradually become disrupted during the procedure. This repetitive procedure will give a high yield of intact chloroplasts. 11. Transfer the pooled, filtered homogenate into the 250-mL centrifugation tube on ice and centrifuge at 1,000 × gmax for 5 min (brake on) at 4°C; this is equivalent to 2,600 rpm in a JA-14 rotor in a Beckman centrifuge.

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12. Decant and discard the supernatant. The pellet obtained should be firmly attached to the tube, and so one can easily turn the tube upside down quickly to remove the supernatant. Resus­ pend the pellet in the residual isolation buffer that is left in the centrifugation tube; do this by swirling the centrifugation tube  on ice in an ice bucket. Add an additional volume of 500–2,000 mL of 1× isolation buffer to facilitate the resuspension. The additional volume added should be proportional to the size of the pellet. 13. Transfer the suspension onto the top of the pre-made Percoll linear gradient (see Subheading 2.3, item 11) using a cut 1-mL pipette tip (cut ~5 mm from the fine end, to increase the aperture size). Pipette slowly to avoid disturbance of the gradient. 14. To separate the intact chloroplasts from broken chloroplasts and other debris, centrifuge in a swing-out rotor at 7,800 × gmax for 10 min (brake off) at 4°C; this is equivalent to 7,000 rpm in a JS-13.1 rotor in a Beckman centrifuge, deceleration set to 1. 15. After centrifugation, remove the tube carefully and place it on ice. The lower green band in the gradient contains intact chloroplasts, whereas the upper band contains broken chloroplasts. First, discard the broken chloroplasts by carefully removing them by pipetting or aspirating (faster). Second, recover the intact chloroplasts carefully using a 1-mL pipette tip (cut at the end, as described above) and transfer them into a precooled 30-mL centrifugation tube on ice. The volume of recovered intact chloroplasts can range from 2 to 8 mL per gradient. 16. Add 1× HMS buffer to the recovered chloroplasts to at least 80% of the tube’s volume, or as much as possible, and invert the tube carefully two to three times to wash off the Percoll. 17. To remove the Percoll, centrifuge the chloroplasts in a swingout rotor at 1,000 × gmax for 5 min (brake on) at 4°C; this is equivalent to 2,500  rpm in a JS-13.1 rotor in a Beckman centrifuge. 18. Gently pour off the supernatant and resuspend the pellet in the remaining HMS buffer (left in the tube after decanting). Add an additional 100–600 mL of fresh 1× HMS buffer (depending on the size of the pellet) to aid resuspension. Resuspend by swirling the tube gently on ice. Do not resuspend by pipetting, and do not dilute too much! Preferably, the suspension is more dark green and thick than light green and thin. A higher chloroplast density may better preserve the chloroplasts’ intactness. Depending on the nature of the experiment to be conducted, the yield of chloroplasts may be counted (see Subheading 3.4); alternatively, chlorophyll content or protein concentration can be measured and recorded (see Note 21).

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3.4. Determining Yield and Intactness of Chloroplasts

When appropriate, the yield of chloroplasts and their intactness can be assessed as follows. This is particularly important for protein import experiments, as we typically add a fixed number of chloroplasts to each assay. Furthermore, it is essential to check the quality of the isolated organelles before proceeding to test protein import. 1. Add 5 mL of isolated chloroplasts to 495 mL of 1× HMS buffer in a 1.5-mL microfuge tube, and then mix gently by flicking the tube to obtain a 1:100 dilution. 2. Pipette ~40–60 mL of the diluted suspension onto the counting chamber of the hemocytometer and place a cover glass on top. 3. Drain the excess suspension with tissue paper. 4. Count the number of chloroplasts in 16 different small squares within the 0.25-mm2 large square, using a phase-contrast microscope with a 10× or 20× objective. The number of chloroplasts per square should average between 10 and 30. If too few or too many chloroplasts are present, adjust the dilution factor (step 1 above) accordingly and repeat the procedure. Intact chloroplasts appear round and bright green, and under phase contrast are surrounded by a bright halo of light. 5. The number of chloroplasts per microliter is calculated as ­follows: n (the average number of chloroplasts per square, ­calculated from the 16 counts in step 4, each square being 0.00625  mm3) × 100 (the dilution factor employed in step 1)/0.00625 (a scaling factor to express the data per microliter, since the volume of a single square is 0.00625 mm3). For example, if one calculates 25 chloroplasts/square, one will have (25 × 100)/0.00625 = 400,000 chloroplasts/mL. 6. Calculate the actual yield of chloroplasts by multiplying the concentration (number of chloroplasts per microliter) by the volume of chloroplast suspension obtained in Subheading 3.3, step 18. A typical good yield is 100–200 × 106 chloroplasts (in 250–500 mL). 7. For one import reaction, we typically use 10 × 106 chloroplasts. In order to use this number of chloroplasts using the concentration given as example in step 5 above, one would have to add 25 mL (=10,000,000/400,000) per import reaction (see Subheading 3.5). 8. Samples prepared using the methodology described here are mostly intact and exhibit minimal contamination from other cellular compartments; see ref. 27.

3.5. Chloroplast Protein Import

Several protocols have been described for conducting chloroplast protein import assays. The method described here is a minimized version for conducting successful import quickly, and is to a large extent similar to the ones initially used for import studies in pea.

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We have not included the optional protease treatment step that can be conducted after import, to further support proper import, as this step is described in detail in Chapter 21, Vol. 1. The presented protocol is suitable for conducting time-course assays, which allow one to monitor import over time and thus compare rates between, for example, wild-type and mutant or transgenic plants. It is important to note that this part of the work must be conducted with appropriate precautions because of its radioactive nature. 1. To run a time course with three time points, prepare a 600-mL import reaction for each genotype in a 1.5-mL microfuge tube. If you are just running one time point (e.g., wild-type chloroplasts to test the import capability of a new protein of interest), then divide the volumes below by three (i.e., start with a 200-mL reaction). Thaw all ingredients just prior to use and keep them on ice. Mix them in the following order: (52.5) mL

10× HMS buffer. One has to know the amount of chloroplasts needed (see below) before calculating the amount of 10× HMS buffer needed (as the chloroplasts are in 1× HMS buffer). Assuming that 75 mL of chloroplasts are to be added, the volume of 10× HMS required = (600 [total reaction volume] − 75 [chloroplast suspension])/10 (dilution factor to reach 1× HMS) = 52.5 mL

(358.5) mL

Deionized water (to take the final volume to 600 mL). The exact volume required depends on the volumes of chloroplast suspension, 10× HMS, and radiolabeled preprotein. Pipette up and down several times to ensure that the 10× HMS buffer is mixed

12 mL

1 M gluconic acid (potassium salt; final concentration 20 mM)

6 mL

1 M NaHCO3 (final concentration 10 mM)

6 mL

20% (w/v) BSA (final concentration 0.2%)

30 mL

200 mM methionine (final concentration 10 mM)

30 mL

100 mM Mg-ATP (final concentration 5 mM)

(30 mL)

Radiolabeled preprotein (not exceeding 10% of the total volume). For a translation product giving a very strong band (see Note 16), 15 mL is enough, while for one giving a weak band, one could use up to 60 mL (see Subheading 3.2)

(75 mL)

Chloroplast suspension containing 30 × 106 chloroplasts. Assuming a concentration of 400,000 chloroplasts/mL, 75 mL should be added. Use a cut tip (avoiding mechanical breakage of the chloroplasts) to load the chloroplast suspension into the reaction tube, thereby initiating the import reaction (see Note 22)

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2. Incubate in white light (100 mmol/m2/s) at 25°C in a water bath for several minutes (see below) (see Fig. 1). It is recommended to flick (or agitate gently) the tubes every third minute to avoid chloroplasts settling to the bottom of the tube, which would otherwise impair import efficiency and affect sampling. 3. To conduct a time course, samples (190 mL) should be withdrawn within the linear range of import, which for pSS is up to ~10  min (3-, 6-, and 10-min time points are suitable). The linear range period might vary for other proteins since they may have a faster or slower import rate (33). Thus, it is advisable to test each individual preprotein prior to initiating such experiments. 4. Immediately upon withdrawal, each 190 mL aliquot should be added to a microfuge tube already filled with 200 mL of icecold import stop solution (see Note 23). Retain all tubes on ice until the time course has been completed. 5. Recover the intact chloroplasts in each aliquot by passing them through a prechilled 35% Percoll cushion in a microfuge. To prepare each cushion, mix 350  mL of Percoll medium with 100 mL of 10× HMS buffer and 550 mL of deionized water in a microfuge tube. Add the import reaction aliquot to the top of the cushion using a cut pipette tip and then centrifuge for 4 min at 4,200 × gmax in a microfuge at 4°C. Proceed to step 7. 6. Alternatively, the procedure may be expedited by omitting step 5 if one is confident that loss of organelle integrity during the import reaction is not an issue. In this case, quickly recover the sample by a short centrifugation burst (~3  s) up to 4,000 × gmax in a microfuge. In tests, we observed no significant decrease in the number of intact chloroplasts following extended incubations (up to 60 min) under import conditions (Student’s t-test, p > 0.5; N. Z. Khan, S. Karim, and H. Aronsson, personal observation). 7. Remove and discard the supernatant with a 200-mL pipette tip. Add 10–15 mL (no more than this, when combined with the pellet, can be loaded into a single well of the SDS-PAGE system used) of denaturation buffer to the remaining pellet and dissolve by pipetting up and down using a 200-mL pipette tip. Boil at 95°C for 90 s and load into an SDS-PAGE system (see Subheading  3.6). If not loading the samples directly, they may be stored at −20°C for analysis on another day. It may be optimal to avoid adding denaturation buffer to the pellets if they are to be stored at −20°C (to better preserve sample integrity); instead, add it to the sample just prior to loading. 8. Prepare a control sample of radiolabeled pSS. Take 10% of the volume of pSS present in each import reaction withdrawal

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volume (in the above example, this would be 0.95 mL) and add 15 mL of denaturation buffer prior to boiling and loading (see step 7 above). The control should be loaded next to the import samples to allow assessment of the protein import efficiency, and to aid orientation and identification of the imported proteins (see Fig. 3c). 3.6. SDS-PAGE for Analysis of Results

1. Load the samples onto Novex precast gels using a MOPS or MES buffer system depending on the size of the product (for 3.5–40 kDa, use MES buffer; for 10–80 kDa, use MOPS buffer). A protein size ladder should also be loaded to aid interpretation of the results (see Note 10). 2. Run the gel at ~20  V/cm for 30–50  min (using the Novex precast gels, this corresponds to 150 V). Stop the gel when the blue dye from the denaturation buffer is ~1 cm from the end of the gel. 3. Demount the gel cassette and remove the gel from the plates. Cut away and discard the wells. 4. Treat the gel with Coomassie stain for 15 min at room temperature using an orbital shaker. 5. Treat the gel in destaining solution for 10 min, prior to incubation in NAMP for 20  min; use an orbital shaker for both steps. 6. Place the gel carefully onto a precut piece of Whatman filter paper that is at least a few centimeters wider/longer than the gel. 7. Soak the Whatman paper around the gel with deionized water. Place the Whatman paper and gel into a gel dryer and cover it with a cling film. Close the gel dryer and dry the gel for 90 min at 80°C. Remove the cling film. 8. In a dark room, place the dried gel inside and at the bottom of a folded piece of A4 paper, and then add a piece of X-ray film on top of the gel, also to the bottom of the folded A4 paper (the A4 paper helps to avoid unwanted movement between the gel and the film during the exposure time). Place the folded paper, gel, and film inside an exposure cassette (see Note 24). 9. Put the film cassette in a dark place at room temperature. The ideal exposure time might vary, but a good protein import experiment should not need more than 2–7 days of exposure (see Note 25). The film cassettes can be stored at −80°C to increase the sharpness of the bands. 10. Remove the film from the gel in a dark room. Place the film in a tray with developer for up to 10 min. Lift the film and let the excess developer drip off before adding the film to a tray with stop solution for 1 min. Lift the film and let the excess stop

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Fig. 3. Time-course analysis of protein import into isolated wild-type and ppi1 mutant chloroplasts. Chloroplasts were isolated from 14-day-old wild-type (a) and ppi1 mutant (b) plants grown on MS plates. Bars indicate 1  cm. (c) Protein import was conducted using [35S]-methionine-labeled pSS and allowed to proceed for 3, 6, and 10 min, before analysis by SDS-PAGE and fluorography. Translation mixture (TM) containing pSS equivalent to 10% of the amount added to each import reaction was analyzed in parallel. The precursor (p) and mature (m) forms of pSS are indicated. (d) To provide an indication of import efficiency, the amount of imported protein in (c) was assessed by measuring the radioactivity associated with each mature band. The data were expressed as percentages of the amount of imported protein in wild-type chloroplasts after 10 min.

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solution drip off before placing it in a tray with fix solution, where it will stay for up to 15  min. Rinse in tap water for 15 min and finally air-dry the film. 11. Scan the film and use ImageQuant software (or similar) to quantify and analyze the results. Alternatively, if a phosphorimager was used to image the results, then scanning is not necessary and the results can be quantified directly. Typical results are shown in Fig. 3.

4. Notes 1. The agar concentration can vary from 0.6 to 0.9% (w/v) depending on what brand is used. If it is too low, the plates will be slushy, and if it is too hard, the MS medium will stick heavily to the plants; both cases will prevent efficient harvesting. Also, the sucrose concentration can be varied depending on the plants used. When working with certain, particularly sick mutants, it may be beneficial to use MS medium supplemented with up to 3% (w/v) sucrose. Stored plates can be used for up to one month. 2. Light slightly below or above this intensity range is also satisfactory, but it is important to measure the light since this can significantly affect growth. This may be of importance if one wishes to compare samples grown on different occasions. 3. For TNT, there are two major types of kit: those based on wheat germ and those based on rabbit reticulocyte (e.g., TNTcoupled wheat germ extract system, and TNT-coupled reticulocyte lysate system; both Promega). There are kits that use linearized plasmids, unlinearized plasmids, or PCR products as the template. There are also kits specific for the T3, T7, and SP6 promoters. However, the kit that uses the PCR product we recommend in this chapter (TNT T7 Quick for PCR DNA; Promega) is only suitable for use with T7 promoters. Unfortunately, obtaining radiolabeled preproteins is not always straightforward and some preproteins might be obtained in higher amount using a wheat germ extract system or a reticulocyte lysate system meant for plasmid templates. Therefore, one has to test empirically to find the conditions that give the best results. One may improve the result of the translation by modifying the incubation time in the water bath (60–90 min), by altering the water bath temperature (27–30°C), by optimizing the translation mixture composition (e.g., reducing/increasing the amount of radioactivity, or changing template or salt concentrations; see Promega TNT handbook), or by editing the

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sequence around the start codon in order to achieve a good Kozak consensus (34). 4. For 10-day-old plants, this is roughly equivalent to ~4,000– 6,000 individuals or ~16 g tissue. More plates are often needed when working with mutant plants with an obvious growth defect or visible chlorosis. 5. The method also works well using a Kinematica Model PT1035 (Kinematica AG, Lucerne, Switzerland) with a large rotor (PTA 20 S) and speed set to 4 on a scale of 11, or Model PT20 with a small rotor (13 mm diameter) at ~40% maximum speed, as described previously (14, 27). A kitchen blender might also be used, but we have identified several problems with the yield of intact chloroplasts using such devices, which might be due to mechanical breakage of the chloroplasts and/or the larger volume when homogenizing the chloroplasts inside the glass beaker of the kitchen blender. 6. Prepare 2× isolation buffer and store in 200  mL aliquots in 400-mL glass bottles at −20°C. A small amount of 2× isolation buffer (13 mL) is needed for preparing each linear Percoll gradient. Before use, thaw overnight at 4°C, mix well, and transfer the needed amount to individual tubes (e.g., for two linear Percoll gradients, 26 mL is needed in total). The remaining 2× isolation buffer (200–26 = 174  mL) is diluted with the same amount of sterile deionized water to obtain 1× isolation buffer; mix well before use. The components of the isolation buffer prevent osmotic lysis of the organelles (sorbitol); stabilize the pH (HEPES); provide a carbon source (NaHCO3), a cofactor for proteins/enzymes (MgCl2), and ions for transporters to maintain metabolic activities (MgCl2, NaHCO3); and minimize the effects of potentially harmful enzymatic reactions (divalent cation cofactors are removed by the chelators EDTA and EGTA) (17, 35–39). Ascorbic acid is an antioxidant component that is sometimes added in chloroplast isolation buffers (40). To be cautious, one might consider adding ascorbic acid to the isolation buffer; however, in our hands, doing so does not detectably improve import efficiency in subsequent assays (H. Aronsson, personal observation). Note that another antioxidant, glutathione, is included in the Percoll gradient step. 7. A Sorvall RC6 centrifuge (Thermo Fisher Scientific, Waltham, MA, USA) can be used as well with rotors SLA-1500 (equivalent to JA-14), SS-34 rotor (equivalent to JA-20), and HB-6 (equivalent to JS-13.1). 8. Gradients can be prepared the day before and stored overnight at 4°C. However, be careful to keep them away from unwanted movements which might cause disturbance of the linear gradient.

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9. Store 1× HMS buffer in 200  mL aliquots in 400-mL glass bottles at −20°C. Store 10× HMS buffer as 1 mL aliquots in microfuge tubes at −20°C. The buffer can be thawed and frozen several times. Mix well after thawing. In this buffer, sorbitol and HEPES prevent osmotic lysis of the organelles and stabilize the pH, respectively; the MgSO4 provides Mg2+ as a cofactor for nucleoside triphosphatases and for thylakoid membrane stabilization, and SO42− for metabolic activities (35, 36, 39, 41, 42). 10. Other SDS-PAGE systems not using precast gels are also satisfactory. 11. If a gel dryer is not available, one can instead blot the proteins to a nitrocellulose membrane to enable fluorography or phosphorimaging. In this case, stain the membrane with Ponceau Red (or Amido Black) to confirm equal loading instead of Coomassie staining the gel. 12. A phosphorimager represents an alternative imaging method that has sensitivity and quantification advantages; if used, the NAMP100 should be omitted. 13. This is only necessary if quantification of the import assay is required. Other similar quantification programs are also available and work fine (e.g., Gene-Tools, Syngene). 14. Sterilize hands with 70% ethanol and avoid touching the seeds directly. Gloves should be avoided since the seeds will stick to the gloves due to static electricity. 15. Depending on the lights, etc., in the growth system to be used, one may need to run tests to find the optimal conditions or growth period, as these can vary between facilities. 16. By assessing the strength of the band after such overnight exposure, one can compare the efficiency between different translation reactions. This is important because one has to add more translation product (to the import reaction) for proteins that produce a weak band than for those that produce a strong band. One may improve the result of a translation reaction by making various modifications (see Note 3). 17. If using newly thawed aliquots of 2× isolation buffer, mix well before use to obtain a homogeneous solution. 18. Water on the finger tips of a glove will facilitate harvesting. Minimizing carryover of MS medium to the next step is crucial since it may interfere with the yield of intact chloroplasts. An optional step is to first put the harvested material in a sieve standing under running cold tap water to wash off any adhering MS medium quickly, prior to adding the plants to the isolation buffer in the 1-L glass beaker. 19. Good homogenization is achieved by moving the 50-mL ­beaker up and down quickly during the homogenization step

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(taking ~3–4  s in total), in each one of the five rounds of homogenization. If working with chlorotic or sick mutants, homogenization times can be reduced depending on the severity of the mutant phenotype. 20. A large amount of tissue will need a larger volume of isolation buffer. However, one should always use a 50-mL beaker as the standard beaker size. Instead of using more rounds, divide the tissue into more beakers. Checking the debris is a good indicator of the likely chloroplast yield. From the first to the last round of homogenization, the debris should go from having several intact pieces of tissue to gradually becoming more like a soup. If the debris contains several pieces of intact material after the last step, then the homogenization has been too mild. On the contrary, if the debris is like a soup already after 1–2 rounds, then the homogenization may have been too rough. In both cases, the final yield of intact chloroplasts will not be optimal. 21. Protein concentration can be determined and calculated using Bradford reagent (e.g., Bio-Rad Protein Assay). Chlorophyll concentration can be determined by measuring the absorbance at 646.8 and 663.8  nm, and calculated as previously described (43). 22. Alternatively, if several microfuge tubes are to be used and compared for import, then it is advisable to make a master mix consisting of 10× HMS buffer, deionized water, K-gluconic acid, NaHCO3, BSA, methionine, and Mg-ATP. Thereafter, add the radiolabeled proteins before finally adding the chloroplasts. This will assure more equal conditions in the test tubes. In addition to buffering capacity and an osmoticum, the import reaction provides various ions to maintain metabolic activity through transporters (K-gluconic acid is used as a K+ source as gluconate is less harmful to ATP/ADP transporters than Cl−; HCO3− as a carbon source and to help keep the solution basic; BSA to preserve biological activity and reduce unwanted protein degradation by acting as a substrate to proteases; cold methionine to prevent the incorporation of radioactive methionine (present in the translation product) into unrelated chloroplast proteins by de novo synthesis in organello during the import reaction; and Mg-ATP as an energy resource for the import reaction to proceed) (37, 44–49). 23. When running a time course, it is advisable to make an appropriately oversized import reaction; e.g., if using a 600-mL import reaction mixture, it is recommended to take out 190 mL at each time point to assure that there is enough sample left for the last sample withdrawal. 24. Optional: To help link radioactive bands on the exposed film with the exact spot on the gel, mix radioactive methionine with ink or charcoal and add small spots at the corners of the gel in

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a characteristic way, prior to exposure. Alternatively, it is useful to put clear scotch tape on the film to attach it to the gel filter paper. When exposure is completed, one can cut the film next to the scotch tape, and later on (after development) bring it back to fit with the cutting in order to orientate the gel. 25. The translation efficiency test is done using 5  mL of radiolabeled protein for a 14–18-h exposure (see Subheading  3.2, step 7). If, for example, one used 10 mL of translation product for each import sample, then one would have to load 1 mL as the 10% control next to the import samples (see Subheading 3.5, step 8). Thus, applying a rough rule of thumb, if it took 14–18 h for 5 mL to be visualized, it should be possible to visualize 1 mL in 5 × 18 h, which is roughly 4 days of exposure. If one wants stronger bands, then one can expose for a longer period.

Acknowledgments Thanks to Dr. Sazzad Karim and Nadir Zaman Khan for valuable comments on the manuscript. This work was supported by the Swedish Research Council VR (H.A.), and the Royal Society and the Biotechnology and Biological Sciences Research Council (BBSRC) (P.J.). References 1. Blobel, G., and Sabatini, D. D. (1971) Ribosome-membrane interactions in eukaryotic cells. In, Biomembranes, Vol. 2 (Manson, L. A., ed.) Plenum, New York, USA, pp. 193–195. 2. Blobel, G., and Dobberstein, B. (1975) Transfer of proteins across membranes. I. Presence of proteolytically processed and unprocessed nascent immunoglobulin light chains on membrane-bound ribosomes of murine myeloma. J. Cell Biol. 67, 835–851. 3. Blobel, G., and Dobberstein, B. (1975) Transfer of proteins across membranes. II. Reconstitution of functional rough microsomes from heterologous components. J. Cell Biol. 67, 852–862. 4. Sandelius, A. S., and Aronsson, H. (2009) The Chloroplast - Interactions with the Environment. Plant Cell Monographs, Vol. 13. SpringerVerlag, Berlin, Germany. 5. Abdallah, F., Salamini, F., and Leister, D. (2000) A prediction of the size and evolutionary origin of the proteome of chloroplasts of Arabidopsis. Trends Plant Sci. 5, 141–142.

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10. Nanjo, Y., Oka, H., Ikarashi, N., Kaneko, K., Kitajima, A., Mitsui, T., Munoz, F. J., Rodriguez-Lopez, M., Baroja-Fernandez, E., and Pozueta-Romero, J. (2006) Rice plastidial N-glycosylated nucleotide pyrophosphatase/phosphodiesterase is transported from the ER-golgi to the chloroplast through the secretory pathway. Plant Cell 18, 2582–2592. 11. Aronsson, H., and Jarvis, P. (2009) The chloroplast protein import apparatus, its components, and their roles. In, The Chloroplast - Interactions with the Environment (Sandelius, A. S., and Aronsson, H., eds.) Springer-Verlag, Berlin, Germany, pp. 89–124. 12. Chua, N. H., and Schmidt, G. W. (1978) Posttranslational transport into intact chloroplasts of a precursor to the small subunit of ribulose1,5-bisphosphate carboxylase. Proc. Natl. Acad. Sci. USA 75, 6110–6114. 13. Highfield, P. E., and Ellis, R. J. (1978) Synthesis and transport of the small subunit of chloroplast ribulose bisphosphate carboxylase. Nature 271, 420–424. 14. Aronsson, H., and Jarvis, P. (2002) A simple method for isolating import-competent Arabidopsis chloroplasts. FEBS Lett. 529, 215–220. 15. Boij, P., Patel, R., Garcia, C., Jarvis, P., and Aronsson, H. (2009) In vivo studies on the roles of Tic55-related proteins in chloroplast protein import in Arabidopsis thaliana. Mol. Plant 2, 1397–1409. 16. Jarvis, P., Chen, L. J., Li, H., Peto, C. A., Fankhauser, C., and Chory, J. (1998) An Arabidopsis mutant defective in the plastid general protein import apparatus. Science 282, 100–103. 17. Somerville, C. R., Somerville, S. C., and Ogren, W. L. (1981) Isolation of photosynthetically active protoplasts and chloroplasts from Arabidopsis thaliana. Plant Sci. Lett. 21, 89–96. 18. Fitzpatrick, L. M., and Keegstra, K. (2001) A method for isolating a high yield of Arabidopsis chloroplasts capable of efficient import of precursor proteins. Plant J. 27, 59–65. 19. Dahlin, C., and Cline, K. (1991) Developmental regulation of the plastid protein import apparatus. Plant Cell 3, 1131–1140. 20. Kovacheva, S., Bedard, J., Patel, R., Dudley, P., Twell, D., Rios, G., Koncz, C., and Jarvis, P. (2005) In vivo studies on the roles of Tic110, Tic40 and Hsp93 during chloroplast protein import. Plant J. 41, 412–428. 21. Kovacheva, S., Bedard, J., Wardle, A., Patel, R., and Jarvis, P. (2007) Further in vivo studies on

the role of the molecular chaperone, Hsp93, in plastid protein import. Plant J. 50, 364–379. 22. Kubis, S., Baldwin, A., Patel, R., Razzaq, A., Dupree, P., Lilley, K., Kurth, J., Leister, D., and Jarvis, P. (2003) The Arabidopsis ppi1 mutant is specifically defective in the expression, chloroplast import, and accumulation of photosynthetic proteins. Plant Cell 15, 1859–1871. 23. Kubis, S., Patel, R., Combe, J., Bedard, J., Kovacheva, S., Lilley, K., Biehl, A., Leister, D., Rios, G., Koncz, C., and Jarvis, P. (2004) Functional specialization amongst the Arabidopsis Toc159 family of chloroplast protein import receptors. Plant Cell 16, 2059–2077. 24. Sjögren, L. L., MacDonald, T. M., Sutinen, S., and Clarke, A. K. (2004) Inactivation of the clpC1 gene encoding a chloroplast Hsp100 molecular chaperone causes growth retardation, leaf chlorosis, lower photosynthetic activity, and a specific reduction in photosystem content. Plant Physiol 136, 4114–4126. 25. Aronsson, H., Boij, P., Patel, R., Wardle, A., Töpel, M., and Jarvis, P. (2007) Toc64/OEP64 is not essential for the efficient import of proteins into chloroplasts in Arabidopsis thaliana. Plant J. 52, 53–68. 26. Aronsson, H., Combe, J., Patel, R., Agne, B., Martin, M., Kessler, F., and Jarvis, P. (2010) Nucleotide binding and dimerization at the chloroplast pre-protein import receptor, atToc33, are not essential in vivo but do increase import efficiency. Plant J. 63, 297–311. 27. Kubis, S. E., Lilley, K. S., and Jarvis, P. (2008) Isolation and preparation of chloroplasts from Arabidopsis thaliana plants. Methods Mol. Biol. 425, 171–486. 28. Garcia, C., Khan, N. Z., Nannmark, U., and Aronsson, H. (2010) The chloroplast protein CPSAR1, dually localized in the stroma and the inner envelope membrane, is involved in thylakoid biogenesis. Plant J. 63, 73–85. 29. Park, S., and Rodermel, S. R. (2004) Mutations in ClpC2/Hsp100 suppress the requirement for FtsH in thylakoid membrane biogenesis. Proc. Natl. Acad. Sci. USA 101, 12765–12770. 30. Sinvany-Villalobo, G., Davydov, O., Ben-Ari, G., Zaltsman, A., Raskind, A., and Adam, Z. (2004) Expression in multigene families. Analysis of chloroplast and mitochondrial proteases. Plant Physiol. 135, 1336–1345. 31. Koo, A. J., Fulda, M., Browse, J., and Ohlrogge, J. B. (2005) Identification of a plastid acyl-acyl carrier protein synthetase in Arabidopsis and its role in the activation and elongation of exogenous fatty acids. Plant J. 44, 620–632.

17  Chloroplast Isolation and Protein Import 32. Flores-Perez, U., Sauret-Gueto, S., Gas, E., Jarvis, P., and Rodriguez-Concepcion, M. (2008) A mutant impaired in the production of plastome-encoded proteins uncovers a mechanism for the homeostasis of isoprenoid biosynthetic enzymes in Arabidopsis plastids. Plant Cell 20, 1303–1315. 33. Aronsson, H., Schottler, M. A., Kelly, A. A., Sundqvist, C., Dörmann, P., Karim, S., and Jarvis, P. (2008) Monogalactosyldiacylglycerol deficiency in Arabidopsis affects pigment composition in the prolamellar body and impairs thylakoid membrane energization and photoprotection in leaves. Plant Physiol. 148, 580–592. 34. Kozak, M. (2002) Pushing the limits of the scanning mechanism for initiation of translation. Gene 299, 1–34. 35. Robinson, S. P. (1985) Osmotic adjustment by intact isolated chloroplasts in response to osmotic stress and its effect on photosynthesis and chloroplast volume. Plant Physiol. 79, 996–1002. 36. Rathnam, C. K. M., and Edwards, G. E. (1976) Protoplasts as a tool for isolating functional chloroplasts from leaves. Plant Cell Physiol. 17, 177–186. 37. Sirevag, R., Buchanan, B. B., Berry, J. A., and Troughton, J. H. (1977) Mechanisms of CO2 fixation in bacterial photosynthesis studied by the carbon isotope fractionation technique. Arch. Microbiol. 112, 35–38. 38. Shaul, O. (2002) Magnesium transport and function in plants: the tip of the iceberg. Biometals 15, 309–323. 39. Neuhaus, H. E., and Wagner, R. (2000) Solute pores, ion channels, and metabolite transporters in the outer and inner envelope membranes of higher plant plastids. Biochim. Biophys. Acta 1465, 307–323. 40. Schulz, A., Knoetzel, J., Scheller, H. V., and Mant, A. (2004) Uptake of a fluorescent dye as a swift and simple indicator of organelle intactness: import-competent chloroplasts from soilgrown Arabidopsis. J. Histochem. Cytochem. 52, 701–704.

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Chapter 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 Abstract Most chloroplast proteins are synthesized in the cytosol as preproteins with N-terminal cleavable transit peptides and are imported into the organelle through the TOC–TIC translocon system. Import involves a complex set of recognition and membrane translocation steps that ensure the fidelity and unidirectional transport of the polypeptide across the double-membrane chloroplast envelope. To understand the mechanism of import, the molecular interactions and energetics of each step must be defined. Here, we describe the methods for capturing intermediates in the import process through the manipulation of the energy state of chloroplasts, and the use of two different chemical cross-linking approaches to examine the molecular interactions that mediate the import process and to assess the assembly state of the translocons. These approaches can be employed to identify sequential protein–protein interactions, and thereby dissect the pathway and roles of import components during protein import into chloroplasts. Key word: Chloroplast, Import, TOC, TIC, GTP, ATP, Cross-link, Early import intermediate

1. Introduction The biogenesis of chloroplasts relies on the import of thousands of nucleus-encoded proteins from the cytoplasm (1). Most chloroplast proteins are synthesized as preproteins with N-terminal cleavable transit peptides. Multimeric translocons embedded in the outer (TOC) and inner (TIC) envelope membranes (2, 3) mediate the recognition of preproteins at the chloroplast surface and their translocation across double-membrane envelope. The core of the TOC translocon consists of two receptor GTPases, Toc159 and Toc34, which associate with a membrane channel Toc75. The TOC

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receptors are required for transit peptide recognition at the ­organelle surface, and they initiate the import process by transferring the ­preprotein to the protein-conducting channel. Components of the TIC complex (Tic20/21, Tic22, and Tic110) associate with the early import intermediates that are inserted into the TOC ­channel (4–7), and inner membrane translocation is powered by a proposed chaperone motor complex, containing Tic40, Hsp93, and Hsp70, at the trans side of the inner envelope membrane (8, 9). A number of discernable steps in import have been defined by manipulating the temperature and energy state of chloroplasts ­during in vitro protein import reactions. The most detailed analysis of the energetics of import and the corresponding intermediates in preprotein translocation has been performed with pea chloroplasts (10). Although these intermediates likely represent basic mechanistic steps that also apply to Arabidopsis, only four discernable steps in import have been examined in Arabidopsis chloroplasts. These include (1) an initial, reversible binding step that is observed in the absence of NTPs and corresponds to the first interaction of the transit peptide with the TOC translocon, (2) a GTP-stimulated binding that has been observed using TOC receptor GTPase mutants, (3) ATP-dependent insertion of the preprotein into the TOC channel (early import intermediate), and (4) subsequent ATP-driven translocation across the inner membrane via the TIC channel and associated molecular chaperones. Although the core components of the TOC and TIC translocons and specific steps in import have been identified (5, 11, 12), defining the molecular mechanism of import requires knowledge of the molecular interactions between the preprotein and import components at each stage. Key questions include the following: (1) what is the molecular basis of transit peptide recognition by the TOC receptors, (2) how do the intrinsic GTPase cycles of the receptors control the transition from initial binding to membrane translocation and thereby regulate the fidelity of import, (3) what is molecular nature of each membrane channel and how are they coupled to provide a continuous translocation pathway, and (4) how is the system of molecular chaperones that associates with the import apparatus organized to provide the unidirectional driving force for translocation? The availability of well-defined import intermediates, coupled with the ability to generate specific mutants in Arabidopsis, now provides a powerful system to gain insight into the molecular details of the import reaction. Recently, we generated Arabidopsis transgenic plants expressing a mutated form of Toc159 with significantly reduced GTPase activity to examine the role of receptor GTPase activity in protein import (13). To characterize the state of import intermediates in the GTPase mutant, we developed chemical cross-link approaches to map preprotein–translocon interactions (13). In a second approach, we used an independent covalent

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cross-linking approach to determine the effects of mutations in Tic110 on the dynamic assembly of protein import complexes (4). In this chapter, we will focus on methods that couple the generation of import intermediates in wild-type and mutant chloroplasts with covalent cross-linking to examine the dynamic assembly of import complexes and identify the roles of core TOC/TIC components at each stage in import.

2. Materials 2.1. Isolation of Chloroplasts from Arabidopsis thaliana

1. MS plates: 2-(4-morpholino)ethane sulfonic acid (MES)KOH, pH 5.7, 0.5× Murashige and Skoog (MS) growth medium, 1% (w/v) sucrose, and 0.8% (w/v) Phytoagar. 2. 150 × 15 mm Petri plates (Falcon). 3. Percoll (GE Healthcare). 4. Miracloth (Calbiochem). 5. Protease Inhibitor Cocktail P9599 (PIC) (Sigma–Aldrich). 6. Ultra-Turrax T 25 (IKA Labortechnik) with a 15-mm probe (or comparable). This is a handheld rotor-stator homogenizer. Similar models produced by Polytron can also be used. 7. Grinding buffer (GB): 50 mM N-(2-hydroxyethyl)piperazineN¢-(2-ethanesulfonic acid) (HEPES)-KOH, pH 7.5, 330 mM sorbitol, 2  mM ethylenediaminetetraacetic acid (EDTA), 1 mM MnCl2, and 1 mM MgCl2. The following reagents are added immediately prior to use: 100 mM ascorbic acid, 0.25% (w/v) bovine serum albumin (BSA), and 0.05% (v/v) PIC. 8. 40% Percoll: 40% (v/v) Percoll, 50  mM HEPES-KOH, pH 7.5, 330 mM sorbitol, 2 mM EDTA, 1 mM MnCl2, and 1 mM MgCl2. Store at −20°C. The following reagents are added immediately prior to use: 0.25% (w/v) BSA, 100 mM ascorbic acid, and 0.05% (v/v) PIC. 9. 85% Percoll: 85% (v/v) Percoll, 50  mM HEPES-KOH, pH 7.5, 330 mM sorbitol, 2 mM EDTA, 1 mM MnCl2, and 1 mM MgCl2. Store at −20°C. The following reagents are added immediately prior to use: 0.25% (w/v) BSA, 100 mM ascorbic acid, and 0.05% (v/v) PIC. 10. Two Percoll step gradients: 1  mL of 85% Percoll overlayed with 4 mL of 40% Percoll in 15 round-bottomed 15-mL glass Corex centrifuge tubes. 11. HS buffer: 50  mM HEPES-KOH, pH 7.5, and 330  mM ­sorbitol. Store at −20°C. 12. 80% (v/v) acetone.

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13. 250-mL Nalgene polypropylene screw-capped centrifuge bottles. 14. Sorvall RC-5 superspeed centrifuge or comparable. 15. Sorvall GSA fixed-angle and HB-4 swinging bucket centrifuge rotors or comparable. 16. Microcentrifuge. 17. Vortex mixer. 2.2. Energy Depletion of Isolated Chloroplasts

1. Nigericin (Calbiochem). This is dissolved in 80% (v/v) ethanol to a concentration of 40 mM. Store at −20°C. 2. Apyrase (Sigma–Aldrich). This is dissolved in HS buffer to a concentration of 100 u/mL (units/microliter). Store at −80°C. 3. Glycerate (Sigma–Aldrich). This is dissolved in HS buffer to a concentration of 100 mM immediately prior to use.

2.3. Preparation of preSSU by Coupled In Vitro Transcription/ Translation

1. The cDNA encoding the precursor to the small subunit of ribulose-1,5-bisphosphate carboxylase/oxygenase (preSSU) from Arabidopsis (At5g38430) cloned downstream of the T7 polymerase promoter in the pET21a plasmid. 2. TNT-coupled reticulocyte lysate system (Promega). 3. 10 mCi/mL [35S]-methionine (Perkin Elmer). 4. 40  u/mL (units/microliter) RNase inhibitor (New England Biolabs). 5. Hydrated Sephadex G-25 (Pharmacia Biotech Inc.). This is equilibrated with HS buffer prior to use.

2.4. Generation of Protein Import Intermediates and Chemical CrossLinking to Map Translocon–Preprotein Interactions

1. 10× MMX (chloroplast import master mix): 250 mM potassium acetate (KOAc) and 50 mM magnesium acetate (MgOAc). Store at −20°C. 2. 35% Percoll: 35% (v/v) Percoll, 330 mM sorbitol, and 50 mM HEPES-KOH, pH 7.5. Store at −20°C. 3. ATP-NaOH, pH 7.5, is dissolved in HS buffer to a final ­concentration of 100 mM and stored at −20°C. 4. GTP-NaOH, pH 7.5, is dissolved in HS buffer to a final ­concentration of 100 mM and stored at −20°C. 5. In vitro-translated [35S]preSSU (see Subheading 3.3). 6. SDS-PAGE sample buffer: 350  mM Tris base, 5% (w/v) sodium dodecyl sulfate (SDS), 80 mM dithiothreitol (DTT), 7.5% (v/v) glycerol, and 0.1% saturated bromophenol blue. Prepare immediately prior to use. 7. Dithio-bismaleimidoethane (DTME, Thermo Fisher Scientific). This is dissolved in dimethylsulfoxide (DMSO) to a final concentration of 50 mM immediately prior to use. 8. 2% (w/v) SDS.

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9. Dilution buffer: 50 mM Tris–HCl, pH 7.5, 1.1% (w/v) Triton X-100, 167  mM NaCl, 5.6  mM EDTA, and 1% (v/v) PIC. Prepare immediately prior to use. 10. Protein-A Sepharose CL-4B (GE Healthcare) is equilibrated with dilution buffer prior to use. 11. Washing buffer A: 50 mM HEPES-KOH, pH 7.5, 0.1% (w/v) Triton X-100, 150  mM NaCl, and 0.1% (v/v) PIC. Prepare immediately prior to use. 12. Microcentrifuge. 13. Phosphorimager. 2.5. Chemical CrossLinking to Map Translocon Assembly and Dynamics

1. 50  mM dithiobis[succinimidyl propionate] (DSP) in DMSO (Thermo Fisher Scientific). Prepare immediately prior to use. 2. TES buffer: 50  mM N-(Tris[hydroxymethyl]methyl)glycine (Tricine)-KOH, pH 7.5, 2  mM EDTA, 150  mM NaCl, 1% (w/v) Triton X-100, and 1% (v/v) PIC. Prepare immediately prior to use. 3. Protein-A Sepharose CL-4B (GE Healthcare) equilibrated with TES buffer. Prepare immediately prior to use. 4. Washing buffer B: 50  mM Tricine-KOH, pH 7.5, 2  mM EDTA, 150 mM NaCl, 0.1% (w/v) Triton X-100, and 0.1% PIC. Prepare immediately prior to use. 5. Microcentrifuge.

3. Methods 3.1. Isolation of Chloroplasts from Arabidopsis thaliana

Traditionally, pea chloroplasts have been used as a model system for assaying chloroplast preprotein import. Peas offer the advantages of large amounts of material that can be grown quickly and easily, and rapid and simple isolation of intact, import-competent chloroplasts (14, 15). However, the tools available in Arabidopsis thaliana for the investigation of genetic, biochemical, and cellular processes make it desirable to use this model system to analyze chloroplast targeting (4, 16, 17). The small size of Arabidopsis and the need to use young rapidly growing tissues for chloroplast isolation present challenges in adapting the Arabidopsis system for the in vitro analysis of chloroplast import. A number of methods for chloroplast isolation from Arabidopsis have been published, but many of them require expensive and time-consuming methods, or result in chloroplasts with low levels of import activity. We have found that the following method adapted from two previously published ­procedures (18, 19) provides excellent yields of highly importactive chloroplasts (13). This method also significantly reduces

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degradation of the Toc159, a common problem with other ­methods used for isolation of Arabidopsis and pea chloroplasts. 1. Spread sterilized Arabidopsis seeds on MS plates (0.1 g seeds per plate) and incubate the plates at 4°C for 24–48 h. Transfer the plates to a growth chamber (16-h day period, 22°C) and allow growth for 14 days (see Note 1). 2. Harvest plants by shaving the green shoots from the surface of the agar plates with a razor and place the shoots directly in 100 mL of ice-cold GB. Plants harvested from up to 2 plates (~10 g plant material wet weight) can be added to 100 mL of GB (see Note 2). 3. Homogenize the plant suspension for <1 min using a UltraTurrax T25 rotor-stator homogenizer with a 15-mm probe (or comparable) at 8,000 rpm. 4. Filter the solution by gravity through two layers of Miracloth into a 250-mL centrifuge bottle held on ice. 5. Centrifuge the solution for 8  min at 1,000 × g at 4°C in a Sorvall GSA fixed-angle rotor. Decant the supernatant and discard, taking care to leave the loose pellet behind. 6. Resuspend the pellet by slow swirling in 10 mL of fresh GB. Ensure that the suspension remains cold and that no large clumps remain. Do not shake or introduce air bubbles. 7. Layer one-half of the suspension onto one each of two Percoll step gradients. Use a large bore pipette to transfer all solutions containing plant extracts to avoid sheering the chloroplasts (see Note 3). 8. Centrifuge the gradients at 7,700 × g for 15 min at 4°C in an Sorvall HB-4 rotor using slow acceleration and deceleration (see Note 4). 9. Remove and discard the GB and the 40% Percoll layer by aspirating from the top of the gradient. Be careful not to disturb the 40–85% Percoll interface. This layer contains the intact chloroplasts. Collect the intact chloroplasts at the 40–85% Percoll interface by slow pipetting and transfer into 10 mL of cold HS buffer in a 15-mL glass Corex centrifuge tube. The layer of chloroplasts can be somewhat viscous. The chloroplasts can be washed out of the pipette by very slowly pipetting up and down with HS buffer. 10. Mix the solution by covering the tube with Parafilm and gently inverting twice. Centrifuge the suspension at 1,000 × g for 6  min at 4°C in an Sorvall HB-4 swinging bucket rotor. Carefully discard the supernatant. Take care not to pour off the pellet because the pellet is very loose. 11. Completely resuspend the pellet by gentle swirling with 400 mL of cold HS buffer. Do not shake vigorously or use a pipette.

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12. Quantify the yield of chloroplasts by measuring the ­chlorophyll content. Dilute 5  mL of the chloroplast suspension into 995 mL of 80% acetone. Mix the acetone solution vigorously using a vortex mixer and centrifuge for 1  min at maximum speed at room temperature in a microcentrifuge to remove cell debris. The absorbance of the supernatant is measured at 652 nm and chlorophyll content measured by the following formula: Chlorophyll (in mg/mL) is calculated as (A652/36) × 200 (dilution factor). Dilution factor = total volume/ volume sample added. 13. Once the chlorophyll content has been determined, the purified chloroplasts are diluted to a final concentration of 1 mg chlorophyll/mL with HS buffer and kept on ice in a covered container until use (see Note 5). 3.2. Energy Depletion of Isolated Chloroplasts

To control the energy state of in vitro import reactions, it is essential to deplete chloroplasts of endogenous ATP pools, preclude light-driven ATP synthesis, and remove external GTP. In general, we have found incubation of isolated chloroplasts in the dark at room temperature for a short period sufficient to deplete endogenous ATP for assays to generate early import intermediates. In cases where it is desirable to examine very detailed steps in import (10, 20), chloroplasts are further treated with nigericin, an ionophore that dissipates the light-generated proton gradient at thylakoid membranes (21), and with glycerate to ensure the depletion of existing stromal ATP. Glycerate is the substrate for glycerate kinase (22, 23). To examine energy-independent binding of preproteins to TOC translocons, apyrase is added to degrade external NTPs to NDPs and NMPs (23). 1. Isolated chloroplasts (see Subheading 3.1) are incubated in the dark for 10 min at 23–26°C to deplete internal ATP prior to use in the generation of all import intermediates. 2. To maximize ATP depletion of isolated chloroplasts (see Subheading 3.1), glycerate is added to a final concentration of 10 mM (22) and nigericin to a final concentration of 400 nM prior to the incubation in step 1 above. 3. If chloroplasts are to be used to measure energy-independent preprotein binding, apyrase (2  units/100  mL) is added to the isolated chloroplasts prior to the incubation in step 1 above.

3.3. Preparation of preSSU by Coupled TranscriptionTranslation

[35S]-methionine-labeled in  vitro-translated preproteins are synthesized using cDNAs cloned into plasmids downstream of a T7 RNA polymerase promoter. The pET21 series of vectors are convenient as they can be used for both in vitro translation and E. coli expression. A commercially available reticulocyte lysate system

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(Promega) allows the in vitro synthesis of the protein by coupling transcription and translation. The reaction is carried out in the presence of a radioactive [35S] methionine in order to label the newly synthesized protein for detection by phosphorimaging or autoradiography. Both the wheat germ and the rabbit reticulocyte lysate systems have been used for protein synthesis and import into chloroplasts. We routinely use the rabbit reticulocyte lysate system because the yield and quality of the translation products exceed those synthesized using the wheat germ system. The protocol described here is according to the manufacturer’s instructions for the TNT-coupled rabbit reticulocyte lysate system (Promega) and previous publications (14). We and others have used preSSU as the model substrate for binding and import experiments. 1. Rapidly thaw all transcription-translation reagents and place on ice immediately before performing the reaction. 2. Place a 1.5-mL microcentrifuge tube on ice and add the following reagents in the following order: 25  mL TNT rabbit reticulocyte lysate, 2 mL TNT reaction buffer, 4 mL [35S] methionine (1,000 Ci/mmol at 10 mCi/mL), 1 mL T7 TNT RNA polymerase, 1 mL amino acid mix minus methionine, and 1 mL RNase inhibitor. 3. Add 1 mg of pET21a-preSSU plasmid (4). The volume of plasmid should be kept to a minimum (£1 mL). (For isolation of plasmid, see Note 6). 4. Add nuclease-free water to bring the final volume to 50 mL. 5. Incubate the reaction at 30°C for 90 min without shaking. 6. After the translation reaction, the translation mixture is diluted with 450  mL of ice-cold HS and gel-filtered using Sephadex G-25 to remove free nucleotides. (For the method for gel filtration, see Note 7). Alternatively, free nucleotides can be removed free by dialysis (24) (see Note 8). 3.4. Generation of Protein Import Intermediates and Chemical CrossLinking to Map Translocon–Preprotein Interactions

To examine the components involved in preprotein binding and the formation of early import intermediates, we have employed a chemical cross-linking approach using the homobifunctional cysteine cross-linker, DTME (13). preSSU contains five cysteines at positions 55, 96, 132, 145, and 167. After the cross-linking reaction, detergent-solubilized chloroplasts are immunoprecipitated with antibodies to TOC/TIC components to identify components that interact with the preprotein at each stage. The results of a typical cross-linking reaction in which preSSU is bound to chloroplasts in the presence of GTP is shown in Fig.  1. Immunoprecipitation of the samples with anti-Toc159 or antiToc33 antibodies under denaturing conditions demonstrates that the method can distinguish interactions between preSSU and the two TOC receptors under these conditions.

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Fig. 1. GTP-dependent binding of preSSU to components of the TOC translocons. Energydepleted, in vitro-translated [35S]preSSU was incubated with isolated chloroplasts in the presence of 0.1 mM GTP to promote GTP-dependent binding. The homobifunctional crosslinker DTME was added to cross-link the bound preprotein to TOC components. The ­chloroplasts were detergent solubilized under denaturing conditions and proteins immunoprecipitated with Toc159 or Toc33 affinity-purified antibodies. The immuno­ precipitates were analyzed by SDS-PAGE and phosphorimaging to quantify co-­ immunoprecipitated [35S]preSSU or by immunoblotting to detect Toc159 or Toc33. 3.4.1. Generation of Protein Import Intermediates

1. Energy-independent binding: Dilute isolated, energy-depleted chloroplasts corresponding to 200  mg of chlorophyll (see Subheading 3.2, step 2) with HS buffer and 0.1 volume 10× MMX (50 mL) to a final volume of 500 mL. Initiate the binding reaction by adding 10–50 mL of energy-depleted in vitro translation products (equivalent to at least 25,000 cpm) in HS buffer and incubate at 26°C for 5 min in the dark (14). 2. GTP-stimulated binding: Dilute ATP-depleted chloroplasts corresponding to 200 mg of chlorophyll (see Subheading 3.2, step 1) with HS buffer and 0.1 volume 10× MMX (50 ml) to a final volume of 500 mL. Add GTP to the chloroplast suspension at a final concentration of 0.1 mM and incubate the mixture at 26°C for 5 min in the dark. Initiate the binding reaction by adding 10–50  mL of energy-depleted in  vitro translation products (equivalent to at least 25,000 cpm) in HS buffer and incubate at 26°C for 5 min in the dark (14). 3. Early import intermediate: Dilute ATP-depleted chloroplasts corresponding to 200  mg of chlorophyll (see Subheading  3.2, step 3) with HS buffer and 0.1 volume 10× MMX (50 mL) to a final volume of 500 mL. Add GTP to the chloroplast suspension to a final concentration of 0.1 mM and ATP to a final concentration of 0.1 mM. Incubate the mixture at 26°C for 5 min in the

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3.4.2. Chemical Crosslinking to Map Translocon– Preprotein Intermediates

dark. Initiate the reaction by adding 10–50 mL of energy-depleted in vitro translation products (equivalent to at least 25,000 cpm) in HS buffer and incubate at 26°C for 5 min in the dark. 1. Initiate cross-linking in energy-independent and ­GTP-dependent binding reactions (see Subheading  3.4.1) with the addition of 2.5 mM DTME. 2. For the early import intermediate (see Subheading 3.4.1), the chloroplasts are isolated through a 35% Percoll cushion to remove excess translation product prior to initiating cross-linking. Layer the reaction over 1 mL of 35% Percoll cushion in HS buffer in a microcentrifuge tube. Centrifuge the sample for 1 min on low speed in the microcentrifuge. Wash the chloroplast pellet twice with 1 mL of HS buffer. Resuspend the chloroplasts in 475 mL of HS buffer and add 25 mL of DTME to a final concentration of 2.5 mM. 3. Incubate the cross-linking reactions on ice in the dark for 45 min. 4. Dilute the reactions tenfold with ice-cold HS buffer, collect the chloroplasts by centrifugation for 1 min at high speed in a microcentrifuge, and wash once with HS buffer before immunoprecipitation.

3.4.3. Immunoprecipitation of Cross-Linked Import Intermediates

1. For immunoprecipitation under denaturing conditions, dissolve chloroplasts (150 mg chlorophyll) (see Subheading 3.4.2) in 80 mL of 2% SDS containing 2% PIC by vigorous mixing. 2. Heat the samples for 5  min at 97°C and cool to room temperature. 3. Dilute the samples with 720 mL of dilution buffer and incubate on ice for 5 min. 4. After a 30-min centrifugation at 18,000 × g at 4°C, divide the supernatant into two equal volumes and incubate with 2 mg/ mL affinity-purified Toc159 A-domain antibody or Toc33 antibody for 16 h at 4°C. 5. Add 20  mL of packed protein A–Sepharose resin in dilution buffer to each sample and incubate for 2 h at 4°C with ­constant rotation. 6. Wash the Sepharose five times with washing buffer A and ­analyze by SDS-PAGE. 7. The amounts of precipitated receptors and bound preSSU are determined by immunoblotting and phosphorimaging, respectively (Fig. 1).

3.5. Chemical CrossLinking to Map Translocon Assembly and Dynamics

To examine the translocon assembly and dynamics, we have employed a chemical cross-linking approach using the homobifunctional amine cross-linker, DSP (4, 5, 25). Although the interactions of several of the core TOC–TIC components (Toc75, Toc159,

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Toc33, and Tic110) appear to be quite stable, the ­association of these proteins with other components of the import apparatus (e.g., the chaperone machinery) is dynamic. As a result, detection of these interactions requires stabilization with covalent cross-linkers. After the cross-linking reaction, detergent-solubilized chloroplasts are immunoprecipitated with antibodies to atTic110 to identify components that interact with atTic110. An example of the stabilization of the Tic40-Hsp93 motor complex with Tic110 using DSP crosslinking is shown in Fig. 2. DSP has the advantage of containing a reducible disulfide that allows cleavage of the cross-links for resolution of individual polypeptides after immunoprecipitation. 1. For cross-linking, isolated chloroplasts (see Subheading  3.1) are incubated in the dark for 30 min at 26°C to deplete internal ATP and reducing potential. If energy-dependent interactions are of interest, the energy state of the chloroplasts can be adjusted accordingly by the exogenous addition of nucleoside triphosphates (NTPs) (see Subheading 3.4.1). 2. Resuspend chloroplasts corresponding to 200  mg of chlorophyll in 475 mL of HS buffer and add 25 mL of DSP to a final concentration of 0.25–1 mM on ice for 15 min in the dark. For the stabilization of Tic40 and Hsp93 with the translocons, we use a maximum of 1 mM DSP.

Fig.  2. Stabilization of the association of Hsp93 and Tic40 with TOC–TIC complexes. Detergent extracts from isolated chloroplasts that had been treated with DSP at the concentrations indicated (lanes 2–4) or DMSO as a control (lane 1) were subjected to immunoprecipitation with anti-Tic110 antibodies. The immunoprecipitates were immunoblotted with the antibodies indicated at the right of the figure.

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3. Quench the reaction by incubating with glycine at a final concentration of 50 mM on ice for an additional 15 min. 4. Collect the chloroplasts by centrifugation for 1  min at high speed in a microcentrifuge and wash the pellet once with HS buffer before fractionation. 5. Lyse the chloroplasts by suspension in HS buffer to a concentration of 0.5–1 mg chlorophyll/mL and dilute with five volumes of 2 mM EDTA to induce osmotic lysis. Mix the lysate vigorously and incubate on ice for 10 min. 6. Collect the membrane fraction by centrifugation at 18,000 × g for 30 min at 4°C in a microcentrifuge. 7. For immunoprecipitation, dissolve the total membranes (200 mg of chlorophyll) from DSP cross-linked chloroplasts in 200 mL TES buffer for 10 min on ice. 8. After a 30-min centrifugation at 18,000 × g at 4°C in a microcentrifuge, incubate the supernatant with 2  mg/mL affinitypurified Tic110 antibody for 16 h at 4°C. 9. Add 20 mL of packed protein A–Sepharose resin in TES buffer to each sample and incubate for 2  h at 4°C with constant rotation. 10. Wash the Sepharose five times with washing buffer B and elute the bound proteins directly with 50 mL of SDS-PAGE buffer. 11. Analyze the immunoprecipitates by SDS-PAGE and immunoblot with the antibodies to TOC/TIC components shown in Fig. 2.

4. Notes 1. Although chloroplasts can be isolated from older plants to ­provide higher yields, the protein import activity decreases with the age of the plants. We have found that 14-day-old plants grown under these conditions yield reasonable amounts of highly import-active chloroplasts. 2. Temperature and the time of isolation are the most important factors that will determine the quality of the isolated chloroplasts. It is essential that all reagents for the isolation be kept ice-cold and that all procedures are performed on ice or in a cold room. The chloroplast isolation procedure should not take >1 h from the time of tissue homogenization to isolation of the intact chloroplasts. 3. High acceleration/deceleration rates disrupt step gradients. After centrifugation of the Percoll step gradients, two distinct bands of very dark green material will be obvious at the GB–40% Percoll interface (broken chloroplasts and cell debris) and the 40–85% Percoll interface (intact chloroplasts).

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4. Larger volume gradients can be used if more than five plates of plants are to be processed. This will give a better resolution of fractions on the step gradients. To scale up the Percoll gradients, layer 10 mL of 40% Percoll over 5 mL of 85% Percoll in a 30-mL round-bottomed centrifuge tube (e.g., Corex 30-mL centrifuge tube). The gradients can be prepared in advance and kept on ice before harvesting the plants. 5. The typical yield of chloroplasts from two plates of 8–10 g tissue is 200–400 mg chlorophyll. 6. To isolate plasmids for coupled transcription-translation, we typically use a commercially available plasmid isolation kit (Qiagen Plasmid Midi kit). This system invariably results in high-quality plasmid lacking contaminants that can interfere with the transcription or translation reactions. 7. A volume of 1 mL of Sephadex G-25 reconstituted in water is pipetted into the barrel of a 1-cc syringe (Becton, Dickinson & Company). The tip is plugged with glass wool to allow solutions but not the Sephadex from passing through the syringe. The syringe is inserted into a 15-mL conical centrifuge tube and the Sephadex is washed by the application of five volumes of HS buffer. The column is centrifuged for 1 min at 1,000 × g in a tabletop clinical centrifuge between each application of buffer to force out the excess liquid. A microcentrifuge tube with cap removed is placed at the bottom of the 15-mL tube and the syringe inserted so that the tip is inserted into the microfuge tube. The translation mixture is layered onto the Sephadex and the syringe is centrifuged for 1 min at 1,000 × g at 4°C. The filtered translation mix is recovered in the microfuge tube and can be stored at −80°C until use for at least 2 weeks. 8. As an alternative method to gel filtration for the removal of free nucleotides, diluted translation mixtures can be dialyzed in 10,000 molecular weight cut-off dialysis tubing (SpectroPor 1 or 2) and incubated with 100 volumes of HS buffer at 4°C. The buffer should be changed two times with fresh HS buffer every 1 h and the dialysis should not exceed 4 h. References 1. Jarvis, P. (2004) Organellar proteomics: chloroplasts in the spotlight. Curr. Biol. 14, R317–R319. 2. Jarvis, P. (2008) Targeting of nucleus-encoded proteins to chloroplasts in plants. New Phytol. 179, 257–285. 3. Inaba, T., and Schnell, D. J. (2008) Protein trafficking to plastids: one theme, many variations. Biochem. J. 413, 15–28.

4. Inaba, T., Alvarez-Huerta, M., Li, M., Bauer, J., Ewers, C., Kessler, F., and Schnell, D. J. (2005) Arabidopsis Tic110 is essential for the assembly and function of the protein import machinery of plastids. Plant Cell 17, 1482–1496. 5. Chen, K. Y., and Li, H. M. (2007) Precursor binding to an 880-kDa Toc complex as an early step during active import of protein into chloroplasts. Plant J. 49, 149–158.

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6. 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. 7. Kouranov, A., Chen, X., Fuks, B., and Schnell, D. J. (1998) Tic20 and Tic22 are new components of the protein import apparatus at the chloroplast inner envelope membrane. J. Cell Biol. 143, 991–1002. 8. Shi, L. X., and Theg, S. M. (2010) a stromal heat shock protein 70 system functions in protein import into chloroplasts in the moss Physcomitrella patens. Plant Cell 22, 205–220. 9. Kovacheva, S., Bedard, J., Patel, R., Dudley, P., Twell, D., Rios, G., Koncz, C., and Jarvis, P. (2005) In vivo studies on the roles of Tic110, Tic40 and Hsp93 during chloroplast protein import. Plant J. 41, 412–428. 10. Inoue, H., and Akita, M. (2008) Three sets of translocation intermediates are formed during the early stage of protein import into chloroplasts. J. Biol. Chem. 283, 7491–7502. 11. Schnell, D. J., Kessler, F., and Blobel, G. (1994) Isolation of components of the chloroplast protein import machinery. Science 266, 1007–1012. 12. Kouranov, A., and Schnell, D. J. (1997) Analysis of the interactions of preproteins with the import machinery over the course of protein import into chloroplasts. J. Cell Biol. 139, 1677–1685. 13. Wang, F., Agne, B., Kessler, F., and Schnell, D. J. (2008) The role of GTP binding and hydrolysis at the atToc159 preprotein receptor during protein import into chloroplasts. J. Cell Biol. 183, 87–99. 14. Smith, M. D., Fitzpatrick, L. M., Keegstra, K., and Schnell, D. J. (2002) In vitro analysis of chloroplast protein import. In, Current Protocols in Cell Biology (Bonifacino, J. S., Dasso, M., Harford, J. B., Lippincott-Schwartz, J., and Yamada, K. M., eds.) John Wiley and Sons, Inc., New York, USA, pp. 11.16.11-11.16.21. 15. Chua, N. H., and Schmidt, G. W. (1978) Posttranslational transport into intact chloroplasts of a precursor to the small subunit of ribulose1,5-bisphosphate carboxylase. Proc. Natl. Acad. Sci. USA 75, 6110–6114. 16. Constan, D., Patel, R., Keegstra, K., and and Jarvis, P. (2004) An outer envelope membrane

component of the plastid protein import ­apparatus plays an essential role in Arabidopsis. Plant J. 38, 93–106. 17. Kubis, S., Patel, R., Combe, J., Bedard, J., Kovacheva, S., Lilley, K., Biehl, A., Leister, D., Rios, G., Koncz, C., and Jarvis, P. (2004) Functional specialization amongst the Arabidopsis Toc159 family of chloroplast protein import receptors. Plant Cell 16, 2059–2077. 18. Brock, I. W., Hazell, L., Michl, D., Nielsen, V. S., Moller, B. L., Herrmann, R. G., Klosgen, R. B., and Robinson, C. (1993) Precursors of one integral and five lumenal thylakoid proteins are imported by isolated pea and barley thylakoids: optimisation of in vitro assays. Plant Mol. Biol. 23, 717–725. 19. Schulz, A., Knoetzel, J., Scheller, H. V., and Mant, A. (2004) Uptake of a fluorescent dye as a swift and simple indicator of organelle intactness: import-competent chloroplasts from soilgrown Arabidopsis. J. Histochem. Cytochem. 52, 701–704. 20. Akita, M., and Inoue, H. (2009) Evaluating the energy-dependent “binding” in the early stage of protein import into chloroplasts. Meth. Enzymol. 466, 43–64. 21. Theg, S. M., Bauerle, C., Olsen, L. J., Selman, B. R., and Keegstra, K. (1989) Internal ATP is the only energy requirement for the translocation of precursor proteins across chloroplastic membranes. J. Biol. Chem. 264, 6730–6736. 22. Olsen, L. J., Theg, S. M., Selman, B. R., and Keegstra, K. (1989) ATP is required for the binding of precursor proteins to chloroplasts. J. Biol. Chem. 264, 6724–6729. 23. Olsen, L. J., and Keegstra, K. (1992) The binding of precursor proteins to chloroplasts requires nucleoside triphosphates in the intermembrane space. J. Biol. Chem. 267, 433–439. 24. Lee, J., Wang, F., and Schnell, D. J. (2009) Toc receptor dimerization participates in the initiation of membrane translocation during protein import into chloroplasts. J. Biol. Chem. 284, 31130–31141. 25. Akita, M., Nielsen, E., and Keegstra, K. (1997) Identification of protein transport complexes in the chloroplastic envelope membranes via chemical cross-linking. J. Cell Biol. 136, 983–994.

Chapter 19 Isolation of Arabidopsis Thylakoid Membranes and Their Use for In Vitro Protein Insertion or Transport Assays Thomas Bals and Danja Schünemann Abstract This chapter focuses on the techniques of chloroplast isolation; their fractionation into envelopes, stroma, and thylakoids; and their further use for in vitro protein transport assays. In addition to the isolation of thylakoids, this chapter also describes the experimental steps of both protein translocation across the thylakoid membrane and protein integration into the membrane. Protein translocation and integration can be analysed by the radioactive labelling of substrate proteins using an in vitro transcription and translation system. The translocated or integrated proteins can then be detected by autoradiography. Our protocol allows the analysis of these transport systems in wild-type Arabidopsis or mutants that lack or overexpress soluble or membrane transport factors that could be of potential interest. Key words: Chloroplast isolation, Thylakoid membrane, Protein transport, cpSec, cpTat, cpSRP, In vitro transcription and translation, Protein translocation assay, Protein integration assay

1. Introduction Thylakoid membranes of chloroplasts contain essential protein complexes that play a crucial role in the process of photosynthesis. The biogenesis of these multi-subunit complexes requires coordinated protein synthesis, targeting, and assembly of both nucleusencoded and chloroplast-encoded proteins. The vast majority of thylakoid proteins are encoded in the nucleus, synthesised as precursors in the cytoplasm, and post-translationally imported into the chloroplast. In the stroma, precursor proteins become proteolytically processed by a stromal peptidase. Imported proteins are further directed to the internal thylakoid membrane system. To date, four different sorting mechanisms of nucleus-encoded thylakoid proteins have been described (1–3). The cpSec pathway

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and the cpTat pathway are mainly involved in the transport and translocation of lumenal proteins. The insertion of nucleus-encoded proteins into the thylakoid membrane is mediated via the chloroplast signal recognition particle (cpSRP) pathway and the spontaneous pathway (Fig.  1). All known lumenal proteins contain a bipartite transit sequence, which codes for the transport across the chloroplast envelopes and additional translocation across the thylakoid membrane. After translocation into the thylakoid lumen, the transit sequence is cleaved off by a thylakoidal processing peptidase. Most integral thylakoid membrane proteins that have been Translocation across the thylakoid membrane cpTat-pathway OE23

cpSec-pathway OE33

stroma PMF

ATP

thylakoid membrane

Hcf 106

cpSecA

Insertion into the thylakoid membrane cpSRP-pathway LHCP

spontaneous CFoII

cpSRP GTP cpFtsY

cp Tha4 TatC

cpSecE cpSecY

cpTat translocase

cpSec translocase

Alb3

LHCP

CFoII

lumen

OE23

OE33

Fig. 1. Targeting pathways for nucleus-encoded proteins to the thylakoids of chloroplasts. Four different targeting mechanisms have been described for the post-translational translocation of proteins from the stroma to the thylakoid membrane or across the membrane into the thylakoid lumen. The translocation of lumenal proteins is mediated by the cpSec pathway (e.g., OE33) and the cpTat pathway (e.g., OE23). Protein transport via the cpSec pathway depends on the stromal targeting factor cpSecA and ATP as energy source. The cpSec translocase is composed of the integral membrane proteins cpSecE and cpSecY. The protein translocation across the thylakoid membrane occurs in an unfolded state. In contrast to the cpSecdependent transport mechanism, the proteins of the cpTat pathway are transported in a folded state. The cpTat translocase comprises the integral membrane proteins Hcf106, cpTatC, and Tha4, whereas no stromal targeting factors are required. Instead, it has been shown that transport into the lumen is dependent on a proton motive force (PMF). The insertion of nucleus-encoded proteins into the membrane occurs by the cpSRP pathway and by spontaneous insertion. The integration of members of the LHCP family is facilitated by the stromal cpSRP complex, the cpSRP receptor cpFtsY, GTP as an energy source, and the integral membrane translocase Alb3. In contrast, the spontaneous pathway (e.g., the integration of CF0II) requires no protein assistance or energy components. The N-terminal targeting domains that direct the proteins to the indicated pathways are marked by bold lines.

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analysed are either inserted spontaneously, without the assistance of other protein components and energy requirements, or directed to the membrane by the cpSRP transport machinery. Protein translocation across the thylakoid membrane or ­integration into the membrane requires the assistance of different stromal targeting factors, protein translocases, and energy ­components. The different transport pathways have been further investigated with reconstituted translocation and integration assays using isolated thylakoid membranes, stromal extracts, and radiolabelled substrate proteins generated in cell-free translation extracts. The following text describes the individual, experimental steps for reconstitution assays of protein translocation across the thylakoid membrane and assays of protein integration into the thylakoid membrane. Firstly, the protein substrates for each transport pathway have to be prepared by in vitro translation coupled with radioactive labelling. The radiolabelled substrates can then be analysed and distinguished from endogenous proteins by autoradiography. In the next step, chloroplasts are freshly isolated and further fractionated into thylakoids and stroma. The thylakoids contain the membrane-embedded translocases, whereas the stromal extract provides the stromal targeting factors. In this chapter, the translocation of the 33-kDa oxygen-­ evolving protein (OE33) and the 23-kDa oxygen-evolving protein (OE23) across the thylakoid membrane, as well as the integration of the light-harvesting chlorophyll-binding protein (LHCP; Lhcb1 isoform) into the membrane, is exemplarily described. To ensure the correct localisation of the test proteins, it is essential to use the precursor form of the substrate proteins, which includes the transit peptide that directs the protein to either the cpTat or the cpSec pathway. The LHCP contains an internal targeting sequence that directs it to the thylakoid membrane. The methods can be adapted to additional proteins to be tested in the translocation or integration assays.

2. Materials 2.1. Arabidopsis Thylakoid Isolation

1. 20  g freshly harvested rosette leaves (4–6 weeks old) of Arabidopsis thaliana (Col-0). Plants should be grown on soil with 8.5-h light (100 mE/m2/s) (22°C)/15.5-h dark (19.5°C) cycle. 2. Waring blender. 3. Plastic funnel. 4. 50/200-mL centrifugation tubes/centrifuge beakers with equivalent rotors (e.g., Beckman JA 25.50 and JLA 16.250). 5. Miracloth (Calbiochem).

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6. Soft paintbrush. 7. Spectrophotometer. 8. Homogenisation buffer (HB1): 50  mM 4-(2-hydroxyethyl) piperazine-1-ethanesulfonic acid (HEPES)-KOH, pH 8.0, 330  mM sorbitol, 10  mM ethylenedinitrilo tetraacetic acid (EDTA), 5 mM ascorbic acid, and 0.05% (w/v) bovine serum albumin (BSA). This should be freshly prepared. 9. SH buffer: 50 mM HEPES-KOH, pH 8.0, and 330 mM sorbitol. Store at −20°C. 10. HM buffer: 10 mM HEPES-KOH, pH 8.0, and 5 mM MgCl2. Store at −20°C. 2.2. Arabidopsis Chloroplast Isolation and Fractionation

1. 150  g freshly harvested rosette leaves (4–6 weeks old) of Arabidopsis thaliana (Col-0). Plants should be grown on soil with an 8.5-h light (100  mE/m2/s) (22°C)/15.5-h dark (19.5°C) cycle. 2. Waring blender. 3. Plastic funnel. 4. 200-mL centrifuge beaker with equivalent rotor (e.g., Beckman JLA 16.250). 5. Ultracentrifuge. 6. Ultracentrifuge beaker (12/40  mL) with equivalent rotors (e.g., Beckman SW 40/SW 28). 7. Miracloth (Calbiochem). 8. Soft paintbrush. 9. Spectrophotometer. 10. Vacuum pump. 11. Percoll (Sigma). 12. Homogenisation buffer (HB2): 450  mM sorbitol, 20  mM Tricine-KOH, pH 8.4, 10 mM EDTA, 10 mM NaCO3, and 0.1% (w/v) BSA. This should be freshly prepared. 13. Resuspension buffer (RB buffer): 300  mM sorbitol, 20  mM Tricine-KOH, pH 7.6, 5  mM MgCl2, and 2.5  mM EDTA. Store at −20°C. 14. TE buffer: 10 mM Tris–HCl, pH 8.0, and 1 mM EDTA. Store at room temperature. 15. HM buffer: 10 mM HEPES-KOH, pH 8.0, and 5 mM MgCl2. Store at −20°C.

2.3. Pea Chloroplast Isolation and Fractionation

1. 25  g freshly harvested pea leaves (Pisum sativum, Kelvedon Wonder) (9–10 days old). Plants should be grown on soil with an 8.5-h light (100 mE/m2/s) (22°C)/15.5-h dark (19.5°C) cycle. 2. Waring blender.

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3. Plastic funnel. 4. 50/200-mL centrifugation tubes/centrifuge beaker with equivalent rotors (e.g., Beckman JA 25.50 and JLA 16.250). 5. Miracloth (Calbiochem). 6. Soft paintbrush. 7. Spectrophotometer. 8. Vacuum pump. 9. 40% (v/v) Percoll in 50  mM HEPES-KOH, pH 8.0, and 330 mM sorbitol. Store at −20°C. 10. 80% (v/v) Percoll in 50  mM HEPES-KOH, pH 8.0, and 330 mM sorbitol. Store at −20°C. 11. Grinding buffer: 50 mM HEPES-KOH, pH 8.0, 330 mM sorbitol, 2 mM EDTA, 1 mM MgCl2, 1 mM MnCl, 0.05% (w/v) BSA, and 0.1% (w/v) ascorbic acid. Freshly prepared. 12. SH buffer: 50 mM HEPES-KOH, pH 8.0, and 330 mM sorbitol. Store at −20°C. 13. 10 mM HEPES-KOH, pH 8.0. Store at −20°C. 2.4. Measurement of Chlorophyll Concentration

1. Isolated chloroplasts/thylakoids. 2. 80% acetone (v/v) in distilled water. 3. Spectrophotometer. 4. Cuvette.

2.5. In Vitro Transcription and Translation

1. ATP, CTP, GTP, and UTP (each 100 mM, Fermentas). 2. Diguanosine triphosphate, sodium (GpppG, 25 U, Amersham). 3. 5× Transcription buffer (Fermentas). 4. Diethylpyrocarbonate (DEPC)-treated water (Fermentas). 5. RNase inhibitor (40 U/mL, Fermentas). 6. SP6 RNA polymerase (20 U/mL, Fermentas). 7. 10 M ammonium acetate. 8. Phenol (pH 4.0)/chloroform (1:1, v/v) mixture. 9. 100% ethanol. 10. 70% (v/v) ethanol. 11. Wheat germ extract (Promega). 12. Amino acid mixture (minus methionine) (1 mM each, Promega). 13. Radiolabelled amino acid [35S]-methionine (10 mCi/mL, 37 TBq/mmol) (Hartmann). 14. 1 M potassium acetate (Promega). 15. Water bath, set at 25°C.

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16. SpeedVac. 17. Vortex. 18. Heating blocks, set at 37, 68, and 98°C. 2.6. Thylakoid Translocation Assays

1. Arabidopsis thylakoids (1 mg chlorophyll/mL in HM buffer). 2. Pea stroma (equivalent to 3  mg chlorophyll/mL in HM buffer). 3. 100 mM ATP. 4. HM buffer: 10 mM HEPES-KOH, pH 8.0, and 5 mM MgCl2. Store at −20°C. 5. [35S]-radiolabelled in vitro translation products. 6. 1 mM carbonyl cyanide 3-chlorophenylhydrazone (CCCP) in 95% (v/v) ethanol. 7. Water bath (set at 25°C) with illumination (100 mmol/m2/s). 8. Thermolysin (10 mg/mL, Sigma). 9. 1 M CaCl2. 10. 0.5 M EDTA, pH 8.0. 11. Heating block, set at 98°C.

2.7. Thylakoid Integration Assay

1. Pea thylakoid membranes (1 mg chlorophyll/mL in SH buffer) 2. Pea stroma (equivalent to 3  mg chlorophyll/mL in 10  mM HEPES-KOH, pH 8.0). 3. 10 mM HEPES, pH 8.0. Store at −20°C. 4. SH buffer: 50 mM HEPES-KOH, pH 8.0, and 330 mM sorbitol. Store at −20°C. 5. 100 mM GTP. 6. Methionine (100 mM in 10 mM HEPES, pH 8.0). 7. Apyrase (1 U/mL in 10 mM HEPES, pH 8.0; Sigma). 8. [35S]-radiolabelled in vitro translation product. 9. Water bath, set at 25°C. 10. Thermolysin (10 mg/mL, Sigma). 11. 1 M CaCl2. 12. 0.5 M EDTA, pH 8.0. 13. 0.2 N NaOH. 14. Heating block, set at 98°C.

2.8. SDSPolyacrylamide Gel Electrophoresis

1. Hoefer Mighty Small minigel system (plate size: 10 × 8 cm). 2. 1.5 M Tris–HCl, pH 8.8. Store at room temperature. 3. 1 M Tris–HCl, pH 6.8. Store at room temperature. 4. 20% (w/v) sodium dodecyl sulfate (SDS). Store at room temperature.

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5. Acrylamide solution (30%), mix (29:1). Store at 4°C. 6. N,N,N¢,N¢-tetramethylethylenediamine (TEMED). Store at 4°C. 7. Ammonium persulphate (25% (w/v) stock solution in distilled water). Store in 200 mL aliquots at 4°C or at −20°C for longterm storage. 8. Stacking buffer (300 mL): 50 mL of 30% acrylamide solution (29:1), 1.5 mL of 20% (w/v) SDS, and 18.8 mL of 1 M Tris– HCl, pH 6.8. Store at 4°C. 9. Isopropanol. Store at room temperature. 10. 0.4% (w/v) bromophenol blue solution (in distilled water). Store in 200 mL aliquots at 4°C or at −20°C for long-term storage. 11. Running buffer (5×): 125 mM Tris (pH not adjusted), 960 mM glycine, and 0.1% (w/v) SDS. Store at room temperature. 12. Pre-stained molecular weight marker (e.g., Pre-stained Protein Ladders from Fermentas). 13. SDS sample buffer (4×): 200  mM Tris–HCl, pH 6.8, 40% (v/v) glycerol, 8% (w/v) SDS, 8% (v/v) b-mercaptoethanol, and 0.03% (w/v) bromophenol blue. Store in aliquots at 4°C or at −20°C for long-term storage. 2.9. Autoradiography

1. Amplify fluorographic reagent (Amersham). 2. Fixing solution (isopropanol:water:acetic acid, 25:65:10). 3. Whatman paper. 4. Plastic wrap. 5. Gel vacuum dryer. 6. X-ray film (Kodak). 7. Radiography cassette. 8. Developer and fixer solutions for developing autoradiographs in dark room.

3. Methods 3.1. Arabidopsis Thylakoid Isolation ( see Note 1)

1. Harvest 20 g of Arabidopsis rosette leaves (see Note 2). 2. The following steps have to be carried out in a refrigerated room, set at 4°C. 3. Homogenise the leaves twice for 3 s at full speed in 400 mL of cold homogenisation buffer HB1 using a Waring blender. 4. Filter the homogenate through two layers of Miracloth into a cold beaker and apply gentle pressure on the cut leaf material to obtain a higher chloroplast yield.

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5. Decant the filtrate into two pre-cooled 200-mL centrifuge beakers and centrifuge for 5 min at 1,500 × g and 4°C using a fixed-angle rotor. 6. Carefully decant and discard the supernatant. The remaining supernatant can be removed by pipetting. Resuspend each pellet in 40 mL of cold SH buffer using a soft paintbrush. Transfer the resuspended suspension into pre-cooled 50-mL centrifugation tubes and centrifuge again for 5 min at 1,500 × g and 4°C. 7. Carefully decant and discard the supernatant. The remaining supernatant can be removed by pipetting. Resuspend the pellets in 15 mL of cold HM buffer using a soft paintbrush. 8. Incubate the suspension on ice for at least 5 min to lyse the chloroplasts. 9. Centrifuge the lysed chloroplasts for 5  min at 1,500 × g and 4°C using a fixed-angle rotor. Resuspend the pellet in 10 mL of cold HM buffer and determine the chlorophyll concentration (see Subheading 3.4). 10. Centrifuge again for 5 min at 1,500 × g and 4°C and adjust the chlorophyll concentration to 1 mg of chlorophyll/mL in HM buffer. Store the thylakoids on ice for direct use or in aliquots at −20°C for long-term storage. 3.2. Arabidopsis Chloroplast Isolation and Subfractionation into Stroma, Envelopes, and Thylakoids (according to Refs. 4, 5)

1. Generate four Percoll gradients by mixing 15 mL of cold 100% Percoll and 15 mL of cold 2× RB buffer per gradient. Gradients are formed by ultracentrifugation for 20 min at 53,000 × g and 4°C in a swing-out bucket rotor. Reduce the deceleration (brake slow) (see Note 3). Store gradients on ice until use. 2. Harvest 150 g of Arabidopsis rosette leaves (see Note 2). 3. The following steps have to be carried out in a refrigerated room. 4. Homogenise the leaves twice for 3 s at full speed in 2 × 500 mL of cold homogenisation buffer HB2 using a Waring blender (see Note 4). 5. Filter the homogenate through two layers of Miracloth into a cold beaker and apply gentle pressure on the cut leaf material to obtain a higher chloroplast yield. 6. Decant the filtrate into pre-cooled 200 mL centrifuge beakers and centrifuge for 5 min at 1,500 × g and 4°C in a fixed-angle rotor. 7. Carefully decant and discard the supernatant. The remaining supernatant can be removed by pipetting. Resuspend each pellet in 5 mL of cold RB buffer using a soft paintbrush and pool the resuspended chloroplasts. 8. Carefully overlay 5 mL of the resuspended chloroplasts on the top of each cold, preformed Percoll gradient and centrifuge for

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6 min at 11,000 × g and 4°C in a swing-out bucket rotor (brake slow) (see Notes 3 and 5). 9. Aspirate the upper broken chloroplasts and the Percoll gradient down to the lower intact chloroplasts. Collect the intact chloroplasts by pipetting and wash them with three volumes of cold RB buffer (centrifugation for 5 min at 1,200 × g and 4°C in a swing-out bucket rotor) (see Note 6). 10. Carefully decant and discard the supernatant. Resuspend the chloroplasts in 2  mL of cold RB buffer and determine the chlorophyll concentration (see Subheading 3.4). 11. Dilute and centrifuge the chloroplasts again; adjust the chlorophyll concentration to 2 mg of chlorophyll/mL in TE buffer and incubate on ice for 30 min. 12. For further fractionation of the lysed chloroplasts, generate two sucrose step gradients. Prepare all sucrose solutions in TE buffer. Start with the denser solution (1.2  M sucrose) and overlay the solutions on top of each other in the following order into ultracentrifugation tubes: 2.5 mL of 1.2 M sucrose, 3.5 mL of 1 M sucrose, and 3.5 mL of 0.46 M sucrose. 13. Carefully load 2 mL of the lysed chloroplasts onto the sucrose gradient and ultracentrifuge for 2 h at 41,000 × g and 4°C in a swing-out bucket rotor. 14. Collect the stromal proteins (upper 2 mL) from the top of the gradient and aspirate the gradient down to the 0.46 M/1 M sucrose interface to collect the envelope membranes (see Note 7). Aspirate the rest of the gradient down to the thylakoid pellet at the base of the tube. Dilute the envelopes to a volume of 11 mL with TE buffer and pellet them by ultracentrifugation for 1  h at 114,000 × g and 4°C in a swing-out bucket rotor. Carefully aspirate the supernatant and resuspend the pellet in a few microlitres (20–50 mL) of TE buffer. Store the envelopes on ice for direct use or at −20°C for long-term storage (see Note 8). 15. Resuspend the thylakoid pellet in at least 15 mL of cold HM buffer; transfer the thylakoid suspension into centrifugation tubes and centrifuge for 5 min at 1,500 × g and 4°C. Adjust the chlorophyll concentration to 1 mg of chlorophyll/mL in HM buffer. Store the thylakoids on ice for direct use or at −20°C for long-term storage. 3.3. Pea Chloroplast Isolation and Subfractionation into Stroma and Thylakoids

1. Generate Percoll step gradients by overlaying 8  mL of cold 80% Percoll solution with 12 mL of cold 40% Percoll solution. Store gradients on ice until use. 2. Harvest 25 g of pea leaves (see Note 2). 3. The following steps have to be carried out in a refrigerated room.

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4. Homogenise the leaves twice for 3 s at full speed in 400 mL of cold grinding buffer using a Waring blender (see Note 3). 5. Filter the homogenate through two layers of Miracloth into a cold beaker and apply gentle pressure on the cut leaf material to obtain a higher chloroplast yield. 6. Decant the filtrate into a pre-cooled 200-mL centrifuge beaker and centrifuge for 5 min at 2,400 × g and 4°C. 7. Carefully decant and discard the supernatant. The remaining supernatant can be removed by pipetting. Resuspend each pellet in 1 mL of cold SH buffer using a soft paintbrush and pool the resuspended chloroplasts. 8. Carefully pipette the resuspended chloroplasts on the top of the cold, preformed Percoll gradient and centrifuge for 10 min at 7,700 × g and 4°C (see Note 5). 9. Aspirate the upper broken chloroplasts and the Percoll gradient down to the lower intact chloroplasts. Recover the intact chloroplasts that accumulate at the interface between 40 and 80% Percoll by pipetting and wash them with five volumes of cold SH buffer (see Note 6). 10. Pellet the chloroplasts by centrifugation for 2 min at 3,000 × g and 4°C. 11. Carefully decant and discard the supernatant. Resuspend the chloroplasts in 1  mL of cold SH buffer and determine the chlorophyll concentration (see Subheading 3.4). 12. Add 40  mL of cold SH buffer, centrifuge the chloroplasts again, and adjust the chlorophyll concentration to 3  mg of chlorophyll/mL in HM buffer (for translocation assays) or in 10  mM HEPES-KOH, pH 8.0 (for integration assays). Incubate the resuspended chloroplasts for 10  min on ice for chloroplast lysis. 13. Centrifuge the lysed chloroplasts for 5 min at 21,000 × g and 4°C in a microfuge. Collect the stromal proteins from the supernatant and retain them for later use. 14. Resuspend the thylakoid pellet in 1–2  mL of SH buffer and centrifuge for 5  min at 1,500 × g and 4°C in a microfuge. Remove the supernatant. 15. Resuspend the thylakoid pellet in SH buffer and adjust the chlorophyll concentration to 1 mg of chlorophyll/mL. 3.4. Measurement of Chlorophyll Concentration (according to Ref. 6)

1. Mix 5–20 mL of the isolated chloroplasts/thylakoids with 80% acetone to a final volume of 1 mL in a 1.5-mL microfuge tube and vortex thoroughly. Centrifuge for 2 min at 16,000 × g and room temperature in a microfuge, and transfer the supernatant to a fresh tube.

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2. Immediately measure the chlorophyll absorption of the ­supernatant at 663 and 645  nm in a spectrophotometer. Use 80% acetone as a blank control. 3. Calculate the chlorophyll concentration as follows: [mg chlorophyll/mL] = (A645nm × 20.2 + A663nm × 8.02) × dilution. 3.5. In Vitro Transcription and Translation

1. Thaw and keep all components for the transcription steps on ice. 2. Prepare the NT solution by mixing 5 mL of GTP (100 mM), 50 mL of CTP (100 mM), 50 mL of ATP (100 mM), and 50 mL of UTP (100 mM). Adjust the volume to 1 mL with distilled water. Add 233  mL of this mixture to 25  U of diguanosine triphosphate (a cap analogue nucleotide). Store the NT solution in aliquots at −20°C. 3. Prepare the reaction mix using the following components: 10 mL of 5× transcription buffer, 5 mL of NT solution, 23 mL of DEPC-treated water, 1 mL of RNase inhibitor, and 1 mL of SP6 RNA polymerase. 4. Adjust 5 mg of the DNA template and distilled water to a final volume of 10 mL (see Note 9). 5. Mix the 10 mL of DNA with 40 mL of the reaction mix and incubate at 37°C for 30 min in a heating block. 6. Add 50 mL of 10 M ammonium acetate and 100 mL of DEPCtreated water (to a final concentration of NH4OAc of 2.5 M). 7. Add 200 mL of phenol (pH 4.0)/chloroform (1:1) and vortex for 10 s. Centrifuge for 2 min at 21,000 × g in a microfuge at room temperature to achieve phase separation. 8. Transfer the aqueous upper phase to a fresh tube for further use. Add 400 mL of cold 100% ethanol and incubate for 30 min at −20°C. 9. Centrifuge for 15 min at 21,000 × g and 4°C in a microfuge to pellet the RNA. Discard the supernatant. 10. Wash the pellet by adding 300  mL of cold 70% ethanol and centrifuging for 15 min at 21,000 × g and 4°C. 11. Discard the supernatant and dry the pellet in a SpeedVac for 1–2 min. 12. Resuspend the pellet in 60  mL of DEPC-treated water and store the RNA on ice for direct use or at −80°C for long-term storage. 13. Before starting the translation steps, incubate the RNA for 10 min at 68°C, and immediately store on ice (see Note 10). 14. Thaw and keep all components for the translation steps on ice.

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15. Firstly, centrifuge the thawed wheat germ extract for 5 min at 4,500 × g and 4°C in a microfuge. Transfer the supernatant to a new tube and keep on ice until further use (see Note 11). 16. The following steps have to be carried out in a radionuclide laboratory (see Note 12). 17. Prepare the reaction mix for each translation with the following components: 20 mL of wheat germ extract, 4 mL of amino acid mixture (minus methionine), 12  mL of RNA, 5  mL of potassium acetate, 1  mL of RNase inhibitor, and 8  mL of [35S]-methionine (see Note 13). 18. After the addition of all the components, gently mix by pipetting and incubate for 90 min at 25°C in a water bath. 19. After incubation, mix 1 mL of the in vitro translation product with 10 mL of 4× SDS sample buffer and heat it for 2 min at 98°C in a heating block. Spin briefly, load the sample on a SDS gel, and confirm that the translation is successful by autoradiography (see Subheadings 3.8 and 3.9) (see Note 14). 3.6. Translocation of OE33 and OE23 (according to Ref. 7)

1. Prepare Arabidopsis thylakoids and pea stroma as described in Subheadings 3.1–3.3 (see Note 15). 2. The following steps have to be carried out in a radionuclide laboratory (see Note 12). 3. Resuspend the thylakoids (45 mg chlorophyll) in either 45 mL of pea stroma including 4 mM ATP (for OE33, cpSec-dependent protein translocation) or 45 mL of HM buffer (for OE23, cpTat-dependent protein translocation) in a centrifugation tube. Perform the control reactions in the absence of stromal extract (for OE33) or by the addition of 0.75  mL of 1  mM CCCP (for OE23) (see Note 16). 4. Add 5 mL of in vitro-translated [35S]-OE33 or [35S]-OE23 (or the [35S]-test protein) to the thylakoid suspension, mix briefly, and incubate for 30 min at 25°C under illumination (100 mmol/ m2/s) in a water bath. 5. Add 1 mL of thermolysin and 0.5 mL of 1 M CaCl2, and incubate for 1 h on ice (see Note 17). 6. After the thermolysin treatment, centrifuge the thylakoids for 1–2 min at full speed in a microfuge, discard the supernatant, and wash the pellet with 1 mL of HM buffer including 20 mL of 0.5 M EDTA (see Note 17). 7. Centrifuge again and discard the supernatant. Resuspend the thylakoid pellet in 25  mL of 4× SDS sample buffer and boil the sample for 2 min at 98°C in a heating block to solubilise the proteins. Centrifuge the sample. 8. Load 20 mL of the sample on a 15% SDS-polyacrylamide gel and separate proteins by SDS-PAGE (see Subheading 3.8).

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Fig.  2. Translocation of OE33 and OE23 across the thylakoid membrane and integration of LHCP into the membrane. Translocation assays (lane 2) were performed using Arabidopsis thylakoids, in vitro translation products of the precursors of OE33 (iOE33, intermediate OE33) or OE23 (pOE23, precursor OE23) (lane 1), stromal pea extract, and ATP (for OE33, cpSec-dependent protein translocation). Control reactions were performed in the absence of stroma (for iOE33) or in the presence of CCCP (for pOE23) (lane 3). The integration assay (lane 2) was performed using pea thylakoids, pea stromal extract, in vitro-translated precursor LHCP (pLHCP) (lane 1), and GTP. The control assay was performed in the presence of apyrase (lane 3). Thermolysin treatment was performed to digest the non-translocated or non-integrated precursor proteins. The translocation and processing of the precursors into the mature form (mOE33 and mOE23) and the integration of LHCP were analysed by SDS-PAGE and autoradiography. Thermolysin treatment of the thylakoid membranes results in a degradation product of LHCP (LHCP*) caused by the removal of an N-terminal peptide that is not protected by the bilayer.

9. Analyse the translocation of the radioactive protein(s) by ­autoradiography (see Subheading  3.9). An example of the translocation assay is shown in Fig. 2. 3.7. Integration of LHCP into Thylakoid Membranes (according to Refs. 8, 9)

1. Prepare pea thylakoids and pea stroma as described in Subheadings 3.1–3.3 (see Note 15). 2. The following steps have to be carried out in a radionuclide laboratory (see Note 12). 3. Prepare the reaction mix using the following reagents: 47.7 mL of 10 mM HEPES, pH 8.0, 6.2 mL of 4× SH buffer, 0.15 mL of 100  mM GTP, 10  mL of 100  mM methionine, 1  mL of MgCl2, and 30  mL of stroma (3  mg chlorophyll/mL). As a negative control, add 1 mL of apyrase (see Note 18) to a second sample (See Note 19). 4. Pellet the isolated thylakoid membranes (20 mg chlorophyll) by centrifugation for 1 min at full speed and at 4°C in a microfuge. 5. Resuspend the thylakoids in the prepared reaction mixtures and add 5 mL of in vitro-translated [35S]-Lhcb1 (or the [35S]-test protein). 6. Mix briefly by pipetting and incubate for 30 min at 25°C in a water bath. 7. Add 2 mL of thermolysin (10 mg/mL) and 1 mL of 1 M CaCl2, mix by pipetting, and incubate for 1 h on ice (see Note 17). 8. Add 2 mL of 0.5 M EDTA (final concentration 10 mM) and 100 mL of 0.2 N NaOH, mix by pipetting, and incubate again for 5 min on ice (see Notes 17 and 20). 9. Centrifuge the sample for 1 min at 4°C and at full speed in a microfuge, and discard the supernatant. Wash the pellet with 1 mL of cold SH buffer containing 10 mM EDTA, and centrifuge again.

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10. Discard the supernatant and resuspend the pellet in 20 mL of 4× SDS sample buffer. 11. Solubilise the membranes for 10 min at room temperature and boil the sample for 2 min at 98°C in a heating block. Spin to collect the sample again. 12. Load 20 mL of the sample on a 15% SDS-polyacrylamide gel and separate the proteins by SDS-PAGE (see Subheading 3.8). 13. Analyse the integration of the radioactive protein(s) by autoradiography (see Subheading  3.9). An example of the integration assay is shown in Fig. 2. 3.8. SDS-PAGE

1. Clean the plates of the gel system with ethanol and distilled water. Dry the plates and assemble the gel unit. 2. Prepare the separating solution for four minigels (0.75-mm thick, 15%) by mixing 8 mL of 30% acrylamide solution (29:1), 4  mL of 1.5 M Tris–HCl, pH 8.8, 4  mL of distilled water, 80  mL of 20% SDS, 20  mL of TEMED, and 60  mL of 25% ammonium persulphate. 3. Quickly pour the separating gel (ca. 3.2 mL per gel) and leave space for the stacking gel. 4. Overlay the separating solution with isopropanol and let the gel polymerise for about 15 min at room temperature. 5. Pour off the isopropanol and rinse the space for the stacking gel with distilled water. Dry the residual water and insert the comb. 6. Prepare the stacking solution for four minigels (0.75-mm thick) by mixing 6 mL of stacking buffer, 6 mL of TEMED, 48 mL of bromophenol blue solution, and 48 mL of 25% ammonium persulphate. 7. Quickly pour the stacking gel by pipetting. Let the gel polymerise for about 15 min at room temperature. 8. Remove the comb and assemble the gel system. 9. Pour running buffer (1×) into the upper and lower chambers of the gel system and wash the wells of the stacking gel with running buffer by pipetting. 10. Load up to 20 mL of sample in each well and spare one well for the pre-stained molecular weight marker (see Note 21). 11. Connect the gel unit to a power supply and let each gel run at 30 mA and at room temperature until the dye front runs out of the gel.

3.9. Visualisation of Radiolabelled Proteins by Autoradiography

1. After electrophoresis, disassemble the gel unit. Remove and discard the stacking gel. 2. Transfer the separating gel into fixing solution and incubate for 30 min at room temperature on a shaker.

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3. Discard the fixing solution and incubate the gel in amplifying solution for 30  min at room temperature on a shaker (see Note 22). 4. Place the gel on a wet piece of Whatman paper avoiding the formation of air bubbles and rupture of the SDS gel. Carefully cover the SDS gel with a plastic wrap while avoiding the formation of air bubbles and folding of the wrap (see Note 23). 5. Dry the gel on a gel vacuum dryer for 2  h at 80°C (see Note 24). 6. Fix the dried gel in a radiography cassette using adhesive tape. 7. This next step has to be carried out in a dark room. Place the X-ray film directly on the fixed gel. 8. Expose the gel to the X-ray film overnight at −70°C (see Note  25). 9. After exposition, develop the film: submerse the film in developer for 1–2 min, rinse it in water, and then place into fixer until the film background appears clear. Rinse the film under running water and let it dry.

4. Notes 1. In this procedure, thylakoids are not isolated by the fractionation of chloroplasts purified by a Percoll gradient (see Subheading  3.2). Therefore, this thylakoid fraction may be contaminated by envelope membranes and other cell components. If this does not interfere with your experiments, this method is recommended since it leads to a higher yield of isolated thylakoids than the method described in Subheading 3.2. 2. Before harvesting, keep the plants overnight in the dark to reduce the amount of leaf starch. A high starch content leads to fewer intact chloroplasts during preparation (centrifugation steps). 3. Disconnecting the brake (or setting the brake to slow) avoids mixing of the gradient. 4. The homogenisation step is critical for not destroying the chloroplasts with intervals that are too long. However, intervals that are too short lead to a low chloroplast yield. 5. Broken chloroplasts remain on the top of the gradient (upper band), whereas intact chloroplasts sediment through the gradient (lower band). 6. The washing step is crucial to remove the Percoll from the chloroplast suspension.

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7. Chloroplast envelopes do not contain chlorophylls and appear as a yellow band in the gradient. 8. The Arabidopsis stroma and envelopes are not used for the translocation or integration assays; thus, they can be stored for other experiments. The isolated thylakoids are used for the translocation assays (see Subheading 3.6). 9. The pGem4SS6.5 vector is used for in  vitro transcription (10). 10. This step is required for RNA denaturation, which enhances the translation process. 11. Do not let the wheat germ extract warm up; keep it on ice until it thaws and needs to be used. The efficiency of in vitro translation is reduced when the wheat germ extract is frozen and thawed again. 12. [35S] is a beta emitter and has to be used according to the radiation safety instructions. 13. In our experience, at least two to three methionines should be present in the translated protein sequence to ensure adequate radiolabelling of the protein during the in  vitro translation process. 14. Coupled in vitro transcription and translation kits (e.g., Roche, Promega) can also be used. 15. For the translocation, Arabidopsis or pea thylakoids can be used. However, according to the current literature, the integration assay does not function with Arabidopsis thylakoids. For the translocation and the integration assay, it is also ­possible to use Arabidopsis stroma instead of pea stroma. However, because of a better yield, it is somewhat easier to isolate pea stroma. 16. The cpSec-dependent translocation can also be inhibited by the addition of 8  mM (final concentration) sodium azide, a SecA inhibitor (11). CCCP abolishes the pH gradient across the thylakoid membrane. 17. Thermolysin hydrolyses the N-terminal amide bonds of hydrophobic amino acid residues in proteins. The treatment of thylakoids with thermolysin results in the digestion of proteins that are not translocated across or integrated into the thylakoid membrane. Peptides of integrated proteins that are exposed to the “stromal” surface of the thylakoid membrane, or peripheral membrane-bound proteins, can be degraded by the thermolysin treatment. Thermolysin is a metalloendopeptidase containing four calcium ions. Therefore, it is recommended to supplement the buffer for protease digestion with CaCl2. After digestion, the chelating agent EDTA is added to sequester the Ca2+, resulting in a reduction of thermolysin activity.

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18. Apyrase catalyses the hydrolysis of ATP and other nucleoside triphosphates. In this assay, apyrase is used as the negative control reaction. The GTP hydrolysis by the apyrase prevents LHCP from integrating into the thylakoid membrane. 19. The stromal extract can be replaced by adding in  vitro-­ translated or recombinant cpSRP transport components (cpSRP43, cpSRP54, and cpFtsY) (12, 13). 20. In addition to thermolysin treatment, the thylakoid membranes are washed with 0.1 M NaOH to remove residual, nonintegrated proteins. Resistance to extraction with 0.1 M NaOH is characteristic for integral membrane proteins. 21. Use a pre-stained molecular weight marker, or cut one corner of the gel for orientation. 22. The enhancer solution increases the sensitivity of the autoradiography. The Amplify™ fluorographic reagent can increase sensitivity of detection and significantly reduce exposure times. 23. The plastic wrap prevents both the radioactive contamination of the dryer cover and the gel sticking to the cover. 24. Vacuum should be applied until the gel is completely dry; otherwise, it will crack and be destroyed. 25. If necessary, the exposure time can be extended or shortened.

Acknowledgements This work was supported by a grant from the Deutsche Forschungsgemeinschaft (SFB 480 B11) (Schünemann). We thank Klaus Hagemann for his graphical contribution. References 1. Schünemann, D. (2007) Mechanisms of protein import into thylakoids of chloroplasts. Biol. Chem. 388, 907–915. 2. Aldridge, C., Cain, P., and Robinson, C. (2009) Protein transport in organelles: Protein transport into and across the thylakoid membrane. FEBS J. 276, 1177–1186. 3. Cline, K., and Dabney-Smith, C. (2008) Plastid protein import and sorting: different paths to the same compartments. Curr. Opin. Plant Biol. 11, 585–592. 4. Kunst, L. (1998) Preparation of physiologically active chloroplasts from Arabidopsis. In, Methods in Molecular Biology, Vol. 82 (Martinez-Zapater, J. and Salinas, J., eds.) Humana Press Inc., Totowa, NJ, USA, pp. 43–48.

5. Li, H. M., Moore, T., and Keegstra, K. (1991) Targeting of proteins to the outer envelope membrane uses a different pathway than transport into chloroplasts. Plant Cell 3, 709–717. 6. Arnon, D. I. (1949) Copper enzymes in isolated chloroplasts. polyphenoloxidase in Beta vulgaris. Plant Physiol. 24, 1–15. 7. Hulford, A., Hazell, L., Mould, R. M., and Robinson, C. (1994) Two distinct mechanisms for the translocation of proteins across the thylakoid membrane, one requiring the presence of a stromal protein factor and nucleotide triphosphates. J. Biol. Chem. 269, 3251–3256. 8. Hoffman, N. E., and Franklin, A. E. (1994) Evidence for a stromal GTP requirement for the integration of a chlorophyll a/b-binding

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polypeptide into thylakoid membranes. Plant Physiol. 105, 295–304. 9. Cline, K. (1986) Import of proteins into chloroplasts. Membrane integration of a thylakoid precursor protein reconstituted in chloroplast lysates. J. Biol. Chem. 261, 14804–14810. 10. Franklin, A. E., and Hoffman, N. E. (1993) Characterization of a chloroplast homologue of the 54-kDa subunit of the signal recognition particle. J. Biol. Chem. 268, 22175–22180. 11. Oliver, D. B., Cabelli, R. J., Dolan, K. M., and Jarosik, G. P. (1990) Azide-resistant mutants of Escherichia coli alter the SecA protein, an azidesensitive component of the protein export

machinery. Proc. Natl. Acad. Sci. USA 87, 8227–8231. 12. Tu, C. J., Schünemann, D., and Hoffman, N. E. (1999) Chloroplast FtsY, chloroplast signal recognition particle, and GTP are required to reconstitute the soluble phase of light-harvesting chlorophyll protein transport into thylakoid membranes. J. Biol. Chem. 274, 27219–27224. 13. Yuan, J., Kight, A., Goforth, R. L., Moore, M., Peterson, E. C., Sakon, J., and Henry, R. (2002) ATP stimulates signal recognition particle (SRP)/FtsY-supported protein integration in chloroplasts. J. Biol. Chem. 277, 32400–32404.

Chapter 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 Abstract Chloroplasts have one of the most complicated structures among organelles. They have three membrane systems, the outer and inner envelope membranes and the thylakoid membrane, which enclose three aqueous spaces: the intermembrane space between the two envelope membranes, the stroma, and the thylakoid lumen. Each of the chloroplast’s sub-organellar compartments houses a distinct set of proteins that perform distinct functions. Determining the sub-organellar location of a protein in the chloroplast is vital for understanding or verifying the function of the protein. Here, we present protocols for determining the sub-organellar location of a chloroplast protein. The protein of interest is synthesized and labeled with [35S]methionine by an in vitro translation system, and imported into isolated chloroplasts. The location of the protein is then identified by fractionation of the chloroplasts through differential and sucrose stepgradient centrifugations. The various sub-chloroplast fractions are analyzed by SDS-PAGE and autoradiography, so no specific antibody against the protein of interest is required. For membrane proteins, an alkaline extraction protocol is provided to further determine whether the protein is a peripheral or an integral membrane protein. The fractionation and extraction procedures presented can also be used in conjunction with immunoblotting, if an antibody against the protein of interest is available, enabling analyses of endogenous proteins. Key words: Chloroplast, Protein import, Sucrose gradient centrifugation, Fractionation, Alkaline extraction

1. Introduction Identifying the location of a protein is critical for understanding the function of the protein. Chloroplasts perform many important functions such as photosynthesis, fatty acid and carotenoid biosyntheses, and nitrogen and sulfur assimilations. These reactions are

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_20, © Springer Science+Business Media, LLC 2011

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carried out by proteins specifically localized in one or several of the six sub-organellar compartments that make up chloroplasts. Techniques such as fluorescence microscopy, immunogold ­electron microscopy, or fractionation (1–3) are all often used to determine the sub-organellar location of a protein within chloroplasts. Among these techniques, fractionation can offer quantitative localization data and is also relatively easy and quick to perform. Fractionation is normally followed by SDS-PAGE and immunoblot to reveal the presence of a protein in a certain fraction. However, specific antibodies against a protein of interest are often not available. Therefore, an alternative, sometimes even preferable, method for determining the location of chloroplast proteins, if the protein of interest is encoded by the nuclear genome, is to perform in vitro import. Nucleus-encoded chloroplast proteins are usually synthesized in the cytosol as higher molecular weight precursors with N-terminally localized transit peptides that govern their posttranslational import into chloroplasts. This import system has been reconstituted in vitro using isolated chloroplasts. Therefore, if the full-length cDNA (i.e., the full-length precursor with all the necessary localization information) is available, the protein can be synthesized, labeled with [35S]methionine, and imported into chloroplasts using the in  vitro reconstitution system. After fractionation, the protein can be visualized by SDS-PAGE and autoradiography. In some situations, it is necessary to use Arabidopsis chloroplasts for the import and fractionation experiments; for example, if the sub-organellar location of a protein needs to be investigated in a particular Arabidopsis mutant. Protocols for growing Arabidopsis seedlings for chloroplast isolation and import assays are provided here and in other chapters of this book. In higher plants, however, the sub-chloroplast location of a protein is usually conserved and it is, therefore, not necessary to use Arabidopsis. If the use of Arabidopsis is not required, it is recommended that chloroplasts are isolated from pea seedlings to allow easier chloroplast isolation and higher protein import efficiency (4, 5). The fractionation protocol presented here is modified from a well-established protocol for the isolation of highly purified chloroplast outer and inner envelope membranes (6). We reduced the scale so the current protocol can be used after an import reaction, which normally only uses up to a few hundred microliters of chloroplasts. Re-isolated intact chloroplasts after the import reaction are resuspended in a hypertonic solution to induce partial separation of the inner and outer envelope membranes (Fig.  1). Chloroplasts are then lysed by two freeze-thaw cycles. After diluting the hypertonic solution, the thylakoid fraction is collected as the pellet fraction of a low speed spin. The supernatant is separated by a sucrose step gradient into stroma, outer envelope membrane,

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Reisolate intact chloroplasts after import and resuspend chloroplasts with 0.6 M sucrose/TE

Lyse chloroplasts by freeze-thaw

Adjust to 0.2 M sucrose/TE

Centrifuge at low speed Supernatant: load onto a sucrose step gradient

Pellet (thylakoid)

Centrifugation Wash with TE

Centrifuge to collect the thylakoid fraction

0.75 mL 0.2 M Sucrose

Stromal fraction

1.1 mL 0.46 M Sucrose 1.1 mL 0.8 M Sucrose 1.1 mL 1M Sucrose

Outer envelope membrane fraction Inner envelope membrane fraction Collect and dilute with TE Centrifuge to collect the outer envelope membrane and inner envelope membrane fractions

Fig. 1. Schematic outline of sucrose step-gradient fractionation after import. TE, Tricine-EDTA buffer.

and inner envelope membrane fractions. If the protein localizes in one of the three membrane fractions, it is often necessary to perform alkaline extractions to determine whether the protein is peripherally associated with the membrane or is truly an integral membrane protein. Because the amount of outer and inner envelope membrane obtained from the small-scale fraction may be too low for further alkaline extraction, a separate import experiment should be performed and the chloroplasts directly extracted by an alkaline solution after import to determine the nature of membrane association of the protein.

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2. Materials 2.1. [35S]MethionineLabeled Precursor Preparation

1. [35S]methionine (Perkin Elmer, NEG709A EASYTAG™). 2. TNT®-coupled wheat germ extract system or TNT®-coupled reticulocyte lysate system (Promega). 3. RNasin ribonuclease inhibitor (Promega). 4. 1× Import buffer (IB): 50 mM 4-(2-hydroxyethyl)piperazine1-ethanesulfonic acid-KOH (HEPES-KOH), pH 8.0, and 0.33 M sorbitol. To prepare 1 L of 2× IB, dissolve 23.8 g of HEPES and 120 g of sorbitol in 700 mL of H2O. Adjust pH to 8.0 with 5 N KOH and add H2O to 1 L. Store at −20°C. 5. 50 mM methionine with 2 mM 1,4-dithiothreitol (DTT) in 2× IB. Store at −20°C.

2.2. Growth of Arabidopsis Seedlings

1. Arabidopsis thaliana seeds. 2. 95% (v/v) ethanol. 3. Bleach solution: mix three volumes of H2O and one volume of bleach (Clorox or 6% [v/v] sodium hypochlorite) and add 1 drop of Tween® 20. 4. Sterile H2O. 5. Sterile 0.1% (w/v) agar. 6. Murashige and Skoog medium (MS medium): dissolve 4.44 g of Murashige and Skoog salt mix with Gamborg’s vitamins, 20 g of sucrose, and 1 g of 2-(N-morpholino)ethanesulfonic acid (MES) hydrate in 900 mL of H2O. Adjust pH to 5.7 with ~300 mL of 5 N KOH and add H2O to make up the volume to 1 L. Add 3 g of Gelrite® (Caisson) (see Note 1) and autoclave at 121°C for 20 min. Use about 100 mL for each 15 × 2.5 cm plate. 7. Surgical tape.

2.3. Chloroplast Isolation

1. 1× Arabidopsis grinding buffer (AGR): 50 mM HEPES-KOH, pH 8.0, 2  mM ethylenediaminetetraacetic acid (EDTA), 0.33 M sorbitol, and 0.5% bovine serum albumin (BSA) (see Note 2). To prepare 1 L of 2× AGR, dissolve 23.8 g of HEPES and 1.48 g of EDTA in 700 mL of H2O, and adjust pH to 8.0 with 5 N KOH. Then add 120 g of sorbitol and 10 g of BSA. Add water to 1 L and store at −20°C. 2. Percoll™. Store at 4°C. 3. 100% Acetone. 4. Miracloth. 5. Funnel. 6. Domestic blender or Polytron.

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1. 100  mM Mg-ATP in 1× IB. Aliquot into small volumes for single use and store at −20°C. 2. 40% Percoll: for 100 mL, mix 40 mL of Percoll™, 40 mL of 2× AGR, and 20 mL of 1× AGR. Store at −20°C. 3. 1× Tricine-EDTA buffer (TE): 10 mM Tricine-KOH, pH 7.5, and 2  mM EDTA. To prepare 500  mL of 10× TE, dissolve 8.959 g of Tricine and 3.722 g of EDTA in 400 mL of H2O. Adjust pH to 7.5 with 5 N KOH and add H2O to 500 mL. 4. 0.2, 0.46, 0.6, 0.8, and 1  M sucrose in 1× TE: to prepare 250  mL of 1.2  M sucrose in 1× TE, dissolve 102.69  g of sucrose in 100 mL of H2O, add 25 mL of 10× TE, and bring the volume up to 250  mL with H2O. Dilute with 1× TE to make the 0.2, 0.46, 0.6, 0.8, and 1 M sucrose solution. Store at −20°C. 5. Beckman SW 60 Ti rotor and 4-mL thin-wall polyallomer open-top tubes (Beckman part number 328874). 6. SpeedVac. 7. Protein extraction buffer: 500  mM Tris–HCl, pH 8.45, 8% (w/v) dodecyl sulfate sodium salt (SDS), and 1 mM EDTA. 8. An SDS-compatible protein concentration determination kit such as the BCA™ protein assay kit (Thermo).

2.5. Alkaline Extraction of Chloroplasts After Import

1. Alkaline solution: 100 mM Na2CO3, pH 11.5. 2. 20% (w/v) trichloroacetic acid (TCA) solution.

3. Methods 3.1. [35S]MethionineLabeled Precursor Preparation

We usually use a coupled transcription-translation system, such as the TNT-coupled wheat germ extract systems or the TNT-coupled reticulocyte lysate systems (Promega) to synthesize [35S]methionine-labeled precursor proteins for the import reactions (see Note 3). To synthesize 100 mL of precursor proteins, mix 25 mL of TNT wheat germ extract or reticulocyte lysate, 2 mL of TNT reaction buffer, 1 mL of the RNA polymerase that drives the transcription of the precursor-protein cDNA, 1 mL of 1 mM amino acid mixture without methionine, 1 mL of RNasin ribonuclease inhibitor, 1–2 mg of the plasmid DNA with the cDNA encoding the precursor protein (see Note 4), and 6–12 mL [35S]methionine, and add nucleasefree H2O to a final volume of 50 mL. Incubate at 30°C for 2 h for the TNT-coupled wheat germ extract systems, or 1.5  h for the TNT®-coupled reticulocyte lysate systems. Add 50 mL of 50 mM cold methionine (see Note 5) in 2× import buffer with 2  mM DTT. Store at −80°C.

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3.2. Growth of Arabidopsis Seedlings

1. Arabidopsis seeds (8–10  mg, about 500 seeds, for one MS plate) are mixed with 10  mL of 95% ethanol for 1  min in a 15-mL tube. 2. Decant the ethanol solution, add 10  mL of the bleach solution, and rotate the tube for 15 min. 3. Decant the bleach solution and wash seeds with 10 mL of sterilized H2O three times. Do the last two washes in a laminar flow hood. 4. Resuspend the seeds in 0.1% soft agar and sow the seeds on one MS medium plate. Allow the soft agar to dry with the lid slightly skewed in the hood. Seal the plate with surgical tape and put the plate at 4°C for 1–2 days for stratification. 5. Transfer the plate to a growth chamber (16-h light/8-h dark photoperiod and 50–80  mE/m2/s light intensity) for about 21–24 days.

3.3. Chloroplast Isolation

1. Mix 15 mL of Percoll and 15 mL of 2× AGR in a transparent 50-mL high-speed centrifuge tube. Centrifuge at 38,700 × g for 30 min at 4°C. 2. Harvest the entire aerial parts of the Arabidopsis plants with scissors and place tissue (about 4 g from one MS plate of about 500 seedlings) into a 450-mL beaker sitting on ice containing 120 mL of chilled 1× AGR. When all the tissues are harvested, homogenize the tissues using a domestic blender by two 5-s pulses of blending. Filtrate the homogenate through two layers of Miracloth held in place by a funnel, into a prechilled 250-mL flask sitting on ice. Squeeze the Miracloth to get as much of the filtrate as possible. 3. Transfer the filtrate into four prechilled 50-mL centrifuge tubes and centrifuge at 3,000 × g for 3 min at 4°C. Decant the supernatant and resuspend each pellet with 1–2.5  mL of 1× AGR. 4. Pool the resuspension and load onto the Percoll gradient from step 1. Do not load more than 10  mL on each gradient. Centrifuge in a prechilled swinging-bucket rotor at 7,700 × g for 10 min at 4°C with the brake set on slow or no brake. 5. Remove and discard the upper green band, which is formed by broken chloroplasts. Retrieve the lower green band, which is formed by intact chloroplasts, into a new 50-mL centrifuge tube containing some chilled 1× AGR. Add 1× AGR to bring the volume up to about 30  mL. Centrifuge at 1,500 × g for 5 min at 4°C. 6. Decant the supernatant and resuspend the chloroplast pellet in 20 mL of 1× AGR. Put the tube on ice in the dark. To measure the chlorophyll concentration, mix 40  mL of the chloroplast

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suspension with 160 mL of water and 800 mL of acetone. Mix thoroughly and centrifuge for 5 min at 3,000 × g at room temperature. Transfer the supernatant to a 1-mL cuvette and measure A652. Use 80% acetone as the blank. Calculate the total amount of chlorophyll obtained in mg using the formula: (A652/36) × 500. A normal yield is around 0.5–0.75  mg per plate. 7. Pellet chloroplasts by centrifuging at 1,500 × g for 5  min at 4°C. Decant the supernatant and resuspend chloroplasts with 1× IB to 1 mg chlorophyll/mL. 3.4. Protein Import and Fractionation of Chloroplasts by a Sucrose Step Gradient

1. For a 750-mL import reaction, in a 1.5-mL microfuge tube mix 250 mL of chloroplasts at a concentration of 1 mg chlorophyll/mL, 22.5 mL of 100 mM Mg-ATP, and 50–100 mL of [35S]methionine-labeled precursor proteins, and add 1× IB to make the volume to 750 mL (see Note 6). 2. Incubate for 25 min at room temperature and tap occasionally to mix. 3. Load the import reaction mixture onto two tubes of 1-mL prechilled 40% Percoll solution (375 mL of import mixture per tube) and centrifuge in a swinging-bucket rotor at 2,900 × g for 6 min at 4°C to re-isolate the intact chloroplasts. 4. Discard the supernatant. Resuspend each pellet with 0.5 mL of 1× IB and combine the chloroplast suspension into one tube. 5. Aliquot 50 mL of the chloroplast suspension to a new 1.5-mL microfuge tube and label as the “total import” sample. Centrifuge the “total import” sample, as well as the rest of the chloroplasts in the original tube (the “fractionation” sample), for 3 min at 3,000 × g, 4°C. 6. Remove and discard the supernatant. Add 15 mL of the protein extraction buffer to resuspend the “total import” sample and store at −80°C. Use 250  mL of 0.6  M sucrose in 1× TE to resuspend the chloroplast pellet of the “fractionation” sample and incubate at 4°C for 10 min. Transfer the sample to −80°C, freeze for 30 min, thaw at 37°C in a water bath for 1 min, and re-freeze the sample at −80°C. The procedure can be stopped at this step. 7. Set up a sucrose step gradient consisting of 1.1  mL of 1  M sucrose, 1.1  mL of 0.8  M sucrose, and 1.1  mL of 0.46  M sucrose in a 4-mL tube for the Beckman SW 60 Ti rotor (see Note 7). We usually use the thin-wall polyallomer ultracentrifuge tubes. All the sucrose solutions are made in 1× TE as described in step 4 of Subheading 2.4. Mark the two interfaces of the three sucrose solutions on the tube wall by drawing a few dots or short broken lines with a permanent maker pen. The outer and inner membrane vesicles will fractionate to the

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0.46  M/0.8  M and 0.8  M/1  M interfaces, respectively. However, their amount will be too small to be visible by color. The sucrose interfaces tend to blur after the centrifugation. 8. Thaw the “fractionation” sample from step 6 in a 37°C water bath for 1 min. Add 500 mL of 1× TE and mix well. Centrifuge for 3 min at 3,000 × g, at 4°C. 9. Transfer the supernatant to a new tube and store on ice for loading onto the sucrose step gradient later. Wash the pellet twice by resuspending in 1  mL of 1× TE and centrifuge for 3 min at 3,000 × g, at 4°C. Resuspend the pellet with 250 mL of the protein extraction buffer and store at −80°C. This is the “thylakoid” fraction. 10. Load the supernatant from step 9 onto the top of the sucrose step gradient from step 7. 11. Centrifuge the sucrose gradient at 50,000 rpm (336,840 × gmax) in a Beckman SW 60 Ti rotor at 4°C for 1 h. 12. Transfer 200 mL of solution from the top of the gradient to a new 1.5-mL microfuge tube and store on ice. This is the “stroma” fraction. Collect 400 mL of the “outer envelope membrane” fraction from the 0.46 M/0.8 M sucrose interface to a 1.5-mL polyallomer microfuge tube and store on ice. Remove and discard some of the 0.46 M sucrose solution carefully. Do not disturb the 0.8 M/1 M sucrose interface. Collect 400 mL of the “inner envelope membrane” fraction from the 0.8  M/1  M sucrose interface to a new 1.5-mL polyallomer microfuge tube and store on ice. 13. Add 1 mL of 1× TE to the two envelope membrane fractions and mix well. Centrifuge at 100,000 × g for 45 min at 4°C. 14. Carefully remove the supernatant with a pipette. Dry the pellets in a SpeedVac and resuspend the pellets in 12 mL of the protein extraction buffer. 15. Quantify the protein concentration of each fraction with an SDS-compatible protein concentration determination kit like the BCA protein assay kit. The stroma fraction is in a different buffer (0.2 M sucrose/TE) from other fractions (the protein extraction buffer) and may need a separate standard series diluted in the same buffer. 16. Analyze all fractions by SDS-PAGE followed by autoradiography or immunoblotting. 17. OEP14 is a protein of unknown function located in the chloroplast outer envelope membrane (7). Tic40 is a co-chaperone located in the chloroplast inner envelope membrane (8). Fractionation of Arabidopsis chloroplasts after import of these two proteins using the protocols described above is shown in Fig. 2. OEP14 and Tic40 were preferentially localized in the

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Fig.  2. Localization of OEP14 and Tic40 by fractionation after import into Arabidopsis ­chloroplasts. In vitro-translated [35S]methionine-labeled OEP14 and Tic40 precursor (prTic40) were incubated with isolated Arabidopsis chloroplasts under import conditions. After import, chloroplasts were fractionated into the outer envelope membrane (OM), the inner envelope membrane (IM), the stroma (S), and the thylakoid (Thy) fractions. Samples were analyzed by SDS-PAGE followed by autoradiography (OEP14 and Tic40) or immunoblotting (Tic110, cpHsc70, and CAB). 1 mg of proteins was loaded in the OM, IM, S, and Thy lanes and about 6 mg of proteins was loaded in the total chloroplasts before fractionation (T). TR, in vitro-translated protein before the import reaction. Due to different salt concentrations, Tic40 ran slightly slower in the total chloroplast sample (T) than in samples after fractionation.

outer and inner envelope membranes, respectively. Marker proteins for the inner envelope membrane (Tic110, (9)), the stroma (cpHsc70, (10)), and the thylakoid (the chlorophyll a/b binding protein of photosystem II, CAB), were analyzed by immunoblots to verify the effectiveness of the fractionation. 3.5. Alkaline Extraction of Chloroplasts After Import

1. For a 150-mL import reaction (see Note 6), in a 1.5-mL microfuge tube mix 50 mL of chloroplasts at a concentration of 1  mg chlorophyll/mL, 4.5  mL of 100  mM Mg-ATP, and 10–15 mL of [35S]methionine-labeled precursor proteins, and then add 1× IB to 150 mL. 2. Incubate for 25 min at room temperature and tap occasionally to mix. 3. Load the import reaction mixture onto 1 mL of prechilled 40% Percoll solution and centrifuge in a swinging-bucket rotor at 2,900 × g for 6 min at 4°C to re-isolate the intact chloroplasts. 4. Discard the supernatant. Resuspend the pellet with 1 mL of 1× IB. 5. Aliquot 50 mL of the chloroplast suspension to a new 1.5-mL microfuge tube as the “total import” sample. Centrifuge the “total import” sample and the rest of the chloroplasts (the

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“extraction” sample) in the original tube at 4°C for 3 min at 3,000 × g. Remove and discard the supernatant. Add 15 mL of the protein extraction buffer to resuspend the “total import” sample and store at −80°C. 6. Resuspend the pellet in the “extraction” sample in 200 mL of the alkaline solution and incubate on ice for 30 min. 7. Transfer the suspension to a new 1.5-mL polyallomer microfuge tube. Centrifuge at 100,000 × g for 45 min at 4°C. 8. Transfer the supernatant (the soluble fraction) to a new 1.5mL microfuge tube. Wash the pellet (the membrane fraction) with 0.5  mL of the alkaline solution and centrifuge at 100,000 × g for 15  min at 4°C. Discard the supernatant and resuspend the pellet in 20 mL of the protein extraction buffer. 9. Precipitate the soluble fraction by adding an equal volume of 20% TCA solution and incubate on ice for 30 min. 10. Collect the precipitate by centrifuging at 14,000  rpm in a microfuge at 4°C for 30 min. 11. Wash the pellet with 500 mL of prechilled 100% acetone and centrifuge at 14,000 rpm in a microfuge for 5 min at 4°C. 12. Decant the supernatant, let the residual acetone dry out in a fume hood, and resuspend the precipitate with 20 mL of the protein extraction buffer. 13. Analyze 5 mL (10 mL after adding 2× SDS-PAGE sample buffer) of all three fractions (total, soluble, and membrane) by SDS-PAGE and autoradiography. Because the membrane and the soluble fractions are resuspended in an equal volume of the extraction buffer, loading an equal volume into the gel will represent an equal proportion of the soluble and membrane fractions, making direct comparison easier. An example of the alkaline extraction experiment is shown in Fig. 3. Hsp93 is a soluble stromal protein but a fraction of it associates with the inner envelope membrane due to interactions with the protein import machinery (11). However, when chloroplasts were subjected to alkaline extraction, almost all the Hsp93 molecules were found in the soluble fraction, indicating that Hsp93 indeed only associates peripherally with the membranes. Some precursor form of Hsp93 (prHsp93) molecules were still present after the import reaction. These prHsp93 had not been fully imported and were still in the envelope membranes. Therefore, they fractionated mostly with the membrane fraction. Tic40 is an integral inner envelope membrane protein. When chloroplasts were subjected to alkaline extraction, almost all of the Tic40 molecules were found in the membrane fraction (Fig. 3).

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Fig.  3. Alkaline extraction of Arabidopsis chloroplasts after import of Tic40 and Hsp93 precursor proteins. In vitro-translated [35S]methionine-labeled Tic40 precursor (prTic40) and Hsp93 precursor (prHsp93) were incubated with isolated Arabidopsis chloroplasts under import conditions. After import, reisolated intact chloroplasts were subjected to alkaline extraction and separated into the soluble (Sol) and membrane (Mem) fractions by centrifugation. Samples were analyzed by SDS-PAGE followed by autoradiography. TR, in vitro-translated protein before the import reaction. T, total chloroplasts after import. All samples for each precursor were run on the same gel, but the TR and T samples of Tic40 were run on different parts of the same gel.

4. Notes 1. When using other kinds of agar, the concentration of the agar has to be optimized. Seedlings may be difficult to harvest and/ or seedling growth may be affected if the agar is too soft or too hard. 2. Omitting the cations in the original grinding buffer designed for pea chloroplasts and adding EDTA and BSA increase the yield of chloroplast isolation from Arabidopsis. 3. For unknown reasons, some precursors have a better chloroplast import efficiency if synthesized by an uncoupled system; i.e., plasmid DNA is first transcribed into RNA and the RNA is then used to program protein synthesis in the wheat germ extract or reticulocyte lysate translation system. For the uncoupled protocols, see ref. 5. 4. The amount of plasmid DNA used should be optimized for each precursor construct. 5. The 50 mM cold methionine is used to dilute the [35S]methionine so that chloroplast-encoded proteins synthesized by the isolated chloroplasts will not be visible on the autoradiograph. 6. It is recommended that the novice use at least 250 mL of chloroplasts for the fractionation. It is difficult to recover sufficient

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amounts of the outer and inner envelope membranes for ­further analyses using smaller amounts of chloroplasts. For alkaline extraction, 50 mL of chloroplasts is sufficient. 7. To set up a sucrose gradient, place a 4-ml thin-wall polyallomer tube in a tube rack. Pipette 1.1 mL of the 1 M sucrose into the tube and mark the position of the upper surface of 1 M sucrose on the tube wall with a permanent marker pen by drawing a few short broken lines. Then, use a Gilson P200 Pipetman and pipette 200 mL each time, to gently layer 1.1 mL of the 0.8 M sucrose solution onto the 1 M sucrose solution. Mark the position of the upper surface of 0.8  M sucrose as described above. Finally, gently layer 1.1  mL of the 0.46  M sucrose onto the 0.8  M sucrose solution, using the same procedure. References 1. Li, H. M., Kaneko, Y., and Keegstra, K. (1994) Molecular cloning of a chloroplastic protein associated with both the envelope and thylakoid membranes. Plant Mol. Biol. 25, 619–632. 2. Li, H., and Chen, L. J. (1997) A novel chloroplastic outer membrane-targeting signal that functions at both termini of passenger polypeptides. J. Biol. Chem. 272, 10968–10974. 3. 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. 4. Smith, M. D., Schnell, D. J., Fitzpatrick, L., and Keegstra, K. (2003) In vitro analysis of chloroplast protein import. Curr. Protoc. Cell Biol. 11,11.16.1-11.16.21. 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. Keegstra, K., and Yousif, A. E. (1986) Isolation and characterization of chloroplast envelope membranes. Methods Enzymol. 118, 316–325.

7. Li, H.-m., and Chen, L.-J. (1996) Protein targeting and integration signal for the chloroplastic outer envelope membrane. Plant Cell 8, 2117–2126. 8. Chou, M. L., Fitzpatrick, L. M., Tu, S. L., Budziszewski, G., Potter-Lewis, S., Akita, M., Levin, J. Z., Keegstra, K., and Li, H. m. (2003) Tic40, a membrane-anchored co-chaperone homologue in the chloroplast protein translocon. EMBO J. 22, 2970–2980. 9. Kessler, F., and Blobel, G. (1996) Interaction of the protein import and folding machineries in the chloroplast. Proc. Natl. Acad. Sci. USA 93, 7684–7689. 10. Su, P. H., and Li, H. M. (2008) Arabidopsis stromal 70-kD heat shock proteins are essential for plant development and important for thermotolerance of germinating seeds. Plant Physiol. 146, 1231–1241. 11. Nielsen, E., Akita, M., Davila-Aponte, J., and Keegstra, K. (1997) Stable association of chloroplastic precursors with protein translocation complexes that contain proteins from both envelope membranes and a stromal Hsp100 molecular chaperone. EMBO J. 16, 935–946.

Chapter 21 Studying Arabidopsis Envelope Protein Localization and Topology Using Thermolysin and Trypsin Proteases John Froehlich Abstract Chloroplasts are metabolically important organelles that perform many essential functions within plant cells. The chloroplasts can be subdivided into six distinct sub-compartments to which a protein may be ultimately targeted. These sub-compartments are defined as the outer envelope membrane (OEM), the inner envelope membrane (IEM), the thylakoid membrane, and three aqueous sub-compartments – the intermembrane space (IMS), the stroma, and the thylakoid lumen. The process by which proteins are targeted to the chloroplastic envelope membrane remains a challenging question in cell biology. Our understanding of protein targeting to the OEM is very limited, whereas targeting of membrane proteins to the IEM appears to utilize at least two targeting pathways called the stop-transfer and the conservative sorting (or post-import) pathways. Furthermore, once a membrane protein arrives at the envelope membrane, our understanding of how it achieves its final topology remains limited. One method that can be used to determine the topology of an envelope membrane protein is to apply the “dual protease” strategy. This approach involves several steps: first, performing an in vitro import assay; second, applying a “dual protease” protection assay using thermolysin and trypsin; and finally, isolating and analyzing chloroplastic subcellular fractionations (i.e., total membrane and soluble fractions). By using this multistep approach, one can gain critical information regarding the final topology of an OEM or IEM protein. Likewise, the “dual protease” approach may help in elucidating the possible targeting pathway that a membrane protein utilizes prior to its insertion into the envelope membrane. Key words: Envelope membrane, Protein targeting, Transmembrane domain, Protease, Topology, Thermolysin, Trypsin

1. Introduction One of the most recognized roles of a chloroplast is to carry out the process of photosynthesis within a plant cell. However, chloroplasts are also involved in many other metabolic processes such as the biosynthesis of fatty acids, aromatic and nonaromatic

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amino acids, isoprenoids, and tetrapyrroles. To ensure the ­functionality of many of these diverse metabolic pathways, thousands of proteins must first be posttranslationally imported into chloroplasts and then finally assembled into active metabolic complexes. One common feature of all chloroplasts is that most of their proteins are encoded in nuclear genes, synthesized on cytosolic ribosomes – generally as larger precursor proteins, and transported into the organelle via a translocation apparatus located in the envelope membranes (1, 2). Approximately 3,000 proteins are estimated to be targeted to the chloroplasts. However, the overall number of membrane proteins that are located in the chloroplasts is considerably smaller. For instance, the plant membrane protein database ARAMEMNON (3) predicts that over 500 integral membrane proteins reside within chloroplasts. The majority of these membrane proteins are components of the four major photosynthetic complexes: photosystem I, photosystem II, the cytochrome b6f complex, and the ATP synthase complex, which are localized to the thylakoid membrane. However, information obtained from a variety of envelope membrane proteomic studies have identified approximately 30–40 outer envelope membrane (OEM) proteins (4) and between 100 and 120 inner envelope membrane (IEM) proteins (4–8). Understanding how membrane proteins are specifically directed to either the OEM or the IEM and achieve their final topology remains a challenging problem in cell biology. While molecular biology approaches can often reveal the function of a membrane protein by using gene knockout strategies, different yet complementary cell biology approaches can help decipher the subcellular localization and topology of a given membrane protein. Indeed, once the topology of a given membrane protein has been established, this information can be used to make reasonable predictions about that protein’s possible function and/or mechanism of insertion into either the OEM or the IEM. Consequently, by employing a combination of different techniques, such as in vitro import assays, “dual protease” protection assays using thermolysin and trypsin, and fractionation studies, the topology of several OEM and IEM proteins has been determined (Fig. 1a, b). For example, various laboratories have used different facets of the “dual protease” assay to determine the topology of the following OEM proteins: Toc75 (9), Toc34 (10), OM14 (11), HPLS (12), and DGD1 (13), and of the following IEM proteins: AtArc6 (14), AtTGD2 (15), AtAPG1 (16, 17), and AtTic40 (18–20). Thus, by applying the “dual protease” protection assay as outlined in this chapter, one will have the method and the tools to determine the subcellular localization and the topology of either an outer or an inner envelope membrane protein.

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a

b TP-1-67

TM-615-635

Arc6

801aa TP-1-22 TM-96-117

TGD2

Thermolysin Trypsin

Toc33 N

382 aa

Cytosol

TP-1-22TM-96-117

GFP

TGD2-GFP TP-1-50

TM-304-326

APG1

620 aa

OEM C

N

N

C

IMS

C

338 aa

IEM

BiTP-1-69 TM-107-129

Tic40

448 aa TM-267-283

Toc33

310 aa

Stroma

C C Tic40

APG1

N TGD2

N Arc6

Fig.  1. Features and topology of selected outer and inner envelope membrane proteins. (a) Comparison of different ­well-characterized OEM and IEM proteins is presented. TP, transit peptide; BiTP, Bipartite transit peptide; TM, predicted transmembrane domain; aa, amino acid; GFP, green fluorescence protein. (b) The topology of various OEM and IEM proteins shown in (a) as determined by various laboratories is presented (10, 14–20). The protease thermolysin is represented by large scissors icon, while trypsin is represented by smaller black scissor icon. C,C-terminal end and N, N-terminal end of membrane protein. Toc33 is an outer envelope membrane protein that serves as a control to ensure that the protease activity of both thermolysin and trypsin is functioning properly. Under normal conditions, Toc33 is protease sensitive when treated with thermolysin and trypsin (10). The hashed lines at the IEM represent regions of the inner envelope membrane that have been partially disrupted by trypsin, resulting in trypsin gaining access to the IMS. OEM, outer envelope membrane; IMS, Intermembrane space; IEM, inner envelope membrane.

2. Materials 2.1. Import Assays

1. Import buffer (2×): 0.1 M N-2-hydroxyethylpiperazine-N’-2ethanesulfonic acid (HEPES)-KOH, pH 8.0, and 0.66  M sorbitol. 2. 100 mM Mg-adenosine-5’-triphosphate (ATP) stock solution: 0.507 g of Mg-ATP is dissolved in 10 mL of 1× import buffer, then aliquoted in 1.0 mL fractions, and stored at −20°C. 3. 40% Percoll cushion: 40% (v/v) Percoll in 1× import buffer, pH 8.0. 4. Lysis buffer: 25  mM HEPES-KOH, pH 8.0, and 4.0  mM MgCl2. 5. Thermolysin stock solution: thermolysin is prepared in 5 mM CaCl2/1× import buffer to generate a 1 mg/mL stock solution. This stock solution is prepared fresh on the day of use (see Note 1). 6. Trypsin stock solution: trypsin is prepared in 1× import buffer to give a 1 mg/mL stock solution. This stock solution is prepared fresh on the day of use (see Note 2). 7. Trypsin inhibitor solution: soybean trypsin inhibitor is prepared in 1× import buffer to give a 1 mg/mL stock solution. This stock solution is prepared fresh on the day of use.

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8. Thermolysin quench solution: 60 mM ethylenediaminetetraacetic (EDTA) is prepared in 1× import buffer. 2.2. Translation Reaction Kits

1. Translation reaction kits: TNT®-coupled reticulocyte lysate (Promega, Madison, WI, USA) or TNT®-coupled wheat-germ extract (Promega) is compatible with the import assay. Transcription/translation kits can be purchased from Promega. These kits are available in three configurations for the translation of genes cloned downstream from the T7, T3, or SP6 RNA polymerase promoters. Detailed translation protocols are included with each specialized kit. 2. Proteins to be translated can be radiolabeled with either [35S]-methionine (PerkinElmer, NEG009T; PerkinElmer, Norwalk, CT, USA) or [3H]-leucine (PerkinElmer, NET460A) according to Promega’s TNT®-coupled transcription/translation system protocols. Alternatively, the Perkin Elmer/NEN EasyTag™ L-[35S]-methionine, 1  mCi (37 MBq), stabilized aqueous solution (NEG709A) can be used for all the transcription/translation systems previously listed (see Notes 3 and 4). 3. RNasin® ribonuclease inhibitor (40 U/mL) (Promega). 4. Translation product dilution buffer: prepare either 50  mM “cold” L-methionine dissolved in 1× import buffer, or 50 mM “cold” L-leucine dissolved in 1× import buffer (see Note 5).

2.3. SDSPolyacrylamide Gel Electrophoresis

1. Resolving buffer (4×): 1.5  M Tris–HCl, pH 8.7, and 0.4% (w/v) sodium dodecyl sulfate (SDS). Store at 4 degrees. 2. Stacking buffer (4×): 0.5 M Tris–HCl, pH 6.8, and 0.4% (w/v) SDS. Store at 4 degrees. 3. Thirty percent acrylamide/bis-acrylamide solution (37.5:1 with 2.6% C) (Caution: acrylamide is a neurotoxin when unpolymerized, so care should be taken to minimize contact with this solution). 4. N,N,N,N¢-tetramethylethylenediamine (TEMED). 5. 10% (w/v) ammonium persulfate solution: 1.0  g of ammonium persulfate is dissolved in 10  mL of distilled water and immediately frozen in single-use (500 mL) aliquots at −20°C. 6. Isopropanol (2-propanol): reagent grade isopropanol is used to overlay an SDS-PAGE gel while it is polymerizing. 7. Running buffer (10×): 250 mM Tris, 1.92 M glycine, and 1% (w/v) SDS. Do not attempt to pH this solution. Store at room temperature. 8. Sample buffer (2×): 20% (v/v) glycerol, 10% (v/v) 2-mercaptoethanol, 4% (w/v) SDS, 0.125  M Tris–HCl, pH 6.8, and 0.01% (w/v) bromophenol blue.

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9. Pre-stained molecular weight markers: e.g., Precision Plus Protein dual color markers (Bio-Rad, Hercules, CA, USA). 10. Fixing solution: 10% (v/v) acetic acid and 30% (v/v) methanol, made up to a final volume of 1 L with distilled water. 2.4. Fluorography Solutions

1. Two 500  mL bottles of reagent grade dimethyl sulfoxide (DMSO) will be required for this protocol. Designate the first bottle as DMSO-Wash 1 and the second bottle as DMSOWash 2. 2. 20% 2,5-diphenyloxazole (PPO)-DMSO solution: 20% (w/v) PPO is dissolved directly in DMSO. Hence, prepare 500 mL by dissolving 100 g of PPO directly into a 500-mL bottle of DMSO. This bottle will become the final wash solution and is designated as the 20% PPO-DMSO wash.

2.5. Equipment and Consumables

1. Eppendorf Germany).

5810R

centrifuge

(Eppendorf,

Hamburg,

2. Eppendorf swinging-bucket A-4-81 rotor. 3. Bench-top microfuge (e.g., Eppendorf 5415C). 4. Kodak BioMax film (Eastman Kodak Company, Rochester, NY, USA). 5. Kodak X-ray film holder cassette (8 × 10 in.). 6. Hoeffer SE400 single gel system or Hoeffer SE600 dual gel system (GE Healthcare, Piscataway, NJ, USA). 7. Hoeffer slab gel dryer (Model GD2000). 8. Saran Wrap, colorless plastic wrapping film (S. C. Johnson & Son, Inc., USA). 9. Standard 1.5-mL Eppendorf tubes. 10. Gilson pipettors and tips (Gilson, Middleton, WI, USA).

3. Methods 3.1. Isolation of Arabidopsis Chloroplasts 3.2. Preparation of Radiolabeled Translation Product for use in Import Assays

The isolation of Arabidopsis chloroplasts is described in Chapter 17, Vol. 1. 1. Obtain purified plasmid (final concentration 1  mg/mL) containing your cDNA of choice downstream from a T7, T3, or SP6 RNA polymerase promoter. This template should be free of ethanol, calcium, RNase, and salt. DNA isolated from most commercially available DNA purification kits or the standard alkaline lysate method (21) will work with TNT® reactions.

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2. The following translation protocol described below utilizes the TNT®-coupled reticulocyte lysate system; however, the protocol can also utilize the TNT®-coupled wheat-germ extract. Gradually thaw all the reagents to room temperature and then maintain them on ice while the components of the translation mixture are being assembled. Assemble translation reactions in a standard 1.5-mL Eppendorf tube as detailed below; reagents in bold are provided by the translation kit: 37.5 mL

TNT® Rabbit Reticulocyte Lysate

3 mL

TNT® Reaction Buffer

1.5 mL

TNT® RNA Polymerase (either SP6, T3, or T7)

1.5 mL

Amino Acid Mixture, 1 mM (Minus Methionine)

3 mL

[35S]-Methionine (1,000 Ci/mmol at 10 mCi/mL)

1.5 mL

RNasin® Ribonuclease Inhibitor (40 U/mL)

7.5 mL

DNA Template (1 mg/mL)

19.5 mL

Nuclease-Free Water

While the recipe above describes a translation reaction using [35S]-methionine, essentially the same translation recipe can be used when using [3H]-leucine as the radio label. 3. Incubate the translation reaction (total volume [Vt] = 75  mL) for 60–90 min at either 30°C or alternatively at room temperature (Caution: you are now working with radioactive material) (see Note 6). 4. Upon completion of the translation reaction, dilute the translation mixture with an equal volume (i.e., 75 mL) of translation product dilution buffer (from item 4 in Subheading  2.2). When labeling with [35S]-methionine, dilute with 50  mM “cold” l-methionine in 1× import buffer. Alternatively, when labeling with [3H]-leucine, dilute with 50 mM “cold” l-leucine dissolved in 1× import buffer. 5. Use the diluted translation product (Vt = 150 mL) immediately for several import assays; alternatively, it can be stored at −80°C until needed (see Note 7). 3.3. Large-Scale Import Assay

Typically, a large-scale import assay (Vt = 450 mL) is performed followed by a “dual protease” protection assay. The large-scale reaction will be subsequently divided into three 150-mL aliquots to be either mock treated (−) or treated (+) with either thermolysin or trypsin. Refer to Fig. 2a for a schematic outline of the experimental design.

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Fig. 2. Schematic outline of the overall flow of a “dual protease” protection assay and representative results from analyses of several outer and inner envelope membrane proteins. (a) Schematic outline of the overall flow of a “dual protease” protection assay described in this chapter. Specific details of each step: (1) Arabidopsis chloroplasts isolation, (2) translation reaction, (3) import assay, (4) “dual protease” protection assay, (5) fractionation, and (6) SDS-PAGE and fluorography are provided in various sections of this method chapter. OEM, outer envelope membrane; IMS, Intermembrane space; IEM, inner envelope membrane. (b) Representative “dual protease” protection assay and fractionation analysis results for several outer and inner envelope membrane proteins. [35S]-labeled Arc6, TGD2, Tic40, or Toc33 was imported into chloroplasts and then either not treated (−) or treated (+) with thermolysin or trypsin. After quenching the protease reaction, intact chloroplasts were recovered by sedimentation through a 40% Percoll cushion and recovered intact chloroplasts were fractionated into a crude membrane (P) and soluble (S) fraction. All samples were analyzed by SDS-PAGE and fluorography. TP, 10% of translation product added to an import reaction is shown; pr, precursor; m, mature; (*) Protease protected fragment of Arc6. For more details on the import and protease treatment of TGD2-GFP, Awai et al. (15). Toc33, an outer envelope membrane protein serves as a control to ensure that both thermolysin and trypsin protease activities are functioning correctly (i.e., Toc33 is digested by both proteases) (10).

1. Assemble a large-scale import assay in a 1.5-mL Eppendorf tube as follows (Vt of the import assay is 450 µL): 150 mL

Isolated Arabidopsis chloroplasts (1 mg chlorophyll/mL)

150 mL

Diluted translation product (from step 5 in Subheading 3.2)

18 mL

100 mM Mg-ATP/1× import buffer

132 mL

1× import buffer

2. Incubate the import reaction at room temperature, under normal room light conditions, for 30 min. 3. Remove 150  mL aliquots sequentially from the large-scale import reaction and place them into three new 1.5-mL Eppendorf tubes.

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3.4. “Dual Protease” Protection Assay: Posttreatment of Import Assays with Thermolysin and Trypsin

1. Treat the three separate 150-mL import assay aliquots as follows: Tube #1 (Mock [−] protease treatment control): 150 mL of import reaction and 100 mL of 1× import buffer are added, to give a final volume of 250 mL. Tube # 2 (thermolysin-treated sample): 150  mL of import reaction, 25  mL of thermolysin stock solution (1 mg/mL, freshly prepared in 5 mM CaCl2/1× import buffer), and 75  mL of 1× import buffer are added, to give a final volume of 250 mL. Tube #3 (Trypsin Treated sample): 150  mL of import reaction, 25  mL of trypsin stock solution (1 mg/mL, freshly prepared in 1× import buffer), and 75 mL of 1× import buffer are added, to give a final volume of 250 mL. 2. Incubate all reactions on ice for 30 min. 3. To selectively quench each protease reaction, add the following to the tubes indicated: Tube # 1: add 50  mL 1× import buffer (mock quench); Tube # 2: add 50  mL of thermolysin quench solution (60 mM EDTA/1× import buffer); and Tube # 3: add 50 mL of trypsin inhibitor solution (1 mg/mL in 1× import buffer). 4. Quench all reactions on ice for an additional 5 min. 5. Carefully apply each reaction (Vt = 300 mL) to the top of a separate 1.0 mL 40% (v/v) Percoll cushion in 1× import buffer (see Fig. 2a); except the thermolysin digestion reaction (Tube #2) which should be applied to a 40% (v/v) Percoll cushion containing 5 mM EDTA. 6. Centrifuge the reactions using an Eppendorf 5810R centrifuge at 3,000 rpm (1,811 × g) for 5 min at 4°C using a swingingbucket Eppendorf A-4-81 rotor. Intact chloroplasts will pellet to the bottom of the Percoll cushion, while broken chloroplasts and excess rabbit reticulocyte lysate translation mix (red in color) will remain at the top of the Percoll cushion. 7. Using a Gilson Pipetman, carefully remove the entire Percoll cushion from the green pellet located at the bottom of the tube (Caution: dispose of radioactivity into a proper waste container) (see Note 8). 8. Resuspend and wash the pellet of intact chloroplasts in 1.0 mL of 1× import buffer and centrifuge again using an Eppendorf 5810R centrifuge at 2,000  rpm (805 × g) for 5  min at 4°C using a swinging-bucket Eppendorf A-4-81 rotor. 9. The washed pellet of intact chloroplasts is now ready for lysis and crude fractionation.

3.5. Fractionation of (−/+) ProteaseTreated Chloroplasts

1. Resuspend each washed chloroplast pellet in 200  mL of lysis buffer and incubate on ice for 15–20 min. 2. Centrifuge the lysed chloroplasts at 14,000 g for 30 min using a bench-top microfuge.

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3. Carefully remove the supernatant (approximately 200 mL) and precipitate by adjusting to 80% acetone by adding 800  mL of reagent grade acetone that has been prechilled on ice. The supernatant is precipitated on ice for 20–30 min (as proteins in the supernatant begin to precipitate, the solution will gradually turn white). Finally, the precipitate is recovered by centrifugation at 14,000 × g for 5  min using a bench-top microfuge. Solubilize the resulting white pellet in 40 mL of 2× sample buffer. 4. Directly solubilize the pellet from step 2 above (green in color, representing a crude total membrane fraction) in 40 mL of 2× sample buffer. 5. Heat all samples to 95°C for 5 min and then immediately place them on ice. All samples can be directly analyzed by SDSPAGE, or stored at −80°C for later analysis. 3.6. SDS-PAGE

1. Analyze the import/“dual protease” protection assays by SDSPAGE using either a Hoeffer SE-400 single gel system or a Hoeffer SE-600 dual gel system (or an equivalent large electrophoresis system). The recipe described below is sufficient to prepare one SDS-PAGE gel. Clean the glass plates (16 × 18 cm) using a standard dish-washing detergent solution and allow to air-dry. After the plates have dried, wash them again with 100% ethanol and allow to air-dry. 2. To resolve a wide range of radiolabeled proteins and/or potential protein fragments with varying molecular weights, prepare a standard 13% gel with a thickness of 0.75  mm by mixing 5.0 mL of 4× resolving buffer with 9.0 mL of 30% acrylamide/ bis solution, 6.0 mL of distilled water, and 8.0 mL TEMED. (Caution: acrylamide is a neurotoxin when unpolymerized, so care should be taken to minimize contact with this solution). Mix the gel solution gently by swirling. To initiate polymerization of the gel, add 200 mL of a 10% (w/v) ammonium persulfate solution to the gel solution, mix, and pour into the gel plate sandwich. Continue pouring the gel solution until it reaches a level 4 cm from the top of the gel plate; this will leave enough space for a stacking gel. Finally, overlay the resolving gel with isopropanol. Under the conditions described here, the gel should polymerize in about 10–15  min at room temperature. 3. Polymerization of the gel is complete when a solid interface between the resolving gel and the isopropanol layer has formed. You can now pour off the isopropanol. Rinse the top of the gel twice with distilled water to ensure that all of the residual isopropanol has been removed. 4. Prepare a 5% stacking gel with the following composition: 1.25 mL of 4× stacking buffer, 0.83 mL of 30% acrylamide/bis solution, 2.92 mL water, and 5 mL TEMED. Mix gently. Add

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50  mL of 10% ammonium persulfate solution and mix once again. Pour the stacking gel mixture on top of the polymerized resolving gel immediately and insert the comb (we normally use a 20-well comb of 0.75-mm thickness). The stacking gel will polymerize within 5–10 min at room temperature. 5. Prepare 1× running buffer by diluting 100 mL of the 10× running buffer stock solution with 900 mL of distilled water in a 1-L graduated cylinder. Cover the 1-L graduated cylinder with Parafilm and slowly invert several times to mix the buffer. This is a sufficient amount of running buffer to run either one gel on the Hoeffer SE-400 single gel system or two gels using the Hoeffer SE-600 dual gel system. 6. After the stacking gel has set, carefully remove the comb and use distilled water from a spray water bottle to rinse out the wells completely. Gently spray the wells to avoid distorting or damaging the shape of the wells. 7. Add 1× running buffer to the upper and lower chambers of the gel unit and also to all of the wells. From each import/“dual protease” protection assay, load 20 mL of sample to each well. (Recall that this is only half of the entire sample. Retain a portion of all of the samples just in case the SDS-PAGE gel needs to be re-run). In a separate lane, load the Precision Plus prestained molecular weight markers. Finally, in any unused lanes, merely load 20 mL of 1× sample buffer. 8. Complete the assembly of the gel unit and connect to a power supply. Begin running the gel at 20 mA (per gel) through the stacking gel and then increase to 30–40 mA (per gel) through the resolving gel. The running time for the gel under these conditions is approximately 2.5 h. Stop the gel before the blue tracking dye runs off the gel. 9. After the run is completed, use a plastic gel wedge and carefully separate the two gel plates. Usually, the gel will adhere to one plate or the other. Take the glass plate that the gel has adhered to and place it into a large Pyrex dish (gel side up). Add sufficient fixing solution to submerge the plate/gel complex completely. By gently tilting the dish back and forth several times, the gel will gradually detach itself from the glass plate. Now, the gel is ready for subsequent fluorography workup. 3.7. Fluorography

The fluorography technique can be used to reduce exposure times drastically for SDS-PAGE gels containing 3H-, 14C-, or 35S-labeled samples. There are many procedures to perform fluorography. Below is a protocol that is relatively cheap and quick to perform (see Note 9). 1. After your SDS-PAGE gel run has been completed, soak the gel in fixing solution. Make sure that the gel is floating in the fixing solution. Soak the gel with continuous rocking for 10 min.

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2. Remove the fixing solution. Dispose of the fixing solution in a proper “Radioactive” organic waste container. 3. Soak the gel in DMSO-Wash 1. Again, make sure that the gel is floating in the DMSO-Wash 1. Soak the gel with continuous rocking for 10–15 min at room temperature. 4. Return the DMSO-Wash 1 back into its original bottle. 5. Add fresh DMSO-Wash 2 solution to the gel. Once again, make sure that the gel is floating in the DMSO-Wash 2. Soak the gel with continuous rocking for 10–15 min at room temperature. 6. Return the DMSO-Wash 2 back into its original container. 7. Final wash: soak the gel in a 20% PPO-DMSO solution. Make sure that the gel is floating in the 20% PPO-DMSO solution. Soak the gel with continuous rocking for 10–15 min at room temperature. Then return the 20% PPO-DMSO solution back into its original bottle. All of the fluorography wash solutions can be reused several times until the PPO starts to precipitate in the 20% PPO-DMSO solution. When this occurs, it will serve as an indicator that you will need to prepare all new fluorography solutions. 8. Add running tap water to the dish containing your treated gel. The gel will immediately turn completely white as the PPO within the gel precipitates immediately (do not panic!!). Continue washing the gel to remove excess white PPO precipitate. Then soak your gel in fresh distilled water for 5 min to allow the gel to re-swell. 9. Create a “drying sandwich” composed of (in order) Saran wrap, your gel, and filter paper. 10. Place “drying sandwich” onto gel drying apparatus (Saran wrap-side up) and dry under heat (60°C) with vacuum for 1 h. These times and conditions are based on a 14 × 16 cm gel of 0.75-mm thickness. Thicker gels will require longer drying times. 11. In a dark room, place dried gels into a Kodak X-ray exposure holder cassette (8 × 10 in.) along with a single piece of Kodak BioMax film. The cassette is placed at −80°C for an overnight exposure or until one is ready to develop the film.

4. Notes 1. Thermolysin has proven to be a valuable protease to probe the surface of chloroplasts (22). Indeed, the conditions necessary to digest various chloroplastic OEM proteins that are exposed to the cytosol have been well established (22). Thermolysin is a 34.6-kDa thermostable extracellular metalloendopeptidase

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that is commercially prepared from Bacillus thermoproteolyticus. It requires one zinc ion for enzyme activity and four calcium ions for structural stability. Since thermolysin requires calcium ions, its protease activity can be easily quenched with EDTA. Thermolysin hydrolyzes protein bonds on the N-terminal side of hydrophobic amino acid residues and has a pH optimum of 8.0. Under proper conditions (i.e., concentrations of thermolysin between 50 and 150 mg/mL), thermolysin will not significantly penetrate beyond the OEM. This allows one to digest specifically OEM proteins that are exposed toward the cytosol without causing any major damage to the integrity of chloroplasts. Thus, after thermolysin treatment, intact chloroplasts can be easily recovered by sedimentation through a 40% (v/v) Percoll cushion and further analyzed by SDS-PAGE and fluorography. This critical property of thermolysin makes it a suitable protease to investigate the topology of both outer and inner envelope membrane proteins (see Fig. 1b). For example, we often use Toc33 as an OEM control protein to ensure that the protease activity of thermolysin is working. In this scenario, after thermolysin treatment, Toc33 is completely digested (see Fig. 2b) (10), whereas, the IEM control protein Arc6 is not affected by thermolysin. From this simple thermolysin digestion experiment, one can conclude that since Toc33 is sensitive to thermolysin digestion, it must be localized to the OEM with the majority of the protein extending into the cytosol. Thus, by performing a standard thermolysin digestion assay on intact chloroplasts, the topology of an OEM protein can be revealed. Finally, unlike Toc33, other OEM proteins that are partially sensitive to thermolysin can often leave a detectable membrane bound radiolabeled peptide fragment behind after protease treatment. Comparing the size of this protected radiolabeled fragment with the size of the full length radiolabeled protein not treated with the protease often allows one to make predictions about the topology of a given membrane protein located at the OEM. 2. Trypsin is a small 23.5-kDa protein and a member of the serine protease S1 family. Trypsin is commercially isolated from bovine pancreas and has an optimum pH of 8.0. Trypsin cleaves peptide chains mainly at the carboxyl side of the amino acids lysine or arginine, except when either is followed by proline. The activity of trypsin is not affected by the inhibitor tosyl phenylalanyl chloromethyl ketone (TPCK); however, it is easily quenched with the addition of trypsin inhibitor which forms a 1:1 stoichiometric dead-end complex with trypsin. Like thermolysin, trypsin has also been used extensively to probe the surface of intact chloroplasts and to determine the topology of countless envelope membrane proteins (9–20). Under proper

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conditions (i.e., concentrations of trypsin between 100 and 200 mg/mL), trypsin has an additional advantage over thermolysin in that trypsin can partially disrupt the integrity of the OEM and thus gain access to the intermembrane space (IMS) where it can digest regions of OEM proteins or IEM proteins that extend into the IMS (see Fig. 1b). However, even though trypsin can partial disrupt the OEM, intact chloroplasts can still be recovered by sedimentation through a 40% (v/v) Percoll cushion and further analyzed. For example, the IEM control protein Arc6 is insensitive to thermolysin digestion (see Figs. 1b, 2b); however, during trypsin digestion, the small portion of trypsin that gains access to the IMS can partially digest the C-terminal domain of Arc6 which extends into the IMS (see Fig. 2b) (14). Thus, by employing the “dual protease” strategy using a combination of both trypsin and thermolysin, the topology of Arc6 was eventually determined (14). Unfortunately, the “dual protease” strategy was unable to determine the topology of several problematic IEM proteins initially. For instance, the topology of both TGD2 and APG1 could not be determined after using the “dual protease” approach and thus required additional steps to determine their respective topologies (15, 16) (see Fig. 1a and b). In the case of TGD2, preliminary evidence indicated that since TGD2 contained a large C-terminal domain that extended into the IMS, it should be readily sensitive to trypsin treatment (15). However, this was not the case. Consequently, an alternative approach was devised to finally determine TGD2’s topology. In this new approach, GFP was fused to the C-terminal end of TGD2, thus creating an even larger protein domain that extended into the IMS. This modification resulted in the TGD2–GFP fusion protein becoming sensitive to trypsin protease treatment, and thus allowed Awai et al. (15) to conclude that the C-terminal end of TGD2 did in fact extend into the IMS. In the case of APG1, Viana et al. (16) were forced to employ a more intensive strategy to elucidate its topology. Their strategy required the isolation of IEM vesicles and subsequent trypsin digestion treatments to determine the topology of APG1 finally (11). Usually, one does not need to undertake such extraordinary steps in order to determine the topology of an envelope membrane protein. However, these two examples illustrate that the “dual protease” digestion assay can be easily modified and/or coupled to many creative alternative strategies in order to determine the topology of recalcitrant IEM proteins. 3. One important and essential decision that must be made when pursuing the “dual protease” strategy is to determine how you will radiolabel your membrane protein prior to adding it to an import assay. In vitro translation kits can be conveniently used

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to radiolabel membrane proteins using either [35S]-methionine or [3H]-leucine. However, when deciding which radiolabeled amino acid to use, it is essential to look at the amino acid sequence of your membrane protein and determine both the number and the distribution of either methionines or leucines throughout your protein. In addition, you must confirm that either methionines or leucines are strategically located before and after any transmembrane domains (TMDs) present in your membrane protein. As a first choice, labeling with [35S]-methionine will produce a radioactive protein with a very high specific activity. However, if your protein has either a low number of methionine residues or the methionine residues are poorly distributed throughout the protein, then [35S]-methionine may not be a suitable choice for labeling your membrane protein. Alternatively, one can label a membrane protein using [3H]-leucine. In general, proteins usually contain a much higher number of leucine residues per protein when compared to methionine residues. Furthermore, the presence of a large number of leucine residues usually ensures that those leucines are widely distributed throughout the protein. Unfortunately, [3H]-leucine-labeled proteins generally have a lower specific activity and require longer exposure times during the fluorography procedure to see labeled peptide fragments. For the “dual protease” strategy to function in helping to determine the topology of a membrane protein, you have to make sure that any membrane bound fragments generated after a protease treatment will be radiolabeled and thus be detectable by subsequent fluorography visualization. It is the generation and detection of different size radiolabeled membrane fragments that will aid you in reconstructing the topology of your particular membrane protein. 4. The TNT® Coupled Reticulocyte Lysate or the TNT® Coupled Wheat-germ Extract Transcription/Translation kits can be used to conveniently prepare radio-labeled precursor protein. Both transcription/translation systems can be used to synthesize fulllength radio-labeled proteins between 10 and 180 kDa in size. Likewise, both transcription/translation systems are compatible with our import assays and DO NOT cause premature lysis of chloroplasts. In our hands, we find that the TNT® Coupled Reticulocyte Lysate is more robust than the TNT® Coupled Wheat-germ translation system in synthesizing radio-labeled protein. Thus, we routinely use this system. However, we have come across unique occasions in which a particular plant cDNA is translated more efficiently when using the Wheat-germ system rather than the Rabbit Reticulocyte Lysate system. 5. Once the translation product (TP) has been synthesized, it is essential to dilute the TP with either “cold” (i.e., nonradioactive) methionine or leucine depending on what radioactive amino

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acid was used to label your protein. The reason for this dilution step is as follows: Regardless of the radiolabeled amino acid used (i.e., [35S]-methionine or [3H]-leucine), the majority of the radiolabeled amino acid added to the translation ­reaction is not incorporated into newly synthesized labeled TP. Consequently, the unincorporated [35S]-methionine or [3H]-leucine can be readily taken up by isolated chloroplasts and subsequently utilized by the chloroplastic translation machinery to synthesize endogenous radio label proteins. The generation of endogenous radiolabeled proteins creates a significant background problem and makes interpretation of fluorograms practically impossible. Thus, to avoid the incorporation of either free [35S]-methionine or [3H]-leucine into background endogenous chloroplastic proteins, large amounts of nonradioactive methionine or leucine is added to the TP mix. The addition of nonradioactive amino acids essentially dilutes out the low levels of free residual [35S]-methionine or [3H]-leucine, thus preventing their incorporation into endogenous chloroplastic proteins. This dilution step will also ensure that the only radioactive band(s) visualized on a fluorogram will have originated only from the added radiolabeled translation product. 6. Cautionary note: Once the translation product is synthesized and then added to an import assay, all subsequent steps (3.2–3.6) must be treated with caution since each step involves the handling of radioactive material. Likewise, all radioactive waste (i.e., both solid and liquid) generated during the course of the experiment must be disposed of in an approved “Radioactive Waste” container. 7. After synthesis and dilution of the TP is completed, one can either use the TP immediately or store the TP at −80°C until needed. The TP is still import competent even after storage for up to 1 week; however, long-term storage and/or multiple freeze-thawing of the TP should be avoided at all cost. 8. Cautionary note: The top portion of the Percoll cushion (red in color when using rabbit reticulocyte lysate) contains essentially radiolabeled precursor proteins that were not imported into chloroplasts and free radiolabeled amino acids (either [35S]-methionine or [3H]-leucine). Hence, the top portion of the Percoll cushion is highly radioactive and must be removed carefully and disposed of in the appropriate “Radioactive Waste” container. 9. Cautionary note: When performing fluorography, one must remember that all of the wash solutions should be treated as “Radioactive Solutions.” Consequently, appropriate steps should be taken to ensure that radioactivity is not being spread throughout the laboratory during the wash procedure.

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Designating a restricted, “Radioactive Work Area” will help minimize the chance of spreading radioactivity in unwanted areas throughout the laboratory.

Acknowledgments I wish to thank Drs. Jon Glynn and Kathy Osteryoung for the generous gift of the prARC6/pBluescript plasmid and Dr. Christoph Benning for the gift of the prTGD2/pGEM-TEasy plasmid. The author is funded by DOE Grant no. DE-FG02-91ER20021 to Ken Keegstra. References 1. Gutensohn, M., Fan, E., Frielingsdorf, S., Hanner, P., Hou, B., Hust, B., and Klosgen, R. B. (2006) Toc, Tic, Tat et  al.: structure and function of protein transport machineries in chloroplasts. J. Plant Physiol. 163, 333–347. 2. Jarvis, P., and Robinson, C. (2004) Mechanisms of protein import and routing in chloroplasts. Curr. Biol. 14, R1064-R1077. 3. Schwacke, R., Schneider, A., van der Graaff, E., Fischer, K., Catoni, E., Desimone, M., Frommer, W. B., Flugge, U. I., and Kunze, R. (2003) ARAMEMNON, a novel database for Arabidopsis integral membrane proteins. Plant Physiol. 131, 16–26. 4. van Wijk, K.J. (2004) Plastid proteomics. Plant Physiol. Biochem. 42, 963–977. 5. Marmagne, A., Salvi, D., Rolland, N., Ephritikhine, G., Joyard, J., and BarbierBrygoo, H. (2006) Purification and fractionation of membranes for proteomic analyses. Methods Mol. Biol. 323, 403–420. 6. Froehlich, J. E., Wilkerson, C. G., Ray, W. K., McAndrew, R. S., Osteryoung, K. W., Gage, D. A., and Phinney, B. S. (2003) Proteomic study of the Arabidopsis thaliana chloroplastic envelope membrane utilizing alternatives to traditional two-dimensional electrophoresis. J. Proteome Res. 2, 413–425. 7. Ferro, M., Salvi, D., Brugiere, S., Miras, S., Kowalski, S., Louwagie, M., Garin, J., Joyard, J., and Rolland, N. (2003) Proteomics of the chloroplast envelope membranes from Arabidopsis thaliana. Mol. Cell. Proteomics 2, 325–345. 8. Rolland, N., Ferro, M., Seigneurin-Berny, D., Garin, J., Douce, R., and Joyard, J. (2003) Proteomics of chloroplast envelope membranes. Photosynth. Res. 78, 205–230.

9. Sveshnikova, N., Grimm, R., Soll J., and Schleiff, E. (2000) Topology studies of the chloroplast protein import channel Toc75. Biol. Chem. 381, 687–693. 10. Chen, D., and Schnell, D. J. (1997) Insertion of the 34-kDa chloroplast protein import ­component, IAP34, into the chloroplast outer membrane is dependent on its intrinsic GTPbinding capacity. J. Biol. Chem. 272, 6614–6620. 11. Li, H.-M., Moore, T., and Keegstra, K. (1991) Targeting of proteins to the outer envelope membrane uses a different pathway than transport into chloroplasts. Plant Cell 3, 709–717. 12. Froehlich, J. E., Itoh, A., and Howe, G. A. (2001) Tomato allene oxide synthase and fatty acid hydroperoxide lyase, two cytochrome P450s involved in oxylipin metabolism, are targeted to different membranes of chloroplast envelope. Plant Physiol. 125, 306–317. 13. Froehlich, J. E., Benning, C., and Dörmann, P. (2001) The digalactosyldiacylglycerol (DGDG) synthase DGD1 is inserted into the outer envelope membrane of chloroplasts in a manner independent of the general import pathway and does not depend on direct interaction with monogalactosyldiacylglycerol synthase for DGDG biosynthesis. J. Biol. Chem. 276, 31806–31812. 14. 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. 15. Awai, K., Xu, C., Tamot. B., and Benning, C. (2006) A phosphatidic acid-binding protein of the chloroplast inner envelope membrane

21  Studying Arabidopsis Envelope Protein Localization and Topology… involved in lipid trafficking. Proc. Natl. Acad. Sci. USA 103, 10817–10822. 16. Viana, A. A., Li, M., and Schnell, D.J. (2010) Determinants for stop-transfer and post-import pathways for protein targeting to the chloroplast inner envelope membrane. J. Biol. Chem. 285, 12948–12960. 17. Dreses-Werringloer, U., Fischer, K., Wachter, E., Link, T. A., and Flugge, U. I. (1991) cDNA sequence and deduced amino acid sequence of the precursor of the 37-kDa inner envelope membrane polypeptide from spinach chloroplasts. Its transit peptide contains an amphiphilic alpha-helix as the only detectable structural element. Eur. J. Biochem. 195, 361–368. 18. Tripp, J., Inoue, K. , Keegstra, K., and Froehlich, J. E. (2007) A novel serine/proline-rich domain in combination with a transmembrane domain is required for the insertion of AtTic40 into the

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inner envelope membrane of chloroplasts. Plant J. 52, 824–838. 19. Li, M., and Schnell, D. J. (2006) Reconstitution of protein targeting to the inner envelope membrane of chloroplasts. J. Cell Biol. 175, 249–259. 20. Chou, M.-L., Fitzpatrick, L. M., Tu, S.-L., Budziszewski, G., Potter-Lewis, S., Akita, M., Levin, J. Z., Keegstra, K., and Li, H.-M. (2003) Tic40, a membrane-anchored co-chaperone homolog in the chloroplast protein translocon. EMBO J. 22, 2970–2980. 21. Sambrook, J., Fritsch, E. F., and Maniatis, T. (1989) Molecular Cloning: A Laboratory Manual, 2nd edn. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, NY, USA. 22. Cline, K., Werner-Washburne, M., Andrews, J., and Keegstra K. (1984) Thermolysin is a suitable protease for probing the surface of intact pea chloroplasts. Plant Physiol. 75, 675–678.

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Index

A aadA���������������������������������������������������������134, 135, 143, 144 ABRC. See Arabidopsis Biological Resource Centre Accumulation response............ 87, 88, 90–93, 95, 97, 98, 100 Alkaline extraction................................................... 339–350 Amyloplast...................................... 29, 73, 74, 103–111, 113 Antibody (antibodies)..................... 33, 36–37, 41, 46, 48–50, 52, 53, 55, 60–62, 67–70, 228, 230, 232, 236, 239, 314–318, 340 Antigen retrieval................................................36, 41, 47, 52 Apyrase.............................................. 310, 313, 326, 333, 337 Arabidopsis Biological Resource Centre (ABRC)........ 11, 30 ARAMEMNON.......253, 258, 260, 266–271, 273–274, 352 AT_CHLORO................................................ 253, 260, 262 ATP synthase........................................................... 247, 352 ATP synthesis................................................................... 313 Autofluorescence........35–37, 47–52, 63, 66, 67, 89, 161, 162 Autoradiography........174, 178, 212, 220, 222, 224, 314, 323, 327, 332–335, 337, 340, 346–349 Avoidance response....................... 87, 88, 90–93, 95–98, 100

B Basic Local Alignment Search Tool (BLAST).............................................. 267, 271–272 Bioinformatics...........................................250, 253, 255, 258 Biolistic (gun, particle delivery system, particle bombardment)...................73–83, 134, 138–143, 145 BLAST. See Basic Local Alignment Search Tool Blot dot (or slot)..........................................172–178, 180, 188 Northern.............. 172, 184–186, 214–215, 218, 221–222 Southern..................................................................... 144 Western (or immunoblot)......... 34, 36, 48, 60–62, 67–70, 227–228, 230, 232, 233, 235–236, 239, 315–318, 340, 346, 347

C Carbonyl cyanide 3-chlorophenylhydrazone (CCCP)..... 326, 332, 333, 336

CCCP. See Carbonyl cyanide 3-chlorophenylhydrazone cDNA�����������������������186, 221, 269, 285, 289–291, 310, 313, 340, 343, 355, 364 Cetyltrimethylammonium bromide (CTAB).......... 156, 164, 167, 168 CFP. See Cyan fluorescent protein Chemical cross-linkers DSP����������������������������������������������������������� 311, 316–318 DTME....................................................... 310, 314–316 Chemiluminescence (or ECL)..... 62, 69, 166, 203, 206, 228, 236, 239 ChloroP..................................... 248–250, 260–262, 271, 272 Chlorophyll fluorescence.................... 5, 8, 10, 15, 16, 27, 30, 31, 48, 50, 52, 63, 66, 67, 76, 80, 83, 89, 100, 161, 162 Chloroplast DNA (cpDNA) analysis................................................151–169, 185, 196 preparation, extraction................................ 154, 162–163 Chloroplast Function Database.......................................... 11 Chloroplast isolation..........................34, 152–153, 157–159, 166, 172–174, 176, 179, 230, 231, 283–286, 291–293, 300, 309, 311–313, 318, 324–325, 328–330, 340, 342, 344–345, 349 Chloroplast movement............................................... 87–101 Chromatography affinity����������������������������������������������������������������������� 187 size exclusion. See Gel filtration Chromoplast............................................................... 73, 113 Confocal microscopy.......... 28–31, 37–38, 42, 50–51, 74–77, 79–81, 93 Coomassie stain..........204, 227, 233–235, 239, 288, 297, 301 cpDNA. See Chloroplast DNA or Plastid DNA (ptDNA) Cre/loxP................................................................... 134, 135 Cross-linking.....................35, 36, 43, 60, 156, 166, 175, 189, 206, 221, 308–311, 314–318 CTAB. See Cetyltrimethylammonium bromide Cyan fluorescent protein (CFP)............................. 75–77, 81

D DAPI. See 4’,6-Diamidino–2-phenylindole

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, © Springer Science+Business Media, LLC 2011

369

Chloroplast Research in Arabidopsis 370  Index

  

Databases ARAMEMNON���������������������� 253, 258, 260, 266–271, 273–274, 352 AT_CHLORO.......................................... 253, 260, 262 Chromoplast 2010........................................................ 11 Chloroplast Function Database.................................... 11 Genevestigator............................................................ 268 insertional mutants....................................... 4, 11–13, 16 organelle................................ 11, 253, 256, 263–271, 352 Plant Proteome DataBase (PPDB)............253, 256–258, 260, 262 proteomics...........................253–254, 256–258, 263–266 Salk Institute Genomic Analysis Laboratory (SIGnAL)................................ 12, 13, 16 SUBA (SUBA II)........ 253, 254, 260, 263–266, 272–273 The Arabidopsis Information Resource (TAIR)....................... 12, 16, 254, 263, 265, 268, 272 Degradation assay, in organello................226–227, 229, 230, 232–234, 238 Detergent.......................21, 40, 236–237, 314, 315, 317, 359 Dexamethasone (DEX)............................ 137–139, 141–144 4’,6-Diamidino–2-phenylindole (DAPI).........153, 159–161, 164, 168–169 Dithio-bismaleimidoethane (DTME)............. 310, 314–316 Dithiobis[succinimidyl propionate] (DSP)...... 311, 316–318 DNA analysis............................................12, 16, 144, 151–169 cpDNA................................ 151, 152, 154–159, 162–169 preparation, extraction...... 6, 13–14, 62–63, 81, 154, 156, 163, 164, 167, 174–176, 285, 289–290, 355 ptDNA....................................................... 133–135, 144 Dot blot.....................................................172–178, 180, 188 1D-PAGE (or 1D SDS-PAGE)............................. 227, 230, 232–235. See also One-dimensional PAGE 2D-PAGE. See Two-dimensional PAGE DpH (delta pH, pH gradient)........................... 246, 322, 336 DSP. See Dithiobis[succinimidyl propionate] DTME. See Dithio-bismaleimidoethane

E Electron microscopy..............................77, 89, 113–131, 340 Electrophoresis agarose������ 6, 14, 154, 155, 162–164, 167, 207, 214, 215, 221, 223, 285, 290 denaturing...................184, 191, 194–195, 214, 221, 227, 228, 230, 232–234, 236–237, 287–288 1D-PAGE (or 1D SDS-PAGE)....................... 227, 230, 232–235 native����������������������������������������������������������������� 189, 206 polyacrylamide........................ 63, 68, 184, 191, 194–196, 202, 205–207, 210–211, 216–217, 232, 237, 285, 326–327, 332, 334, 354–355 RNA��������������������������� 183–192, 199–207, 214, 221–222 SDS-PAGE....... 63, 68, 69, 204, 211, 212, 216–220, 223,

237, 285, 287–288, 290, 291, 296–299, 310, 315, 333, 334, 347, 349, 354–355, 357, 359–360, 362 two-dimensional PAGE..............228, 230, 232, 236–237 urea����������������������194–195, 202, 205, 207, 228, 236–237 Electrophoresis mobility shift assay (EMSA).................. 189, 201–203, 205–207 Embedding..........................34–36, 40–41, 43–47, 52, 54, 55, 114–117, 121–123, 127, 130 EMS. See Ethyl methanesulfonate EMSA. See Electrophoresis mobility shift assay Enhancer mutants........................................................ 3, 4, 6 Envelope inner envelope membrane (IEM).......244–246, 254, 270, 308, 340–341, 346–350, 352, 353, 357, 362–363 outer envelope membrane (OEM).....62–63, 66, 67, 254, 340–341, 346, 347, 352, 353, 357, 361–363 protein topology..................254–258, 268–271, 351–366 proteomics.......................................................... 244, 260 purification, fractionation.... 328–329, 339–350, 352, 357 Ethyl methanesulfonate (EMS)....................5–7, 15, 40, 229 Etioplast..................................................................... 73, 119

F Fatty acid...................................................282, 339, 351–352 Fixation (fixative) aldehyde.........................20–21, 35, 36, 39, 43, 47–50, 61, 67, 116–118, 127, 129, 153, 159–161 primary................................................117, 119, 120, 129 secondary.............................................117, 119, 120, 128 tertiary��������������������������������������������������������������� 117, 120 Flow cytometry................................................153–154, 158, 161–162 Fluorescence microscopy.....19–56, 60, 63, 66–68, 70, 74–77, 80–81, 100, 153, 154, 159–163, 340 Fluorography.....297–299, 301, 334–335, 355, 357, 360–362, 364, 365 Forward genetics (forward-genetic)................................ 4–11

G Gel filtration (size exclusion chromatography)................................... 188, 314, 319 Gel shift assay (gel mobility shift, GMS, EMSA)........... 189, 201–203, 205–207 Genetics (genetic screens) forward..................................................................... 4–11 modifiers................................................................. 3, 4, 6 reporter based....................................................... 5, 8–10 reverse�����������������������������������������������������4, 6, 11–14, 184 Genevestigator.................................................................. 268 Genotyping.................................................................. 12–14 GFP. See Green fluorescent protein Glycerate.................................................................. 310, 313 Gradient centrifugation Percoll��������� 152, 157–159, 168, 173, 176, 179, 231, 286,

Chloroplast Research in Arabidopsis 371 Index     

291–293, 300, 309–310, 312, 318–319, 325, 328–330, 344–345 sucrose���������213, 218, 220, 329, 340–341, 345–346, 350 Gravitropism (gravitropic)........................................ 103–111 Green fluorescent protein (GFP)....... 5, 8–10, 16, 29, 30, 33, 36–37, 61–63, 66, 67, 70, 74–77, 79, 81, 82, 89, 253, 254, 263–265, 353, 357, 363

H Homogenization....................... 157, 173, 174, 176, 179, 212, 218, 226, 231, 283, 284, 288, 291, 292, 300–302, 312, 318, 324, 327, 328, 330, 334, 335 Homogenizer kitchen or domestic blender....................... 300, 342, 344 pestle (and mortar, or microtube).................6, 13, 14, 53, 153, 156, 157, 167, 192, 213, 220 rotor-stator (Polytron, Ultra-Turrax, Kinematica)..... 227, 286, 292, 300, 309, 312, 342 Waring blender.................... 152, 323–324, 327, 328, 330 Homoplastomic........................................................ 134, 144 Horseradish peroxidase (HRP).......................... 62, 228, 239 HPLC������������������������������������������������������194, 232, 235, 237 Hybridization....155–156, 158, 165–168, 172–180, 184–185, 188, 214, 215, 218, 221–222

I ImageJ������� 20, 42, 51, 53, 101, 107, 109, 153, 154, 161, 165 ImageQuant....................................................5, 10, 288, 299 Image acquisition and/or analysis................. 5, 20, 24, 31, 51, 101, 104, 105, 107, 109, 110, 127 Immunoblot (or Western blot)....... 34, 36, 48, 60–62, 67–70, 227–228, 230, 232, 233, 235–236, 239, 315–318, 340, 346, 347 Immunocytochemistry.......................................60, 61, 67, 70 Immunofluorescence microscopy.................................. 33–56 Immunoprecipitation (coimmunoprecipitation)................ 60, 187–188, 190, 314, 316–318 In-gel digestion (trypsin).................................................. 363 Insertional mutagen........................................................ 3–17 In situ hybridization........................................................... 36 Intermembrane space........................270, 353, 357, 362–363 In vitro transcription......... 173–174, 177, 190–191, 204–205 in vitro transcription/translation, TNT............285, 289–291, 299–300, 310, 313–314, 323, 325–326, 331–332, 336, 342, 343, 354–356, 363–365 In vivo labeling......................................... 211–212, 217–218 Iodine�����������������������������������������������������������21, 28, 104, 105

lTP. See Luminal transit peptide Lugol��������������������������������������������������21, 104–106, 109, 110 Lumen���������������244, 246, 248, 250, 252–254, 270, 272, 322 LumenP�������������������������������������������� 248, 250, 252, 262, 272 Luminal transit peptide (lTP).......... 246, 248, 250, 252, 262, 272, 321–323

M Macrocarrier..................................................78, 80, 138, 141 Mass spectrometry.............230, 232–237, 253, 254, 256, 258, 263–265 Microarray........................................................ 172, 184, 188 Microcarrier...........................................78, 80, 138, 140–142 Microscopy confocal........28–31, 37–38, 42, 50–51, 74–77, 79–81, 93 electron............................................77, 89, 113–131, 340 epifluorescence.............................................75, 79, 80, 93 fluorescence.......19–56, 60, 63, 66–68, 70, 74–77, 80–81, 100, 153, 154, 159–163, 340 immunocytochemisty............................60, 61, 67, 68, 70 immunofluorescence............................................... 33–56 TEM��������������������������������������������������118, 120–125, 127 Morphology, plastid���������������������������������������������������� 31, 77 Murashige and Skoog (Murashige–Skoog, MS medium)������������������� 5, 7–10, 16, 21, 78–79, 82, 96–97, 100, 104, 137–138, 210, 216–217, 223, 283–284, 288–289, 298–299, 301, 309, 312, 342, 344 Mutagenesis................................................................... 3–17 Mutant................ 3–17, 22, 25, 26, 36–37, 48, 52, 53, 60, 70, 76–77, 90, 91, 95, 104, 109–111, 136, 143, 165, 178, 179, 184, 186–188, 200, 210, 216, 218, 228–230, 232, 235, 237–239, 282, 283, 295, 298–302, 308, 309, 340

N Native electrophoresis (native PAGE)...................... 189, 206 NCBI���������������������������������������������������������������������� 268, 272 NEP. See Nuclear-encoded plastid RNA polymerase Nigericin................................................................... 310, 313 NIH image. See ImageJ Northern blot........................... 172, 184–186, 214–215, 218, 221–222 Nottingham Arabidopsis Stock Centre (or Centre) (NASC)............................................................... 5, 9, 11, 16 Nuclear-encoded plastid RNA polymerase (NEP)........... 172

L

O

Leucoplast����������������������������������������������������������������� 73, 113 Light-harvesting chlorophyll-binding protein (LHCP)...... 9, 218, 246, 322, 323, 333–334, 337 Lipid����������������������������������������� 237, 255, 268, 269, 271, 273

One-dimensional PAGE denaturant...........................................227, 230, 233–234 transfer........................................................................ 235 use������������������������������������������������������������������������������ 232

Chloroplast Research in Arabidopsis 372  Index

  

P Particle gun. See Biolistic PCR amplification of template, probe.........172, 174, 204, 285, 289–290, 299 genotyping (genomic PCR).................................... 12–14 quantitative (qPCR)....155–158, 163, 164, 168, 172, 184, 186 rapid amplification of cDNA ends (RACE)...... 172, 186, 193, 195 RT-PCR............................................................. 185, 186 PEG. See Polyethylene glycol Pentatricopeptide repeat (PPR).........192, 200–201, 203, 204 PEP. See Plastid-encoded plastid RNA polymerase Percoll cushion�����������������������159, 296, 316, 345, 353, 357, 358, 362, 363, 365 gradient........................152, 157–159, 168, 173, 176, 179, 231, 286, 291–293, 300, 309–310, 312, 318–319, 325, 328–330, 344–345 pH gradient (DpH, delta pH)........................... 246, 322, 336 Phenol�������������������������������173, 177, 180, 213, 220, 325, 331 Phenylmethylsulfonyl flurpide (PMSF)...........154, 162, 163, 201, 212, 222 Photorelocation.......................................................... 87–101 Photosynthesis photosynthetic efficiency................................................8 photosynthetic pigments....19, 20, 26, 28, 35, 48, 50, 179 photosynthetic protein complexes.............................. 352 Photosystem.......................200, 209, 217, 218, 247, 347, 352 Phototropin����������������������������������������������������������������������� 91 Plant growth conditions....................4, 7–10, 14, 15, 28, 60, 69, 79–81, 104–108, 186, 210, 216, 229, 283–284, 288–289, 299, 301, 309, 312, 342, 344, 349 medium. see Murashige and Skoog Plant Proteome DataBase (PPDB)...253, 256–258, 260, 262 Plant regeneration.....................................134, 135, 137, 144 Plasmid.............. 60, 62–63, 65, 78, 79, 81–82, 135–136, 138, 140–143, 204, 285, 289–290, 299, 310, 313–314, 319, 343, 349, 355–356 Plastid DNA (ptDNA)...............................75, 133–135, 144 Plastid-encoded plastid RNA polymerase (PEP)..... 171–172 Plastome....................................................133–135, 171, 172 PMF. See Proton motive force PMSF. See Phenylmethylsulfonyl flurpide Poisoned-primer extension assay.............. 184–187, 192–196 Polyethylene glycol (PEG).................. 35, 60, 61, 65, 75, 173 Polysome assembly, isolation and purification.......... 209–224 Post-translation modification................................... 271, 273 PPDB. See Plant Proteome DataBase PPR. See Pentatricopeptide repeat Predotar......................248–250, 254, 259, 260, 262–263, 272

PredSL..............................................248–250, 252, 262, 272 Preparation (purification, isolation) of chloroplasts..........152–153, 157–159, 168, 173, 176, 179, 231–232, 237–238, 285–287, 291–294, 300, 309–313, 318–319, 324–325, 328–330, 335, 342, 344–345, 349 DNA�������� 6, 13–14, 62–63, 81, 154, 156, 163, 164, 167, 174–176, 285, 289–290, 355 envelope membranes... 324, 328–329, 336, 340–341, 343, 345–347, 349–350 polysomes....................................212–213, 220–221, 223 proteins...... 61, 67–68, 188, 201, 203–204, 212, 218–219, 343, 345–348 ribosomes.....................................212–213, 220–221, 223 RNA��� 173, 177, 179–180, 192, 202, 204–205, 212–213, 220–221, 325–326, 331 stroma������������������������ 218–219, 232, 324–325, 328–330, 336, 340–341, 343, 345–347 thylakoids................... 218–219, 232, 323–325, 327–330, 335–336, 340–341, 343, 345–347, 349–350 preSSU (pSS).............285, 290, 291, 296–298, 310, 313–316 Protease inhibitors.............................................. 61, 222, 309 Proteases, chloroplast Clp�����������������������225, 226, 229, 230, 233, 234, 237, 238 Deg�������������������������������������������������������������������� 225, 237 FtsH������������������������������������������������������������������ 225, 237 Lon�������������������������������������������������������������������� 225, 237 Protein complexes (or multiprotein complexes)�������������� 77, 189, 190, 200, 209, 218, 226, 244, 245, 308, 309, 317, 321, 322, 352 degradation............. 34–35, 210, 216–219, 223, 225–239, 291, 302, 311–312, 333, 336, 361–363 extraction............61, 67–68, 212, 218–219, 343, 347–349 import�������������������60, 62, 66, 69–70, 243–247, 281–303, 307–319, 339–366 insertion or integration................245–247, 308, 321–337 interactions.................... 60, 187–192, 199–207, 307–319 labeling................211–212, 217–218, 223, 285, 289–291, 299–300, 310, 313–314, 323, 325–326, 331–332, 336, 342, 343, 354–356, 363–365 localization����������������� 33–56, 66–68, 243–274, 339–365 precipitation������������������������� 60, 163, 187–188, 236–237, 314–318, 348, 359 purification.......................... 188, 201, 203–204, 236–237 quantification.......212, 219, 229, 230, 235–237, 239, 299, 301, 315, 346 synthesis..............209–224, 285, 289–291, 299–300, 310, 313–314, 323, 325–326, 331–332, 336, 342, 343, 354–356, 363–365 targeting prediction............................................ 243–274 topology����������������������������������������������243–274, 351–365 translocation..........60, 62, 66, 69–70, 243–274, 281–303, 307–319, 321–337, 339–366, 370

Chloroplast Research in Arabidopsis 373 Index     

turnover...............................................218, 223, 225–239 Protein A.......................................................... 311, 316, 318 Protein import early import intermediate........................... 308, 313–316 energetic manipulation....................................... 307–319 mutants.......................................... 60, 283, 298, 308, 340 time-course................................................. 284, 295, 298 Proteomics chloroplast.......................................................... 281, 283 databases..............................253–254, 256–258, 263–266 envelope...................................................................... 352 Proton gradient................................................................. 313 Proton motive force (PMF).............................................. 322 Protoplast.........................20, 25–26, 32, 59–70, 75, 282, 283 pSS. See preSSU ptDNA. See Plastid DNA or Chloroplast DNA (cpDNA) Pulse-chase (pulse labelling).............211–212, 215–218, 223, 228, 229 Pulsed-field gel electrophoresis.................155, 159, 164–165

R Radiolabelling (radiolabeling)..................173–174, 176–178, 193–194, 202, 204–205, 211–212, 217–218, 223, 285, 289–291, 299–300, 310, 313–314, 323, 325–326, 331–332, 336, 342, 343, 354–356, 363–365 Recombination, homologous.................................... 134, 135 Red fluorescent protein (RFP)..........................30, 63, 66, 67 Resin embedding........................... 116–118, 121–123, 128, 130 epoxy����������������������������������� 115, 116, 118, 121–123, 128 Spurr’s (low viscosity, modified)...........115–118, 121, 128 Reticulocyte lysate.......60, 285, 289–291, 299, 310, 313–314, 342, 343, 349, 354, 356, 358, 364, 365 Reverse genetics (reverse-genetic).................4, 6, 11–14, 184 RFP. See Red fluorescent protein Ribosome (ribosomal)................ 74, 143, 187, 209, 213, 215, 216, 218, 229, 352 RNA analysis�������������� 171–180, 183–196, 199–207, 212–215, 220–222 binding protein (RBP).................184, 187–192, 199–207 editing�������������������������������������������������183, 184, 199–207 interference (RNAi).................................................... 229 messenger (mRNA).................................... 215–216, 218 preparation, extraction........ 173, 177, 179–180, 192, 202, 204–205, 212–213, 220–221, 325–326, 331 processing........................................... 183–196, 199–201 splicing....................................................... 183–196, 201 Roots culture�������������������������������������������������136–139, 141–143 imaging������������������������������ 21, 28–32, 34, 73, 74, 76–77, 103–111, 118–119 gravitropism........................................................ 103–111

staining................................................... 28–29, 104–106 Rubisco.......................................62, 73, 285. See also preSSU Run-on assay............................................................ 171–180

S Salk Institute Genomic Analysis Laboratory (SIGnAL)................................................... 12, 13, 16 Salk (SALK) lines.................................................. 12, 13, 16 SDS-polyacrylamide gel electrophoresis (PAGE)............. 63, 68, 69, 204, 211, 212, 216–220, 223, 237, 285, 287–288, 290, 291, 296–299, 301, 310, 315, 316, 318, 332–334, 340, 346–349, 354, 357, 359–360, 362 Sec (cpSec)........................ 246, 248, 321–323, 332, 333, 336 Sectioning cryosectioning.......................35, 40, 43, 44, 48, 50, 52, 54 microtomy.......................................40–41, 44, 46–47, 55 optical�������������������������������������������������������������� 30, 38, 81 ultramicrotomy............................ 114, 122, 123, 128, 129 Seed sowing����������������������������� 16, 28, 229, 278, 284, 289, 344 starch�������������������������������������� 28, 75, 104–106, 109–110 sterilization�����������������������7, 79, 104, 105, 139, 216, 223, 283, 288–289, 344 SIGnAL. See Salk Institute Genomic Analysis Laboratory SignalP...................................... 248, 251, 254, 262, 270, 272 Solubilization....................213, 220, 311, 314, 315, 317–318, 332, 334, 359 Sonication.......................................... 61, 63, 67–68, 201, 203 Southern blot.................................................................... 144 Spectinomycin...................................134, 136, 139, 142–144 Splicing..............................................183–187, 192–194, 201 Spurr’s resin...............................................115–118, 121, 128 SRP (spSRP).....................................246–247, 322–323, 337 Starch analysis....................................................28, 29, 103–111 grain (granule)...........................................28, 29, 74, 105 removal reduction....................................... 238, 291, 335 sedimentation..................................................... 103–111 staining....................................28, 29, 103–106, 109–110 Statocyte................................................................... 103, 104 Statolith.............................................................. 28, 103–111 Steedman’s wax............................................35, 36, 40, 44, 52 Stock centre (stock center) ABRC������������������������������������������������������������������� 11, 30 NASC�������������������������������������������������������������5, 9, 11, 16 RIKEN������������������������������������������������������������ 11, 13, 16 Streptomycin��������������������������������������������134, 139, 142–144 Stroma����������������������67, 73–75, 83, 127, 218–219, 226, 230, 232–234, 236–238, 244–246, 254, 270, 313, 321–323, 326, 328–330, 332, 333, 336, 337, 340–341, 346–348, 353 Stromules��������������������������������������������������������������������� 73–83 SUBA (SUBA II)������������� 253, 254, 260, 263–266, 272–273

Chloroplast Research in Arabidopsis 374  Index

  

Sucrose as cryoprotectant............................................... 35, 40, 43 gradient..........................64, 213, 218, 220, 329, 340–341, 343, 345–347, 350 in growth media...................5, 8–10, 21, 61, 78, 104, 108, 137–139, 283, 299, 309, 342 Suppressor mutants.......................................................... 4, 6 SYBR Green..................... 153–156, 159–162, 164, 168, 169 Systems (biology,-based approaches)........................ 4, 11, 12 SYTO 42/45............................................ 153–154, 159–162

T TAIR. See The Arabidopsis Information Resource TargetP...................... 248–250, 252, 254, 259–263, 271–272 Tat (cpTat)..........246, 248, 252, 253, 262, 272, 322–323, 332 T-DNA.....................................................12–14, 16–17, 229 TEM. See Transmission electron microscopy Tetrapyrroles, biosynthesis intermediates................. 351–352 The Arabidopsis Information Resource (TAIR)......... 12, 16, 254, 263, 265, 268, 272 Thermolysin...................... 326, 332, 333, 336, 337, 351–365 Thylakoid isolation...................... 218–219, 232, 323–325, 327–330, 335–336, 340–341, 343, 345–347, 349–350 lumen��������������������������������� 244, 246, 248, 250, 252–254, 270, 272, 322 membrane������� 73, 83, 218, 219, 230, 232, 238–239, 244, 246–247, 252–254, 258, 262, 270, 301, 313, 321–337, 340–341, 352 D pH (delta pH, pH gradient)��������������������246, 322, 336 protein transport or insertion..............246–253, 321–337, 339–350 TMHMM. See TransMembrane prediction using Hidden Markov Models TNT. See Transcription and translation TOC/TIC TIC���������������������� 60, 244–246, 307–309, 314, 316–318 Tic proteins�������������������������������245, 308–309, 317–318, 346–349, 352–353, 357 TOC����������������������������� 60, 244–246, 307–309, 313–318 Toc proteins��������������������������� 63, 66, 245, 307–308, 312, 314–317, 352–353, 357, 362 Topology, protein experimental determination������������������������������� 351–366 prediction����������������������������������������������������������� 243–274 Transcript..................................................174, 178, 183–196 Transcriptional regulation.................................136, 171–172, 174, 178 Transcription and translation (TNT)...................... 285, 289, 290, 299–300, 310, 313–314, 319, 342, 343, 354–356, 364

Transcription, chloroplast measurement...................................................... 171–180 run-on assay................................................ 172, 174–179 Transcriptome.................................................................. 172 Transfection������������������������������������������������������62, 65, 66, 75 Transient expression..................................59–70, 73–83, 145 Transit peptide chloroplast transit peptide (cTP)................29, 59, 63, 69, 70, 74, 75, 81–83, 244–245, 248, 252, 282, 307–308, 340, 353 dual targeting peptide (dTP)...................... 245, 249, 252 luminal transit peptide (lTP)...... 246, 248, 250, 252, 262, 272, 321–323 prediction............................................................ 243–274 Translation, chloroplast in vivo labeling.............................211–212, 215, 217–218 measurement...................................................... 209–224 polysome isolation....... 210, 212–213, 216, 218, 220–223 Translocon........................... 60, 244, 246, 307–311, 313–318 Transmembrane domains alpha helices........................................................ 255–256 beta barrels............................................................ 257258 prediction.............................254–258, 266–271, 273–274 TransMembrane prediction using Hidden Markov Models (TMHMM)................................................. 254, 256 Transmission electron microscopy (TEM)............... 113–131 Transplastomic..............................................74, 75, 133–145 Transposon............................................................. 12–13, 16 Trizol������������������������������������������������������������������������������� 192 Trypsin..................................................................... 351–366 Two-dimensional PAGE (2D-PAGE).............228, 230, 232, 236–237

U Ultracentrifugation (ultracentrifuge)................213, 220, 324, 328, 329, 345

W Wax embedding................34–36, 40, 41, 44–47, 50, 52, 54–55 Steedman’s.............................................35, 36, 40, 44, 52 Western blot (or immunoblot)........ 34, 36, 48, 60–62, 67–70, 227–228, 230, 232, 233, 235–236, 239, 315–318, 340, 346, 347 Wheat germ........................60, 285, 289, 299, 314, 325, 332, 336, 342, 343, 349, 354, 356, 364

Y Yellow fluorescent protein (YFP)..................... 74–77, 81, 82 YFP. See Yellow fluorescent protein