Photosynthesis Research Protocols

Methods

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Molecular Biology™

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



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Photosynthesis Research Protocols Second Edition

Edited by

Robert Carpentier Département de Chimie Biologie, Université du Québec à Trois-Rivières, Trois-Rivières, Québec, Canada

Editor Dr. Robert Carpentier Département de Chimie Biologie Université du Québec à Trois-Rivières Trois-Rivières, Québec Canada [email protected]

ISSN 1064-3745 e-ISSN 1940-6029 ISBN 978-1-60761-924-6 e-ISBN 978-1-60761-925-3 DOI 10.1007/978-1-60761-925-3 Springer New York Dordrecht Heidelberg London Library of Congress Control Number: 2010938370 © 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. While the advice and information in this book are believed to be true and accurate at the date of going to press, ­neither the authors nor the editors nor the publisher can accept any legal responsibility for any errors or omissions that may be made. The publisher makes no warranty, express or implied, with respect to the material contained herein. Printed on acid-free paper Humana Press is part of Springer Science+Business Media (www.springer.com)

Preface Photosynthesis is one of the most important biological phenomena on Earth. The capture of sunlight by photosynthetic organisms supplies most of the energy required to develop and sustain life on the planet. Not only is photosynthesis at the heart of plant bioenergetics, it is also fundamental to plant productivity and biomass. Photosynthetic carbon fixation and oxygen evolution directly intervene into many environmental aspects, such as the global atmospheric CO2 level and global climate. Therefore, it is not surprising that a large effort is devoted to photosynthesis research. Photosynthesis, in itself, is of great interest to a multidisciplinary field of research involving agriculture, biochemistry, biotechnology, botany, cell biology, environmental sciences, forestry, plant genetics, plant molecular biology, photobiology, photophysics, photoprotection, plant physiology, plant stress, etc. This book is thus intended as a source of information for scientists working on any of the numerous aspects of photosynthesis. Several biochemical methods of isolation, treatment, and analysis have been developed to fulfil the needs of photosynthesis research. This 2nd edition of Photosynthesis Research Protocols aims at presenting a detailed description of a broad range of general and fundamental methods that are commonly used by plant biochemists, physiologists, and molecular biologists. It retains most of the methods presented in chapters of the first edition, but several new interesting methods are added. This includes coverage of methods related to the most abundant protein on earth, ribulose-1,5-bisphosphate carboxylase/oxygenase. Each technique is described by an expert, and the methods presented can serve as basic protocols for new photosynthesis researchers as well as for experienced scientists seeking to use a new type of preparation or method. The book is especially valuable to the beginner in the field of photosynthesis since each technique is described in simple terms, requiring no previous knowledge of the method. In the Note section of each chapter, appears some further hints and tips which are not provided in regular research papers. I would like to acknowledge and congratulate our series editor, John Walker, for his idea of writing the 1st edition of Photosynthesis Research Protocols. Such a book was badly missing from our shelves. This first version had great success, and we hope this 2nd edition does the same. I also want to thank Johanne Harnois for her great help in preparing the final layout and arrangement of the chapters. Finally, I wish to express my deep gratitude to all the contributors for agreeing to participate. Thanks to their considerable effort, this 2nd edition of Photosynthesis Research Protocols constitutes a major reference book in many laboratories.

Québec, Canada

Robert Carpentier

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Contents Preface . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Contributors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .   1 Isolation of Photosystem II-Enriched Membranes and the Oxygen-Evolving Complex Subunit Proteins from Higher Plants . . . . . . Yasusi Yamamoto, Jing Leng, and Jian-Ren Shen   2 Isolation of Photosystem I Submembrane Fractions . . . . . . . . . . . . . . . . . . . . . . . Johanne Harnois, Najoua Msilini, and Robert Carpentier   3 Isolation of Photosystem II Reaction Center Complexes from Plants . . . . . . . . . . Michael Seibert and Rafael Picorel   4 Methods for the Isolation of Functional Photosystem II Core Particles from the Cyanobacterium Synechocystis sp. PCC 6803 . . . . . . . . . . . . . . . Dmitrii V. Vavilin   5 Purification and Crystallization of Oxygen-Evolving Photosystem II Core Complex from Thermophilic Cyanobacteria . . . . . . . . . . . . Jian-Ren Shen, Keisuke Kawakami, and Hiroyuki Koike   6 Isolation of Cytochrome b6f Complex from Grana and Stroma Membranes from Spinach Chloroplasts . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Elz˙ bieta Romanowska   7 Purification and Crystallization of the Cyanobacterial Cytochrome b6 f Complex . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Danas Baniulis, Huamin Zhang, Taisiya Zakharova, S. Saif Hasan, and William A. Cramer   8 Purification of Plastocyanin and Cytochrome c6 from Plants, Green Algae, and Cyanobacteria . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . José A. Navarro, Manuel Hervás, and Miguel A. De la Rosa   9 Isolation and Identification of Chloroplast Lipids . . . . . . . . . . . . . . . . . . . . . . . . . Norihiro Sato and Mikio Tsuzuki 10 Isolation and Purification of CP43 and CP47 Photosystem II Proximal Antenna Complexes from Plants . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Rafael Picorel, Miguel Alfonso, and Michael Seibert 11 Preparation of Native and Recombinant Light-Harvesting Chlorophyll-a/b Complex . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Wolfgang Rühle and Harald Paulsen 12 Isolation and Characterization of Lamellar Aggregates of LHCII and LHCII-Lipid Macro-assemblies with Light-Inducible Structural Transitions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Ilian Simidjiev, Zsuzsanna Várkonyi, Petar H. Lambrev, and Gyo˝zo˝   Garab 13 Protein Targeting Across and into Chloroplast Membranes . . . . . . . . . . . . . . . . . Shari M. Lo and Steven M. Theg

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14 Proteomic Analysis of Thylakoid Membranes . . . . . . . . . . . . . . . . . . . . . . . . . . . . Venkateswarlu Yadavalli, Sreedhar Nellaepalli, and Rajagopal Subramanyam 15 Thylakoid Phosphoproteins: Identification of Phosphorylation Sites . . . . . . . . . . . Anne Rokka, Eva-Mari Aro, and Alexander V. Vener 16 Direct Detection of Free Radicals and Reactive Oxygen Species in Thylakoids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Éva Hideg, Tamás Kálai, and Kálmán Hideg 17 Assay of Photoinhibition and Heat Inhibition of Photosystem II in Higher Plants . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Nobuyoshi Nijo, Björn Lundin, Miho Yoshioka, Noriko Morita, and Yasusi Yamamoto 18 Photosystem II Reconstitution into Proteoliposomes and Methodologies for Structure–Function Characterization . . . . . . . . . . . . . . . . David Joly, Sridharan Govindachary, and Mário Fragata 19 Physical and Chemical Immobilization Methods of Photosynthetic Materials . . . . Lise Barthelmebs, Robert Carpentier, and Régis Rouillon 20 Identifying Chloroplast Biogenesis and Signalling Mutants in Arabidopsis thaliana . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Verónica Albrecht, Gonzalo M. Estavillo, Abby J. Cuttriss, and Barry J. Pogson 21 Expression of Genes in Cyanobacteria: Adaptation of Endogenous Plasmids as Platforms for High-Level Gene Expression in Synechococcus sp. PCC 7002 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Yu Xu, Richard M. Alvey, Patrick O. Byrne, Joel E. Graham, Gaozhong Shen, and Donald A. Bryant 22 Construction of Gene Interruptions and Gene Deletions in the Cyanobacterium Synechocystis sp. Strain PCC 6803 . . . . . . . . . . . . . . . . . . . Julian J. Eaton-Rye 23 A Simple Method for Chloroplast Transformation in Chlamydomonas reinhardtii . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Vellupillai M. Ramesh, Scott E. Bingham, and Andrew N. Webber 24 Rapid Isolation of Intact Chloroplasts from Spinach Leaves . . . . . . . . . . . . . . . . . David Joly and Robert Carpentier 25 Mechanical Isolation of Bundle Sheath Cell Strands and Thylakoids from Leaves of C4 Grasses . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Elz˙ bieta Romanowska and Eugeniusz Parys 26 Isolation of Ribulose-1,5-Bisphosphate Carboxylase/Oxygenase from Leaves . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . A. Elizabete Carmo-Silva, Csengele Barta, and Michael E. Salvucci 27 Quantifying the Amount and Activity of Rubisco in Leaves . . . . . . . . . . . . . . . . . David S. Kubien, Christopher M. Brown, and Heather J. Kane 28 Purification of Rubisco Activase from Leaves or after Expression in Escherichia coli . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Csengele Barta, A. Elizabete Carmo-Silva, and Michael E. Salvucci

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29 Rubisco Activase Activity Assays . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 375 Csengele Barta, A. Elizabete Carmo-Silva, and Michael E. Salvucci 30 Quantification of Rubisco Activase Content in Leaf Extracts . . . . . . . . . . . . . . . . 383 Wataru Yamori and Susanne von Caemmerer Index . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 393

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Contributors Verónica Albrecht  •  School of Biochemistry and Molecular Biology, The Australian National University, Canberra ACT, Australia Miguel Alfonso  •  Estación Experimental de Aula Dei, Consejo Superior de Investigaciones Científicas, Zaragoza, Spain Richard M. Alvey  •  Department of Biochemistry and Molecular Biology, The Pennsylvania State University, University Park PA, USA Eva-Mari Aro  •  Department of Biology, University of Turku, Turku, Finland Danas Baniulis  •  Department of Biological Sciences, Lilly Hall of Life Sciences, Purdue University, West Lafayette IN, USA Csengele Barta  •  U.S. Department of Agriculture, Arid-Land Agricultural Research Center, Maricopa AZ, USA Lise Barthelmebs  •  Centre de Phytopharmacie, Université de Perpignan, Perpignan, France Scott E. Bingham  •  Department of Plant Biology, Arizona State University, Tempe AZ, USA Christopher M. Brown  •  Department of Biology, University of New Brunswick, Fredericton NB, Canada Donald A. Bryant  •  Department of Biochemistry and Molecular Biology, The Pennsylvania State University, University Park PA, USA Patrick O. Byrne  •  Department of Biochemistry and Molecular Biology, The Pennsylvania State University, University Park PA, USA Susanne von Caemmerer  •  Molecular Plant Physiology Group, Research School of Biology, Australian National University, Canberra ACT, Australia A. Elizabete Carmo-Silva  •  U.S. Department of Agriculture, Arid-Land Agricultural Research Center, Maricopa AZ, USA Robert Carpentier  •  Département de chimie biologie (GRBV), Université du Québec à Trois-Rivières, Trois-Rivières QC, Canada William A. Cramer  •  Department of Biological Sciences, Lilly Hall of Life Sciences, Purdue University, West Lafayette IN, USA Abby J. Cuttriss  •  School of Biochemistry and Molecular Biology, The Australian National University, Canberra ACT, Australia Miguel A. De la Rosa  •  Instituto de Bioquímica Vegetal y Fotosíntesis, Universidad de Sevilla y Consejo Superior de Investigaciones Científícas, Sevilla, Spain Julian J. Eaton-Rye  •  Biochemistry Department, University of Otago, Dunedin, New Zealand Gonzalo M. Estavillo  •  School of Biochemistry and Molecular Biology, The Australian National University, Canberra ACT, Australia

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Mário Fragata  •  Département de chimie biologie (GRBV), Université du Québec à Trois-Rivières, Trois-Rivières QC, Canada ˝zo ˝ Garab  •  Institute of Plant Biology, Biological Research Center, Gyo Hungarian Academy of Sciences, Szeged, Hungary Sridharan Govindachary  •  Département de chimie biologie (GRBV), Université du Québec à Trois-Rivières, Trois-Rivières QC, Canada Joel E. Graham  •  Department of Biochemistry and Molecular Biology, The Pennsylvania State University, University Park PA, USA Johanne Harnois  •  Département de chimie biologie (GRBV), Université du Québec à Trois-Rivières, Trois-Rivières QC, Canada S. Saif Hasan  •  Department of Biological Sciences, Lilly Hall of Life Sciences, Purdue University, West Lafayette IN, USA Manuel Hervás  •  Instituto de Bioquímica Vegetal y Fotosíntesis, Universidad de Sevilla y Consejo Superior de Investigaciones Científícas, Sevilla, Spain Éva Hideg  •  Institute of Plant Biology, Biological Research Center, Hungarian Academy of Sciences, Szeged, Hungary Kálmán Hideg  •  Department of Organic and Medicinal Chemistry, University of Pécs, Pécs, Hungary David Joly  •  Département de chimie biologie (GRBV), Université du Québec à Trois-Rivières, Trois-Rivières QC, Canada Tamás Kálai  •  Department of Organic and Medicinal Chemistry, University of Pécs, Pécs, Hungary Heather J. Kane  •  Department of Biology, University of New Brunswick, Fredericton NB, Canada Keisuke Kawakami  •  Graduate School of Natural Science and Technology, Okayama University, Okayama, Japan Hiroyuki Koike  •  Department of Biosciences, Faculty of Science and Engineering, Chuo University, Tokyo, Japan David S. Kubien  •  Department of Biology, University of New Brunswick, Fredericton NB, Canada Petar H. Lambrev  •  Institute of Plant Biology, Biological Research Center, Hungarian Academy of Sciences, Szeged, Hungary Jing Leng  •  Graduate School of Natural Science and Technology, Okayama University, Okayama, Japan Shari M. Lo  •  Department of Plant Biology, University of California – Davis, Davis CA, USA Björn Lundin  •  Graduate School of Natural Science and Technology, Okayama University, Okayama, Japan Noriko Morita  •  Graduate School of Natural Science and Technology, Okayama University, Okayama, Japan Najoua Msilini  •  Département de chimie biologie (GRBV), Université du Québec à Trois-Rivières, Trois-Rivières QC, Canada José A. Navarro  •  Instituto de Bioquímica Vegetal y Fotosíntesis, Universidad de Sevilla y Consejo Superior de Investigaciones Científícas, Sevilla, Spain

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Sreedhar Nellaepalli  •  Department of Biochemistry, School of Life Sciences, University of Hyderabad, Hyderabad, India Nobuyoshi Nijo  •  Graduate School of Natural Science and Technology, Okayama University, Okayama, Japan Eugeniusz Parys  •  Department of Plant Physiology, Warsaw University, Warszawa, Poland Harald Paulsen  •  Institut für Allgemeine Botanik, Johannes-Gutenberg-Universität, Mainz, Germany Rafael Picorel  •  Estación Experimental de Aula Dei, Consejo Superior de Investigaciones Científicas, Zaragoza, Spain Barry J. Pogson  •  School of Biochemistry and Molecular Biology, The Australian National University, Canberra ACT, Australia Vellupillai M. Ramesh  •  Department of Plant Biology, Arizona State University, Tempe AZ, USA Anne Rokka  •  Department of Biology, University of Turku, Turku, Finland Elz˙ bieta Romanowska  •  Department of Plant Physiology, Warsaw University, Warszawa, Poland Régis Rouillon  •  Centre de Phytopharmacie, Université de Perpignan, Perpignan, France Wolfgang Rühle  •  Institut für Allgemeine Botanik, Johannes-Gutenberg-Universität, Mainz, Germany Michael E. Salvucci  •  U.S. Department of Agriculture, Arid-Land Agricultural Research Center, Maricopa AZ, USA Norihiro Sato  •  School of Life Science, Tokyo University of Pharmacy and Life Sciences, Tokyo, Japan Michael Seibert  •  National Renewable Energy Laboratory, Basic Sciences Center, Golden CO, USA Gaozhong Shen  •  Department of Biochemistry and Molecular Biology, The Pennsylvania State University, University Park PA, USA Jian-Ren Shen  •  Graduate School of Natural Science and Technology, Okayama University, Okayama, Japan Ilian Simidjiev  •  Institute of Plant Biology, Biological Research Center, Hungarian Academy of Sciences, Szeged, Hungary Rajagopal Subramanyam  •  Department of Biochemistry, School of Life Sciences, University of Hyderabad, Hyderabad, India Steven M. Theg  •  Department of Plant Biology, University of California – Davis, Davis CA, USA Mikio Tsuzuki   •  School of Life Science, Tokyo University of Pharmacy and Life Sciences, Tokyo, Japan Zsuzsanna Várkonyi   •  Institute of Plant Biology, Biological Research Center, Hungarian Academy of Sciences, Szeged, Hungary Dmitrii V. Vavilin  •  Genencor, A Danisco Division, Palo Alto CA, USA Alexander V. Vener  •  Division of Cell Biology, Linköping University, Linköping, Sweden Andrew N. Webber  •  Department of Plant Biology, Arizona State University, Tempe AZ, USA

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Yu Xu  •  Department of Biochemistry and Molecular Biology, The Pennsylvania State University, University Park PA, USA Venkateswarlu Yadavalli  •  Department of Biochemistry, School of Life Sciences, University of Hyderabad, Hyderabad, India Yasusi Yamamoto  •  Graduate School of Natural Science and Technology, Okayama University, Okayama, Japan Wataru Yamori  •  Molecular Plant Physiology Group, Research School of Biology, Australian National University, Canberra ACT, Australia Miho Yoshioka  •  Graduate School of Natural Science and Technology, Okayama University, Okayama, Japan Taisiya Zakharova  •  Department of Biological Sciences, Lilly Hall of Life Sciences, Purdue University, West Lafayette IN, USA Huamin Zhang  •  Department of Biological Sciences, Lilly Hall of Life Sciences, Purdue University, West Lafayette IN, USA

Chapter 1 Isolation of Photosystem II-Enriched Membranes and the Oxygen-Evolving Complex Subunit Proteins from Higher Plants Yasusi Yamamoto, Jing Leng, and Jian-Ren Shen Abstract We describe methods to isolate highly active oxygen-evolving photosystem II (PSII) membranes and core complexes from higher plants, and to purify subunits of the oxygen-evolving complex (OEC). The membrane samples used as the material for various in vitro studies of PSII are prepared by solubilizing thylakoid membranes with the nonionic detergent Triton X-100, and the core complexes are prepared by further solubilization of the PSII membranes with n-dodecyl-b-d-maltoside (b-DDM). The OEC subunit proteins are dissociated from the PSII-enriched membranes by alkaline or salt treatment, and are then purified by ion-exchange chromatography using an automated high performance liquid chromatography system. Key words: Photosystem II, Oxygen evolution, Membrane preparation, Oxygen-evolving complex, Protein purification, Ion-exchange chromatography

1. Introduction Highly active oxygen-evolving photosystem II (PSII) membranes are the starting material for isolation of various membrane proteins related to the water-oxidation activity of PSII. Preparation of these membranes was first reported in the early 1980s (1–3), which promoted the structural and functional analyses of PSII. The principle technique used is the solubilization of thylakoids with nonionic detergents to obtain PSII-enriched grana thylakoid membrane particles. In previous studies, the detergents Triton X-100 and digitonin were used to separate the grana-stacked regions from the stroma-exposed regions of the thylakoids.

Robert Carpentier (ed.), Photosynthesis Research Protocols, Methods in Molecular Biology, vol. 684, DOI 10.1007/978-1-60761-925-3_1, © Springer Science+Business Media, LLC 2011

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However, Triton X-100 was shown to be superior in yielding highly active oxygen-evolving PSII particles. The PSII membranes obtained using the methods described above have a large amount of light-harvesting chlorophyll proteins (LHCII) with an antenna size of around 250 chlorophylls per reaction center. For some studies, it is necessary to remove these LHCII proteins and purify the PSII core complexes so that they contain almost no LHCII apo-proteins. The major LHCII subunits can be removed by treating PSII membranes with n-heptyl-b-D-thioglucoside, giving rise to a partially purified PSII complex that retains the minor LHCII subunits and the three oxygen-evolving complex (OEC) subunits (4). PSII core complexes can be further purified by solubilizing the PSII membranes with n-dodecyl-b-d-maltoside (b-DDM), followed by separation with sucrose density gradient centrifugation (5). The PSII complexes thus obtained are essentially devoid of LHCII and contain 35–50 chlorophyll a molecules per reaction center. The presence of OEC subunits in PSII was first reported from PSII-enriched oxygen-evolving samples (1). The OEC is composed of three extrinsic proteins, namely, OEC33, 24, and 18, encoded by their respective nuclear genes psbO, psbP, and psbQ. These proteins have apparent relative molecular masses of 33, 24, and 18 kDa, respectively, when separated by SDS-polyacrylamide gel electrophoresis. Among these proteins, OEC33 stabilizes the catalytic Mn cluster. The exact functions of OEC24 and 18 are not known yet, but it has been suggested that they are involved in regulating the concentrations of Ca2+ and Cl− that control the function of the catalytic Mn cluster. Proteins equivalent to the OEC subunit proteins of higher plants are present in PSII of green algae, while in PSII of cyanobacteria and red algae, the two small subunits are replaced by cyt c550 and a 12 kDa protein (6, 7). The red algal PSII contains an additional 20  kDa extrinsic protein (8). The three OEC subunit proteins of higher plants were first isolated by isoelectric focusing (pIs of OEC33, 24, and 18 are 5.1, 6.5, and 9.2, respectively) (9). However, these proteins are released from PSII-enriched membranes relatively easily by urea, salt, or alkaline treatment, and then can be separated by ion-exchange chromatography. A method to isolate large amounts of OEC33 was also reported (10). The isolated proteins have been used for various biochemical and biophysical assays of the oxygen-evolving system associated with PSII, demonstrating the crucial roles of these proteins in maintaining the activity of the water-splitting reaction. These proteins have also served as model systems to analyze protein transport across the thylakoid membrane during chloroplast biogenesis (11, 12). More recently, cloned OEC subunit proteins overexpressed in Escherichia coli were crystallized, which allowed for structural studies of the proteins by X-ray diffraction analysis (13, 14). In this paper, we

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describe the isolation of PSII membranes and core complexes and purification of the three OEC subunits from higher plants.

2. Materials 2.1. Plant Materials

Spinach and pea leaves are the most widely used materials for the isolation of PSII-enriched membranes from higher plants. Spinach can be purchased in local markets. Peas are usually grown in a plant growth chamber at 20°C.

2.2. Buffers for the Isolation of Active Oxygen-Evolving PSII Membranes (BBY Membranes)

There are several types of PSII-enriched membranes that are prepared from higher plants. The most commonly used are the so-called BBY membranes (2) and the membranes prepared by Kuwabara and Murata (KM membranes) (3). All buffer solutions should be stored at 4°C.

2.2.1. BBY Membranes

1. BBY-1: 20 mM Tricine–NaOH, pH 8.0, 0.4 M NaCl, 0.2% BSA, 2 mM MgCl2. 2. BBY-2: 20 mM Tricine–NaOH, pH 8.0, 0.15 M NaCl, 0.2% BSA, 5 mM MgCl2. 3. BBY-3: 40 mM MES–NaOH, pH 6.5, 0.4 M sucrose, 10 mM NaCl, 5 mM MgCl2. 4. BBY-3T (BBY-3 + Triton X-100): 14% (w/v) Triton X-100, add BBY-3 solution to 20 mL. To dissolve Triton X-100, use a water bath or a hot-plate with a stirrer. Cool and store the solution at 4°C after preparation. 5. BBY-4: 20 mM MES–NaOH, pH 6.0, 0.4 M sucrose, 15 mM NaCl, 5 mM MgCl2, 30% (v/v) ethyleneglycol.

2.2.2. KM Membranes

1. K-1: 50 mM KH2PO4–NaOH, pH 6.9, 0.1 M sucrose, 0.2 M NaCl. 2. K-2: 50 mM KH2PO4–NaOH, pH 6.4, 0.3 M sucrose, 0.1 M NaCl. 3. SMN: 40 mM MES–NaOH, pH 6.50, 0.4 M sucrose, 10 mM NaCl. 4. K-3 (the same as BBY-4): 20  mM MES–NaOH, pH 6.0, 0.4  M sucrose, 15  mM NaCl, 5  mM MgCl2, 30% (v/v) ethyleneglycol. 5. TX-100: 20% (w/v) Triton X-100. Mix the Triton X-100 into solution using a hot water bath or a hot-plate with a stirrer, and add distilled water to 50 mL. Cool and store at 4°C after preparation.

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2.3. Buffers for the Purification of Oxygen-Evolving PSII Core Complexes

1. C-1: 30 mM MES–NaOH, pH 6.0, 10 mM MgCl2, 3 mM CaCl2. 2. C-2: 0.3 M sucrose and 0.03% b-DDM in buffer C-1. 3. C-3: 0.7 M sucrose and 0.03% b-DDM in buffer C-1. 4. C-4: 50% polyethylene glycol (PEG) 2,000. 5. C-5: 25% glycerol in buffer C-1. 6. C-6: 20% b-DDM dissolved in water.

2.4. Buffers for the Isolation of the OEC Subunit Proteins 2.4.1. Buffers for Isolation of OEC24 and 18

OEC24 and 18 are released from PSII membranes by a 1.0 M NaCl-wash, and are then purified by column chromatography. 1. NaCl-1: 20 mM MES–NaOH, pH 6.0, 0.4 M sucrose, 1.0 M NaCl, 5 mM MgCl2. 2. A-1: 30 mM citric acid. Add distilled water to 500 mL, and adjust pH to 4.0 with NaOH. 3. B-1: 30 mM citric acid, 1.0 M NaCl. Add distilled water to 500 mL, and adjust pH to 4.0 with NaOH.

2.4.2. Buffers for the Isolation of OEC33

OEC33 is released from NaCl-washed PSII membranes (depleted of OEC24 and OEC18) by a Tris-wash and is then purified by column chromatography. 1. Tris-washing: 1.0 M Tris–HCl, pH 8.5, 5 mM MgCl2, 0.4 M sucrose. 2. A-2: 30 mM Tris–HCl, pH 9.0. 3. B-2: 30 mM Tris–HCl, pH 9.0, 1.0 M NaCl.

2.5. Other Chemicals and Materials Required for the Preparation of Membrane and Core Samples and for Isolation of OEC Proteins

1. Cheese cloth to filter spinach leaf homogenate.

2.6. Instruments

1. Blender.

2. Acetone (80%) to determine chlorophyll concentration. 3. Black sample tubes. 4. Liquid nitrogen. 5. Dialysis tubing.

2. High-speed refrigerated centrifuge with angled rotors. 3. Ultra-speed centrifuge with swing and angled rotors. 4. UV-visible spectrophotometer. 5. Brushes for resuspending membrane and core precipitates after centrifugation. 6. Stirrer and a hot-plate with a stirrer. 7. Vortex mixer. 8. Cation exchange column Bio-Scale S (BioRad) or similar.

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9. Anion exchange column Bio-Scale Q (BioRad) or similar. 10. Biologic chromatography system (BioRad) or an equivalent FPLC system. 11. Ultrafiltration cell (Millipore) connected to a high pressure nitrogen or helium gas cylinder (for dialysis and concentration of proteins).

3. Methods 3.1. Isolation of Highly Active OxygenEvolving PSII Membranes 3.1.1. BBY Membranes

The following steps are carried out on ice or at 4°C. Light should be avoided as much as possible during the preparation if samples are to be used as materials for photochemical measurements or for photoinhibition studies. Green safe light is used for the preparation in the dark. 1. Rinse spinach leaves with tap water. 2. Cool the leaves on ice or in a refrigerator. 3. Homogenize 100  g spinach leaves with 300  mL of BBY-1 solution in a precooled blender. 4. Filter the homogenate through cheese cloth. If foaming occurs, store the homogenate in a large container at 4°C until the foam disappears. 5. Centrifuge the filtrate at 6,000 × g for 10 min. 6. Suspend the precipitate in 80 mL BBY-1 solution and centrifuge at 400 × g for 30  s. Resuspend the precipitate gently using a paintbrush (see Note 1). 7. Centrifuge the supernatant at 35,000 × g for 7 min. 8. Suspend the precipitate in 40 mL BBY-2 solution and centrifuge at 35,000 × g for 7 min. 9. Suspend the precipitate in 7 mL BBY-3 solution. 10. Measure the volume of the suspension with a precooled graduated cylinder (A mL). 11. Determine the chlorophyll concentration in the suspension (B mg chlorophyll/mL) (see Subheading 3.1.3). 12. Adjust the chlorophyll concentration to 2.5 mg chlorophyll/ mL by adding {(A × B/2.5) – A} mL BBY-3 solution. 13. Add a ¼ volume of BBY-3T solution to the suspension. Add slowly, stirring with a pipette and avoiding foaming as much as possible. The final chlorophyll concentration is 2  mg chlorophyll/mL, and the ratio of TX-100 to chlorophyll (w/w) is 14:1. To change the ratio of TX-100 and chlorophyll, for example, to 20:1, adjust the concentration of TX-100 in the BBY-3T solution to 20%.

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14. Stir slowly for 10 min. 15. Centrifuge the suspension at 35,000 × g for 15 min. 16. Suspend the precipitate in 40 mL BBY-3 solution and centrifuge at 3,000 × g for 3 min. 17. Centrifuge the supernatant at 35,000 ×g for 20  min. If the resulting supernatant is green, resuspend the precipitate from step 15 in BBY-3 solution, and centrifuge the suspension at 35,000 × g for 20 min. 18. Suspend the precipitate in a small volume of BBY-4 solution. 19. Determine the chlorophyll concentration. 20. Store the sample in small plastic sample tubes (see Note 2). 21. Freeze the samples in liquid nitrogen, and store at −80°C. 3.1.2. KM Membranes

1. Rinse spinach leaves with tap water. 2. Cool the leaves on ice. 3. Homogenize leaves in a blender with 250 mL K-1 solution per 100 g leaves. 4. Filter the homogenate through four layers of cheese cloth. 5. Centrifuge the filtrate at 4,000 × g for 5 min. 6. Suspend the precipitate in K-1 solution, and centrifuge the suspension at 500 × g for 30 s (see Note 1). 7. Centrifuge the supernatant at 4,000 × g for 10 min. 8. Suspend the precipitate in K-2 solution. 9. Measure the volume of the suspension with a precooled graduated cylinder (A mL). 10. Determine the chlorophyll concentration (B mg chlorophyll/ mL) (see Subheading 3.1.3). 11. Adjust the chlorophyll concentration to 2.5 mg chlorophyll/ mL by adding {(A × B/2.5) – A} mL K-2 solution. 12. Add a 1/4 volume of 20% TX-100 solution to the suspension with stirring. 13. Centrifuge at 35,000 × g for 15 min. 14. Suspend the precipitate in SMN, and centrifuge at 1,500 × g for 2 min. This step may be omitted. 15. If step 14 is conducted, centrifuge the suspension again at 35,000 × g for 15 min. 16. Suspend the precipitate in a small volume of K-3 solution. 17. Determine the chlorophyll concentration. 18. Transfer the samples to sample tubes (see Note 2). 19. Freeze the samples quickly in liquid nitrogen, and store at −80°C.

Isolation of Photosystem II-Enriched Membranes 3.1.3. Determination of Chlorophyll Concentration

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1. Transfer 5 or 10  mL samples to test tubes with a glass microsyringe. 2. Add 2.5 mL 80% acetone and mix with a vortex mixer. 3. Centrifuge at 1,300 × g for 3–5 min. 4. Measure absorption at 645, 663, and 720  nm. Calculate chlorophyll concentration using the following formulas and the reported chlorophyll absorption coefficients (15): Total chlorophyll (mg/mL) = (8.05 × A663 + 20.29 × A645)/ (X dilution) Chlorophyll a (mg/mL) = (12.7 × A663 – 2.59 × A645)/(X dilution) Chlorophyll b (mg/mL) = (22.9 × A645 – 4.67 × A663)/(X dilution) Subtract A720 from A663 and A645 to correct for nonspecific absorbance. The X dilution value, for example, is 1/500 when 5  mL sample is used.

3.2. Purification of Oxygen-Evolving PSII Complexes

Partially purified PSII complexes can be obtained by solubilizing PSII  membranes with n-heptyl-b-D-thioglucoside, followed by ­differential centrifugation (4). Here, we describe the isolation of highly purified PSII core complexes from spinach BBY membranes. 1. Centrifuge BBY membranes at 35,000 × g for 15  min, then suspend the membranes in buffer C-1 to a chlorophyll concentration of 1 mg/mL. 2. Add b-DDM stock solution to a final concentration of 0.8%, and gently stir the solution for 30 min in the dark at 4°C. 3. Load the b-DDM-treated membranes onto a linear sucrose density gradient (0.3–0.7 M sucrose in C-1 solution containing 0.03% b-DDM). Centrifuge at 160,000 × g for 16 h at 4°C. 4. The result of sucrose density gradient centrifugation is shown in Fig. 1. From the top to the bottom of the tube, the four clear bands correspond to LHCII monomer, LHCII trimer, PSII monomer, and PSII dimer. 5. Collect the PSII monomer and dimer fractions, and dilute each fraction twofold with buffer C-1. 6. Add PEG 2,000 to a final concentration of 12.5%, and centrifuge at 100,000 × g for 20 min. 7. Resuspend the precipitated PSII monomers and dimers in buffer C-5, and store the purified PSII complexes in liquid nitrogen until use. The PSII complexes purified by this method are free from LHCII apo-proteins. The OEC33 subunit is retained in the complexes, but OEC24 and OEC18 have been removed. The complexes have an oxygen-evolving activity of 700–1,200 mmoles O2/mg chl/h.

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Fig. 1. Sucrose density gradient centrifugation pattern of spinach BBY membranes solubilized by 0.8% b-DDM. Sample was centrifuged at 160,000 × g for 16 h on a 0.3–0.7 M sucrose density gradient. For other details, see text.

3.3. Isolation of the OEC Subunit Proteins

OEC24 and OEC18 are released from PSII membranes by 1.0  M NaCl-washing and are further purified by column chromatography.

3.3.1. Isolation of OEC24 and OEC18

1. Thaw PSII membranes, and then suspend in NaCl-1 buffer to a chlorophyll concentration of 1.0 mg/mL. 2. Incubate for 30 min on ice in the dark. 3. Centrifuge the PSII membranes at 35,000 × g for 15 min. 4. Collect the supernatant containing OEC24 and OEC18 proteins. The supernatant should be colorless; if not, centrifuge again at 200,000 × g for 60 min, and then collect the supernatant. If it is not to be used immediately, the supernatant should be stored at −80°C. 5. Immediately before loading onto the column, dilute the NaCl-washed supernatant sixfold with A-1 buffer to ensure that the NaCl concentration in the supernatant is low enough for the OEC24 and OEC18 proteins to bind to the column (see Note 3). 6. Load the NaCl-washed supernatant onto a Bio-Scale S cation-exchange column (BioRad) that has been equilibrated with A-1 buffer.

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7. Elute OEC24 and OEC18 with a linear 350–650  mM NaCl gradient using buffers A-1 and B-1. Under these elution conditions, OEC24 elutes first, followed by OEC18 (see Note 4). 8. If necessary, eluted OEC24 and OEC18 are concentrated by ultrafiltration. The proteins are dialyzed against an appropriate buffer. Store the purified OEC24 and OEC18 at −80°C. It is important that the concentration of NaCl is 1.0  M in the  supernatant prior to column chromatography, since a high NaCl concentration protects OEC24 and OEC18 from digestion by proteases that are released from PSII membranes by the NaCl-wash. 3.3.2. Isolation of OEC33

OEC33 is released by Tris-washing of PSII membranes that are depleted of OEC24 and OEC18 by NaCl-washing and is then purified by column chromatography. 1. Thaw PSII membranes, and suspend in NaCl-1 buffer to a chlorophyll concentration of 1.0 mg/mL to release OEC24 and OEC18. 2. Incubate for 30 min on ice in the dark. 3. Centrifuge the PSII membranes at 35,000 × g for 15 min. 4. Suspend PSII membrane pellet in Tris-washing buffer to a chlorophyll concentration of 1.0 mg/mL; incubate for 30 min on ice in the dark. 5. Centrifuge the mixture and collect the supernatant containing OEC33. The supernatant should be colorless; if not, centrifuge at 200,000 × g for 60 min, and collect the supernatant. Store at −80°C if not used immediately for purification by column chromatography. 6. Dialyze the Tris-washing supernatant against a large volume of A-2 buffer for 3 h. Longer dialysis should be avoided to prevent proteolysis of OEC33. 7. Load the Tris-washed supernatant onto a Bio-Scale Q anionexchange column (BioRad) that has been equilibrated with A-2 buffer. 8. Elute OEC33 with a linear 100–600  mM NaCl gradient using buffers A-2 and B-2. Under these elution conditions, OEC33 may appear in two major fractions; collect both fractions for future use. 9. If necessary, concentrate eluted OEC33 by ultrafiltration. Dialyze the protein against an appropriate buffer. Store the purified OEC33 at −80°C until use.

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4. Notes 1. Watercolor paintbrushes are useful for suspending precipitates. 2. If necessary, black sample tubes should be used to block light. 3. It is important to keep the NaCl concentration in the supernatant to 1.0  M prior to column chromatography, since a high NaCl concentration protects OEC24 and OEC18 from digestion by proteases that are released from PSII membranes by the NaCl-wash. 4. A small amount of OEC24 may be found in the OEC18 fraction; if this is the case, rerun the OEC18 fraction on the column to separate the residual OEC24 from OEC18. References 1. Yamamoto, Y., Doi, M., Tamura, N., and Nishimura, M. (1981) Release of polypeptides from highly active O2-evolving photosystem-2 preparation by Tris treatment. FEBS Lett 133, 265–268. 2. Berthold, D. A., Babcock, G. T., and Yocum, C. F. (1981) A highly resolved, oxygen-evolving photosystem II preparation from spinach thylakoid membranes. FEBS Lett 134, 231–234. 3. Kuwabara, T., and Murata, N. (1982) Inactivation of photosynthetic oxygen evolution and concomitant release of three polypeptides in the photosystem II particles of spinach chloroplasts. Plant Cell Physiol 23, 533–539. 4. Enami, I., Kamino, K., Shen, J.-R., Satoh, K., and Katoh, S. (1989) Isolation and characterization of Photosystem II complexes which lack light-harvesting chlorophyll a/b proteins but retain three extrinsic proteins related to oxygen evolution from spinach. Biochim Biophys Acta 977, 33–39. 5. Smith, P.J., Peterson, S., Masters, V. M., Wydrzynski, T., Styring, S., Krausz, E., and Pace, R. J. (2002) Magneto-optical measurements of the pigments in fully active photosystem II core complexes from plants. Biochemistry 41, 1981–1989. 6. Shen, J.-R., Ikeuchi, M., and Inoue, Y. (1992) Stoichiometric association of extrinsic cytochrome c 550 and 12 kDa protein with a highly purified oxygen-evolving photosystem II core complex from Synechococcus vulcanus. FEBS Lett 301, 145–149. 7. Enami, I., Murayama, H., Ohta, H., Kamo, M., Nakazato, K., and Shen, J.-R. (1995) Isola­ tion and characterization of a Photosystem II complex from the red alga Cyanidium caldarium: Association oif cytochrome c-550 and a

8.

9.

10.

11.

12.

13.

14.

15.

12 kDa protein with the complex. Biochim Biophys Acta 1232, 208–216. Enami, I., Kikuchi, S., Fukuda, T., Ohta, H., and Shen, J.-R. (1998) Binding and functional properties of four extrinsic proteins of photosystem II from a red alga, Cyanidium caldarium, as studied by release-reconstitution experiments. Biochemistry 9, 2787–2793. Yamamoto, Y., Shimada, S., and Nishimura, M. (1983) Purification and molecular properties of 3 polypeptides released from a highly active O2-evolving photosystem-II preparation by Tris-treatment. FEBS Lett 151, 49–53. Yamamoto, Y., Hermodson, M.A., and Krogmann, D.W. (1986) Improved purification and N-terminal sequence of the 33-kDa protein in spinach PS II. FEBS Lett 195, 155–158. Cline, K., Henry, R., Li, C., and Yuan J. (1993) Multiple pathways for protein transport into or across the thylakoid membrane. EMBO J 12, 4105–4114. Hashimoto, A., Ettinger, W., Yamamoto, Y., and Theg, S. M. (1997) Assembly of newly imported oxygen-evolving complex subunits in isolated chloroplasts: sites of assembly and mechanism of binding. Plant Cell 9, 441–452. Ifuku, K., Nakatsu, T., Kato, H., and Sato, F. (2004) Crystal structure of the PsbP protein of photosystem II from Nicotiana tabacum. EMBO Rep 5, 362–367. Calderone, V., Trabucco, M., Vujicic´, A., Battistutta, R., Giacometti, G. M., Andreucci, F., Barbato, R., and Zanotti, G. (2003) Crystal structure of the PsbQ protein of photosystem II from higher plants. EMBO Rep 4, 900–905. Mackinney, G. (1941) Absorption of light by  chlorophyll solutions. J Biol Chem 140, 315–322.

Chapter 2 Isolation of Photosystem I Submembrane Fractions Johanne Harnois, Najoua Msilini, and Robert Carpentier Abstract In this chapter, we describe a method to prepare photosystem I (PSI) submembrane fractions derived from the chloroplast stroma lamellae of spinach chloroplasts. These preparations retain the cytochrome b6 /f complex and a pool of about 11 plastoquinones per P700. The PSI submembrane fractions are thus able to perform both cyclic and linear electron transport reactions from various artificial electron donors to oxygen or methylviologen. They are useful to study both PSI and cytochrome b6 /f complex activities in a nearly native form without interference from photosystem II. Key words: Photosystem I, Cytochrome b6 /f complex, Digitonin, Stroma lamellae

1. Introduction In this chapter, a simple method is described to prepare photosystem I (PSI) submembrane fractions from spinach leaves. Stable and well coupled PSI-enriched vesicles, mainly derived from the chloroplast stroma lamellae, are obtained by mild digitonin treatment of spinach chloroplasts using the method first described by Peters et al. (1) with some modifications (2, 3). Optimal conditions for chloroplast solubilization were established at a digitonin/chlorophyll ratio of 1 (w/w) and a chlorophyll concentration of 0.2 mM, resulting in a little loss of native components (1). In particular, plastocyanin is easily released at a higher digitonin/ chlorophyll ratio. The PSI submembrane fractions retain plastocyanin, the cytochrome b6 /f complex and a pool of about 11 plastoquinones per P700 (1). However, these membranes are devoid of PSII activity and proteins (3). They are able to sustain strong electron transport reactions from various artificial electron donors to oxygen or methylviologen (2, 4, 5) and to maintain cyclic electron transport with the photoinduced reduction and oxidation Robert Carpentier (ed.), Photosynthesis Research Protocols, Methods in Molecular Biology, vol. 684, DOI 10.1007/978-1-60761-925-3_2, © Springer Science+Business Media, LLC 2011

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of the cyt b6 /f complex (1, 5). The endogenous pool of plastoquinones that is retained also takes part in PSI photochemistry (6, 7). Hence, these preparations are useful not only to study PSI reactions in the absence of PSII, but should also be appropriate to study the function of the cyt b6 /f complex in a nearly native form.

2. Materials 1. Fresh spinach leaves from the local market. 2. Homogenization buffer A: 50  mM Tricine–KOH, pH 7.8, 10 mM KCl, 10 mM NaCl, 5 mM MgCl2, 400 mM sorbitol, and 1mM phenylmethyl sulfonyl fluoride (PMSF) (see Notes 1–5). 3. Hypotonic buffer B: 20 mM Tricine–KOH, pH 7.8, and 10 mM MgCl2. 4. Isotonic buffer C: 20  mM Tricine–KOH, pH 7.8, 20  mM KCl, 20 mM NaCl, and 500 mM sorbitol (see Notes 1–3). 5. Incubation buffer D: 20 mM Tricine–KOH, pH 7.8, 10 mM KCl, 10 mM NaCl, 5 mM MgCl2, and 250 mM sorbitol (see Notes 1–3). 6. Resuspension buffer E: 20 mM Tricine-KOH, pH 7.8, 10 mM KCL, 10 mM NaCl, and 5 mM MgCl2. 7. Digitonin. 8. 80% (v/v) acetone in distilled water. 9. Miracloth (Calbiochem). 10. Wheaton tissue grinder (55 mL). 11. Waring blender with sharp blades. 12. Vortex mixer. 13. Bench-top centrifuge.

3. Methods 3.1. Photosystem I Submembrane Fractions Preparation

1. Weigh 200 g of deveined spinach leaves. 2. Clean the spinach leaves in cold distilled water and dry on absorbent paper. 3. Cut the leaves in small pieces and place them with 300 mL of homogenization buffer A in the Waring blender (see Note 6). 4. Add 1 mM PMSF. 5. Homogenize for about 1 min using the pulse mode. 6. Filter the slurry through two layers of Miracloth. 7. Centrifuge the filtrate 5 min at 3,500×g at 4°C.

Isolation of Photosystem I Submembrane Fractions

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8. Resuspend the pellets in 50 mL of hypotonic buffer B using a paintbrush and homogenized with a 55 mL Wheaton tissue grinder. 9. Icubate the solution for 2–3 min on ice in the dark. 10. Add 50 mL of isotonic buffer C (see Note 7). 11. Centrifuge for 5 min at 3,500×g at 4°C. 12. Resuspend the new pellets in the incubation buffer D to obtain a chlorophyll concentration of 2  mg/mL after the further addition of 0.2% digitonin (see Note 8) and Subheading 3.2. 13. Incubate the solution for 30  min in the dark at 4°C with gentle stirring. 14. Add more incubation buffer D to obtain a threefold dilution. 15. Centrifuge for 30 min at 42,000×g at 4°C. 16. Pool the supernatants and centrifuge for 1 h at 150,000×g at 4°C. 17. Resuspend the final pellets (PSI preparation) in a small volume of the resuspension buffer E, and then dilute to a chlorophyll concentration of 2–3 mg/mL. 18. Chlorophyll is determined in 80% acetone as described in Subheading 3.2. 3.2. Determination of Chlorophyll Concentration

1. Add 10  mL of photosynthetic membrane to 5  mL acetone 80% in a conical tube and mix carefully using a vortex mixer (see Note 9) 2. Centrifuge in the bench-top centrifuge for a few minutes to remove precipitated proteins. 3. Verify the exact volume (5  mL) and adjust if necessary to compensate for evaporation. 4. Measure the absorbance at 647 and 664 nm. 5. Taking the dilution of the membranes in the acetone solution into account, the chlorophyll concentration (mg/mL) in the membrane preparation is calculated from the following equation: 0.5 [17.76 (A 647) + 7.38 (A664)], in which A 647 and A664 represent the absorbancies at the respective wavelengths (see Note 10).

4. Notes 1. This buffer should be prepared just before use. 2. Sorbitol is used to keep the medium isotonic; sugars are also known to help in the protection of biological membranes against denaturation.

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3. Sorbitol can be prepared in advance as a concentrated solution (2 M) and kept at −20°C. It is diluted to the required concentration during the preparation of the buffer. 4. PMSF is used as an inhibitor of proteases. It should be prepared as a concentrated solution (1 M) in dioxane and diluted to the proper concentration. 5. The PMSF should be added directly in the blender just before grinding the leaves. 6. The chamber of the blender and all the solutions used must be ice cold when used. During the preparation, care must be applied to always keep the material in a cold (near 4°C) environment. 7. Add isotonic buffer at the same of the hypotonic buffer. 8. The membranes should be first resuspended in a small volume for chlorophyll determination. Then, the final volume required to obtain 2 mg chlorophyll/mL is calculated. The volume of incubation buffer D to be added to adjust the final concentration is used to prepare a digitonin solution containing the amount of detergent required to obtain 0.2% in the final volume. It may be necessary to heat this solution for about 5  min to improve the solubility of digitonin. Cool down the solution, and add it progressively to the membrane suspension with gentle stirring on ice. 9. The method of Arnon (8) was used for many years and is now used with the corrections of Porra et al. (9). The sample of photosynthetic membranes should be added to the acetone while mixing. This is necessary to minimize the amount of chlorophyll that remains bound to the precipitated proteins. 10. Several replicates should be done to obtain a better estimation of the chlorophyll concentration. If the concentration is above 6 mg/mL, it is better to prepare a first dilution just above the required final concentration and to determine the chlorophyll concentration in this predilution before final dilution. References 1. Peters, F.A.L.J., Van Wielink, J.E., Wong Fong Sang, H.W., De Vries, S., and Kraayenhof, R. (1983) Studies on well coupled photosystem I-enriched subchloroplast vesicles: content and redox properties of electron-transfer components. Biochim Biophys Acta 724, 159–165. 2. Boucher, N., Harnois, J., and Carpentier, R. (1989) Heat-stress stimulation of electron flow in a photosystem I submembrane fraction. Biochem Cell Biol 68, 999–1004.

3. Rajagopal, S., Bukhov, N.G., and Carpentier, R. (2003) Control of energy dissipation and photochemical activity in photosystem I by NADP-dependent reversible conformational changes. Biochemistry 42, 11839–11845. 4. Boucher, N, and Carpentier, R. (1993) Heatstress stimulation of oxygen uptake by Photosystem I involves the reduction of superoxide radicals by specific electron donors. Photosynth Res 35, 213–218.

Isolation of Photosystem I Submembrane Fractions 5. Velitchkova, M.Y., and Carpentier, R. (1994) Variable thermal dissipation in a photosystem I submembrane fraction. Photosynth Res 40, 263–268. 6. Rajagopal, S., Egorova, E, Bukhov, N.G., and Carpentier, R. (2003) Quenching of excited states of chlorophyll molecules in submembrane fractions of Photosystem I by exogenous quinones. Biochim Biophys Acta 1606, 147–152. 7. Joly, D. and Carpentier, R. (2007) Regulation of energy dissipation in Photosystem I by the redox

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state of the plastoquinone pool. Biochemistry 46, 5534–5541. 8. Arnon, D.I. (1949) Copper enzymes in isolated chloroplasts. Polyphenoloxydase in B. vulgaris. Plant Physiol 42, 1–15. 9. Porra, R.J., Thompson, W.A., and Kriedemann, P.E. (1989) Determination of accurate extinction coefficients and simultaneous equations for essaying chlorophylls a and b extracted with four different solvents: verification of the concentration of chlorophyll standards by atomic spectroscopy. Biochim Biophys Acta 975, 384–394.

Chapter 3 Isolation of Photosystem II Reaction Center Complexes from Plants Michael Seibert and Rafael Picorel Abstract Methods to isolate and purify 6- and 5-Chl D1/D2/Cyt b559 photosystem II (PSII) reaction center (RC) complexes from plants are presented, and the advantages and disadvantages of each procedure are discussed. One of the simpler 6-Chl procedures and a procedure for isolating 5-Chl complexes are described in detail. Furthermore, a rapid procedure that produces relatively large amounts of less pure 6-Chl material (i.e., more nonpigmented protein) is also described. Criteria to assess the purity of PSII RC preparations are presented, and problems associated with each of the isolation procedures are discussed. Key words: Intrinsic protein, Isolation, Pigment-protein complex, Photosystem II, Reaction center, Purification, Chromatography

1. Introduction The reaction center (RC) complex of photosystem II (PSII) was first isolated by Nanba and Satoh (1), 19 years after the bacterial reaction center was purified. Both are integral membrane protein complexes, and the latter was the first such complex for which an X-ray crystal structure was obtained (2). The isolation of the PSII RC, containing the D1 and D2 polypeptides and Cyt b559, started a long series of events that have led to many advances in understanding functional aspects of PSII and its structure at the molecular level (3, 4). Although the isolated PSII RC itself has not been crystallized well enough that a detailed crystal structure could be obtained, X-ray crystal structures of PSII core complexes, which contain the RC, have been obtained for cyanobacteria, though not plants (5, 6). Nevertheless, this chapter illustrates some of the

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more common procedures used to isolate the PSII RC from plants and the pitfalls that researchers might face in doing so. The procedures described have been optimized for spinach, pea, and sugar beet.

2. Materials 1. Plant leaves, such as spinach (see Note 1), pea, and sugar beet, but modifications of the procedures have worked with wheat and Spirodella. 2. Warring blender and cheesecloth. 3. Cold room or cold box. 4. Preparative centrifuge, centrifuge rotors capable of generating forces of up to 33,000 g, and centrifuge tubes (see Note 2). 5. Spectrophotometer for determining chlorophyll concentrations and absorption spectra. 6. Detergent (Triton X-100 and n-dodecyl-b-D-maltoside (b-DM)), buffers, sucrose, and common salts. 7. Anion-exchange resin, TSK-GEL Toyopearl DEAE 650(S) (see Note 3). 8. Chromatographic columns (2.5 × 9  cm, 1.6 × 10  cm, and 1 × 10 cm). 9. Fast-Flow Chelating Sepharose (Cat. No. 17-0575-01); (Amersham [Pharmacia], Uppsala, Sweden) 10. Pumps and fraction collector or an FPLC. 11. Linear gradient maker. 12. Dialysis tubing. 13. Centrifugal filter devices concentrators: Centriprep YM-50 50,000 MW cut-off (Amicon Bioseparations, Bedford, MA) or Centricon-50 50,000 MW cut-off (Amicon, Beverely, MA). 14. Buffer K-1: 50 mM Na/K phosphate, pH 7.4, 100 mM sucrose, and 200 mM NaCl. 15. Buffer K-2: 50 mM Na/K phosphate, pH 6.9, 300 mM sucrose, and 50 mM NaCl. 16. Buffer K-3: 40 mM Na/K phosphate and pH 6.9. 17. Buffer K-4: 20 mM Mes-NaOH, pH 6.5, 400 mM sucrose, 15 mM NaCl, and 5 mM MgCl2. 18. Buffer A: 50 mM Tris–HCl, pH 7.2, and 30 mM NaCl. 19. Buffer B: 50 mM Tris–HCl, pH 7.2, 30 mM NaCl, and 0.05% (w/v) Triton X-100.

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20. Buffer C: 50 mM Tris–HCl, pH 7.2, and 0.05% (w/v) b-DM. 21. Buffer D: 50 mM MES, pH 6.5, and 30 mM NaCl. 22. Buffer E: 50 mM MES, pH 6.5, 30 mM NaCl, and 1% (w/v) Triton X-100. 23. Buffer F: 50  mM MES, pH 6.5, and 0.05% (w/v) Triton X-100. 24. Buffer G: 50 mM MES, pH 6.5, and 0.05% (w/v) b-DM. 25. Buffer H: 50  mM Na2HPO4, pH 6.5, 50  mM NaCl, 0.2% (w/v) Triton X-100, and 1.2 mM b-DM. 26. Buffer I: 50 mM MES, pH 6.5, 5 mM imidazol, and 2 mM b-DM.

3. Methods The methods outlined below describe how to isolate PSII membrane fragments which are required for isolating and purifying PSII RC material and how to isolate standard 6-chlorophyll (Chl) per center (2 pheophytins) PSII RC preparations using anionicexchange chromatography. Also, how to isolate 5-Chl per RC preparations using immobilized metal affinity chromatography (IMAC) and determine the purity of the preparations. In addition to the second step above, a procedure is also described to rapidly isolate large amounts of 6-Chl-type preparations when the presence of nonpigmented protein contaminants in the RCs is not a major issue. 3.1. Isolation of PSII Reaction Center Complexes

Isolated PSII membrane fragments are the starting material for the further isolation and purification of PSII RC material. In 1981 and 1982, three groups reported the use of different detergentsolubilization procedures to isolate active PSII membranes (i.e., membranes that still exhibited water-splitting function) from broken thylakoid preparations (7–9). Dunahay et al. (10) reported a detailed comparison of the purity of the different types of PSII membrane fragments. It is important for the purpose of the RC-isolation protocols that the PSII membranes are not contaminated with photosystem I (PSI) because PSI complexes if present can under some conditions elute with the PSII RC complex. See the Chapter by Yasusi Yamamoto for more information about the preparation of PSII membrane fragments. The following protocol is used in our laboratories, and it is a slight modification of the so-called KM procedure (8, 10). However, the BBY procedure (7, 10) can also be used.

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3.1.1. Procedure for Isolating PSII Membrane Fragments

1. The following steps should be carried out at 4°C in dim room light (steps 2 and 3) and in the dark (steps 4–11). The buffers referred to in this section, and those below, are described in Subheading 2. 2. In a Warring blender, grind 1 Kg of deribbed spinach in icecold K-1 buffer (1:2 w/v). Repeat 4–5 times for 10 s each time with about 30 s in between each grind until the green suspension does not contain leaf debris larger than a few millimeters on a side. Filter the suspension through 16 layers of cheesecloth. 3. Centrifuge the filtrate at 500 g for 1 min to remove any cell debris that passes through the cheesecloth. 4. Pour the supernatant into appropriate sized centrifuge tubes and pellet at 12,300 g for 25 min. 5. Discard the supernatant (it contains free Chl, cell cytoplasmic material, proteases, etc.), resuspend the pellet in a smaller volume fresh K-1 buffer with a paint brush, pour into centrifuge tubes, and pellet at 12,300 g for 25 min. 6. Resuspend the pellet in K-2 buffer first with a paint brush and then with a homogenizer and run a Chl analysis. Adjust the suspension to around 3.5 mg Chl/ml. The Chl a/b ratio should be around 2.8–3.0, if the procedure is successful at this point. 7. Add 20% (w/v) Triton X-100 to the suspension with gentle stirring to a final ratio of 25:1 (w/w) Triton/Chl (see Note 1). Incubate with gentle stirring for 7  min in the cold. The suspension should turn a deeper shade of green. 8. Pour the suspension into clean centrifuge tubes and pellet at 32,900 g for 15 min. 9. Discard the supernatant, resuspend the pellet in K-3 buffer first with a paint brush and then with a homogenizer, and pellet at 737× g for 1.5 min to remove starch. 10. Pour the supernatant into clean centrifuge tube and pellet at 32,900 g for 15 min. 11. Resuspend the pellet in K-4 first with a paint brush and then with a homogenizer and run a Chl analysis. A Chl a/b ratio of 1.8–2.2 indicates a successful preparation at this point. The membranes can be stored at −80°C in the freezer or under liquid N2 for up to 1 year until use.

3.1.2. Procedures for Isolating and Purifying 6-Chl PSII Reaction Center Complex

This procedure is based on the original by Nanba and Satoh (1), which produced highly labile material (11, 12). Stabilization of the material was accomplished by exchanging the nonionic detergent b-DM (or lauryl maltoside) for the nonionic detergent Triton X-100 (11, 13). However, even after purification and

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stabilization, if the material is to be left unfrozen in room light for more than a few minutes, the researcher should work at 4°C and an oxygen-scrubbing system added to the material (13). 3.1.2.1. Standard Procedure for Preparing Highly Purified RCs (0.05% Triton X-100)

This protocol produces a highly purified PSII RC preparation, but it can take up to several days to complete (around 72 h with the starting material suggested below, depending on how much material is to be isolated (see Note 4). The following steps should be carried out at 4°C in the dark: 1. Unfreeze 62 mg Chl of PSII-enriched membrane fragments and pellet at 32,900× g for 15 min. 2. Resuspend the pellet in 48 ml (final volume) of buffer A. 3. Add 12 ml 20% (w/v) Triton X-100 (60 ml total final volume at 1 mg Chl/ml and 4% w/v Triton) slowly with gentle stirring and incubate for 1 h also with gentle stirring. 4. Centrifuge at 32,900 g for 1 h and keep the supernatant. 5. Load the supernatant onto a prepacked (4°C) TSK-GEL Toyopearl DEAE 650(S) anion exchange column (1.6 × 10 cm) that had been pre-equilibrated with buffer B. 6. Wash the column overnight with the same buffer at a flow rate of 2.5  ml/min until the eluate exhibits a 417:435  nm peak ratio of about 1.16 (this ratio is approximately the ratio of the final RCs as purified). This ratio is a better measure of final purity than the absorbance of the eluate at 670  nm, which depends on the column flow rate. The wash step takes about 30  h. Higher amounts of starting material will take longer periods of time to wash. 7. Exchange b-DM for Triton-X100 by washing the column with the same buffer except with 0.1% (w/v) b-DM substituted for the Triton. Continue to wash until the absorption band at 280  nm (residual Triton) has an absorbance lower that 0.1 (1-cm path). 8. Apply 140 ml of a 30–200 mM NaCl linear gradient (in the same buffer and detergent) at a flow rate of 1 ml/min. Collect 3 ml fractions of the eluate and save. The green band containing the RC complex elutes at 90–120 mM salt. Note that two peaks may come off the column, but the second is much smaller than the first. If so, use the peak that comes off first since it is purer (see Note 5). 9. Take the absorption spectrum of each fraction, combine those with a red-peak maxima at 675 nm or higher, and desalt by dialyzing against buffer C for 1 h with one buffer exchange. Desalting is required if the samples are to be used for low temperature spectroscopy since high salt will interfere with the formation of a clear glass.

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10. If desired, the material can be concentrated using centrifugal filter devices concentrators, if small volumes are to be concentrated. When the desired concentration is achieved, freeze the sample in liquid N2 and store under liquid N2. The samples can also be stored at −80°C in a freezer if they are to be used within a short period of time or on dry ice if they are to be shipped. 3.1.2.2. Modified Procedure for Faster Preparation of RCs with High Yield but with Some Nonpigmented Protein Contaminants (1% Triton X-100)

This procedure can be used if large amounts of material are needed, and contamination from nonpigmented proteins is not a significant issue. The advantage of this protocol over the one described above is that the isolation procedure takes much less time (see Subheading “Modified Procedure for Faster Preparation of RCs with High Yield but with Some Non-pigmented Protein Contaminants (1% Triton X-100),” steps 2–12, take around 20 h with the suggested starting material) (see Note 6). The following steps should be carried out at 4°C in the dark: 1. Unfreeze 62 mg Chl of PSII-enriched membrane. 2. Centrifuge the suspension at 32,900 g for 15 min. Resuspend the pellet in buffer D (see Note 7) to a final volume of 48 ml. 3. Add 12 ml 20% (w/v) Triton X-100 (60 ml total final volume at 1 mg Chl/ml and 4% w/v Triton) slowly with gentle stirring and incubate for 2 h also with gentle stirring. 4. Pellet at 32,900× g for 1 h. 5. Load the supernatant onto a TSK-GEL Fractogel DEAE 650(S) column (2.5 × 9  cm) that had been pre-equilibrated with buffer E. 6. Wash the column with the same buffer for 2 h at a flow rate of 2 ml/min, and then apply 170 ml of a 60–400 mM NaCl linear gradient in the same buffer but with 0.05% (w/v) Triton X-100 at a flow rate of 1 ml/min. 7. Pool the 3 ml fractions with a clear Qx Pheo band at around 543 nm. 8. Dilute the pooled eluate eightfold with buffer F and then load it onto a smaller TSK-GEL Fractogel DEAE 650(S) column (1.6 × 2.5 cm) pre-equilibrated with buffer E (see Note 8). 9. Wash the column overnight with the same buffer at a flow rate of 1 ml/min. 10. When the eluate shows a 417:435  nm peak ratio of about 1.16 (the wash step takes about 14 h), exchange b-DM for Triton X-100 by further washing with 45  ml of the same buffer, but with 0.1% (w/v) b-DM substituted for the Triton. 11. Elute the RC complex with 140  ml of a 60–350  mM NaCl linear gradient in the same buffer with 0.1% b-DM at a flow rate of 1 ml/min (the green band elutes at about 170 mM salt).

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12. Take the spectra of each 3 ml fraction, combine those with a red-peak maximum at 675  nm or higher and desalt by dialyzing against buffer G for 1 h with one buffer exchange (see Note 9). 13. If desired, the material can be concentrated using centrifugal filter devices concentrators. When the desired concentration is achieved, freeze the sample in liquid N2 and store under liquid N2. The samples can also be stored at −80°C in a freezer, if they are to be used within a short period of time or on dry ice if they are to be shipped. 3.1.3. Procedure for Isolating and Purifying 5-Chl PSII Reaction Center Complex

This procedure was developed by Vacha et al. (14). This preparation is directly eluted with b-DM, but may be less stable than the 6-Chl preparations since one of the proximal Chls is missing. Note that an oxygen-scrubbing system can also be added to this preparation to improve stability. The entire isolation procedure takes 30 h with the amount of starting material suggested below. The following steps should be carried out at 4°C in the dark. 1. Unfreeze 31 mg Chl of PSII-enriched membrane fragments and treat with 2 M CaCl2 for 10 min with gentle stirring. 2. Centrifuge the suspension at 32,900 g for 15 min and resu­ spend the pellet in 50 mM MES (pH 6.5) at a Chl concentration of 2 mg Chl/ml. 3. Solubilize by adding 1 volume of 30% (w/v) Triton X-100 to 3 volumes of treated membranes, and incubate for 2 h with gentle stirring. 4. Centrifuge at 40,000 g for 30 min to remove any nonsolubilized material. 5. Load the supernatant onto a Cu (II) affinity column (1 × 10 cm) at a flow rate of 4 ml/min. The column material is prepared by washing Fast-Flow Chelating Sepharose with distilled water and then degassing the material under vacuum to remove O2 bubbles that could interfere with the chromatography. At this point, the material is packed into the 1 × 10 cm column. The column is prepared at room temperature, washed extensively with distilled water and then with 10 ml of 0.1 M CuSO4. The excess Cu is removed by washing the column with 100 ml of distilled water, and finally the column is equilibrated with 50 ml of buffer H at 4°C. The use of both detergents seems to improve the stability of the preparation. 6. Wash the loaded column at a flow rate of 2.5 ml/min with buffer H until the 417:435  nm peak ratio of the eluate is about 1.16. 7. Elute the column with buffer I at a flow rate of 1 ml/min, and collect 3.0 ml fractions.

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8. Take the absorption spectra of each fraction and pool those with a red-peak maximum at 677 nm or higher. 9. If desired, the material can be concentrated using centrifugal filter devices concentrators, if small volumes are to be concentrated. When the desired concentration is achieved, freeze the sample in liquid N2 and store under liquid N2. The samples can also be stored at −80°C in a freezer, if they are to be used within a short period of time or on dry ice if they are to be shipped. 3.2. Purity of the Preparations 3.2.1. UV-VIS Absorption Spectra

3.2.2. Pigment Quantitation

Room temperature absorption spectra of 6-Chl RCs, obtained using either low (0.05% w/v) or high (1% w/v) Triton X-100 concentration in the column-washing buffers, exhibit a red-band maximum at 675.5  nm resulting from the Qy transition of Chl and Pheo, a 435 nm band due to Chl, and a 417 nm band mainly due to Pheo (but also to the Soret bands of Chl and Cyt b559). The distinct band at 543 nm corresponds to the Pheo Qx transition and the bands between 450 and 525 nm to b-carotene. The spectrum of a 5-Chl RC preparation is very similar to that of the 6-Chl RC material with the exception that the red-band peak is at 677  nm, and the absorption at around 670  nm is smaller on a relative basis. Determination of the Chl a, Pheo a, and b-carotene contents in all three PSII RC preparations can be done by spectral analysis of the pigments extracted in 80% (v/v) acetone following the method described in (15). The pigments (see Note 10) are extracted at room temperature under dim light, sonicated for 1 min, and centrifuged for 3 min in a microfuge to remove insoluble material. Complete pigment extraction is assured by repeating the extraction procedure twice and checking that no pigments (except Cyt b559) remain in the pellet. Note that the extract needs to have an optical density of between 0.3 and 0.6 absorption units in the Qy absorption region, to accurately calculate the pigment concentrations below from the absorption values found in the lower absorption regions. The following equations are used to determine the pigment concentrations where c and A are given in mM and cm−1, respectively.



cChl = 11.577 A663 – 76.994 A535–551 + 0.624 A480



cPheo = 0.020 A663 + 132.505 A535–551 – 1.150 A480



cCar = –0.146 A663 – 4.054 A535–551 + 8.311 A480

3.2.3. Polypeptide Content Determination by SDS-PAGE

Coomassie Brilliant Blue staining of a gel (SDS-PAGE containing 12.5% acrylamide) demonstrates that all three types of RCs contain the same polypeptides. The heaviest staining bands are

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at around 33 and 30  kDa, which correspond to D2 and D1, respectively, depending on the presence of urea in the gel. A weakly staining band at 55–60 kDa corresponds to a cross-linked D1-D2 heterodimer, and the low molecular weight band (9.5 kDa) corresponds to the a-subunit of the Cyt b559.

4. Notes 1. If market spinach is used, we have found that material purchased in Denver between the months of April and October produce good RC material. If spinach is purchased during the winter months, it can be difficult to (a) get membranes unless the Triton/Chl ratio is lowered or (b) separate RC and antenna complexes during RC isolation. 2. The centrifugation times may vary with conditions. The times given are the longest times that we have found necessary. 3. TSK-GEL Toyopearl DEAE 650(S) can be stored at room temperature or at 4°C; however, it should not be frozen. The user is cautioned that the material is an eye and skin irritant, and it is extremely flammable if dried. Refer to the instructions for preparing the resin prior to packing a column and for regenerating the material after use. 4. Increasing the column flow rate can shorten the timeconsuming, column-washing steps (although more buffer will be used), if precautions are taken to be sure that everything online (column, resin, fraction collector, tubing, pumps, for example) has the capability to maintain the higher flow rate. Note that the viscosity of a solution containing detergent is quite dependent on the temperature and thus at 4°C the viscosity is much higher than that at room temperature where all the chromatography tools and method specifications are normally checked. 5. Higher levels of the D1-D2 heterodimer band at 55–60 kDa in the SDS-PAGE indicate a more damaged RC preparation. The ideal situation is a nondetectable level of the heterodimer. 6. The RC preparations obtained with high Triton X-100 concentration (1% w/v) tend to have less b-carotene content. 7. The use of buffers at pH 6.5 (compared to higher pHs) results in a more transparent (less turbid) final preparation of RC. 8. When Taurine at a concentration of 1.5% (w/v) is added to the column-washing buffers, the RC-6 Chl preparation obtained with high Triton X-100 concentration (1% w/v) has less nonpigmented protein contaminant, but the yield of the isolated material is lower.

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9. The 6-Chl RC preparation obtained with high Triton X-100 concentration (1% w/v) in the column-washing buffer can be further purified, if necessary, using a sucrose density gradient. To make the gradient, a solution of 50 mM MES, pH 6.5, 500 mM sucrose, and 0.1% b-DM is placed in 10 ml centrifuge tubes. Tubes containing this solution are frozen at −20°C and are then allowed to slowly thaw at 4°C. This procedure will form a sucrose gradient. The tubes are then loaded with contaminated RC material, placed in a swinging bucket rotor (e.g., a Beckman Ti SW41 rotor), and centrifuged overnight at 35,000 rpm at 4°C. A major green band appears at the middle of the gradient, and this is the pure RC material. We emphasize that the contaminating material is nonpigmented since the pigment content of the preparation is 6 Chls/2 Pheos before and after the sucrose density step. Any green pigmentation in the pellet appears to be aggregated RC. 10. Recent studies (16, 17) reveal differences in the spectroscopy of the pigments associated with P680 in PSII core versus PSII RC preparations due to perhaps structural changes in the reaction center accompanied by the loss of QA. This observation provides new opportunities for the use of the preparations described in this paper for characterizing the effects of structure on the spectroscopy of the reaction center pigments.

Acknowledgments The authors would like to thank all of their current and past collaborators for helping us make good use of the preparations described in this article. This work was supported by the Chemical Sciences, Geosciences, and Biosciences Division, Office of Science, U.S. Department of Energy (MS), and by the MICINN of Spain (Grant BFU2005-04722-C02-01 and AGL2008-00377) to RP. References 1. Nanba, O. and Satoh, K. (1987) Isolation of a photosystem II reaction center consisting of D1 and D2 polypeptides and cytochrome b-559. PNAS USA 84,109–112. 2. Deisenhofer, J., Epp. O., Miki, K., Huber, R., and Michel H. (1985) Structure of the protein subunits in the photosynthetic reaction center of Rhodopseudomonas viridis at 3 Å resolution. Nature 318, 618–624.

3. Dekker, J. P. and Van Grondelle, R. (2000) Primary charge separation in photosystem II. Photosynth Res 63, 195–208. 4. Diner, B. A. and Rappaport, F. (2002) Structure, dynamics, and energetics of the primary photochemistry of photosystem II of oxygenic photosynthesis. Ann Rev Plant Biol 53, 551–580. 5. Kamiya, N. and Shen, J.-R. (2003) Crystal structure of oxygen-evolving photosystem II

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6.

7.

8.

9.

10.

11.

from Thermosynechococcus vulcanus at 3.7 Å resolution. Proc Natl Acad Sci USA 100, 98–1003. Guskov, A., Kern, J., Gabdulkhakov, A., Broser, M., Zouni, A., and Saenger, W. (2009) Cyanobacterial photosystem II at 2.9 Å resolution and the role of quinines, lipids, channels and chloride. Nat Struct Biol 16(2), 334–342. Berthold, D. A., Babcock, G. T., and Yocum, C. F. (1981) A highly resolved, oxygen-evolving photosystem II preparation from spinach thylakoid membranes. FEBS Lett 134, 231–234. Kuwabara, T. and Murata, N. (1982) Inactivation of photosynthetic oxygen evolution and concomitant release of three polypeptides in the photosystem II particles of spinach chloroplasts. Plant Cell Physiol 23, 533–539. Yamamoto, Y., Doi, M., Tamura, N., and Nishimura, M. (1981) Release of polypeptides from highly active O2-evolving photosystem 2 preparation by Tris treatment. FEBS Lett 133, 265–268. Dunahay, T. G., Staehelin, L. A., Seibert, M., Ogilvie, P. D., and Berg, S. P. (1984) Structural, biochemical and biophysical characterization of four oxygen-evolving photosystem II pre­ parations from spinach. Biochim Biophys Acta 764, 179–193. Seibert, M., Picorel, R., Rubin, A. B., and Connolly, J. S. (1988) Spectral, photophysical, and stability properties of isolated photosystem II reaction center. Plant Physiol 87, 303–306.

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12. Chapman, D. J., Gounaris, D., and Barber, J. (1988) Electron-transport properties of the isolated D1-D2-cytochrome b559 photosystem II reaction center. Biochim Biophys Acta 933, 423–431. 13. McTavish, H., Picorel, R., and Seibert, M. (1989) Stabilization of isolated PSII reaction center complex in the dark and in the light using polyethylene glycol and an oxygen-scrubbing system. Plant Physiol 89, 452–456. 14. Vacha, F., Joseph, D. M., Durrant, J. R., Telfer, A., Klug, D. R., Porter, G., and Barber, J. (1995) Photochemistry and spectroscopy of a five-chlorophyll reaction center of photosystem II isolated by using a Cu affinity column. Proc Natl Acad Sci USA 92, 2929–2933. 15. Eijckelhoff, C. and Dekker J. P. (1997) A routine method to determine the chlorophyll a, pheophytin a, and b-catotene contents of isolated photosystem II reaction center complexes. Photosynth Res 52, 69–73. 16. Hillmann, B., Brettel, K., van Mieghem, F. J. E., Kamlowski, A., Rutherford, A. W., and Schlodder E (1995) Charge recombination in Photosystem II. 2. Transient Absorbance difference spectra and their temperature dependence. Biochemistry 34, 4814–4827. 17. Smith, P.J., Peterson, S., Masters, V. M., Wydrzynski, T., Styring, S., Krausz, E., and Pace, R. J. (2002) Magneto-optical measurements of the pigments in fully active photosystem II core complexes from plants. Biochemistry 41, 1981–1989.

Chapter 4 Methods for the Isolation of Functional Photosystem II Core Particles from the Cyanobacterium Synechocystis sp. PCC 6803 Dmitrii V. Vavilin Abstract This chapter contains the description of several methods used for the isolation of functional photosystem II (PS II) core particles from wild type, photosystem I-less, and CP47 histidine-tagged cells of the cyanobacterium Synechocystis sp. PCC 6803. The presented protocols cover cultivation of photosystem I-containing and photosystem I-less cells, isolation of thylakoid membranes, purification of PS II core particles using a weak cation exchange or metal affinity column chromatography, and characterization of the final preparation. These isolation procedures yield highly active oxygen-evolving PS II particles and can be easily adapted for obtaining preparations from Synechocystis mutants with genetically modified photosystem II. Key words: Photosystem II, Synechocystis 6803, Isolation protocol, Anion exchange chromatography, Metal affinity chromatography, Oxygen evolution

1. Introduction Photosystem II (PS II) is a multisubunit pigment–protein complex that uses the energy of light to mediate electron transfer from water to plastoquinone in thylakoid membranes of plants, algae, and cyanobacteria (for recent reviews see refs. 1, 2). PS II is composed of more than 25 protein subunits, which include the reaction center proteins D1 and D2, the pigment-binding proteins CP43 and CP47, the a and ß subunits of cytochrome b559, and several Mn-stabilizing proteins of the oxygen-evolving complex. These proteins together with some other polypeptides of unknown function constitute the “core” of PS II.

Robert Carpentier (ed.), Photosynthesis Research Protocols, Methods in Molecular Biology, vol. 684, DOI 10.1007/978-1-60761-925-3_4, © Springer Science+Business Media, LLC 2011

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The unicellular photosynthetic cyanobacterium Synechocystis sp. strain PCC 6803 serves as an important model organism to study PS II structure and function (3, 4). Synechocystis sp. PCC 6803 has a number of advantages over many other eucaryotic and prokaryotic photosynthetic organisms. The strain is naturally transformable with a high efficiency and can integrate foreign DNA into its genome by double homologous recombination, thus enabling facile introduction of mutations into the genes encoding the PS II subunits. Moreover, Synechocystis sp. PCC 6803 can grow photoheterotrophically, in the absence of PS II or/and of photosystem I (PS I) activities, allowing for the construction and study of PS II mutants that normally have lethal phenotype. During the decades of intensive research, a number of protocols have been developed to isolate functional PS II complexes from Synechocystis (5–10). Because the PS I complex is at least five times more abundant in cyanobacterial thylakoids as compared to PS II, it has been a challenge to separate PS II from PS I while preserving the oxygen evolution activity of the former complex. Tang and Diner (5) were the first to obtain a PS II preparation, which was highly active in oxygen evolution and which contained less than 4% of contaminating PS I. The procedure, that took at least 12 h from cell harvesting to completion, included a purification step on a weak anion exchange column. A few years later, Bricker and coauthors (6) described an isolation of highly active PS II particles using a histidine-tagged mutant of CP47. Introduction of six histidines at the N-terminus of CP47 of Synechocystis had no significant effect on PS II function and allowed for the completion of the isolation procedure in less than 7  h using metal-affinity chromatography. The final preparation was highly depleted from many contaminating proteins, including PS I reaction center proteins (6). Isolation of PS II particles completely free from contaminating PS I was accomplished by using PS I-less cells (11) as a starting material together with minor adjustments of the purification protocol reported by Tang and Diner (7). The column chromatography protocols for the isolation of PS II particles from the wild type, his-tagged, and PS I-less strains of Synechocystis sp. PCC 6803 are described below (see Note 1).

2. Materials 2.1. Cyanobacterial Growth Medium

1. Stock solutions of K2HPO4 (3.05 g in 100 mL H2O), Na2CO3 (2.00 g per 100 mL H2O), ferric ammonium citrate (0.60 g per 100 mL H2O), Na2EDTA (0.25 M, pH 8.0), 1 M TESNaOH buffer, pH 8.2, and 1  M glucose (glucose solution

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should be filter-sterilized by passing through a 0.45  mM filter). 2. Stock solution of trace minerals: 2.86  g H3BO3, 1.81  g MnCl2 · 4H2O, 0.22 g ZnSO4 · 7H2O, 0.39 g Na2MoO4 · 2H2O, 0.079 g CuSO4 · 5H2O, and 0.049 Co(NO3)2 · 6H2O, 1.12 mL Na2EDTA from stock. Completely dissolve trace minerals one by one in approximately 900 mL H2O, add Na2EDTA, and bring final volume to 1 L. 3. Stock solution of macronutrients: 150  g NaNO3, 7.5  g MgSO4 · 7H2O, 3.6  g CaCl2 · 2H2O, and 100  mL of trace minerals stock solution. Dissolve salts one by one in approximately 800  mL of deionized Millipore Q grade water, add trace minerals, and bring the final volume to 1 L. 4. Modified BG-11 medium: Take approximately 900  mL of deionized Millipore Q grade water, add 10 mL of macronutrients, 10  mL TES buffer (not required for growing wild type Synechocystis strain), 1 mL of K2HPO4, 1 mL of Na2CO3, and 1  mL of ferric ammonium citrate from corresponding stock solutions, adjust to 1 L by adding water. Sterilize the medium by autoclaving on the same day. Filter-sterilized glucose (10 mM final concentration) is added to the autoclaved medium before starting the Synechocystis culture. The autoclaved BG-11 medium and sterile stock of glucose can be stored at room temperature for several months (the brownish pellet of iron salts formed in the medium after autoclaving does not affect cell growth). Other stock solutions should be stored at 4°C. 2.2. Thylakoid Isolation

1. BeadBeater or Mini-BeadBeater (Biospec Products). 2. Glass beads, 0.1 mm in diameter. 3. Buffer A (thylakoid buffer): 50  mM MES-NaOH, pH 6.0, 10 mM MgCl2, 5 mM CaCl2, and 25% (v/v) glycerol. Store at 4°C. 4. Buffer B (thylakoid buffer with protease inhibitors and DNase): 1 mM aminocaproic acid, 1 mM dimethyl sulfoxide, 1  mM phenylmethanesulfonyl fluoride (PMSF), 50  mg/L DNase I (from bovine pancreas, type IV, (e.g., Sigma)) in buffer A. Prepare immediately before use because of a short lifetime of PMSF in water. PMSF and benzamidine hydrochloride are initially dissolved in dimethyl sulfoxide. Aminocaproic acid and DNase are dissolved in water or buffer. 100  mM stock solutions of protease inhibitors can be stored at −20°C. Note that PMSF is toxic by inhalation, in contact with skin, and if swallowed; this chemical can be replaced with less toxic and more stable 4-(2-aminoethyl) benzenesulfonyl fluoride hydrochloride, AEBSF.

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2.3. PS II Solubilization and Column Chromatography

1. Chromatography column packed with DEAE-Toyopearl 650S (Toso Haas) weak cation exchanger or with Ni-NTA Superflow (Qiagen) affinity resin. Ni-resin from other manufacturers can be used as well. 2. Buffer C: 10 mM CaCl2 in buffer A. 3. Detergent stock solution: 10% (w/v) dodecyl b-d-maltoside in water. Can be stored at room temperature for at least several days. 4. Elution buffer D1: 20 mM MgSO4 and 0.03% w/v dodecyl b-d-maltoside in buffer C. 5. Elution buffer D2: 0.03% w/v dodecyl b-d-maltoside in buffer C. 6. Elution buffer D3: 0.04% w/v dodecyl b-d-maltoside in buffer A. 7. Elution buffer D4: 0.04% w/v dodecyl b-d-maltoside and 50 mM histidine in buffer A. All buffers listed in this section are stored at 4°C.

2.4. Quality Analysis

1. Buffer E: 50 mM MES-NaOH, pH 6.5, 10 mM NaCl, 5 mM MgCl2, and 20 mM CaCl2. 2. Potassium ferricyanide. 3. 2, 6-dichloro-p-benzoquinone. 4. Reverse phase C-18 HPLC column (e.g., Spherisorb S5ODS1; 250 × 4.6 mm). 5. Buffer F: 40  mM HEPES-NaOH buffer, pH 7.5, 10  mM NaCl, 5 mM MgCl2, 20 mM CaCl2, and 0.03% (w/v) dodecyl b-d-maltoside.

3. Methods 3.1. Cell Cultivation

Synechocystis cells are typically grown at 28–30°C in liquid BG-11 medium (12), which should be supplemented with 5–10  mM glucose and 10 mM TES-NaOH buffer (pH 8.2) for the cultivation of the PS I-less mutant (see Note 2) or mutants with functionally impaired PS II. A small amount of material (£0.5 L) can be grown in sterile flasks on a shaker. Larger volumes require continuous pumping of filter-sterilized air through the culture. Wild type (PS I-containing) cells are generally grown at light intensity of 50–100  mmol photons/m2s. The light intensity for growing PS I-less mutants should not exceed 5 mmol photons/ m2s. The doubling time of wild type cells typically varies from 9 to 13  h, whereas PS I-less mutants generally double every 20–24 h.

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Cells are harvested by centrifugation at their late logarithmic growth phase at OD730~ 0.8–1.0 (see Notes 3 and 4). The protocols presented below are based on a 20-L volume of cyanobacterial culture. A 20-L carboy typically yields 17–67  nmol of PS II particles containing 0.6–2.4 mg of chlorophyll. 3.2. Isolation of Thylakoid Membranes

This subsection outlines procedures for cell disruption and isolation of thylakoid membranes. All steps starting from here should be performed at 4°C at very low light or in darkness (see Notes 5 and 6). 1. Wash harvested cells by suspending them in 100 mL of buffer A. Pellet cells by centrifugation for 5 min at 6,000g. 2. Suspend the pellet in 50–100 mL of buffer B (see Notes 7–9). Load the cell suspension into a prechilled bead beater chamber filled with 0.1 mm glass beads to give a 1:2 to 1:1 (v/v) ratio of glass beads to the cell suspension. 3. Break cells with up to 15 break cycles, where each cycle consists of 15 s of homogenization followed by 2–4 min of cooling. After breakage, separate cell homogenate from the beads by decantation; wash the beads several times with buffer B to recover additional homogenate. The final volume of cell homogenate should be about 200 mL. 4. When isolating his-tagged PS II particles, steps 5–7 of this section can be skipped. Otherwise, continue from step 5 of this subsection. 5. Remove unbroken cells and glass beads remaining in the suspension by centrifugation (5 min, 1,000g). 6. Carefully aspirate or decant the supernatant into clean ultracentrifuge tubes (avoid taking the loose pellet). Pellet thylakoids by centrifugation for 20 min at 120,000g (see Note 10). 7. Suspend the thylakoids in 200 ml of buffer C and repellet by centrifugation for 20 min at 120,000g (see Note 10) in order to wash the material from remaining phycobilins and other soluble proteins. Discard the supernatant.

3.3. Solubilization of Thylakoids and Column Chromatography 3.3.1. Photosystem I-Containing Cells

The Subheadings 3.3.1, 3.3.2, and 3.3.3 contain the procedures for solubilization and chromatography purification of PS II from wild type (PS I-containing), PS I-less, and his-tagged cells, respectively. 1. Pre-equilibrate a weak anion exchange column (5.5 × 17 cm) containing DEAE-Toyopearl 650S resin with 500  mL of buffer D1. 2. Thoroughly suspend thylakoids in buffer C first with a paintbrush, and then use a Potter-Elvehjem tissue grinder to a final chlorophyll concentration of 1 mg/mL (see Note 11).

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3. Add dropwise a 10% stock solution of dodecyl b-d-maltoside to the thylakoid suspension to give a final detergent concentration of 1%. Proceed with the extraction for 10 min at 4°C with gentle stirring. 4. Pellet unsolubilized material by centrifugation (20  min, 40,000 g). 5. Apply the supernatant containing the solubilized material onto the column at a flow rate of 3–4 mL/min. 6. Wash the column with 500–1,000  mL of the equilibration buffer D1 at a flow rate of up to 10 mL/min under continuous monitoring of the absorption spectrum of the eluant. The wash with buffer D1 elutes in the order: free chlorophyll and carotenoids (fractions having green and orange colors), cytochromes (fractions with a pink color characterized by an absorbance peak around 400 nm), residual phycobiliproteins (greenish-yellow color with a wide absorption peak around 620 nm), and monomeric PS I complexes (fractions with an intensive green color characterized by an absorption maximum around 679 nm). PS II, which is characterized by a red absorption maximum at about 673.5 nm, is eluted right after the monomeric PS I. 7. After the red absorption maximum of the eluant decreased from ~679  nm to below 674.5  nm, apply a 500-mL linear gradient of 20–30  mM MgSO4 in buffer D1. Continue to wash the column with buffer D1 containing 30 mM MgSO4 until the red absorption maximum of the eluant does not shift above 675 nm due to the beginning of elution of PS I aggregates, characterized by an absorption peak around 679 nm. 8. Spool fractions containing PS II with the absorbance maxima ranging from 673.5 to 674.0 nm. 3.3.2. Photosystem I-Less Cells

1. Pre-equilibrate a weak anion exchange column (2.5 × 15 cm) containing DEAE-Toyopearl 650S resin with 200  mL of buffer D2. 2. Thoroughly resuspend thylakoids in buffer B first with a paintbrush, and then use a Potter-Elvehjem tissue grinder to a final chlorophyll concentration of 0.1 mg/mL (see Note 12). 3. Add dropwise a 10% stock solution of dodecyl b-d-maltoside to the thylakoid suspension to give a final detergent concentration of 0.4%. Proceed with the extraction for 30  min at 0°C with gentle stirring. Pellet unsolubilized material by centrifugation (20 min at 40,000 g). 4. Apply the supernatant containing the solubilized material onto the column at a flow rate of 2.5–3.0 mL/min.

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5. Wash the column with 100 mL of the equilibration buffer D2 at a flow rate of 6–7 mL/min. This wash elutes free chlorophyll and carotenoids. 6. Apply a 500-mL linear gradient of 0–50 mM MgSO4 in buffer D2, and then continue to wash the column with buffer D2, containing 50 mM MgSO4. Collect fractions during and after the salt gradient. 7. Pool PS II fractions eluted at the end of the MgSO4 gradient and during the wash with 50 mM MgSO4 which satisfy the following criteria: (1) the position of the peak of the Qy absorption band is in the range from 672.5 to 674.0 nm; (2) the ratio of b-carotene absorption at ~490 nm to chlorophyll a absorption at ~673  nm is below 0.55; and (3) no wellresolved absorption peak is observed at 420  nm (these fractions appear at the end of the PS II elution). 3.3.3. His-tagged Photosystem II

1. Pre-equilibrate Ni2+ affinity column (2.5 × 10 cm) containing 25 mL of Ni-NTA Superflow resin with 75 mL of buffer D3. 2. Under continuous gentle stirring, bring cell homogenate to a concentration of 1% dodecyl maltoside by dropwise addition of a 10% stock of the detergent. Without additional incubation period pellet unsolubilized material, unbroken cells, and residual glass beads by centrifugation (20 min at 40,000g). 3. Apply the supernatant containing the solubilized material onto the column at a flow rate of 2–3 mL/min. 4. Wash the column with 75  mL of buffer D3 at flow rate of 4–5 mL/min. 5. Elute PS II particles from the column with buffer D4 and collect fractions. Pool the fractions with an intense green color. These fractions should satisfy the criteria listed in Subheading 3.3.2.7.

3.4. Concentration and Storage of Photosystem II Particles

PS II particles can be concentrated by using centrifugal concentrators equipped with a 30 or 50 kDa MWCO filter (e.g., Vivaspin-20, Sartorius). This approach is rather time-consuming although it allows for the additional purification of the sample from low molecular weight contaminating proteins. Alternatively, PS II can be precipitated with polyethylene glycol (PEG), as described below. 1. Mix the eluted PS II particles with an equal volume of 25% PEG-8000 in buffer C containing 0.03% w/v dodecyl b-dmaltoside and incubate the mixture for 30 min on ice. 2. Harvest PS II by centrifugation (30 min, 40,000g). 3. Discard the supernatant, and resuspend the pellet in a small volume (~2 mL) of buffer C containing 0.03% w/v dodecyl b-d-maltoside.

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4. Put small aliquots in 0.6 mL microcentrifuge tubes, freeze in liquid nitrogen, and store in darkness at −80°C. 3.5. Quality Analysis

The quality of the purified PS II core particles can be assayed by measuring the functional activity (oxygen evolution), pigment composition, and contamination with PS I.

3.5.1. Oxygen Evolution

Oxygen evolution rates are measured using a Clark-type oxygen electrode under continuous illumination of samples at 25°C. PS II core complexes are suspended in buffer E to chlorophyll concentration of 2–10 mg/mL. Potassium ferricyanide (1 mM) and 2, 6-dichloro-p-benzoquinone (0.3 mM) are added to the suspension as electron acceptors. Upon illumination with a saturating light (³1,000  mmol photons/m2s), fully active PS II particles evolve oxygen at a rate of about 2,500 mmol O2/mg chlorophyll·h.

3.5.2. Absorbance Spectra

A typical absorbance spectrum of purified PS II particles is shown in Fig. 1. The spectrum is characterized by large chlorophyll a absorption peaks at ~436 and 673–674 nm. Absorption of pheophytin a is generally observed as a small peak or a shoulder near 545 nm. The absorption around 460–500 nm is attributed to carotenoids (mostly b-carotene); the absorption ratio A490/A674 varies from 0.35 to 0.45, but may be larger if cells used to isolate PS II particles contained significant amount of carotenoids. The absorption ratio A625/A674 ~0.22 indicates that the preparation is essentially free of contaminating phycobiliproteins, which are characterized by a broad absorption band peaking at 620–623 nm.

Fig.  1. Typical absorption spectra of isolated PS I (dashed line) and PS II (solid line) particles.

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3.5.3. HPLC Analysis

Pigments are extracted at 4°C by adding nine volumes of cold HPLC grade acetone to one volume of PS II preparation (e.g., 90 mL of acetone to 10 mL of the sample). The mixture is shacked vigorously for 30  s and then centrifuged in a microcentrifuge (3  min at a maximum speed) to precipitate proteins. To avoid pheophetinization of chlorophyll, the extract is immediately injected in a chromatograph for analysis on a reverse phase C-18 HPLC column. The column is eluted at a flow rate of 1 mL/min with an isocratic mobile phase consisting of methanol and ethyl acetate (68:32, v:v). The pigments are detected at 663 nm (chlorophyll a and pheophytin a), 450 nm (b-carotene), and 254 nm (plastoquinone-9). Extinction coefficients for chlorophyll a, pheophytin a, and b-carotene dissolved in the methanol/ethyl acetate mix (68:32, v:v) are 86.9 mM/cm at 663 nm, 49.3 mM/ cm at 663 nm , and 135.0 mM/cm at 450 nm, respectively (13). The extinction coefficient of plastoqunone-9 dissolved in hexane is 19.1  mM/cm at 254  nm (14). A typical PS II preparation contains 33-40 chlorophyll a molecules, ³10 b-carotenes, and ~2 plastoquinones per two pheophytins.

3.5.4. Contamination with Photosystem I

Contamination of the preparation with PS I can be assessed by measuring ascorbate-reduced minus ferricyanide-oxidized absorption difference spectrum using a double-beam spectrophotometer. PS II particles are suspended in buffer F to OD674~1 and then placed in two different cuvettes. P700 is oxidized in the first cuvette by adding ferricyanide (20 mM). A few granules of solid sodium ascorbate and phenazine methosulfate (20 nM) are added to the second cuvette and the difference absorption spectrum is recorded a few minutes later. The upper value for the concentration of P700 can be calculated using the differential absorption coefficient De705–750 = 64 mM/cm (15) and the percentage of chlorophyll molecules from contaminating PS I can be estimated considering that each PS I complex can contain 96 chlorophylls (16).

4. Notes 1. PS II core particles can also be purified by gradient centrifugation in sucrose, or gradient centrifugation can be used as an additional purification step in combination with column chromatography (8–10). One of the major disadvantages of this approach is that it cannot be scaled up to isolate large quantities of the material often required for spectroscopic studies. Also preparations obtained by sucrose density centrifugation are usually inactive in oxygen evolution.

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2. To accelerate cell growth, glucose (5 mM final concentration) can be added to wild type Synechocystis cultures as well. 3. The suspension of wild type Synechocystis cells with OD730 = 1 (measured in a 1-cm cuvette using a Shimadzu UV-150 spectrophotometer) contains 2–5 mg chlorophyll/mL (0.5–1.0 mg chlorophyll/mL for PS I-less cells). Growing cultures to higher optical density results in less efficient cell breakage. 4. Cyanobacterial cells can be harvested by 5-min centrifugation at 3,000g using a conventional centrifuge. Considering the large volume of the starting material, it is highly desirable to use rotors accommodating large-volume buckets that allow concentration of 4–6 L of cell suspension in one run. In this case, the harvesting of 20-L culture can be completed in 1.5–2.0 h. Another option is to use a continuous flow rotor/ centrifuge, or a special filtration apparatus for bacterial cells (e.g., Pellicon tangential ultrafiltration device equipped with 0.4 mm filter, Millipore). 5. Because PS II complexes are easily and irreversibly damaged by illumination, exposure of the material to light should be reduced to a minimum. It is best to perform the isolation in a cold room under a dim green light or in darkness, whenever possible. 6. Freezing of cells or isolated thylakoids in liquid nitrogen for their subsequent storage at low temperature in between the isolation steps results in the partial loss of oxygen-evolving activity of the final preparation (5). Therefore, only the fresh material should be used for the isolation of PS II. If the isolation procedure cannot be completed in 1  day, the minimal loss of oxygen evolution occurs if solubilized PS II particles are left overnight on the column (the column with PS II should be washed with 0.03–0.04% dodecyl maltoside dissolved in the corresponding washing buffer and then stored in complete darkness at 4°C). 7. The pelleted material (cells, thylakoids, or purified PS II particles) is easily suspended with a small paintbrush soaked in ice-cold wash buffer. 8. The optimal pH for the isolation of oxygen evolving PS II particles is about 6.0–6.4. Higher pH (7.2–7.4) cause the partial loss of oxygen evolution. However, the chlorophyll to pheophytin ratio of the final preparation tends to be higher, if the isolation is performed at slightly basic pH. 9. The volume of cell suspension and the amount of added glass beads depend on the type of available bead beater chamber. Variations in the concentration of cell suspension and the amount of glass beads within certain limits do not affect the efficiency of cell breakage.

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10. Since most of thylakoids can be pelleted by 30 min centrifugation at a lower centrifugal force (~40,000g), the isolation protocol can be adapted to use an ordinary centrifuge (e.g., Sorvall RC-5 centrifuge equipped with SS-34 rotor) rather than an ultracentrifuge. 11. Chlorophyll concentration is determined by pigment extraction with acetone or methanol (100% methanol is particularly good for chlorophyll extraction from unbroken cells). Proteins are precipitated from the extract by 3 min centrifugation in a bench microcentrifuge at maximum speed. The specific extinction coefficients for chlorophyll dissolved in 100% methanol (lmax = 665.2 nm) and in 80% acetone (lmax = 663.6 nm) are 80.0 mL mg/cm and 86.0 mL mg/cm, respectively (17). 12. PS I-less cells have nearly fivefold-reduced chlorophyll content as compared to wild type Synechocystis. Therefore, initial solubilization of thylakoid membranes in dodecyl maltoside is performed at a lower chlorophyll concentration. The best results were obtained when final concentration of the detergent was reduced to 0.4%, but the solubilization continued for a longer time (30 min) comparing to wild type cells.

Acknowledgments The author would like to thank Dr. Wim Vermaas for his support and Dr. Ilya R. Vassiliev for the absorption spectrum of isolated PS I complex. References 1. Satoh K. (2008) Protein pigments and the photosystem II reaction center: a glimpse into the history of research and reminiscences. Photosynth Res 98, 33–42. 2. Kern, J. and Renger, G. (2007). Photosystem II: Structure and mechanism of the water: plastoquinone oxidoreductase. Photosynth Res 94, 183–202. 3. Vermaas, W. F. J. (1998) Gene modifications and mutation mapping to study the function of photosystem II. Method Enzymol 297, 293–310. 4. Vermaas, W. (1996) Molecular genetics of the cyanobacterium Synechocystis sp. PCC 6803: Principles and possible biotechnology applications. J Appl Phycol 8, 263–273. 5. Tang, X. S. and Diner, B. A. (1994) Biochemical and spectroscopic characterization of a new oxygen-evolving photosystem II core complex

from the cyanobacterium Synechocystis PCC 6803. Biochemistry 33, 4594–4603. 6. Bricker, T. M., Morvant, J., Masri, N., Sutton, H. M., and Frankel, L. K. (1998) Isolation of a highly active Photosystem II preparation from Synechocystis 6803 using a histidinetagged mutant of CP 47. Biochim Biophys Acta 1409, 50–57. 7. Vavilin, D., Xu, H., Lin, S., and Vermaas, W. (2003) Energy and electron transfer in photosystem II of a chlorophyll b-containing Synechocystis sp. PCC 6803 mutant. Biochemistry 42, 1731–1746. 8. Szabo, I., Rigoni, F., Bianchetti, M., Carbonera, D., Pierantoni, F., Seraglia, R., Segalla, A., and Giacometti, G. M. (2001) Isolation and characterization of photosystem II subcomplexes from cyanobacteria lacking photosystem I. Eur J Biochem 268, 5129–5134.

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9. Rögner, M., Nixon, P. J., and Diner, B. A. (1990) Purification and characterization of photosystem I and photosystem II core complexes from wild type and phycocyanin-deficient strains of the cyanobacterium Synechocystis PCC 6803. J Biol Chem 265, 6189–6196. 10. Xu, H., Vavilin, D., and Vermaas, W. (2001) Chlorophyll b can serve as the major pigment in functional photosystem II complexes of cyanobacteria (2001) Proc Natl Acad Sci USA 98, 14168–14173. 11. Shen, G.-Z., Boussiba, S., and Vermaas, W.F.J. (1993) Synechocystis sp. PCC 6803 strains lacking photosystem I and phycobilisome function. Plant Cell 5, 1853–1863. 12. Rippka, R., Deruelles, J., Waterbury, J. B., Herdmann, M., and Stanier, R. Y. (1979) Generic assignments, strain histories and properties of pure cultures of cyanobacteria. J Gen Microbiol 111, 1–61. 13. Giorgi, L. B., Nixon, P. J., Merry, S. A. P., Joseph, D. M., Durrant, J. R., De Las Rivas,

14. 15. 16.

17.

J., Barber, J., Porter, G., and Klug, D. R. (1996) Comparison of primary charge sepa­ ration in the photosystem II reaction center complex isolated from wild-type and D1-130 mutants of the cyanobacterium Synechocystis PCC 6803. J Biol Chem 271, 2093–2101. Morton, R. A. (1965) Biochemistry of quinones. London, New York: Academic press. Hiyama, T. and Ke, B. (1972) Difference spectra and extinction coefficients of P700. Biochim Biophys Acta 267, 160–171. Jordan, P., Fromme, P., Witt, H. T., Klukas, O., Saenger, W., and Krauss, N. (2001) Threedimensional structure of cyanobacterial photosystem I at 2.5 A resolution. Nature 411, 909–917. Porra, R. J. (2002) The chequered history of the development and use of simultaneous equations for the accurate determination of chlorophylls a and b. Photosynth Res 73, 149–156.

Chapter 5 Purification and Crystallization of Oxygen-Evolving Photosystem II Core Complex from Thermophilic Cyanobacteria Jian-Ren Shen, Keisuke Kawakami, and Hiroyuki Koike Abstract This chapter describes the purification and crystallization of oxygen-evolving photosystem II core dimer complex from a thermophilic cyanobacterium Thermosynechococcus vulcanus. Procedures used for purification of photosystem II from the cyanobacterium involves cultivation of cells, isolation of thylakoid membranes, purification of crude and pure photosystem II core complexes by detergent solubilization, followed by differential centrifugation and column chromatography. The purified core dimer particles were successfully used for crystallization, and the methods and conditions used for crystallization are presented. These purification and crystallization procedures can be applied for another thermophilic cyanobacterium T. elongatus. Key words: Photosystem II, Oxygen evolution, Crystallization, Membrane proteins, Ion-exchange chromatography

1. Introduction Highly active oxygen-evolving photosystem II (PSII) core complexes have been purified from various organisms, and used for various biochemical, biophysical, and structural biological studies. The procedures for purification of PSII core complexes from higher plants and a mesophilic cyanobacterium Synechocystis sp. PCC 6803 have been described in the previous Chapters of this volume. Thermophilic cyanobacteria are unique in that they grow at temperatures much higher (45–60°C) than mesophilic cyanobacteria, so that proteins isolated from thermophilic cyanobacteria are usually much more stable than those from mesophilic cyanobacteria. This has been proven to be a feature important for

Robert Carpentier (ed.), Photosynthesis Research Protocols, Methods in Molecular Biology, vol. 684, DOI 10.1007/978-1-60761-925-3_5, © Springer Science+Business Media, LLC 2011

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the structural studies of PSI and PSII, since crystallization of these large membrane–protein complexes has to be performed at medium temperatures for a long period of time. In particular, the oxygen-evolving activity of PSII is rather unstable, and the extrinsic proteins involved in stabilizing the oxygen-evolving complex are usually easy to be released in PSII from mesophilic organisms. In contrast, the extrinsic proteins are found to be tightly associated with PSII purified from thermophilic cyanobacteria (1, 2), and the oxygen-evolving activity of PSII isolated from a thermophilic cyanobacterium Thermosynechococcus elongatus has been shown to be stable even after 3 weeks incubation at 20°C (3). T. vulcanus and T. elongatus are two thermophilic cyanobacteria very similar with each other in terms of their gene sequences and cellular growth or photosynthetic characteristics. The two cyanobacteria are isolated from two different Japanese hot springs, and they both grow at 50–60°C. This is among the highest temperatures known so far for photosynthetic organisms to be able to grow. Highly purified and active oxygen-evolving PSII has been obtained from both cyanobacteria, and they were successfully used for crystallization and structure analysis (4–7). So far, the crystal structure of PSII has been solved only for these two cyanobacteria, although PSII from an acidophilic, thermophilic red alga Cyanidium caldarium has been crystallized recently (8). Here we have described the purification procedure for PSII core complex from T. vulcanus, which involves the culture of cells, isolation of thylakoid membranes and crude PSII, and purification of pure PSII dimer by ion-exchange chromatography. Subsequently, crystallization procedures and conditions for the purified PSII dimers are described. Similar procedures can be applied for the purification and crystallization of PSII from T. elongatus.

2. Materials 2.1. Basic Materials

1. Strain: Thermosynechococcus vulcanus (or T. elongatus). 2. Growth chamber capable of maintaining temperatures at 50–55°C. 3. High-speed refrigerated centrifuge, ultra-speed refrigerated centrifuge, and angled rotors. 4. Spectrophotometer for determining chlorophyll (chl) concen­ trations. 5. Brushes for suspending membranes and core particles. 6. Temperature-controlled water bath (38°C). 7. Equipment for column chromatography, including pumps and fraction collector, which is capable of making a liner gradient of

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salt concentration and maintained at a low-temperature chamber or cold room. 8. Chromatography column packed with Q sepharose highperformance (GE Healthcare). 9. Crystallization plates, cover slides, and temperature-controlled incubators for crystallization. 10. Microscope for the observation of crystals. 2.2. Groth Medium for T. vulcanus (see Note 1)

First make 1 l each of the following: 1. Stock solution 1: 45 g KH2PO4, 55 g K2HPO4. 2. Stock solution 2: 100 g NaNO3, 100 g KNO3, 30 g MgSO4 ·  7H2O. 3. Stock solution 3: Trace minerals containing 6  g EDTA-2Na, 3.2  g FeCl3 · 6H2O, 0.132  g ZnSO4 · 7H2O, 0.376  g CuSO4 ·  5H2O, 4.4 g (NH4)6Mo7O24 · 4H2O, 0.003 g Co(NO3)2 · 6H2O, 0.8 g MnCl2 · 4H2O, 6.8 g H3BO4. 4. Stock solution 4: 60 g CaCl2 · 2H2O. 5. Final growth medium: 5 ml each of the stock solutions (steps 1–3) and 1 ml of stock solution (step 4). Dilute to 1 l with distilled water. (If the growth medium is to be sterilized, make 1  l of diluted solution containing the stock solutions (steps 1–3) only, and sterilize it. After cooling the solution, add 1 ml of stock solution (step 4), which has been sterilized separately by passing through a 0.22-mm filter. The pH of the final medium should be at 7.0–7.5.)

2.3. Stock Solutions and Buffers for Isolation of Thylakoid Membranes

1. Stock solutions: 1  M MgCl2 · 6H2O, 0.5 M EDTA-2Na, pH 8.0, DNase I (1  mg/ml, stored at −20°C), Lysozyme (stored at 4°C). 2. Buffer B-1: 40 mM KH2PO4–KOH, pH 6.8, 2 l. 3. Buffer B-2: 40 mM KH2PO4–KOH, pH 6.8, 0.4 M mannitol, 2 l. 4. Buffer B-3: 30  mM Hepes–NaOH, pH 7.0, 10  mM MgCl2 · 6H2O, 2 l. 5. Buffer B-4: 30  mM Hepes–NaOH, pH 7.0, 25% glycerol, 10 mM MgCl2 · 6H2O, 1 l.

2.4. Solutions and Buffers for Isolation of Crude PSII Particles

1. Detergent: N,N-dimethyldodecylamine-N-oxide (LDAO) (~30%) (Fluka). 2. Stock solution: 50% polyethylene glycol (PEG) 1,450 (Sigma), 500 ml. 3. Buffer B-5: 5% glycerol, 30  mM Hepes–NaOH, pH 7.0, 500 ml.

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2.5. Solutions and Buffers for Isolation of pure PSII Dimer

1. Detergent: 20% n-dodecyl-b-d-maltoside (b-DDM) (Anatrace Co.). Store at −20°C. 2. Column buffer-A: 30 mM Mes–NaOH, pH 6.0, 3 mM CaCl2, 0.03% b-DDM. 3. Column buffer-B: 30 mM Mes–NaOH, pH 6.0, 3 mM CaCl2, 0.03% b-DDM, 1 M NaCl. 4. Buffer B-6: 30  mM Mes–NaOH, pH 6.0, 20  mM NaCl, 3 mM CaCl2.

2.6. Buffers for Crystallization

1. Crystallization buffer C-1: 20  mM Mes–NaOH, pH 6.0, 20 mM NaCl, 10 mM CaCl2, 40 mM MgSO4, 0.03% b-DDM, 20% glycerol, 4–10% PEG 1,450. 2. Reservoir buffer C-2: 20 mM Mes–NaOH, pH 6.0, 20 mM NaCl, 10 mM CaCl2, 40 mM MgSO4, 0.015% b-DDM, 10% glycerol, 7% PEG 1,450.

3. Methods The following procedures describe growing of the thermophilic cyanobacterial cells, isolation of thylakoid membranes and crude PSII particles, preparation of purified PSII dimers, and growth of 3-dimensional crystals suitable for X-ray crystallographic analysis. 3.1. Growth of Cells and Purification of PSII 3.1.1. Growth of Cells

T. vulcanus can be grown at temperatures up to 60°C; for laboratory use, cells are typically grown at 50–55°C. Small volumes of liquid culture can be grown in sterilized flasks on a shaker. For accelerating the growth, bubbling with air containing 1–5% CO2 is recommended. In the beginning of cultivation with a low cell density, the light intensity should be kept weak, for example, around 5–15  mmol photons/m2 s. The light intensity may be increased gradually when the cell density increases; and the final light intensity may reach to 50–100  mmol photons/m2 s. For a larger volume of culture such as 50 l, sterilization may be difficult, and the growth may be carried out without sterilization. Since the cyanobacterium grows at a high temperature, contamination from other bacteria is rather limited. In fact, we routinely grow the cyanobacterium in a 50 l scale without sterilization of the medium. For the growth of the larger volume, continuous circulation of the medium with a strong water pump is required to avoid the sedimentation and adhesion of cells to the walls of the container, and bubbling with CO2-containing air is strongly recommended. The doubling time of the cells is typically 6–12 h and may be shortened up to 4 h at optimum conditions. Cells harvested from a 50 l culture at their late logarithmic growth stage typically will

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give thylakoids of 400–500 mg chl, and this will give rise to 6–12  mg chl of PSII dimers using the purification procedures described below. 3.1.2. Preparation of Thylakoid Membranes

The cells can be broken either mechanically by glass beads with a beads beater or enzymatically with lysozyme followed by an osmotic shock-treatment. Here we describe the method of cell breakage by enzyme treatment, together with the procedure for preparation of thylakoid membranes. The following steps 1–4 are carried out at room temperature and steps after step 4 are performed at 4°C or on ice. 1. Harvest cells from a 50 l culture and wash them with 1 l of buffer B-1. 2. Suspend the cells in 1 l of buffer B-2, and add EDTA from a 500  mM stock solution to a final concentration of 1  mM. Add lysozyme to a final concentration of 1.2 mg/ml. Incubate the solution at 38°C for 2.5 h with gentle shaking in a water bath. 3. Collect the cells by centrifugation at 12,000g for 5  min at room temperature, and wash the precipitate with 0.8 l of buffer B-2. 4. Suspend the precipitate with a small volume of B-2 (see Note 2). Add 1 l of 20 mM Hepes–NaOH (pH 7.0), which has been pre-heated to 38°C, as quick as possible and mix the solution immediately so as to induce a sufficient osmotic shock to break the cells. Add MgCl2 to a final concentration of 5 mM and 200 ml of DNase I solution of 1 mg/ml. Mix and incubate for a short time (less than 5 min is enough) to allow the DNase treatment to proceed. The following steps should be performed at 4°C or on ice. 5. Centrifuge the solution at 15,000g for 10 min and wash the precipitate with 1 l of buffer B-3. 6. Suspend the final precipitate to buffer B-4 at a chl concentration of 2–3 mg/ml. This is the thylakoid membrane that can be stored at −80°C.

3.1.3. Preparation of Crude PSII Particles

Pure PSII dimers are purified in two steps. First, thylakoid membranes are solubilized with LDAO, and crude PSII particles are obtained by several times of differential centrifugations. Subsequently, the crude PSII particles are solubilized by b-DDM and pure PSII dimers are separated from monomers by ionexchange column chromatography. These procedures are described in this and the following section. These steps should be carried out at 4°C or on ice under dim light conditions. 1. Take 220  mg chl of thylakoid membranes and then wash it with 1 l of buffer B-3 by centrifugation at 9,000g for 10 min.

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2. Suspend the precipitate with buffer B-5 to a final, overall weight of 110 g. 3. Solubilize the precipitated membranes by adding 0.59  g of LDAO (~30%) and stir gently on ice for 5 min (see Note 3). 4. Precipitate the LDAO-treated membranes by centrifugation at 35,000g for 1 h. Suspend the precipitate with buffer B-4 to a final weight of 140 g. 5. Perform the second LDAO-treatment by adding 1.25  g of LDAO and stir gently on ice for 5 min (see Note 4). 6. Remove PSI and other non-PSII components by centrifugation at 100,000g for 1 h. Collect the supernatant, and dilute it twofolds with buffer B-4. Centrifuge at 100,000g for another 1 h to remove residual non-PSII components. 7. Collect the supernatant and add PEG 1,450 to a final concentration of 13%. 8. Centrifuge at 100,000g for 30 min. The precipitate yielded is crude PSII particles. Suspend it in B-4 and store at −80°C or liquid nitrogen. The crude PSII particles obtained above typically have an oxygen-evolving activity of 1,000–2,000  mmoles O2/mg chl/h and a yield of 10–20% based on the chl of the starting thylakoid membranes. A lower oxygen-evolving activity and/or a higher yield suggest contaminations by PSI. In this case, the amount of LDAO added in the second LDAO-treatment step should be reduced by 5–15%. On the other hand, a lower yield suggests an insufficient extraction of PSII by the LDAO solubilization, and the amount of LDAO used in the second LDAO-treatment should be increased by 5–15%. 3.1.4. Preparation of pure PSII Dimers and Monomers

The crude PSII obtained above contains a large amount of phycobili-proteins as well as some other contaminating proteins such as PSI and is a mixture of PSII dimer and monomers. This is further purified by ion-exchange column chromatography following solubilization by b-DDM. 1. Solubilize crude PSII in buffer B-4 with 1% b-DDM at 1 mg chl/ml. Stir gently for 30 min on ice in the dark. 2. Filtrate the solubilized crude PSII with a 0.45-mm disc filter and load the sample onto a Q sepharose high-performance column, which has been pre-equilibrated in column buffer-A (see Note 5). 3. Wash the column with 17% column buffer-B (e.g., at 170 mM NaCl) at a flow rate of 2–3 ml/min until the absorbance of the eluate at 280 nm decreases to a sufficiently low and constant level. Most of the phycobili-proteins and residual PSI should be eluted by this washing step.

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Fig.  1. Elution pattern of b-DDM solubilized crude PSII particles from a Q sepharose high-performance column. Crude PSII particles were prepared with the two steps-LDAO solubilization method, and a linear gradient of 170–300 mM NaCl was used to separate PSII monomers and dimers. The eluate was monitored at 280 nm.

4. Elute the column with a linear gradient of NaCl of 170–300 mM. A typical elution pattern is shown in Fig. 1, where the complexes are eluted in the order of PSII monomers, PSII dimers, and PSI trimers. The amount of PSII dimers is usually much larger than that of PSII monomers or PSI trimers. If the amount of PSII monomers is larger than PSII dimers, some of the PSII dimers may have been monomerized during the preparation of crude PSII by LDAO-treatment, and the conditions for LADOsolubilization should be optimized to minimize the amount of PSII monomers. If the amount of PSI is remarkably increased, it suggests a significant contamination of PSI in the crude PSII particles, and conditions for LDAO-solubilization and subsequent differential centrifugations should be optimized to reduce the contamination by PSI. 5. Collect the fraction of PSII monomers and dimers separately and dilute them with two volumes of buffer B-6 respectively. Add PEG 1,450 to a final concentration of 13% and pellet PSII by centrifugation at 100,000g for 20 min. 6. Suspend the PSII monomers and dimers with buffer B-6 at a chl concentration as high as possible and store them in liquid nitrogen. The purified PSII monomers and dimers are stable in B-6; however, for repeated use of the sample, 25% glycerol may be included in the suspending buffer to minimize inactivation during multiple freeze–thawing cycles.

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Fig. 2. SDS-PAGE analysis of PSII monomers (lane 1) and dimers (lane 2). SDS-PAGE was performed with a 16–22% acrylamide gradient gel (11).

The PSII dimers obtained above bind three extrinsic proteins of PsbO, PsbU, and PsbV (cytochrome c-550) (Fig. 2), exhibits an oxygen-evolving activity of 2,500–4,000 mmol O2/mg chl/h, and has a yield of 2–4% based on the chl from the starting thylakoid membranes. If the activity or yield is much lower, the purification procedures need to be improved. PSII monomers have a slightly lower oxygen-evolving activity and much lower yield than those of PSII dimers. 3.2. Crystallization of Purified PSII Dimers

Crystals of PSII dimers can be grown with the hanging drop, sitting drop vapor diffusion method, or the batch method, at 20°C. Here we describe the crystallization of PSII dimers by the hanging drop vapor diffusion method (9). 1. Adjust the concentration of PSII dimers to a chl concentration of 4 mg/ml in buffer B-6. 2. To make the crystallization droplets, mix 4–5 ml of PSII dimers with an equal volume of crystallization buffer C-1 (see Note 6), and set the droplet to a cover slide whose surface has been siliconized in advance.

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Fig. 3. Crystals of PSII dimers. The crystals were grown at 20°C for 5 days and have a dimension of 1.0 × 0.5 × 0.1 mm.

3. Set the hanging drop against a reservoir of 0.5 ml containing reservoir buffer C-2 and allow the crystals to grow for up to 1 week at 20°C. 4. Rhombic crystals appear in 3–4 days to 0.5 × 0.3 × 0.05 mm and reach to a maximum size of 1.0 × 0.7 × 0.1 mm in 1 week (Fig. 3). 5. For X-ray diffraction experiments at 100 K, the crystals are transferred to a cryoprotectant solution containing 25% glycerol, 20% PEG 1,450 (in addition to the other components contained in the crystallization solution) gradually, and are flash frozen with a nitrogen gas stream, and stored in liquid nitrogen.

4. Notes 1. BG11 can also be used for the growth of thermophilic cyanobacterium. In addition, a DTN medium has been used for the growth of T. elongatus (10). The selection of growth media may depend on the cost and easiness of the method with which to make. 2. The volume of buffer B-2 used to suspend the lysozymetreated thylakoid membranes should be kept as small as possible, in order to ensure the effective osmotic shock to take place in the subsequent step.

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3. The amount of LDAO used in the first LDAO-solubilization step should not lead to partial solubilization of the thylakoid membranes. That is, there should be no chl present in the supernatant after centrifugation of the solubilized membranes. If there is chl present in the supernatant, the amount of LDAO used for solubilization should be reduced slightly. 4. The amount of LDAO used in the second solubilization step may vary slightly depending on the growth condition of the cells and conditions of the thylakoid membranes. A suitable amount of LDAO should give rise to a yield of 10–20% of crude PSII based on the chl in the starting thylakoid membranes. Increase the amount of LDAO if the yield is lower than this range and reduce the amount of LDAO if the yield exceeds 20%. 5. Other columns such as Mono Q (GE Healthcare) or DEAE650S (TOSHO) can be used. Mono Q is a strong anion exchange column and can be used to purify PSII with a similar washing and eluting salt concentrations (9). If a DEAE650S column is used, the washing concentration should be reduced to 100  mM NaCl, and a linear gradient of 100– 250 mM NaCl should be used for elution. 6. The optimum concentration of precipitation agent PEG 1,450 may vary depending on the sample condition and should be determined for each sample by varying its concentration from around 4.0 to 6.0%, with a 0.5% difference for each step. References 1. Shen, J.-R., Ikeuchi, M., and Inoue Y. (1992) Stoichiometric association of extrinsic cytochrome c-550 and 12 kDa protein with a highly purified oxygen-evolving photosystem II core complex from Synechococcus vulcanus. FEBS Lett 301, 145–149. 2. Shen, J.-R. and Inoue Y. (1993) Binding and functional properties of two new extrinsic components, cytochrome c-550 and a 12 kDa protein, in cyanobacterial photosystem II. Biochemistry 32, 1825–1832. 3. Sugiura, M. and Inoue, Y. (1999) Highly purified thermo-stable oxygen-evolving photosystem II core complex from the thermophilic cyanobacterium Synechococcus elongatus having his-tagged CP43. Plant Cell Physiol 40, 1219–1231. 4. Zouni, A., Witt, H. T., Kern, J., Fromme, P., Krauss, N., Saenger, W., and Orth, P. (2001) Crystal structure of photosystem II from Synechococcus elongatus at 3.8 Å resolution. Nature 409, 739–743.

5. Kamiya, N. and Shen, J. –R. (2003) Crystal structure of oxygen-evolving photosystem II from Thermosynechococcus vulcanus at 3.7-Å resolution. Proc Natl Acad Sci USA 100, 98–103. 6. Ferreira, K. N., Iverson, T. M., Maghlaoui, K., Barber, J., and Iwata, S. (2004) Architecture of the photosynthetic oxygen-evolving center. Science 303, 1831–1838. 7. Guskov, A., Kern, J., Gabdulkhakov, A., Broser, M., Zouni, A., and Saenger, W. (2009) Cyanobacterial photosystem II at 2.9 Å resolution: role of quinones, lipids, channels and chloride. Nat Struct Mol Biol 16, 334–342. 8. Adachi, H., Umena, Y., Enami, I., Henmi, T., Kamiya, N., and Shen, J.-R. (2009) Towards structural elucidation of eukaryotic photosystem II: Purification, crystallization and preliminary X-ray diffraction analysis of photosystem II from a red alga. Biochim Biophys Acta 1787, 121–128.

Purification and Crystallization of Oxygen-Evolving Photosystem II Core Complex 9. Shen, J.-R. and Kamiya, N. (2000) Crystallization and the crystal properties of the ­oxygen-evolving photosystem II from Synechococcus vulcanus. Biochemistry 39, 14739–14744. 10. Mülenhoff, U. and Chauvat, F. (1996) Gene transfer and manipulation in the thermophilic

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cyanobacterium Synechococcus elongatus. Mol Gen Genet 252, 93–100. 11. Ikeuchi, M. and Inoue, Y. (1988) A new 4.8-kDa polypeptide intrinsic to the PSII reaction center, as revealed by modified SDS-PAGE with improved resolution of low-molecular-weight proteins. Plant Cell Physiol 29, 1233–1239.

Chapter 6 Isolation of Cytochrome b 6f Complex from Grana and Stroma Membranes from Spinach Chloroplasts Elz˙ bieta Romanowska Abstract The cytochrome b 6 f complex is located in the appressed granal membranes and nonappressed stroma thylakoids. The procedure presents isolation of the complex from both types of thylakoids by washing with NaBr, detergent treatment, ammonium sulfate fractionation, and sucrose gradient centrifugation. Optimal concentration of the detergent is lower for grana than for stroma vesicles. The cytochrome b 6f complex from stroma lamellae locates at a higher density in the sucrose gradient than the granal complex. Electrophoretic analyses indicate that both complexes are dimeric and contain four large subunits and at least three small subunits with masses below 4 kDa. Plastocyanin and 15 kDa protein are also identified in the complexes but in variable amounts. Key words: Cytochrome b 6f complex, Grana and stroma membranes (B3 and T3), Isolation ­procedure, spinach, Thylakoids

1. Introduction Cytochrome b 6 f complexes are located in the appressed granal membranes and in the nonappressed stroma thylakoids (1, 2).The cytochrome b 6 f complex occupies a central position in photosynthetic electron transport by linking PSII and PSI in linear electron flow and in PSI-dependent cyclic electron flow (3, 4). This transfer is associated with proton translocation across the thylakoid membrane. Thus cytochrome b 6 f complex contributes to formation of the proton gradient that is used for the synthesis of ATP. The b 6 f complex consists of eight subunits. Four subunits: cytochrome f, cytochrome b 6 , Rieske iron-sulfur protein and subunit IV have assigned functions there are four small ­hydrophobic

Robert Carpentier (ed.), Photosynthesis Research Protocols, Methods in Molecular Biology, vol. 684, DOI 10.1007/978-1-60761-925-3_6, © Springer Science+Business Media, LLC 2011

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subunits additionally, the products of the petG, L, M and N genes (5, 6). The role of these small subunits remains unknown. Possibly they are of structural importance. The cytochrome b 6 f complex from higher plants and from algae is isolated as an active dimer (7) although activities have been reported with a monomeric complex (8). The complex also mediates activation of the LHCII kinase (9). To understand the function of the complex from both types of thylakoids, a structural model is necessary, but a detailed structure of the whole complex is still missing (10, 11). Isolation procedures of cytochrome b 6 f complex from chloroplasts of higher plants have been published (1, 12–15) and also ones from fractionated grana and stroma membranes (16). Here, I present the first protocol involving a separation procedure. It should greatly facilitate the study of structure–function relationship involving b 6 f complexes from different membrane domain in respect to the functional heterogeneity of both types of membranes.

2. Materials 1. Source of the complex: spinach leaves. Spinach plants are grown in a growth chamber under a 10-h photoperiod and a day/night regime at 20°C. Fresh leaves are harvested from 6- to 10-week-old plants. 2. Isolate chloroplasts, and thereafter grana and stroma lamellae (see Notes 1 and 2). Vesicles derived from grana (B3) and stroma membranes (T3) are obtained after sonication of stacked thylakoids in an aqueous dextran-polyethylene glycol two-phase system followed by partitioning by a batch procedure in three steps as described previously (2, 16), and use the B3 and T3 thylakoids for cytochrome b 6 f complex isolation. 3. B3 and T3 thylakoid membranes obtained from 400 g leaves (use 100 g of leaves per isolation; this should give membranes corresponding to ca. 7 mg of chlorophyll) (see Note 4) are stored in liquid nitrogen until use. 4. Muslin. 5. Amicon-30 filters (Millipore). 6. Homogenizer for leaf samples up to 36 g or Waring blender. 7. Centrifugation and ultracentrifugation equipment. 8. Beakers (50 mL, 100 mL, 250 mL, 1 L), cylinders (50 mL, 100 mL), graduated tube with stopper (15–20 ml), crystallizers (500 and 900 mL). 9. Potter homogenizers 1 mL, 3 mL.

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10. Soft paintbrush. 11. 80% (v/v) Acetone. 12. Solid n-octyl-b-d-glucoside (OG), at –20°C. 13. 4 M Solid (NH4)2SO4, stock. 14. Solid Na-ascorbate. 15. Solid dithionite. 16. Solid ferricyanide. 17. 20% (w/v) Na-cholate, stock. 18. 5% Soybean lecithin dispersed by sonication, stock. 19. 10 mM Tris–HCl, pH 8.0. 20. 0.15 M NaCl. 21. Buffer 1: 10 mM Tris–HCl, pH 8.0, 2 M NaBr, 0.4 M sucrose. 22. Buffer 2: 20 mM Tricine–NaOH, pH 8.0, 3 mM KCl, 3 mM MgCl2, 0.2 M sucrose. 23. Buffer 3: 20 mM Tricine–NaOH, pH 8.0, 3 mM KCl, 3 mM MgCl2, 0.5% cholate, 0.4  M (NH4)2SO4 and 25  mM or 35 mM OG for B3 and T3 membranes, respectively. 24. Buffer 4: 30  mM Tris–succinate, pH 6.5, 0.2% cholate and 25 mM or 35 mM OG for B3 and T3 membranes, respectively. 25. Buffer 5: 30 mM Tris–succinate, pH 6.5, 0.2% cholate. 26. Buffer 6: 30 mM Tris–succinate, pH 6.5, 0.5% cholate, 0.1% soybean lecithin and 25 mM or 35 mM OG for B3 and T3 membranes, respectively. 27. Buffer 7: 30 mM Tris–succinate, pH 6.5, 1% Triton X-100. 28. Buffer 8: 20 mM MES–NaOH, pH 6.2, 12.5 nM cyt f in the form of cytochrome b 6 f complex, 10 mM cyt c, and 17 mM plastoquinol-9.

3. Methods 3.1. General Isolation Procedure

1. The following recipe describes the isolation of cytochrome b 6 f complexes from grana and stroma membranes. Fractionation of thylakoid membranes into grana (B3) and stroma (T3) lamellae is performed by sonication of stacked thylakoids in an  aqueous dextran-polyethylene glycol two-phase system ­followed by partitioning by a batch procedure in three steps described previously (2, 16). 2. Wash the cytochrome b 6 f complex derived from grana and stroma thylakoid membranes in high concentration of NaBr to remove peripheral proteins and coupling factors.

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3. Solubilize complex with n-octyl-b-d-glucoside (OG) and sodium cholate. 4. Purifiy cytochrome b 6 f complex by ammonium sulfate fractionation and sucrose density gradient centrifugation. 3.2. Purification of Cytochrome b6 f Complexes from Grana (B3) and Stroma (T3) Membranes

1. Carry out isolation procedure with both types of membranes in parallel (see Note 3). 2. Thaw and resuspend B3 and T3 thylakoid membranes stored in liquid nitrogen in 10 mM Tris–HCl, pH 8.0 (1 mL of buffer per 1 mL of membrane suspension) using paintbrush until suspension appears uniform, stir gently, and after 5 min add 0.15 M NaCl (the same volume as buffer) (see Note 4). 3. Centrifuge at 150,000 × g for 30 min. 4. Resuspend pellets in 20 mL of buffer 1. Stir on ice for 30 min and then add an equal volume of ice-cold water and centrifuge as above. Repeat this step and resuspend pellets in 20 mL of 10  mM Tris–HCl, pH 8.0 (see Note 5). Measure total chlorophyll content in suspension (see Note 6) and centrifuge as above. The pellets (B3W and T3W) contain membranes free from extrinsic proteins (see Note 5 and 7). 5. Resuspend B3W and T3W membranes in buffer 2 to obtain chlorophyll concentration of 3 mg/mL. 6. Dilute the mixtures to chlorophyll concentration of 1.5 mg/ mL, slowly adding an equal volume of buffer 3. Stir on ice for 30 min and centrifuge at 300,000 × g for 1 h (see Note 8). 7. Collect the supernatants (B3OG and T3OG) and slowly add solid ammonium sulfate to reach 45% saturation (2.16 g to 10 mL), making allowance for the 10% saturation ammonium sulfate initially present in the supernatant. 8. Stir the suspensions on ice for 30 min and remove precipitates by centrifugation at 16,000 × g for 10 min. 9. Discard pellets and add ammonium sulfate slowly to the supernatant solution to give 55% saturation (0.65 g to 10 ml), stir on ice for 30 min, and centrifuge at 16,000 × g for 10 min (see Note 9). 10. Discard the supernatant and dissolve the pellet in a minimal volume of buffer 4 and dialyze for 45 min against 1 L of buffer 5 with gentle stirring to remove excess salt. Concentrate solution to about 1 mL with Amicon-30 filters (see Note 10).

3.3. Sucrose Density Gradient

1. Prepare a continuous sucrose gradient of 14–30% in buffer 6 (see Note 11). 2. Load the concentrated sample to the top of gradient (see Note 10).

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Fig.  1. Sucrose density gradient (14–30%) centrifugation of cytochrome b 6f complex from grana (B3) and stroma (T3) membranes. Number of fractions is from the bottom to the top.

3. Centrifuge at 300,000 × g for 20 h. 4. The cytochrome b 6 f complex will form a week brown band at the bottom of gradient (see Note 12). 5. Collect 1 mL fractions by syringe from the top of gradient to individual tubes. 6. Estimate in all fractions the content of cytochrome f and b from reduced minus oxidized difference spectra (Fig. 1). 7. Measure the protein contents in all fractions. 8. Typical preparation of cytochrome b 6 f complex from grana (B3c) and stroma (T3c) membranes for cytochrome f is given in Table  1. Measure the plastoquinol-cytochrome c oxidoreductase activity in the presence of complex (see Subheading 3.4), freeze quickly in small aliquots at 77 K, and store in liquid nitrogen until use. 9. Use complexes for electrophoretic characterization. 3.4. Assays

1. The cytochrome content was determined from difference spectra of the cytochrome b 6 f complex. Oxidize aliquots of cytochrome fractions in buffer 7 with a grain of ferricyanide and divide between sample and reference cuvette (see Note 13). After recording the baseline add a grain of Na-ascorbate to the sample cuvette and take spectrum between 520 and

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Table 1 Purification steps of cytochrome b 6f complex from grana (B3) and stroma (T3) membranes cyt f Purification steps

Chl (mg)

nmol

nmol/mg protein

Thylakoids

127

166

0.5

T3 membranes

40

  38

0.3

B3 membranes

23

  34

0.4

T3 NaBr washed

33

  35

0.6

B3 NaBr washed

19

  31

0.8

OG extraction of T3

0.9

  25.7

1

OG extraction of B3

0.7

  19.2

2

Sucrose gradient band T3

n.d

   2.1

4.7 ± 0.5

Sucrose gradient band B3

n.d

   3

4.8 ± 0.6

580 nm. Calculate the concentration of cytochrome f from the peak height at 554 nm above a line drawn between the isosbestic points at 543.5 and 560  nm using an absorption coefficient of 18 mM−1 cm−1. For determination of cytochrome b content, add ascorbate to the reference cuvette and record second baseline, then add a grain of dithionite to the sample cuvette and take spectrum between 563 and 575 nm. Measure the height of the peak at 563 nm above a line drawn through the minima. The absorption coefficient is 20 mM−1 cm−1. 2. Protein is determined as described in (17, 18). 3. Measure plastoquinol-cytochrome c oxidoreductase activity in 1 mL of buffer 8. Reduction of cytochrome c was initiated by addition of 17 mM plastoquinol-9 and monitored by the absorbance change at 550 nm relative to that at 540 nm. The absorption coefficient is 17 mM−1cm−1. Activity for B3c and T3 c was 5.4 and 3.6 mmol cytochrome c reduced per nanomolar cytochrome f × h, respectively. 3.5. SDS-PAGE

1. Polypeptide patterns obtained after Laemmli (19) are shown in Figs. 2–4. 2. Better resolution of small polypeptides in the cytochrome b 6 f complex can be obtained by 12–22% gradient gels in the presence of urea or by PhastGel high density (Pharmacia) with 20% polyacrylamide (Figs. 2 and 3, respectively).

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Fig.  2. Urea/SDS polyacrylamide gel electrophoresis patterns. Gel concentration was 12–22%. Lane 1 and 6, protein standards; lane 2, cytochrome f; lane 3 and 7, cytochrome b 6f complex from thylakoids; lane 4, cytochrome b 6f complex from T3 membranes; lane 5, cytochrome b 6f complex from B3 membranes. Lanes 1–7, stained with Coomassie blue; lane 8 is identical as lane 7 but gel stained for heme.

Fig. 3. SDS-PAGE on PhastGel high density with 20% polyacrylamide. Lane 1, protein standards; lane 2 and 3, cytochrome b 6f complex from grana lamellae (B3); lane 4 and 5, cytochrome b 6f complex from stroma lamellae. The same amount of protein (1 mg) was loaded in each lane. Silver staining.

3. Better separation of the Rieske and cytochrome b 6 proteins is obtained by Urea/SDS-PAGE gradient gel (12–22%) as shown in Fig. 2. Polypeptide patterns obtained in this system demonstrate that mobilities of the Rieske protein and cytochrome b 6 are reversed relative to those in standard PAGE (Fig. 4). Gel was stained for proteins in Coomassie Brilliant Blue or for heme-associated peroxidase activity by the method described in (20).

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Fig. 4. SDS-PAGE of purified cytochrome b 6f complex. Electrophoresis was performed as described by Laemmli (19) on a linear gradient gel from 12–22% polyacrylamide. Lane 1 and 7, thylakoids; lane 2, B3W membranes; lane 3, cytochrome complex from B3 membranes (B3c); lane 4, protein standards; lane 5, cytochrome complex from T3 membranes (T3c);lane 6, T3W membranes. Coomassie staining.

4. Notes 1. Collect leaves after dark period to avoid starch grains. High content of starch granules in chloroplasts after the light period may lead to their injury. 2. Before starting isolation procedure of cytochrome b 6 f complex from B3 and T3 membranes isolate the complex from whole unfractionate thylakoid membranes for better optymalization of procedure. The final preparation has a cytochrome f concentration of about 15–20  mM and protein kinase activity of 110 pmol 32P/min × mg protein. Flow-chart diagram (Fig.  5) can help in cytochrome b 6 f complex purification. 3. All operations are performed in a cold room (2–4°C). 4. It is preferable to carry out whole isolation procedure of cytochrome b 6 f complex in 1 day (takes 12 h). It is possible only if B3 and T3 thylakoids are isolated earlier. Isolated B3 and T3 thylakoids should be stored until use in liquid nitrogen at a final chlorophyll concentration of 3 mg/mL. For isolation of cytochrome b 6 f complex from B3 and T3 membranes, one should start with thylakoids containing at least 30 mg of Chl.

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Fig.  5. Flow-chart diagram of purification steps of cytochrome bf complex from whole unfractionates thylakoid from spinach chloroplasts.

Such amount of membranes is obtained from about 400 g of leaves (four isolations of B3 and T3 membranes from 3 × 35 g of leaves). Purification of the cyt b 6 f complex from B3 and T3 membranes should be done in parallel.

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5. For efficient removal of extrinsic proteins repeat the NaBr washing at least two times. 6. Add 5 mL of thylakoid suspension to 1,495 mL of 80% acetone into 2 mL centrifuge tube and vortex and centrifuge for 2 min at 2,500 × g. Transfer clear supernatant to cuvette and measure absorbance (A) at 645 and 663 nm with 80% acetone as reference. Calculate the concentration of chlorophyll using the equations: Chl a + b (mg/mL) = 20.2 A645 + 8.02 A663, Chl b (mg/mL) = 22.9 A645 – 4.68 A663 (21). 7. B3W and T3W membranes may be stored at 4°C overnight. 8. Detergent concentration used for membrane extraction must be optimized in pilot experiment. For spinach use octylglucoside (OG) concentration varying from 20 to 50  mM. Use concentration of OG that solubilizes more than 70% of cytochrome f and less than 5% of chlorophyll. Use B3 and T3 membranes from one preparation (from 35 g of leaves) and resuspend the obtained B3W and T3W membranes in buffer 3. Stir on ice for 30 min and centrifuge at 300,000 × g for 1 h or at 132,000 × g for 15  min (in a Beckman air fuge with 200  mL tubes). Determine chlorophyll and cytochrome f concentration in supernatants. Also use cholate (improves extraction by suppressing nonspecific membrane solubilization and prevents aggregation of cytochromes) in the medium. High salt (0.4  M ammonium sulfate) added together with detergent increases efficiency of extraction. 9. The cytochrome complex from T3 membranes at 55% saturation of ammonium sulfate may form aggregates that float instead of precipitating. They can be collected by filtering through cotton wool. 10. Use Amicon-30 filter for concentration of complexes. Centrifuge for 20 min at 5,000 × g. 11. Gradient former should be used to obtain satisfactory gradient linearity. Sample volume should not exceed 5% of the gradient. Use 20-ml tubes for gradient formation. Soybean lecithin is included in the gradient for stabilization of the cytochrome b 6 f complex. Concentration of OG in the gradient is kept the same as during solubilization. Triton X-100 is not included in the gradient because it inhibits the activity of the cytochrome b 6 f complex. 12. Cytochrome b 6 f complex from stroma (T3c) lamellae locates at a higher density than the grana (B3c) complex (Fig. 1). Cytochrome b 6 f complex from grana membranes sometimes is separated into two bands in the gradient. 13. Volume in cuvette is 1,400 mL; use 50 mL of B3c or T3c. Add dithionite and wait 2–3 min to reach full cytochrome b reduction (until no further increase of absorbance is detectable).

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14. For protection of proteins from proteolysis add protease inhibitors: phenylmethylsulfonylfluoride (PMSF), amino-ncaproic acid (EACA), and benzamidine to media at final concentrations of 0.2  mM, 5  mM, and 1  mM, respectively. Prepare stock solutions and add just before use.

Acknowledgments I thank Per-Åke Albertsson for useful discussions and financial support. References 1. Anderson, J.M. (1982) Distribution of the cytochromes of spinach chloroplasts between the appressed membranes of grana stacks and stroma exposed thylakoid regions. FEBS Lett. 138, 62–66. 2. Albertsson, P.-A., Andreasson, E., Svensson, P., and Yu, S.-G. (1991) Localization of cytochrome f in the thylakoid membrane: evidence for multiple domains. Biochim. Biophys. Acta 1098, 90–94. 3. Dietrrich, J. and Kühlbrandt, W. (1999) Purification and two-dimentional crystallization of highly active cytochrome b6 f complex from spinach. FEBS Lett. 463, 97–102. 4. Joliot, P. and Joliot, A. (1992) Electron transfer between Photosystem II and the cytochrome bf complex; mechanistic and structural implications. Biochim. Biophys. Acta 1102, 53–61. 5. Pierre, Y., Breyton, C., Kramer, D., and Popot, J.L. (1995) Purification and characterization of the Cyt b6 f complex from Chlamydomonas reinhardtii. J. Biol. Chem. 270, 29342–29349. 6. Whitelegge, J.P., Zhang, H., Taylor, R., and Cramer, W.A. (2002) Full subunit coverage liquit chromatography electrospray-ionization mass spectrometry (LCMS+) of an oligomeric membrane protein complex: the cytochrome b6 f complex from spinach and the cyanobacterium, M. laminosus. Mol. Cell. Proteomics 1, 816–827. 7. Huang, D., Everly, R.M., Cheng, R.H., Heymann, J.B., Schägger, H., Baker, T.S., and Cramer, W.A. (1994) Charakterization of the Cyt b6 f complex as a structural and functional dimer. Biochemistry 33, 4401–4409. 8. Chain, R.K. and Malkin, R. (1995) Functional activities of monomeric and dimeric forms of the chloroplast Cyt b6 f complex. Photosynth. Res. 46, 419–426.

9. Vener, A., van Kan, P.J., Rich, P.R., Ohad, I.I., and Andersson, B. (1997) Plastochinol at the quinol oxidation site of reduced cytochrome bf mediates signal transduction between light and protein phosphorylation: thylakoid protein kinase deactivation by a single-turnover flash. Proc. Natl. Acad. Sci. U S A 94, 1585–1590. 10. Carrel, C.J., Schlarb, B.G., and Bendall, D.S. (1999) Structure of the soluble domain of cytochrome f fro cyanobacterium Phormidium laminosum. Biochemistry 38, 9590–9599. 11. Zhang, H., Kurisu, G., Smith, J.L., and Cramer, W.A. (2003) A defined proteindetergent-lipid complex for crystallization of integral membrane proteins: the cytochrome b6 f complex of oxygenic photosynthesis. Proc. Natl. Acad. Sci. U S A 100, 5160–5163. 12. Hurt, E. and Hauska, G. (1981) A cytochrome fb6 complex of five polypeptides with plastoquinol-plastocyanin-oxidoreductase activity from spinach chloroplasts. Eur. J. Biochem. 117, 591–599. 13. Hurt, E. and Hauska, G. (1982) Identification of the polypeptides in the cytochrome b6 f complex from spinach chloroplasts with redox-center caring subunits. J. Bioenerg. Biomembr. 14, 1–22. 14. Clark, R.D. and Hind, G. (1983) Isolation of the five-peptide cytochrome b-f complex from spinach chloroplasts. J. Biol. Chem. 258, 10348–10354. 15. Black, M.T., Widger, W.R., and Cramer, W.A. (1987) Large-scale purification of active cytochrome b6 f complex from spinach chloroplasts. Arch. Biochem. Biophys. 252, 655–661. 16. Romanowska E. and Albertsson P.-A. (1994) Isolation and characterization of the cytochrome bf complex from whole thylakoids, grana and stroma lamellae vesicles from spinach chlorolasts. Plant Cell Physiol. 35, 557–568.

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17. Bradford, M. (1976) A rapid and sensitive method for the quantification of microgram quantities of protein utilizing the principle of protein-dye binding. Anal. Biochem. 72, 248–254. 18. Bensadoun, A. and Wenstein, D. (1976) Assay of proteins in the presence of interfering materials. Anal. Biochem. 70, 241–250. 19. Laemmlli, U.K. (1970) Cleavage of structural proteins during the assembly of the head of bacteriophage T4. Nature 227, 680–685.

20. Thomas, J.O., Ryan, D., and Levin, W. (1976) An improved staining procedure for the detection of the peroxidase activity of cytochrome P-450 on sodium dodecyl sulfate polyacryamide gels. Anal. Biochem. 75, 168–176. 21. Hipkins, M.F. and Baker, N.R. (1986) Spectroscopy, in Photosynthesis energy transduction, a practical approach (Hipkins, M.F. and Baker, N.R., eds.), IRL Press, England, pp. 51–101.

Chapter 7 Purification and Crystallization of the Cyanobacterial Cytochrome b6f Complex Danas Baniulis, Huamin Zhang, Taisiya Zakharova, S. Saif Hasan, and William A. Cramer Abstract The cytochrome b6 f complex from the filamentous cyanobacteria (Mastigocladus laminosus, Nostoc sp. PCC 7120) and spinach chloroplasts has been purified as a homo-dimer. Electrospray ionization mass spectroscopy showed the monomer to contain eight and nine subunits, respectively, and dimeric masses of 217.1, 214.2, and 286.5 kDa for M. laminosus, Nostoc, and the complex from spinach. The core subunits containing or interacting with redox-active prosthetic groups are petA (cytochrome f ), B (cytochrome b6), C (Rieske iron-sulfur protein), D (subunit IV), with protein molecular weights of 31.8–32.3, 24.7–24.9, 18.9–19.3, and 17.3–17.5 kDa, and four small 3.2–4.2 kDa polypeptides petG, L, M, and N. A ninth polypeptide, the 35 kDa petH (FNR) polypeptide in the spinach complex, was identified as ferredoxin:NADP reductase (FNR), which binds to the complex tightly at a stoichiometry of approx 0.8/cytf. The spinach complex contains diaphorase activity diagnostic of FNR and is active in facilitating ferredoxin-dependent electron transfer from NADPH to the cytochrome b6 f complex. The purified cytochrome b6 f complex contains stoichiometrically bound chlorophyll a and b-carotene at a ratio of approximately one molecule of each per cytochrome f. It also contains bound lipid and detergent, indicating seven lipid-binding sites per monomer. Highly purified complexes are active for approximately 1 week after isolation, transferring 200–300 electrons/cytf s. The M. laminosus complex was shown to be subject to proteolysis and associated loss of activity if incubated for more than 1 week at room temperature. The Nostoc complex is more resistant to proteolysis. Addition of pure synthetic lipid to the cyanobacterial complex, which is mostly delipidated by the isolation procedure, allows rapid formation of large (³0.2 mm) crystals suitable for X-ray diffraction analysis and structure determination. The crystals made from the cyanobacterial complex diffract to 3.0 Å with R values of 0.222 and 0.230 for M. laminosus and Nostoc, respectively. It has not yet been possible to obtain crystals of the b6 f complex from any plant source, specifically spinach or pea, perhaps because of incomplete binding of FNR or other peripheral polypeptides. Well diffracting crystals have been obtained from the green alga, Chlamydomonas reinhardtii (ref. 10). Key words: Cytochrome b6 f complex, Electron transport, Photosynthetic, Heme cn, Plastoquinone

Abbreviations cn cyt

Heme covalently bound on the electrochemically negative side of the complex Cytochrome

Robert Carpentier (ed.), Photosynthesis Research Protocols, Methods in Molecular Biology, vol. 684, DOI 10.1007/978-1-60761-925-3_7, © Springer Science+Business Media, LLC 2011

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DMSO dimethyl sulfoxide DOPC 1,2-dioleoyl-sn-glycero-3-phosphocholine EDTA Ethylenedinitrilo tetraacetic acid FNR Ferredoxin:NADP+ oxidoreductase MGDG Monogalactosyl-diacylglycerol PG Phosphatidylglycerol Q Plastoquinone bound on the electrochemically negative side of the b6 f complex OG n-octyl-b-d-glucoside PEG 400 Polyethylene glycol MW 400 PEGMME 550 Polyethylene glycol monomethyl ether MW 550 UDM n-undecyl-b-d-maltoside

1. Introduction The hetero-oligomeric cytochrome b6 f complex (plastoquinol: plastocyanin/ cytochrome c6 oxidoreductase) functions in oxygenic photosynthesis to mediate electron transfer from photo­ system II (PSII) to PSI in the linear electron transfer pathway, from H2O to NADPH or around PSI in a cyclic pathway (1–6), generating a trans-membrane electrochemical proton gradient that is used for the synthesis of adenosine 5¢-triphosphate (ATP). The b6 f complex also mediates trans-membrane activation of the stromal-side light-harvesting complex protein kinase (7). The dimeric b6 f complex is composed of eight (8–11) or nine (12) polypeptide subunits. The complex contains three traditional hemes per monomer (two linked to the cytochrome b polypeptide and one covalently attached to cytochrome f  ), one novel covalently bound heme cn in cytochrome b6 (9, 10, 13, 14), and two nonheme Fe in the prosthetic group of the Rieske iron-sulfur protein (ISP). The functions of the chlorophyll a (15–17) and b-carotene (17) whose presence, like that of heme cn, is unique to b6  f compared to bc1 complexes, are presently not known, except for the ability of the carotene to quench the Chl triplet state (18). The core structure and linear electron transport function of the b6 f complex is similar to that of the cytochrome bc1 complex of mitochondria and photosynthetic bacteria, for which crystal structures have been obtained from eukaryotic bovine (19, 20), avian (21), yeast (22), and photosynthetic bacterial (23) sources. In cyanobacteria, the b6 f complex is shared by both photosynthetic and respiratory electron transfer chains (24). The 24-kDa cytochrome b6 subunit, which has four transmembrane a-helices and contains two b-type hemes, together with the 17-kDa subunit IV that has three transmembrane a-helices are functionally and structurally analogous to the N-terminal and C-terminal halves of the cytochrome b subunit in the bc1 complex (25). The 19-kDa Rieske ISP, which has an N-terminal single transmembrane

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a-helix domain and a C-terminal 140 residue soluble extrinsic domain including a linker region connecting these two domains, is structurally and functionally related to the ISP subunit in the bc1 complex (26). The 31-kDa c-type cytochrome f subunit is functionally but not structurally related to cytochrome c1 of the bc1 complex (27, 28). Its b-strand secondary structure is virtually unique and its distal heme ligation by the N-terminal a-amino terminus of cytochrome f is a unique structural signature, conserved from the cyanobacteria (29) through the green algae (30, 31) to plants (27, 28).

2. Materials 1. Sources of the complex: Spinach, thermophilic filamentous cyanobacterium Mastigocladus laminosus and filamentous cyanobacterium Nostoc sp. PCC 7120. Spinach was purchased from a market. Fresh leaves were washed and stored at 4°C. The thermophilic cyanobacterium M. laminosus was grown in 10 L carboys at 55°C to late logarithmic phase, and harvested utilizing a Millipore Pellicon filtration system. Nostoc sp. PCC 7120 was grown in BG-11 media (32) at 30°C under a light intensity of 100–150  mE/m2 s obtained from fluorescent lamps. Cells were harvested at early exponential growth phase (OD730 » 1–2). 2. Cell breakage solution for cyanobacteria cells: 25 mM Hepes– KOH, pH 7.7, 0.4 M sucrose, 10 mM MgCl2, 10 mM CaCl2, 2 mM benzamidine, 2 mM e-amino-caproic acid (see Note 1). 3. Homogenizing solution for spinach leaves (TNS): 50  mM Tris–HCl, pH 7.7, 0.1 M NaCl, 0.2 M sucrose. 4. Propyl-agarose. 5. Solid n-undecyl-b-d-maltoside. Stored at −20°C. 6. Solid n-octyl-b-d-glucoside. Stored at −20°C. 7. 1,2-dioleoyl-sn-glycero-3-phosphocholine (DOPC). Stored at −20°C. 8. Buffer 1: 10 mM Tricine/NaOH, pH 8.0, 2 M NaBr. 9. 10 mM Tricine/NaOH, pH 8.0. 10. Buffer 2: 50  mM Tris–HCl, pH 7.5, 50 mM NaCl, 1  mM EDTA, 1 mM UDM. 11. Buffer 3: 50  mM Tris–HCl, pH 7.5, 50  mM NaCl, 1  mM EDTA, 1 mM UDM, 20% DMSO. 12. Buffer 4: 30  mM Tris–HCl, pH 7.5, 50  mM NaCl, 1  mM UDM.

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13. Buffer 5: 100 mM Tris–HCl, pH 7.5, 100 mM MgCl2 and 30% PEG 400. 14. Buffer 6: 100 mM Tris–HCl, pH 7.5, 200 mM MgCl2, 5 mM CdCl2, 1 mM DOPC, 3 mM UDM and 12% PEGMME 550. 15. TNE buffer: 30 mM Tris–HCl, pH 7.5, 50 mM NaCl, 1 mM EDTA. 16. TMKNE buffer: TNE buffer supplemented with 5  mM MgCl2, 5 mM KCl. 17. Sample buffer for SDS-PAGE: 50  mM Tris–HCl, pH 8.6, 8 M urea, 4% SDS, 10% glycerol, and 4% mercaptoethanol.

3. Methods 3.1. Thylakoid Membrane Preparation

1. Homogenize 400 g of leaves in a 4-L blender for 2 min in 1.5 L TNS.

3.1.1. Spinach

2. Filter the slurry through a wire sieve and then through four layers of cheesecloth. 3. Centrifuge the eluate at 13,000 g for15 min. 4. Suspend pellet in “Material 9;” centrifuge at 13,000 g for 20 min. 5. The pellet was resuspended in 1.2 L of buffer 1. 6. The suspension was stirred on ice for 20 min, an equal volume of cold water added, and then centrifuged at 13,000 g for 15 min. 7. Steps 5 and 6 were repeated, and the thylakoid membranes were then washed with 2.4 L of buffer 2 without UDM.

3.1.2. The Cyanobacterium, M. laminosus

1. Harvest 40 g (wet weight) M. laminosus cells from 40 L culture, broken by shearing twice in a French press at 20,000 psi in 200 mL of cell breakage solution. 2. Sediment broken cell fragments by centrifugation at 6,000 g for 10  min and isolate thylakoid membranes by centrifugation at 250,000 g for 45 min. 3. Resuspend thylakoid membranes and centrifuge in the following solution: 200  mL of 10  mM Tricine/NaOH, pH 8.0; resuspend in 100 mL of buffer 1. 4. Stir the suspension on ice for 20 min. 5. Add an equal volume of cold water and centrifuge again at 300,000 g for 45 min. 6. Repeat steps 3–5. 7. Resuspend the pellet in 200 mL TNE buffer and centrifuge at 300,000 g for 45 min.

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1. 25 g (wet weight) of Nostoc sp. PCC 7120 cells were suspended in 200 mL of cell breakage solution and broken by shearing three times in a French press at 20,000 psi. 2. Thylakoid membranes were isolated and washed as in the procedure for the M. laminosus (Subheading 3.1.2, steps 2–6).

3.2. Purification of the Cytochrome b6f Complex 3.2.1. Selective Solubilization of the Complex from Spinach and M. laminosus (see Note 2)

1. Resuspend thylakoid membranes in TMKNE buffer with protease inhibitors (2 mM benzamidine and 2 mM e-aminocaproic acid) supplemented with 300 mM sucrose to obtain a chlorophyll concentration of 2 mg/mL. 2. Slowly add an equal volume of TMKNE containing 60 mM OG, 0.1% cholate to the stirred membrane suspension. 3. Stir the mixture at 4ºC for 30–45  min and centrifuge at 200,000 g for 45 min. 4. Collect the supernatant, add solid ammonium sulfate to 35% saturation, stir at 4ºC for 15  min, and then centrifuge at 200,000 g for 30 min. 5. Collect the supernatant for further purification (see Note 3).

3.2.2. Solubilization of the Complex from Nostoc sp. PCC 7120

1. Resuspend thylakoid membranes in TNE buffer with protease inhibitors (2  mM benzamidine and 2  mM e-amino-caproic acid) supplemented with 300 mM sucrose to obtain a chlorophyll concentration of 2.2 mg/mL. 2. Stir the membrane suspension on ice and slowly add 0.1 volume of UDM solution to a final concentration of 10  mM UDM and 2 mg/ml chlorophyll (see Note 4). 3. Stir the mixture for 30 min on ice and centrifuge at 200,000 g for 45 min. 4. Repeat steps 4,5 from 3.2; Collect the supernatant for further purification.

3.2.3. Propyl-Agarose Chromatography

1. Pack the propyl-agarose resin into a 2 × 15 cm column, wash sequentially with distilled water, TNE buffer, and equilibrate with 2–3 column volumes of TNE containing 35% (w/v) saturated ammonium sulfate and 1 mM UDM All steps at 4ºC. 2. Load the sample on the column by gravity. 3. Wash the column thoroughly with equilibration buffer until the eluant is colorless. 4. Elute the cytochrome b6 f complex with TNE containing 20% (w/v) saturated ammonium sulfate, 1 mM UDM. The earlier fractions are greenish, and those eluting later greenish-blue (see Note 5). 5. The cytochrome f concentration was determined by the difference spectrum of ascorbate minus ferricyanide, using an extinction coefficient of 25 mM/cm at 554 nm relative to a baseline drawn between the spectral troughs at 538 and 568 nm (33).

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6. Fractions with a cytochrome f concentration greater than 1  mM were pooled and concentrated with a Millipore Centriprep 10. 3.2.4. Sucrose Gradient Ultra-Centrifugation

1. Prepare a continuous sucrose gradient of 10–32% in TNE, 1 mM UDM. 2. Load the concentrated sample from the propyl-agarose column to the top of the gradient at a ratio of approx 1:10 (v/v). 3. Perform centrifugation at 160,000 g for 16 h. 4. Collect the brown band in the middle of the gradient. 5. A second sucrose gradient is needed if the bands are not well separated in the first run. 6. After purification, store the cytochrome b6 f complex on ice in buffer 2.

3.3. SDS-PAGE

1. The sample was solubilized in the sample buffer and then heated at 90°C for 2  min. The 15% SDS-gel was prepared according to Laemmli.(34). 2. The purified cytochrome complexes from spinach chloroplasts and M. laminosus show different patterns on SDS-PAGE. The four large subunits were well separated in the M. laminosus complex (Fig. 1, lane 1). However, in the spinach complex, cytochrome b6 and the Rieske ISP co-migrated into one thick band (Fig. 1, lane 2). A high-molecular-weight band with an Mr value greater than that of cytochrome f was detected in the spinach complex and was characterized as bound FNR (12).

3.4. Visible Difference Absorbance Spectra

1. The cytochrome content was determined from difference spectra of the cytochrome complex. 2. The complex was diluted in buffer 4 to 0.1 mg/mL.

Fig.  1. SDS-PAGE of the cytochrome b6f complexes from M. laminosus (lane 1) and spinach chloroplasts (lane 2 ). The four large (>17 kDa) subunits are labeled.

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Fig. 2. Absolute visible absorbance spectra of the cytochrome b6f complex from spinach chloroplasts (solid line) and M. laminosus (dashed line). The complexes (equivalent to 1 mM cyt f ) were reduced with Na-ascorbate. Peak 1, Soret bands; 2, FNR; 3, FNR and carotenoid; 4, cytochrome f a-band; 5, chlorophyll a Q band. Inset panel shows the difference spectra of cyt f and cyt b6, respectively, as ascorbate minus ferricyanide and dithionite minus ascorbate difference spectra.

3. Visible spectra were measured on a Cary 3 UV-visible ­spectrophotometer with a measuring beam half-band width of 2.0 nm. 4. The absorbance spectrum of the complex was measured as a difference spectrum of ascorbate minus buffer (which ­corrected for the system response of the instrument; Fig. 2, outset panel). 5. Cytochrome f and b6 spectra were measured, respectively, as chemical difference spectra, ascorbate minus ferricyanide and dithionite minus ascorbate (Fig. 2, inset panel). The extinction coefficients that were used are e554 = 25 mM/cm (33) and e563 = 21 mM/cm, respectively. 3.5. Electron Transfer Activity

1. Electron transfer activity was measured according to the methods described (12, 35). The assay mixture contained 5 mM plastocyanin, and 5 nM cytochrome complex in buffer 4. 2. Reduction of plastocyanin was initiated by addition of 25 mM decyl-plastoquinol and monitored by the absorbance change at 600 nm relative to that at 500 nm. 3. The electron transfer activity of the isolated cytochrome complex from M. laminosus is typically 200–300 electrons/cytf s.

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Fig. 3. Electron transfer activity and inhibitor effects. The uninhibited electron transfer activity of the isolated cytochrome complex from M. laminosus is typically 200–300 electrons/cyt f/s, the latter rate obtained at a temperature of 25–27°. In the presence of 2 mM stigmatellin, the activity was inhibited approx 60%, whereas inhibition by 2 mM tridecyl-stigmatellin or 2 mM DBMIB is >90%.

4. In the presence of 2 mM stigmatellin, the activity was inhibited approx 60%, whereas inhibition by 2 mM tridecyl-stigmatellin or 2 mM DBMIB is >90% (Fig. 3). 3.6. Mass Spectrometry

The subunit composition of the cytochrome complex in M.laminosus (36) and Nostoc (8) was analyzed by mass spectrometry. The active dimeric forms of the cytochrome complex from the cyanobacterium (M. laminosus, Nostoc sp. PCC 7120) and spinach chloroplasts contain eight and nine subunits, respectively, which are petA (cytochrome f ), petB (cytochrome b6), petC (Rieske ISP), petD (subunit IV), and the small 3.2–4.2 kDa polypeptides petG, L, M, and N, as listed in Table 1. The ninth subunit in the spinach complex was identified as ferredoxin:NADP+ reductase (FNR).

3.7. Lipid Analysis

Total lipids were extracted from the isolated complex according to the procedure described by Sato and Murata (37). 1. Thin-layer chromatography was performed on a precoated 20 × 20 cm silica gel plate (Merck). 2. 50 mL of extracted lipid solution was applied to the plate. 3. The plate was developed in CHCl3/CH3OH/NH4OH (28%) = 13/7/1.

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Table 1 Masses and subunits of the cytochrome b6 f complex Measured mass (Da) Protein

Spinacea oleracea

Mastigocladus laminosus

Nostoc sp. PCC 7120

–

–

PetH (FNR)

35,314

PetA (Cyt f  )

31,935

32,270

31,769

PetB (Cyt b6)

24,887

24,710

24,757

PetC (Rieske iron-sulfur protein)

18,936

19,295

19,064 (19,107)a

PetD (subunit IV)

17,312

17,528

17,404

PetG

4,198

4,057

4,023

PetM

3,972

3,841

3,574

PetL

3,478

3,530

3,253

PetN

3,197

3,304

3,262

143,227

108,535

Total mass of monomer

107,106 (107,149)a

acetylated ISP

a

4. The lipids were visualized by spraying 50% H2SO4 in methanol, followed by heating at 110°C until the spots are visible. 5. Standard lipids were run to determine the identity of the sample. 6. The lipid content was determined by densitometric analysis of the scanned image using NIH Image 1.54. The major bound lipids found in the less-pure cytochrome complex are MGDG and PG, which are the most abundant lipids in cyanobacterial thylakoid membranes (37). However, in the extensively purified complex, there was less than one bound MGDG lipid per cytochrome f. 3.8. Pigment Analysis

The pigment content of the purified complex was determined as described by Zhang et al. (17). The isolated cytochrome complex contains stoichiometrically bound chlorophyll a and b-carotene at a ratio of approximately one per monomer.

3.9. Crystallization of the Cytochrome b6f Complex

1. Add synthetic DOPC to the complex at a final concentration of 0.1% (w/v) by diluting a 20% (w/v) stock solution stored in buffer 3, so that DOPC: cyt f, mol:mol, =10:1. The cytochrome complex was concentrated with a Millipore Centricon 100 to 20 mg/mL.

3.9.1. M. laminosus

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2. Crystallization droplets were created by the hanging drop vapor diffusion technique at 18°C. Two microliters of concentrated protein (20 mg/mL) in buffer 4 was mixed with an equal volume of buffer 5. 3. Hexagonal crystals with space group P61 grew overnight to 0.3 × 0.15 × 0.1 mm and reached a size of 0.5 × 0.25 × 0.2 mm after 3–4 days. 4. For data collection, crystals were frozen in 30% PEG 400 with 5% 2,3-butanediol. 3.9.2. Nostoc sp. PCC 7120

1. Buffer was exchanged to 100  mM Tris–HCl, pH 7.5, containing 1 mM DOPC lipid, and 3 mM UDM detergent, and the protein was concentrated to 135  mM with a Centricon 100 concentrator. 2. 4 mL droplets for crystallization were set up by the hanging drop vapor diffusion method by diluting the protein solution 1:1 with reservoir solution containing buffer 6. 3. Bipyramidal crystals with the hexagonal space group P6122 were grown at 6°C for 2 days (Fig. 4). 4. For data collection, crystals were transferred to cryo-protectant solutions containing the same components as the reservoir solution except that the PEGMME 550 concentration was 14% and the glycerol concentration was increased in steps of 5%, to 25%, and flash frozen.

Fig. 4. Crystals of the cytochrome b6f complex from Nostoc sp. PCC 7120. Inset panel shows the reddish brown bipyramidal crystals with space group P61 22 that diffracted to 3.0 Å. (bar size, 100 mm). Baniulis et al. (8), reprinted with permission from the Journal of Biological Chemistry.

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4. Notes 1. All the stock solutions should be stored at 4°C in the presence of protease inhibitors: 0.25 mM PMSF, 2 mM benzamidine, 2 mM e-amino-caproic acid. 2. In this chapter we have described the methods for purification of the cytochrome b6 f complex from spinach chloroplasts and a cyanobacterium. For purification of the complex from the green alga, Chlamydomonas reinhardtii, consult Pierre et al. (38). 3. The purified cytochrome complex is subject to proteolysis. Even in the presence of protease inhibitors, the Rieske ISP, cytochrome b6, and subunit IV could be partially proteolysed if left at room temperature for a week or so. After 2 weeks at room temperature, these three subunits were completely cleaved. The complex is stable on ice over several weeks, and it can be frozen in liquid nitrogen and stored at −80°C for a longer period of time without significant loss of activity. 4. Better extraction efficiency could be obtained by increasing concentrations of the UDM detergent to 15–20  mM, although this results in a significant reduction of the protein purity ultimately obtained. 5. After elution, the propyl-agarose column retained a blue color because of bound phycobilin proteins. The column can be regenerated by washing with one volume of distilled water, followed by seven sequential washes with one volume of 25, 50, and 95% ethanol, two volumes of n-butanol, and one volume of 95, 50, and 25% ethanol. The clean resin is kept in 20% ethanol.

Acknowledgments These studies were supported by NIH GM-32383 (WAC). Acknowledgments to the earlier phase of crystallization and structure studies on the cytochrome b6  f complex, which focused solely on the M. laminous b6 f complex, appear in the first description of these Methods (39). For the subsequent studies, we thank E. Yamashita for collaboration on the analysis of the b6  f crystals. References 1. Baniulis, D., Yamashita, E., Zhang, H., Hasan, S.S., and Cramer, W.A. (2008) Structurefunction of the cytochrome b6f complex. Photochem Photobiol 84, 1349-58.

2. Cramer, W.A., Baniulis, D., Yamashita, E., Zhang, H., Zatsman, A.I., and Hendrich, M.P. (2008) Structure, spectroscopy, and function of the cytochrome b6f complex: heme cn and

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3.

4. 5. 6. 7.

8.

9.

10. 11.

12.

13.

14.

15.

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cytochrome b6 f complex as a structural and functional dimer. Biochemistry 33, 4401–9. Pierre, Y., Breyton, C., Lemoine, Y., Robert, B., Vernotte, C., and Popot , J.-L. (1997) On the presence and role of a molecule of chlorophyll a in the cytochrome b6 f complex. J Biol Chem 272, 21901–8. Zhang, H., Huang, D., and Cramer, W.A. (1999) Stoichiometrically bound beta-carotene in the cytochrome b6 f complex of oxygenic photosynthesis protects against oxygen damage. J Biol Chem 274, 1581–7. Kim, H., Dashdorj, N., Zhang, H., Yan, J., Cramer, W.A., and Savikhin, S. (2005) An anomalous distance dependence of intra-protein chlorophyll-carotenoid triplet energy transfer. Biophys. J. Letters 89, L28–30 Huang,. L.S., Cobessi, D., Tung, E.Y., and Berry, E.A. (2005) Binding of the respiratory chain inhibitor antimycin to the mitochondrial bc1 complex: a new crystal structure reveals an altered intramolecular hydrogenbonding pattern. J Mol Biol 351, 573–97. Xia, D., Yu, C.-A., Kim, H., et  al. (1997) Crystal structure of the cytochrome bc1 complex from bovine heart mitochondria. Science 277, 60–6. Zhang, Z., Huang, L., Shulmeister V.M, et al. (1998) Electron transfer by domain movement in cytochrome bc1. Nature 392, 677–84. Solmaz, S.R. and Hunte, C. (2008) Structure of complex III with bound cytochrome c in reduced state and definition of a minimal core interface for electron transfer. J Biol Chem 283, 17542–9. Esser, L., Elberry, M., Zhou, F., Yu, C.A., Yu, L., and Xia, D. (2008) Inhibitor-complexed structures of the cytochrome bc1 from the photosynthetic bacterium, Rhodobacter spha­ eroides. J Biol Chem 283, 2846–57. Kallas, T. (1994) The cytochrome b6 f complex, in The molecular biology of cyanobacteria (Bryant, D.A. ed.), Dordrecht: Kluwer, pp. 259–317. Widger, W.R., Cramer, W.A., Herrmann, R.G., and Trebst, A. (1984) Sequence homology and structural similarity between the b cytochrome of mitochondrial complex III and the chloroplast b6 f complex: position of the cytochrome b hemes in the membrane. Proc Natl Acad Sci USA 81, 674–8. Carrell, C.J., Zhang, H., Cramer, W.A., and Smith, J.L. (1997) Biological identity and diversity in photosynthesis and respiration: structure of the lumen-side domain of the chloroplast Rieske protein. Structure 5, 1613–25.

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Chapter 8 Purification of Plastocyanin and Cytochrome c6 from Plants, Green Algae, and Cyanobacteria José A. Navarro, Manuel Hervás, and Miguel A. De la Rosa Abstract Plastocyanin and cytochrome c6 are widely distributed over the oxygen-evolving photosynthetic organisms. The two proteins are functionally equivalent, but strongly differ in their global electrostatic charge. In fact, they are acidic in eukaryotes, but either neutral or basic in cyanobacteria. Such a difference in their electrostatic features is a critical factor in designing the purification procedure, which must thus be modified and adapted accordingly. This chapter reports the methods for producing (including cell cultures), isolating, and purifying plastocyanin and cytochrome c6 – which greatly differ in their isoelectric point – from a number of eukaryotic and prokaryotic organisms. Key words: Cyanobacteria, Cytochrome c6, Escherichia coli, Green algae, Metalloproteins, Monoraphidium, Nostoc, Plastocyanin, Protein purification, Recombinant protein, Spinach, Synechocystis

1. Introduction Plastocyanin and cytochrome c6 are two soluble metalloproteins located inside the thylakoidal lumen. They have different tertiary structures – the former is a b-barrel protein with a copper atom, whereas the latter is an iron protein consisting of four a-helices and a heme group – but play the same physiological role: the transfer of electrons between the two membrane-embedded complexes cytochrome b6 f and photosystem I. In cyanobacteria, the two proteins play an extra role as alternative electron donors to the terminal respiratory cytochrome c oxidase, thus connecting the photosynthetic and respiratory electron transfer chains (1). Plastocyanin and cytochrome c6 are alternatively synthesized in green algae and cyanobacteria in response to copper availability,

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although plastocyanin is constitutively produced in plants. The molecular mass (ca. 10 kDa) and redox potential (ca. 350  mV, pH 7.0) of these two proteins are near each other. However, their global electrostatic charge varies from one organism to another, but it is the same within each organism. In fact, the two proteins are acidic in eukaryotes (plants and green algae) but neutral or highly basic in prokaryotes (cyanobacteria) (2). The purification procedure of any given protein (either plastocyanin or cytochrome c6) must thus be modified depending on the organism, but it is the same for two proteins isolated from the same source. The variable content in DNA, pigments, and other cellular components of cyanobacteria, green algae, and plants is a crucial point. In this chapter, we describe the methods for the production, isolation, and purification of plastocyanin and cytochrome c6 with different isoelectric points (pI) from a number of organisms: spinach and Monoraphidium (pI 3.5–4), Synechocystis (pI ca. 6), and Nostoc (pI ca. 9). The procedures herein described can be used to purify the proteins from other organisms. Actually, the protocols for the purification of plastocyanin and cytochrome c6 from Monoraphidium and Synechocystis can be likewise used to obtain the proteins from Chlamydomonas and Phormidium, respectively.

2. Materials 2.1. Organisms

1. Spinach (Spinacea oleracea): Plants can be obtained from a local market. Frozen spinach leaves render a very low protein yield. 2. Monoraphidium braunii (formerly Ankistrodesmus braunii): The green alga can be obtained from the Culture Collection of Algae at the University of Göttingen (Albrecht-von-HallerInstitut, Universität Göttingen, Nikloausberger Weg 18, 37073 Göttingen, Germany; www.epsag.uni-goettingen.de/ html/sag.html), with the accession number 48.87. 3. Synechocystis sp. and Nostoc sp. (formerly Anabaena sp.): Both cyanobacteria can be obtained either from the Pasteur Culture Collection (PCC, Institute Pasteur, 28 rue du Dr. Roux, 75724, Paris Cedex 15, France; www.pasteur.fr/recherche/ banques/PCC), with the accession numbers 6803 and 7119, respectively, or from the American Type Culture Collection (ATCC, P.O. Box 1549, Manassas, VA 20108, USA; www. lgcstandards-atcc.org), with the accession numbers 27184 and 29151, respectively. 4. Escherichia coli (E. coli): Transformable strains TG1 (phenotype: supE hsdD5 thiD (lac-proAB) F’ [traD36 proAB+ lacIq

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lacZ DM15]) (3) and RV308 (phenotype: su− Dlacx74 galISII::OP308 strA; ATCC 31608) are used for the expression of spinach plastocyanin. E. coli transformable strain − MC1061 (phenotype: F araD139 D(ara-leu)7696 galE15 galK16 D(lac)X74 rpsL (Strr) hsdR2 (rK− mK+) mcrA mcrB1) and BL21 (DE3) (phenotype F´::Tn10 proA+B+, lacIq, D(lacZ)M15/ompT(lon), hsdSB(rB−mB−), l(DE3)) (4) are used for the expression of cyanobacterial proteins. Alternatively, other strains can be used (5). 2.2. Culture Media

Sterile culture media can be prepared in either glass Roux flasks, glass Erlenmeyer flasks, or glass cylindrical bottles (10 and 20 L) (see Note 1). 1. Monoraphidium braunii: 14.5  mM NaH2PO4, 2.5  mM Na2HPO4, 1  mM MgSO4, 10  mM KNO3, 0.7  mM ZnSO4, 2.5  mM MnCl2, 10  mM Na2MoO4, 0.1  mM CaCl2, 8  mM H3BO3, 8  mM NaCl, and 2.5  mg/L iron (as an Fe(III)EDTA complex) (6) (see Note 2). To make the cells synthesize plastocyanin, 10 mM CuSO4 must be added to the culture medium. 2. Synechocystis and Nostoc (BG11 medium): 0.2 mM Na2CO3, 0.3 mM MgSO4, 0.24 mM CaCl2, 0.2 mM K2HPO4, 17.6 mM NaNO3, 11.9 mM NaHCO3, 28.5 mM citric acid, 6/mg L of Fe(III)-ammonium citrate (17% Fe), 2.4  mM Na2-EDTA, 46 mM H3BO3, 9.1 mM MnCl2, 1.6 mM Na2MoO4, 0.8 mM ZnSO4, and 0.2  mM CoCl2 (7) (see Note 3). To make the cells synthesize either cytochrome c6 or plastocyanin, the content of Fe(III)-ammonium citrate is doubled or 1 mM CuSO4 is added, respectively. 3. Escherichia coli (Luria–Bertani medium, LB): 10  g of tryptone, 5 g of yeast extract, and 10 g of NaCl/L (8). To maintain and amplify the plasmids, ampicillin is added to the LB medium at 100 mg/mL (see Note 4). This medium must be supplemented with 200 mM CuSO4 in case of plastocyanin.

2.3. Purification of Proteins 2.3.1. Spinach Plastocyanin

1. Buffer A: 0.4 M sucrose and 15 mM NaCl in 20 mM Tricine– KOH, pH 8.0. 2. Buffer B: 5 mM Tricine–KOH, pH 8.0. 3. 1 M Tris–HCl buffer, pH 8.0. 4. 10 mM Potassium phosphate buffer, pH 7.0. 5. 10 mM Tris–HCl buffer, pH 8.0. 6. 10 mM Tris–acetate buffer, pH 8.0 (see Note 5). 7. Acetone 80% (v:v) in distilled water. 8. Reagent-grade pyridine.

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9. Protamine sulfate: 20  mg/mL in 20  mM Tricine–KOH, pH 8.0; make fresh as required. 10. MgCl2. 11. NaCl. 12. 0.1 M Potassium ferricyanide in distilled water. 13. Washed DEAE-cellulose. 14. Polybuffer exchanger 94. 15. Polybuffer 74 solution. 16. Amicon pressure filtration cells (250 and 10 mL) with YM-3 membranes. 17. Dialysis membranes (3.5 kDa cutoff, several diameters). 18. Three chromatography columns (6 × 12  cm, 2 × 15  cm, and 1.5 × 20 cm). 19. Gradient glasses (three couples of 1, 0.4, and 0.125 L). 20. Waring blender (1 gallon) or similar apparatus. 21. Teflon-glass homogenizer (50 mL). 22. Branson 250 Sonifier (or equivalent) with a standard tip. 23. Organdy or cheesecloth (a piece of 1 × 1 m). 2.3.2. Monoraphidium Plastocyanin and Cytochrome c6

1. Buffer C: 1  mM phenylmethylsulfonyl fluoride (PMSF), 1  mM benzamidine, and 1  mM e-aminocaproic acid in 20 mM phosphate, pH 7.0 (make fresh as required). 2. 10 mM Tris–HCl buffer, pH 8.0. 3. 10 mM Tris–acetate buffer, pH 8.0 (see Note 5). 4. 0.1 M Streptomycin sulfate: in water, pH 7.0; make fresh as required. 5. Ammonium sulfate. 6. NaOH. 7. NaCl. 8. 0.1 M Potassium ferricyanide in distilled water. 9. Washed DEAE-cellulose. 10. Polybuffer exchanger 94. 11. Polybuffer 74 solution. 12. Amicon pressure filtration cells (250 and 10 mL) with YM-3 membranes. 13. Dialysis membranes (3.5 kDa cutoff, several diameters). 14. Two chromatography columns (2 × 20 cm and 1.5 × 20 cm). 15. Gradient glasses (two couples of 0.4 and 0.125 L). 16. Manton-Gaulin disruptor (or similar apparatus).

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17. Teflon-glass homogenizer (50 mL). 18. Branson 250 Sonifier (or equivalent) with a standard tip. 2.3.3. Synechocystis Plastocyanin and Cytochrome c6

1. Buffer C: 1  mM PMSF, 1  mM benzamidine, and 1  mM e-aminocaproic acid in 20  mM phosphate, pH 7.0 (make fresh as required). 2. 2 mM Tris–HCl buffer, pH 8.0. 3. 5 mM Tris–acetate buffer, pH 8.0 (see Note 5). 4. 20 mM Tricine–KOH buffer, pH 7.5. 5. 0.1 M Streptomycin sulfate: in water, pH 7.0; make fresh as required. 6. Ammonium sulfate. 7. NaOH. 8. NaCl. 9. 0.1 M Potassium ferricyanide in distilled water. 10. Washed DEAE-cellulose. 11. Polybuffer exchanger 94. 12. Polybuffer 74 solution. 13. Amicon pressure filtration cells (250 and 10 mL) with YM-3 membranes. 14. Dialysis membranes (3.5 kDa cutoff, several diameters). 15. Two chromatography columns (2 × 30 cm and 1.5 × 20 cm). 16. A Superdex 75 (Amersham) FPLC column (1.6 × 60 cm) (or equivalent). 17. Gradient glasses (two couples of 0.4 and 0.125 L). 18. Manton-Gaulin disruptor (or similar apparatus). 19. Branson 250 Sonifier (or equivalent) with a standard tip.

2.3.4. Nostoc Plastocyanin and Cytochrome c6

1. Buffer C: 1  mM PMSF, 1  mM benzamidine, and 1  mM e-aminocaproic acid in 20  mM phosphate, pH 7.0 (make fresh as required). 2. 1 mM Phosphate buffer, pH 7.0. 3. 20 mM Tricine–KOH buffer, pH 7.5. 4. 0.1 M Streptomycin sulfate: in water, pH 7.0; make fresh as required. 5. Ammonium sulfate. 6. NaOH. 7. NaCl. 8. 0.1 M Potassium ferricyanide in distilled water. 9. Washed carboxymethyl-cellulose.

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10. Amicon pressure filtration cells (250 and 10 mL) with YM-3 membranes. 11. A chromatography column (2 × 30 cm). 12. A Superdex 75 (Amersham) FPLC column (1.6 × 60 cm) (or equivalent). 13. Gradient glasses (0.4 L). 14. Manton-Gaulin disruptor (or similar apparatus). 15. Branson 250 Sonifier (or equivalent) with a standard tip.

3. Methods The protocols described herein have been modified from a series of previously reported methods for the isolation and purification of plastocyanin and/or cytochrome c6 from spinach (9, 10), Monoraphidium (11, 12), Synechocystis (13–15), and Nostoc (16, 17). Figure 1 shows a flowchart with the different procedures. 3.1. Cell Growth and Collection 3.1.1. Green Algae and Cyanobacteria

3.1.2. Escherichia coli

Grow green algal and cyanobacterial cells in culture rooms at 25 or 30°C, respectively. Routinely maintain cells by 1:10 dilution of stationary cultures (absorbance at 600  nm higher than 1.5) in sterile glass Roux flasks (culture volume, 750 mL) (see Note 6). Use a Roux flask to inoculate a 10-L sterile glass bottle, which in turn serves to inoculate four 20-L glass bottles. All the cultures are kept under continuous irradiation by fluorescent light tubes (50 mE/m2 s) and gas bubbling (air: CO2 mixture, 99:1 [v/v]). Collect cells from 80 L cultures at stationary phase (absorbance at 600 nm, 1.6–1.8) by centrifugation at 14,000 × g in either a continuous flow system or large volume rotor. Freeze centrifuge tubes with cell pellets overnight for efficient recovery of the pellets. Typical cell yields are around 1–2  g (wet weight)/L. The pellets can be kept at −20°C until use. For high-yield protein purification, E. coli cells are transformed by following standard molecular biology methods (10, 14, 15, 17). Inoculate 100  mL of transformed cells in a 250-mL glass Erlenmeyer flask containing 100 mL of LB medium. After 12 h of growth at 37°C under continuous stirring in an orbital shaker (200  rpm), the culture is used to inoculate three 5-L glass Erlenmeyer flasks containing 2 L of LB medium each. Leave cells to grow under similar temperature and shaking conditions to reach the stationary phase (approximately 18  h; absorbance at 600  nm higher than 1.2) and collect upon centrifugation at 14,000 × g for 5 min. For systems under control of the T7 promoter, isopropyl-b-D-thio-galactopyranoside (IPTG) at a final

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Algal and Cyanobacterial Proteins

Spinach Plastocyanin

cell collection and breakdown

leaves breakdown

centrifugation at 3,000g

chloroplasts isolation and breakdown

supernatant sonication centrifugation at 12,000g

centrifugation at 30,000g pellet resuspension, pyridine addition and sonication

streptomycin precipitation of the supernatant E. coil periplasmic fractions (cyanobacterial proteins)

centrifugation at 12,000g

protamine sulfate precipitation ammonium sulfate precipitation of the supernatant

centrifugation at 4,500g E. coli periplasmic fractions (eukaryotic proteins)

supernatant

centrifugation at 12,000g pellet resuspension and dialysis

ionic exchange chromatography algal proteins chromatofocusing and salt gradient

ionic exchange chromatography chromatofocusing (Synechocystis proteins) FPLC chromatography

Fig. 1. Flow-chart summarizing the different steps of the procedures for purification of plastocyanin and/or cytochrome c6 from plants, green algae and cyanobacteria.

concentration of 0.5–1 mM is added to the culture 8 h before cell collection (absorbance at 600, ca. 1.0). Freshly collected cells are immediately used for protein purification. All the cultures are made under sterile conditions. 3.2. Cell Breaking Procedures 3.2.1. Spinach Leaves

The starting material consists of 3  kg of depeciolated and denerved spinach leaves that must be washed with cold distilled water and dried in a cold chamber. This is a time-consuming procedure, and so the material should be prepared the day before and kept in a cold room in the dark. 1. 500  g batches of spinach leaves are mixed with 700  mL of buffer A and broken in the Waring blender (30 s of blending at 70% power).

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2. The resulting suspension is filtered through four organdy or cheesecloth layers in a funnel to eliminate unbroken material. 3. To recover as much crude extract as possible, the cell suspension has to be squeezed by hand. 4. The filtered suspension is centrifuged at 300 × g for 1 min to remove whole cells, and the supernatant is again centrifuged at 4,000 × g for 10 min to precipitate chloroplasts. 5. The resulting pellet is resuspended with a glass bar in 400 mL of buffer B and gently homogenized in a teflon-glass homogenizer. 6. The suspension is left to stand for 10 min under gentle stirring in darkness and then centrifuged at 30,000 × g for 20 min. 7. The final pellet is resuspended in 200  mL of distilled cold water and homogenized again. 8. Chlorophyll concentration is determined as described by Arnon (18). (a) 0.1 mL of suspension is dissolved in 20 mL of 80% acetone in water (v/v). (b) After vigorous mixing, insoluble material is removed upon centrifugation at 13,000 × g for 5  min at room temperature. (c) The absorbance at 652 nm of the supernatant is measured, and the chlorophyll content is determined by using an extinction coefficient of 34.5 mg chlorophyll mL−1 cm−1. 3.2.2. Green Algal and Cyanobacterial Cells

1. The initial steps are the same for Monoraphidium, Synechocystis, and Nostoc cells. Mix 250 g of frozen cell paste with 1 L of cold buffer C (freshly prepared, owing to the instability of the protease inhibitors) and keep under vigorous stirring at 4°C for at least 1 h in the dark. 2. Cell disruption is carried out on pressure/depressure cycles by passing the cell suspension 3 times through a Manton-Gaulin high pressure continuous system at 7,000 psi (see Note 7). 3. The resulting suspension is centrifuged at 3,000 × g for 2 min. 4. The supernatant is sonicated in 150-mL aliquots in an ice bath for 1 min with a sonifier (50% power) after adding solid NaCl at 50 mM final concentration. 5. The suspension is centrifuged at 12,000 × g for 20 min, and the pellet is discarded.

3.2.3. Escherichia coli Cells

E. coli transformed cells are collected upon centrifugation of 6-L cultures, and the pellets (see Subheading  3.1.2) are ­resuspended in 20 mL of distilled water. Transformed cells are

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designed to send the overexpressed proteins (either plastocyanin or cytochrome c6) to the periplasmic space, and therefore, the cell suspension must undergo a gentle breaking consisting of two cycles of freezing in liquid nitrogen and thawing at 40°C in a thermostated bath. Upon centrifugation at 12,000 × g for 5  min, the periplasmic fraction is collected from the supernatant. 3.3. Purification of Plastocyanin and Cytochrome c6

3.3.1. Spinach Plastocyanin 3.3.1.1. Differential Precipitation

The purification procedures must be carried out in a cold chamber (4°C), or alternatively, the cell and protein samples are kept in ice containers. To better follow the proteins throughout purification, cytochrome c6 is maintained in its reduced native redox state so that it can be easily recognized by its typical pink-orange color, whereas a few microliters of a 0.1 M potassium ferricyanide stock solution are added to plastocyanin to get the characteristic blue color of its oxidized state (see Note 8). From here thereafter, the same protocol is used for purification of plastocyanin and cytochrome c6 when they are both isolated from the same organism. Purity of cytochrome c6 fractions is determined from the A553/ A275 ratio of its reduced state (see Note 9), whereas purity of plastocyanin samples is determined from the A278/A597 ratio of its oxidized state. Plastocyanin becomes partially autoreduced throughout purification; therefore, its redox state should be checked at every step by measuring the absorbance change at 597 nm in the absence of ferricyanide and after addition of a little amount of oxidant. 1. Chlorophyll and buffer concentration of the suspension obtained in Subheading 3.2.1 is first fitted to 1.5 mg/mL and 10  mM Tris–HCl, pH 8.0, respectively, by adding distilled water and 1 M Tris–HCl, pH 8.0. 2. Add pyridine (0.7 mL to every 100 mL of solution) and gently stir the mixture for 30 min. 3. Sonicate the resulting solution (150 mL aliquots each time) in a sonifier (100% power) for 150 s in a salt-ice bath to avoid sample heating. 4. Add the protamine sulfate solution under vigorous stirring to reach the proportion of 1 mL of protamine solution to every 5 mL of sonicated solution. 5. Stir the resulting preparation for 5 min and allow to rest for 30 min. 6. Add solid MgCl2 under continuous stirring to reach a final concentration of 5 mM. 7. After 10 min with no stirring, the preparation is centrifuged at 14,000 × g for 15 min, and the pellet is discarded.

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3.3.1.2. Anionic Exchange Chromatography

1. Load the supernatant resulting from the previous step onto a DEAE-cellulose column (6 × 12 cm, 300 mL of gel volume, elution flow ca. 40  mL/h) that must be previously equilibrated with the potassium phosphate buffer. 2. After sample loading, wash the column with at least 600 mL of the phosphate buffer to remove unbound material. 3. Elute plastocyanin by applying a 0.01–0.4 M NaCl linear gradient in phosphate buffer (total volume, 2 L). 4. Pool plastocyanin fractions with an A278/A597 ratio lower than 10, fully oxidize with potassium ferricyanide, and dialyze for 24  h against 10  mM Tris–HCl, pH 8.0, supplemented with 10  mM ferricyanide, with at least one dialysis buffer replacement. 5. Load the resulting dialyzed preparation – or alternatively, the supernatant obtained from E. coli transformed cells in Subheading 3.2.3 – onto a second DEAE-cellulose column (2 × 15 cm, 40 mL of gel volume, elution flow ca. 20 mL/h) that must be previously equilibrated with 10 mM Tris–HCl, pH 8.0. 6. After sample loading, wash the column with at least 80 mL of the same Tris–HCl buffer supplemented with 60 mM NaCl to remove weakly bound material. 7. Plastocyanin is eluted by applying a 0.06–0.3 M NaCl linear gradient in Tris–HCl buffer (total volume, 0.4 L). 8. Plastocyanin fractions with an A278/A597 ratio lower than 5 are pooled, dialyzed against the Tris–acetate buffer, and fully oxidized with ferricyanide.

3.3.1.3. Chromatofocusing

1. The solution resulting from the previous step is loaded onto the chromatofocusing Polybuffer exchanger 94 column (1.5 × 20 cm, 30 mL of gel volume, elution flow ca. 10 mL/h) that must be previously equilibrated with the Tris–acetate buffer. 2. Wash the column with 150  mL of a 10-times diluted Polybuffer 74 solution, pH 4.0, and elute plastocyanin by applying a 0.01–0.45  M NaCl linear gradient in 25  mM Tris–acetate, pH 5.0 (total volume, 0.2 L). 3. Fractions with an A278/A597 ratio close to one are pooled, suspended in an adequate buffer upon several cycles of concentration/dilution in the Amicon pressure filtration cell, concentrated, and finally frozen until use.

3.3.2. Monoraphidium Plastocyanin and Cytochrome c6 3.3.2.1. Differential Precipitation

1. Add streptomycin sulfate to the supernatant obtained in Subheading 3.2.2 at the final ratio of 1:10 (v/v) under vigorous stirring. 2. Gently stir the solution for 1 h and allow to rest for 2 h. 3. Discard the precipitated material upon centrifugation at 12,000 × g for 20 min.

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4. Slowly add solid ammonium sulfate to the supernatant, to reach a saturation level of 60% (36.1  g added to every 100 mL), under vigorous stirring and continuous pH testing to keep the pH value close to 7 by adding small amounts of a concentrated NaOH solution. 5. Gently stir the resulting preparation for 1 h and then clear it upon centrifugation at 12,000 × g for 20 min. 6. Add ammonium sulfate to the supernatant as before to reach a saturation level of 98% (27.5 g added to every 100 mL). 7. Upon centrifugation at 12,000 × g for 20 min, resuspend the pellet by means of a glass bar in 200 mL of the Tris–HCl buffer and dialyze against the same Tris–HCl buffer for 48 h with at least two dialysis buffer replacements. 3.3.2.2. Anionic Exchange Chromatography

1. The dialyzed solution is loaded onto the DEAE-cellulose column (2 × 20  cm, 60  mL of gel volume, elution flow ca. 20  mL/h) that must be previously equilibrated with the Tris–HCl buffer. 2. After column washing with at least 120  mL of Tris–HCl buffer, elute plastocyanin or cytochrome c6 by applying a 0.01–0.4 M NaCl linear gradient in Tris–HCl buffer (total volume, 0.6 L). 3. Pool and dialyze plastocyanin or cytochrome c6 containing fractions.

3.3.2.3. Chromatofocusing

1. The solution resulting from the previous step is further purified by chromatofocusing as described under Subheading 3.3.1. 2. The pure protein fractions with either an A278/A597 ratio of approximately 1.5 for plastocyanin or an A553/A275 ratio of 1.15 in case of cytochrome c6 are pooled, suspended in an adequate buffer upon several cycles of concentration/­dilution in the Amicon pressure filtration cell, concentrated, and finally frozen until use.

3.3.3. Synechocystis Plastocyanin and Cytochrome c6 3.3.3.1. Differential Precipitation

1. Streptomycin and two-sequential ammonium sulfate precipitations are as described under Subheading  3.3.2, with the two following exceptions: (a) In case of proteins isolated from Synechocystis, the first ammonium sulfate precipitation is performed at a saturation level of 50% rather than 60%. (b) In case of recombinant proteins expressed in and isolated from E. coli, the streptomycin precipitation step is omitted and so the supernatants obtained at Subheading 3.2.3 are directly subjected to sequential ammonium sulfate precipitation.

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3.3.3.2. Anionic Exchange Chromatography

1. After 98% ammonium sulfate precipitation, resuspend the ­centrifugation pellets with a glass bar in 200 mL of the Tris– HCl buffer, desalt along several cycles of concentration/­ dilution with the same buffer in the Amicon pressure filtration cell, and finally dilute with buffer up to 400 mL final volume. 2. Load the resulting solution onto the DEAE-cellulose column (2 × 30 cm, 80 mL of gel volume, elution flow ca. 20 mL/h) that must be previously equilibrated with the Tris–HCl buffer. 3. Wash the column with at least 80 mL of Tris–HCl buffer. 4. Plastocyanin or cytochrome c6 are eluted from the column by applying a 0–0.2  M NaCl linear gradient in Tris–HCl buffer (total volume, 0.6 L), and the fractions with an A275/ A597 (plastocyanin) or A274/A552 (cytochrome c6) ratio lower than 10 are pooled and dialyzed against the Tris–acetate buffer.

3.3.3.3. Chromatofocusing

3.3.3.4. FPLC Chromatography

The solution resulting from the previous step is further purified by chromatofocusing as described in Subheading  3.3.1, except that plastocyanin or cytochrome c6 are eluted with 10-times diluted Polybuffer 74 solution, pH 4.0. The fractions with an A275/A597 lower than 5 (plastocyanin) or an A274/A552 ratio lower than 3 (cytochrome c6) are pooled and concentrated by pressure filtration in an Amicon cell. 1. The protein solution resulting from the previous step is subjected to FPLC chromatography on a Superdex 75 column (1.6 × 60 cm; elution flow 0.5 mL/min) previously equilibrated with the Tricine–KOH, supplemented with 150 mM NaCl. 2. Plastocyanin or cytochrome c6 are eluted using the same buffer solution. 3. Pure protein fractions with an A275/A597 ratio of approximately 2.2 (oxidized plastocyanin) or an A552/A274 ratio of 1.14 (reduced cytochrome c6) are pooled, concentrated, and frozen until use.

3.3.4. Nostoc Plastocyanin and Cytochrome c6

1. Streptomycin and two-sequential ammonium sulfate precipitations are as described under Subheading 3.3.2.

3.3.4.1. Differential Precipitation

2. In case of recombinant proteins expressed in and isolated from E. coli, the streptomycin precipitation step is omitted and so the supernatants obtained at Subheading 3.2.3 are directly subjected to sequential ammonium sulfate precipitation.

3.3.4.2. Cationic Exchange Chromatography

1. After 98% ammonium sulfate precipitation, the ­centrifugation pellets are resuspended with a glass bar in 200 mL of 1 mM

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sodium phosphate, pH 7.0, and dialyzed against the same buffer for 48 h, with at least two dialysis buffer replacements. 2. Load the dialyzed solution onto the carboxymethyl-cellulose ­column (2 × 30  cm, 80  mL of gel volume, elution flow ca. 20 mL/h) previously equilibrated with the sodium phosphate buffer. 3. Wash the column with at least 160 mL of phosphate buffer supplemented with 5 mM ferricyanide. 4. Plastocyanin or cytochrome c6 are eluted from the column by means of a 1–35 mM phosphate linear gradient, pH 7.0 (total volume, 0.6 L). 5. The fractions with an A275/A597 (plastocyanin) or A274/A552 (cytochrome c6) ratio lower than 3 are pooled and concentrated. 3.3.4.3. FPLC Chromatography

1. The protein solution resulting from the previous step is subjected to FPLC chromatography as described under Subheading 3.3.3. 2. Pure protein fractions with an A275/A597 ratio of approximately 1.2 (oxidized plastocyanin) or an A552/A274 ratio of approximately 1.0 (reduced cytochrome c6) are pooled, concentrated, and frozen until use.

4. Notes 1. When bubbling is required, the gas flow is passed through commercial 0.2-mm filters of different sizes depending on culture volume. Foaming is avoided by adding one or two drops of a suitable antifoaming agent (e.g., silicone) to every liter of culture medium before sterilization. 2. To avoid salt precipitation, the culture medium should be ­prepared from a sterilized and 100-fold concentrated solution containing all the components but the phosphates and Fe(III)EDTA. Solid NaH2PO4 . H2O (2 g/L) and Na2HPO4 . 12H2O (0.9 g/L), as well as Fe(III)-EDTA (0.5 mL/L from a stock solution) are added to the culture medium just before sterilization. The Fe(III)-EDTA stock solution is prepared as follows: 16  g of EDTA and 10.4  g of KOH are dissolved in 186 mL of distilled water, and 13.7 g of FeSO4 . 7H2O are dissolved in 364  mL of distilled water; the two solutions are mixed and vigorously bubbled overnight with air in the dark to fully oxidize iron atoms. 3. To avoid contamination and salt precipitation, the culture medium is prepared from a 100-fold concentrated stock

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solution containing all the components but K2HPO4, NaNO3, and NaHCO3. K2HPO4 (0.2 mL/L from a 1 M stock solution), solid NaNO3 (1.5 g/L), and NaHCO3 (1.0 g/L) are added to the final culture medium just before sterilization. 4. Ampicillin is added from a 1,000-fold concentrated stock solution sterilized by filtration through a 0.25-mm sterile ­filter. The filtered ampicillin solution can be stored at −20°C for several months. Other antibiotics can be alternatively used for the molecular biology constructs and plasmid selection. 5. 10  mM Tris–acetate buffer, pH 8.0, must be prepared by adding small aliquots of concentrated acetic acid to a solution of 10 mM Tris up to reach a pH value of 8.0. 6. For long-term maintenance, the organisms can be kept in either petri dishes with solid media (1.5% (w/v) agar concentration) or 100-mL glass Erlenmeyer flasks with liquid medium and no bubbling. In the first stages of diluted cultures in glass Roux flasks, light intensity should be dimmed to avoid photobleaching by placing a piece of paper between flasks and fluorescent tubes. Every 80-L culture usually takes 4 days to reach the saturation level; after the second day, additional side panels of fluorescent lamps can be placed to enhance cell growing. Cell collection from each 80-L culture takes 4–5 h by continuous flow centrifugation. 7. The cells can alternatively be disrupted in a French press after three cycles at 20,000 psi, but there is a limit in the capacity of the cylinder (ca. 40  mL). Algal (Monoraphidium) and cyanobacterial (Synechocystis and Nostoc) cells can also be broken in a Sorvall Omnimixer (or similar apparatus), for which the wet cell paste is carefully mixed with the same ­volume of 0.2  mm diameter glass beads; after 10 cycles of 30 s shaking with 2 min cooling intervals, the glass beads are sepa­rated upon decantation and the resulting suspension is centrifuged to remove unbroken cells. In case of Synechocystis, however, the cell wall has to be previously degraded by ­adding ­penicillin G (100 mg/L) to the cultures 24 h before collection. 8. Cyanobacterial plastocyanin cannot be followed throughout the first steps of purification because of optical interference of phycobiliproteins, and it is not until the ammonium sulfate fractionation that the copper protein can be detected. 9. Depending on the organism from which cytochrome c6 is isolated, a difference of 1–2 nm in the maximum value of the ultraviolet (274–275  nm) and alpha (552–553  nm) bands can be observed in the absorption spectrum of the reduced form.

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Acknowledgments The authors wish to thank their students, co-workers, and collaborators, listed in the references, for their contributions to this research. Financial support was provided by the Spanish Ministry of Science and Innovation, the European Commission, and the Andalusian Government. References 1. Hervás, M., Navarro, J.A., and De la Rosa, M.A. (2003) Electron transfer between membrane complexes and soluble proteins in photosynthesis. Accounts Chem. Res. 36, 798–805. 2. De la Rosa, M.A., Navarro, J.A., Díaz-Quintana, A., De la Cerda, B., Molina-Heredia, F.P., Balme, A., Murdoch, P.S., Díaz-Moreno, I., Durán, R.V., and Hervás, M. (2002) An evolutionary analysis of the reaction mechanisms of photosystem I reduction by cytochrome c6 and plastocyanin. Bioelectrochem. 55, 41–45. 3. Carter, P. (1986) Site directed mutagenesis. Biochem. J. 237, 1–7. 4. Casadaban, M.J. and Cohen, S.N. (1980) Analysis of gene control signals by DNA fusion and cloning in Escherichia coli. J. Mol. Biol. 138, 179–207. 5. Hippler, M., Drepper, F., Farah, J., and Rochaix, J.-D. (1997) Fast electron transfer from cytochrome c6 and plastocyanin to photosystem I of Chlamydomonas reinhardtii requires PsaF. Biochemistry 36, 6343–6349. 6. Kessler, E., Langner, W., Ludewig, I., and Wiechmann, H. (1963) Bildung von sekundarcarotinoiden bei stickstoffmangel und hydrogenase-aktivitat als taxonomische ­merkmale in der gattung Chlorella, in Studies on Microalgae and Photosynthetic Bacteria (Japanese Society of Plant Physiology, ed.), Tokyo University Press, Tokyo, pp. 7–20. 7. Rippka, R., Deruelles, J., Waterbury, J.B., Herdman, M., and Stainer, R.Y. (1979) Generic assignments, strain stories and properties of pure cultures of cyanobacteria. J. Gen. Microbiol. 111, 1–61. 8. Sambrook, J., Fritsch, E., and Maniatis, T. (1989) Molecular Cloning: A Laboratory Manual, Second ed. Cold Spring Harbor Laboratory Press, Cold Spring Harbor. 9. Yocum, C.F. (1982) Purification of ferredoxin and plastocyanin, in Methods in Chloroplast Molecular Biology (Edelman, M., Hallick, R.B., and Chua, N.-H., eds.), Elsevier Biomedical, Amsterdam, pp. 973–981.

10. Ejdebäck, M., Young, S., Samuelsson, A., and Karlsson, B.G. (1997) Effects of codon usage and vector-host combinations on the expression of spinach plastocyanin in Escherichia coli. Protein Expres. Purif. 11, 17–25. 11. Hervás, M., De la Rosa, M.A., and Tollin, G. (1992) A comparative laser flash photolysis study of algal plastocyanin and cytochrome c552 photooxidation by photosystem I particles. Eur. J. Biochem. 203, 115–120. 12. Campos, A.P., Aguiar, A.P., Hervás, M., Regalla, M., Navarro, J.A., Ortega, J.M., Xavier, A.V., De la Rosa, M.A., and Teixeira, M. (1993) Cytochrome c6 from Monoraphidium braunii: A cytochrome with an unusual heme axial coordination. Eur. J. Biochem. 216, 329–341. 13. Hervás, M., Navarro, F., Navarro, J.A., Chávez, S., Díaz, A., Florencio, F.J., and De la Rosa, M.A. (1993) Synechocystis 6803 plastocyanin isolated from both the cyanobacterium and E. coli transformed cells are identical. FEBS Lett. 319, 257–260. 14. De la Cerda, B., Navarro, J.A., Hervás, M., and De la Rosa, M.A. (1997) Changes in the  reaction mechanism of electron transfer from  plastocyanin to photosystem I in the cyanobacterium Synechocystis sp. PCC 6803 as  induced by site-directed mutagenesis of  the copper protein. Biochemistry 36, 10125–10130. 15. Feio, M.J., Díaz-Quintana, A., Navarro, J.A., and De la Rosa, M.A. (2006) Thermal unfolding of plastocyanin from the mesophilic cyanobacterium Synechocystis sp. PCC 6803 and comparison with its thermophilic counterpart from Phormidium laminosum. Biochemistry 45, 4900–4906. 16. Medina, M., Díaz, A., Hervás, M., Navarro, J.A., Gómez-Moreno, C., De la Rosa, M.A., and Tollin, G. (1993) A comparative laser flash absorption spectroscopy study of Anabaena PCC 7119 plastocyanin and cytochrome c6 photooxidation by photosystem I particles. Eur. J. Biochem. 213, 1133–1138.

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17. Molina-Heredia, F.P., Hervás, M., Navarro, J.A., and De la Rosa, M.A. (1998) Cloning and correct expression in Escherichia coli of the petE and petJ genes respectively encoding plastocyanin and cytochrome c6 from the cyanobacterium

Anabaena sp. PCC 7119. Biochem. Biophys. Res. Commun. 243, 302–306. 18. Arnon, D.I. (1949) Copper enzymes in isolated chloroplasts: polyphenol oxidase in Beta vulgaris. Plant Physiol. 24, 1–15.

Chapter 9 Isolation and Identification of Chloroplast Lipids Norihiro Sato and Mikio Tsuzuki Abstract Glycerolipids of photosynthetic organisms are accounted for largely by thylakoid membrane lipids consisting of chloroplast-specific glycolipids such as monogalactosyl diacylglycerol, digalactosyl diacylglycerol, and sulfoquinovosyl diacylglycerol, and a sole phospholipid, phosphatidylglycerol. In this chapter, methods for characterization of lipids from plant cells are described. The methods include extraction of total lipids from the cells, separation of these lipids into individual lipid classes by thin-layer chromatography, and identification of respective lipid classes by their mobility. We also present methods for the determination of compositions of constituent fatty acids, distribution of fatty acids between sn-1 and sn-2 positions, and determination of contents of individual lipid classes by gas–liquid chromatography. These methods are applicable to isolated chloroplasts or some membrane fractions such as thylakoid membranes. Key words: Fatty acid, Gas–liquid chromatography, Lipase from Rhizopus delemar, Lipid, Methanolysis, Thin-layer chromatography

1. Introduction Lipids in biomembranes interact among one another through hydrophobic bonds, whereas they interact with membrane proteins not only through hydrophobic bonds, but also with electrostatic and hydrogen bonds. The hydrophobic bonds are eliminated by relatively non-polar solvents such as chloroform, and the electrostatic and hydrogen bonds are cut by polar solvents such as methanol. Thus, lipids can be prepared from cells or membrane preparations as total lipids through extraction with a mixture of chloroform and methanol. From the total lipid fraction extracted with such a solvent system, individual lipid classes are separated through thin-layer chromatography (TLC) on the basis of their particular mobility, detected on TLC plates with a non-destructive

Robert Carpentier (ed.), Photosynthesis Research Protocols, Methods in Molecular Biology, vol. 684, DOI 10.1007/978-1-60761-925-3_9, © Springer Science+Business Media, LLC 2011

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reagent such as primulin. Fatty acid methyl esters are prepared from individual lipid classes through transmethylation by methanolysis, and then analyzed with gas–liquid chromatography for the determination of fatty acid compositions. The chromatograph is equipped with a column that separates molecular species of fatty acid methyl esters according to their partition property between gas and liquid phases at high temperatures. Each lipid class can be quantified through the estimation of contents of the constituent fatty acids relative to the content of a particular fatty acid that is originally absent, but is added as an internal standard to the lipid fraction. Fatty acid compositions at the sn-1 and sn-2 positions of glycerol backbones can also be estimated through the determination of fatty acid compositions of both individual lipids and their sn-2-acyl-lysolipids synthesized through specific hydrolysis at the sn-1 positions by a lipase from Rhizopus delemar. Two biosynthetic pathways designated as prokaryotic and eukaryotic pathways operate for glycerolipid synthesis in plants. The prokaryotic pathway, the metabolism of which is confined within plastids, produces prokaryotic lipids, similar to cyanobacterial lipids, containing fatty acids with a chain of 16 carbon atoms at the sn-2 positions. On the contrary, the eukaryotic pathway, through cooperation of three cellular compartments, i.e., plastids, cytoplasm, and endoplasmic reticulum, produces eukaryotic lipids possessing fatty acids with a chain of 18 carbon atoms at the sn-2 positions. Thus, information not only on their molecular structure, but also on their synthetic pathway will be provided by the analysis of positional distribution of fatty acids of individual lipid classes. It has been shown that chloroplast lipids are synthesized by prokaryotic and/or eukaryotic pathways, while extra-chloroplast lipids are constructed exclusively by eukaryotic pathways (1).

2. Materials 1. Methanol. Toxic and flammable liquid. 2. Chloroform. Toxic. 3. Distilled water. 4. Rotary evaporator. 5. TLC plates (20 cm × 20 cm, No. 5721 plate, Merck). 6. TLC chamber. 7. 25–28% ammonia solution. Toxic and corrosive. 8. Primulin reagent: 0.01% primulin in 80% acetone. 9. I2. 10. Dittmer–Lester reagent: dissolve 4.01 g of MoO3 in 100 ml of 25  N H2SO4 by boiling. Cool it at room temperature

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(solution A). Add 180 mg of molybdenum powder to 50 ml of solution A and boil it for 15 min. Cool it at room temperature and recover the supernatant by decantation (solution B). The reagent is stable for several months at room temperature (see “Dittmer–Lester Reagent”). 11. Anthrone reagent: dissolve 0.1 g of anthrone and 2 g of thiourea in 66% H2SO4 (see “Anthrone Reagent”). 12. 5% (w/w) hydrogen chloride methanol solution: Prepared by bubbling of anhydrous methanol with HCl gas being passed beforehand through sulfuric acid for dehydration. Make a fresh solution every 3–4 months. Toxic, corrosive, and flammable liquid. This solution is commercially available. 13. n-Hexane. Toxic and flammable liquid. 14. Gas chromatograph (e.g., GC-14B, Shimadzu, Kyoto). 15. Capillary column (e.g., HR-Thermon 3000B, 0.25 mm × 25 m, Shinwa Chemical Industries, Kyoto). 16. Chromatography data processor (e.g., C-R7A plus, Shimadzu, Kyoto). 17. Lipase from R. delemar (Seikagaku Corporation). 18. Buffer for lipase: 50 mM Tris–HCl (pH 7.2), 0.05% Triton X-100. 19. KCl solution 1% (w/v). 20. Pre-coated silica gel TLC plate (20  cm × 20  cm, No. 5721 plate, Merck).

3. Methods The methods described here are applied for (1) the isolation of total lipids, (2) the separation into individual lipid classes, (3) the detection and identification of lipid classes, (4) the determination of fatty acid compositions of respective lipid classes and quantification of lipid classes by estimation of fatty acid contents, and (5) the analysis of distribution of fatty acids between sn-1 and sn-2 positions of glycerol backbones. 3.1. Lipid Extraction

3.1.1. Isolation of Lipids from Algae or Chloroplasts

Individual lipids can be isolated first as total lipid fraction. Shown below is the method of the isolation according to Bligh and Dyer (2) (see Note 1). 1. Prepare cells of microalgae or chloroplasts as pellets (see Note 2), the volume of which corresponds to 0.5–1.0 ml, in a 50-ml centrifuge tube with a Teflon-lined screw cap (see Note 3). 2. Add 10 ml of methanol and agitate it for 30 s with a vortex mixer.

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3. Add 5 ml of chloroform and agitate it for 2 min with a vortex mixer. 4. Allow it to stand for 10 min. 5. Add 5 ml of chloroform and agitate it with a vortex mixer for 30 s. 6. Add 5 ml of distilled water and agitate it with a vortex mixer for 30 s. The solution will become turbid. 7. Centrifuge at 1,500 × g for 5 min. The solution will separate into three phases, i.e., the upper phase of H2O and methanol, the medium phase of cell debris, and the lower phase of chloroform. 8. Take the lower phase containing lipids with a Pasteur pipette into a flask suitable for the evaporator. Do not contaminate the upper or medium phase. 9. Add 10 ml of chloroform to the remaining upper and medium phases and agitate it for 1 min with a vortex mixer. 10. Centrifuge it at 1,500 × g for 5 min. Recover the lower phase of chloroform and combine it with the lipid solution prepared in step 8. 11. Evaporate the solvent completely with a rotary evaporator under reduced pressure. 12. Add methanol or chloroform/methanol (2:1, by vol.) for complete evaporation if H2O is left. 13. Dissolve the lipids in an appropriate volume of chloroform/ methanol (2:1, by vol.), transfer the solution into a tube with a Tefron-lined screw cap, and store it at –20°C until use. 3.1.2. Isolation of Lipids from Leaves of Higher Plants

1. Cut green leaves (5 g of wet weight) into slices. 2. Add 150  ml of chloroform/methanol (1:2, by vol.) and homogenize it with a homogenizer. 3. Filtrate the slurry through a piece of filter paper and recover the filtrate in a flask. 4. Add 50 ml of chloroform and agitate it for 1 min. 5. Add 100 ml of 1% KCl solution (w/v) and agitate it for 2 min. 6. Allow it to stand for half a day until the turbid solution separates into three phases. 7. Follow steps 8–13 in Subheading 3.1.1.

3.2. Separation into Individual Lipid Classes by TLC

TLC that requires no expensive equipment can separate individual lipid classes simultaneously on a TLC plate and in a short time (a few hours). For analysis of lipids prepared from plant cells, two-dimensional TLC with two distinct solvent systems as to, e.g., pH is performed, since one-dimensional TLC fails to separate some lipid classes.

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1. Activate a pre-coated silica gel TLC plate by heating at 120°C for 1.5 h (see Note 4). 2. Cool it at room temperature. 3. Using a pencil on a TCL plate, outline an oblong spot of approximately 5  mm × 15  mm, which is 2.5  cm away from two neighboring sides of the plate (see Note 5). 4. Apply lipids dissolved in 20–50 ml of chloroform/methanol (2:1) to the spot with a microsyringe. 5. Develop the plate with the first solvent system, chloroform/ methanol/H2O (65:25:4, by vol.), until the front line of the solvent comes up to 1–2 cm away from the upper side of the plate. 6. Dry the plate by heating the backside with a hair dryer. 7. Develop the plate with the second solvent system, chloroform/methanol/25–28% ammonia solution (13:7:1, by vol.), until the front line of the solvent comes up to 1–2 cm away from the upper side of the plate. 8. Dry the plate by heating the backside with a hair dryer. 3.3. Detection and Identification of Respective Lipid Classes

Lipid classes separated on a TLC plate must be detected with a non-destructive reagent for further structural analyses, such as determination of compositions of constituent fatty acids. Primulin and I2 described in Subheading 3.3.1 are amenable to such study, as they are used for the detection of all lipid classes. Zones of the detected lipid classes on the TLC plate are outlined with a pencil for their later identification dependent on their mobility and also on staining by reagents that react with particular polar-head groups (see Subheading 3.3.2) (see Note 6). Figure 1 shows the detected lipid classes prepared from Chlamydomonas reinhardtii cells.

3.3.1. Non-destructive Reagents for Detection of All Lipid Classes

1. Spray primulin reagent onto the plate and dry it at room temperature.

3.3.1.1. Primulin Reagent

2. Illuminate the plate with a long-wave UV light (e.g., 365 nm) to detect blue spots of lipid classes.

3.3.1.2. I2 Vapor

1. Place a lump of I2 in a closed box to fill it with I2 vapor. 2. Allow the TLC plate in the box to stand until brown spots of lipid classes appear.

3.3.2. Destructive Reagents for Detection of Particular Lipid Classes 3.3.2.1. Dittmer–Lester Reagent

Outline lipid zones detected in advance by primulin, and treat the zones with Dittmer–Lester or anthrone reagent for detection of phospholipids and glycolipids, respectively, as described below. 1. Mix equal volumes of solutions A and B, and dilute it threefold with H2O. 2. Spray the reagent onto the plate to detect blue spots of phospholipids.

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Fig. 1. Separation of total lipids of C. reinhardtii cells into individual lipid classes. DGDG, digalactosyl diacylglycerol; DGTS, diacylglyceryltrimethylhomoserine; MGDG, monogalactosyl diacylglycerol; PE, phosphatidylethanolamine; PG, phosphatidylglycerol; PI, phosphatidylinositol; SQDG, sulfoquinovosyl diacylglycerol. 3.3.2.2. Anthrone Reagent

1. Spray the reagent onto the plate and heat it at 120°C to detect purple spots of glycolipids.

3.4. Determination of Fatty Acid Compositions and Contents of Lipids

There is no need to extract respective lipid classes from silica gels of the TLC plates for analysis of their constituent fatty acids. Fatty acid methyl esters are first produced according to the method described in Subheading  3.4.1, and are then analyzed by gas– liquid chromatography as in Subheading 3.4.2.

3.4.1. Preparation of Methyl Ester of Fatty Acids

1. Put 50 ml of an internal standard solution such as arachidic acid (1  mg/ml chloroform) into a test tube with a Teflonlined screw cap and dry it under N2 stream. This step is necessary for quantification of lipids, but is omitted for determination of only fatty acid compositions. 2. Scrape silica gels of lipid zones on TLC with the handle edge of a small spatula, and transfer them to the test tube with a piece of powder paper. Otherwise, transfer lipid fractions such as total lipids dissolved in chloroform/methanol (2:1, by vol.) to the test tube and dry it under N2 stream. 3. Add 2–3 ml of 5% hydrogen chloride methanol solution and close the tube tightly with a screw cap (see Note 7). 4. Heat it at 90°C for 3 h in a heating block with occasional agitation with a vortex mixer and then cool it at room temperature. 5. Add 2–3 ml of n-hexane and agitate it for 30 s with a vortex mixer.

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6. After standing it for separation into two phases, transfer the upper n-hexane phase containing fatty acid methyl esters into a test tube suitable for evaporation. 7. Evaporate the solvent with a rotary evaporator at 30–40°C. 8. Add 2–3 ml of n-hexane to the remaining lower phase in step 6 and agitate it for 30 s with a vortex mixer. 9. After standing it for separation, transfer the upper phase to the test tube in step 7, and evaporate the solvent. 10. Repeat steps 8 and 9. 11. Dissolve the methyl esters in 50 ml of n-hexane. 3.4.2. Analysis of Fatty Acid Methyl Esters with Gas–Liquid Chromatography

1. Set up a gas chromatograph equipped with a capillary column (see Note 8). Temperatures of injector and detector of the flame ionization are 250°C, while that of the column is 180°C. 2. Inject 1 ml of the methyl ester solution. 3. Record and store chromatograms with a chromatography data processor. Figure 2 shows a separation pattern of fatty acid methyl esters prepared from digalactosyl diacylglycetol of Chlorella kesslerii (see Note 9). Peak areas that correspond

Fig. 2. Separation of fatty acid methyl esters prepared from digalactosyl diacylglycerol of C. kesslerii. Fatty acyl groups are denoted by the number of carbon atoms and double bonds, e.g., 18:1(9) = oleoyl. aAn internal standard.

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to weights of fatty acid methyl esters are calibrated by respective molecular weights for determination of molar ratios and compositions of fatty acids. 4. Total fatty acid contents of each lipid are calculated through multiplication of a molar ratio of total constituent fatty acids to an internal standard by the known content of the internal standard. Lipid contents can thus be estimated by division of the total fatty acid contents by two. 3.5. Determination of Fatty Acid Compositions at sn-1 and sn-2 Positions of Individual Lipid Classes

Lipases that liberate fatty acids from lipids are classified into phospholipid-specific lipases and nonspecific lipases. Described below is the method of determination of fatty acid distribution between sn–1 and sn–2 positions with the use of nonspecific lipase from R. delemar that hydrolyzes ester bond at the sn-1 position of diacylglycerol moiety (2). 1. Scrape off silica gels containing a lipid class separated by TLC into a test tube. 2. Add 2–3 ml of chloroform/methanol (2:1, by vol.) and agitate the tube for 2 min for the lipid extraction. 3. Centrifuge at 1,500 × g for 5 min and recover the supernatant into a test tube suitable for evaporation. 4. Evaporate the solvent completely with a rotary evaporator. 5. Add 0.9 ml of 50 mM Tris–HCl, pH 7.2, and 0.05% Triton X-100, and agitate the tube with a vortex mixer. Sonicate the solution for 5 min until the lipids are completely dissolved. 6. Add 0.5 mg of lipase from R. delemar dissolved in 0.1 ml of 50 mM Tris–HCl, pH 7.2. 7. Incubate it at 37°C for appropriate times (e.g., 10  min for monogalactosyl diacylglycerol, 20 min for digalactosyl diacylglycerol, 30–60  min for sulfoquinovosyl diacylglycerol or phosphatidylglycerol, 90 min for phosphatidylethanolamine, and 360 min for phosphatidylcholine) (see Note 10). 8. Add 1 ml of chloroform/methanol (1:1 by vol.) and agitate it for 1 min to extract products by the lipase. 9. Recover the lower phase and evaporate the solvent. 10. Dissolve the lipids in 50  ml of chloroform/methanol (2:1 by vol.) and develop it using one-dimensional TLC, with chloroform/methanol/H2O (65:25:4, by vol.) as the solvent system. 11. Spray primulin reagent to detect sn-2-acyl-lysolipids, original lipids, and released fatty acids. The sn-2-acyl-lysolipids migrate slower than the original lipids, while fatty acids move faster, positioned close to the front line of the solvent. 12. Analyze fatty acid methyl esters of the sn-2-acyl-lysolipids by gas–liquid chromatography for determination of fatty acid

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compositions at the sn-2 positions, according to the method described in Subheading 3.4. Fatty acid compositions at the sn-1 positions can be estimated by comparison of fatty acid compositions between original lipids and sn-2-acyl-lysolipids.

4. Notes 1. Do not use plastic, but glass-made instruments for isolation and storage of lipids, since chloroform and methanol elute plasticizer of plastic instruments. 2. Samples for isolation of total lipids can be stored not at –20°C, but at –80°C, for repression of breakdown of lipid molecules by endogenous lipases. 3. A much larger pellet is needed to increase volumes of reagents for subsequent lipid isolation. 4. Activated TLC plates can be stocked in a desiccator, but activation just before performance of TLC is recommended, especially in places of high humidity. 5. The spot of lipids as an origin on the TLC plate should not be so big as to repress increases in the sizes of respective lipid zones. Large amount of silica gel will perturb subsequent fatty acid analysis by gas–liquid chromatography, since the TLC plate is contaminated with saturated fatty acids such as 16:0 and 18:0. 6. The reagents for the detection of lipids with particular polar head groups destroy lipid molecules, and thus are not useful for their subsequent structural analyses. 7. Be careful not to use a cracked tube, as such tubes explode during analyses. 8. Chromatograph not equipped with a packed column, but with a capillary column, is recommended owing to the high performance in the separation of almost all fatty acid methyl esters, including isomers such as 18:1(9) and 18:1(11). 9. Molecular species of fatty acid methyl esters can be identified through comparison of retention times on chromatograms with fatty acid methyl ester standards, which can be prepared by methanolysis of commercially available fatty acids. 10. Fatty acids at the sn-2 positions of the lysolipids will be transferred to the sn-1 positions and be hydrolyzed by the action of lipase. Thus, fatty acids to be analyzed should be sn-2-acyl-lysolipids, but not released fatty acids, and reaction should be stopped before complete breakdown of original lipids.

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References 1. Bligh, E.G. and Dyer, W.J. (1959) A rapid method of total lipid extraction and purification. Can. J. Biochem. Physiol. 37, 911–917. 2. Fischer, W., Heinz, E., and Zeus, M. (1973) Suitability of lipase from Rhizopus arrhizus

delemar for analysis of fatty acid distribution in dihexosyl diglycerides, phospholipids, and plant sulfolipids. Hoppe-Seyler’s Z. Physiol. Chem. 354, 1115–1123.

Chapter 10 Isolation and Purification of CP43 and CP47 Photosystem II Proximal Antenna Complexes from Plants Rafael Picorel, Miguel Alfonso, and Michael Seibert Abstract A method to isolate and purify CP43 and CP47 pigment–protein complexes from Photosystem (PS) II of higher plants is presented. The method has been developed in spinach, but it may also be valid for other plant species, since there is high PSII core complex homology in all plants. Core complex, obtained from highly enriched PSII membrane fragments (the extrinsic proteins were previously removed by Tris treatment), is used as starting material. The core complex is first treated with the chaotropic agent, LiClO4, and the nonionic detergent, n-dodecyl b-D-maltoside. After dialysis against buffer lacking detergent or chaotropic agent, the solubilized material is separated by anion-exchange chromatography using a TSK Toyopearl DEAE 650s column. CP43 complex does not bind to the column under these conditions and elutes along with free pigments and few other contaminants. When the eluate becomes colorless, the column is subjected to a 0–170-mM LiClO4 linear gradient. The main pigment elution band corresponds to the CP47 complex with some contaminants. To obtain pure preparations of CP43 and CP47 complexes, other chromatographic steps were developed. The CP43 material is passed through a S-Sepharose cation-exchange column at room temperature and then through a Q-Sepharose anionexchange column. After dialysis, the solution is passed through a new Q-Sepharose anion-exchange column at a different pH. The bound material is eluted with a 10–70-mM MgSO4 linear gradient, and the fractions with a prominent peak at 670 nm and a clear shoulder at 683 nm are combined. This constitutes the pure CP43 complex. The CP47 material from the first column is dialyzed, loaded onto a new TSK Toyopearl DEAE 650s column, and eluted with a 0–175-mM LiClO4 linear gradient. The fractions with a peak at 674.8 nm are combined and constitute the pure CP47 complex. Key words: Ion-exchange chromatography, CP43, CP47, Photosystem II, Pigment–protein complexes, Purification, Spectroscopy

1. Introduction Photosystem II is a membrane-bound, pigment–protein complex present in all oxygenic photosynthetic organisms, and it is composed of a large number of protein subunits and cofactors. The oxygen-evolving core complex (OECC), an integral part Robert Carpentier (ed.), Photosynthesis Research Protocols, Methods in Molecular Biology, vol. 684, DOI 10.1007/978-1-60761-925-3_10, © Springer Science+Business Media, LLC 2011

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of PSII, is surrounded by distal antenna complexes, including light-­harvesting complex (LHCII), chlorophyll protein (CP) 29, CP26, and CP24. It can be isolated from plants, algae, and cyanobacteria, and is composed of the D1-D2-cytochrome (Cyt) b559 reaction center complex (the site of primary charge separation), the CP43 and CP47 proximal antenna complexes, and the 33-kDa extrinsic protein. After Tris–HCl treatment of PSII-enriched membrane fragments at basic pH, the membrane-bound OECC loses three extrinsic proteins, and with them, the capability of splitting water (i.e., the isolated OECC material lacking the extrinsic proteins is then called the core complex). The names of the proximal antenna complexes are derived from the apparent molecular mass, determined by sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE), of the individual polypeptide present in each complex. The CP43 and CP47 complexes are encoded by the chloroplast genes, psbC and psbB, respectively. According to the three-dimensional structure of the OECC from cyanobacteria (1–3), CP43 is physically closer to D1 and CP47 to the D2 reaction center polypeptides. Both proximal antennae most probably function as a conduit for excitation energy transfer between the distal antenna complexes and the reaction center. However, other functions have also been attributed to them, including involvement in the assembly of an active oxygenevolving complex (4). The purification of native CP43 and CP47 complexes is necessary to assess their physical, chemical, and spectroscopic properties without contamination from other photosynthetic pigments or pigment–protein complexes. In this chapter, we describe a method to isolate and purify CP43 and CP47 after disruption of the core complex (isolated directly from Tris-treated PSII membrane fragments) by a combination of a chaotropic agent (LiClO4) and a mild detergent (DM), followed by separation using several ion-exchange chromatographic columns. Highly pure pigment–protein complexes result from this procedure.

2. Materials 1. PSII core complex from spinach (see Notes 1 and 2). 2. TSK Toyopearl DEAE 650s, a weak anion-exchange resin. 3. S-Sepharose, a strong cation-exchange resin. 4. Q-Sepharose, a strong anion-exchange resin. 5. A high-speed, refrigerated centrifuge and microfuge. 6. A high-performance liquid chromatography (HPLC) apparatus.

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7. A reversed-phase liquid chromatography column. 8. Unloaded chromatography columns. 9. n-Dodecyl b-D-maltoside (DM). 10. Buffer 1: 50 mM MES, pH 6.0, 150 mM NaCl, and 400 mM sucrose. 11. Buffer 2: 20  mM Bis-Tris, pH 6.0, 4  M LiClO4, and 15% (w/v) DM. 12. Buffer 3: 20 mM Bis-Tris, pH 6.0. 13. Buffer 4: 20 mM Bis-Tris, pH 6.0, and 0.05% (w/v) DM. 14. Buffer 5: 20  mM Bis-Tris, pH 6.0, 20  mM NaCl, 10  mM MgCl2, 1.5% (w/v) taurine, and 0.03% (w/v) DM. 15. Buffer 6: 50 mM Tris–HCl, pH 7.8, and 0.03% (w/v) DM.

3. Methods The basic protocols have been published in Alfonso et al. (5) and Dang et al. (6). All procedures are carried out at 4°C in the dark, unless otherwise specified. 3.1. Isolation of PSII Core Complex

1. Treat highly enriched PSII membrane fragments (60 mg total Chl) from spinach with 0.8 M Tris–HCl, pH 8.0, to remove the three extrinsic proteins associated with O2 evolution (see Notes 1 and 2). 2. Centrifuge at 40,000 × g for 20 min. 3. Resuspend the pellet in buffer 1 at 3 mg Chl/mL and obtain good-quality PSII core complex following the method of Ref. (7). This method is normally used to isolate OECC. The fact that the PSII membranes lack the extrinsic proteins does not affect the ability of this procedure to produce goodquality PSII core complex instead of OECC. 4. Mix the Tris-treated PSII membranes with an equal volume of buffer 2, and incubate for 15  min with occasional hand shaking (see Notes 2 and 3). 5. Dialyze the solubilized material in 10-kDa molecular cut-off dialysis tubing against buffer 3 for 2 h.

3.2. Purification of the CP43 Complex 3.2.1. TSK Toyopearl DEAE 650s, Weak AnionExchange Chromatography

1. Load the dialyzed material onto a TSK Toyopearl DEAE 650s weak anion-exchange column (1.6 × 7.5  cm), previously equilibrated with buffer 4, at a flow rate of 2 mL/min. 2. Wash the column with buffer 4 at a flow rate of 1 mL/min and collect 1-mL fractions of the eluate. The eluate contains CP43, free pigments, and some other contaminants.

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3. When the eluate becomes colorless, subject the column to a 0–175-mM LiClO4 linear gradient in buffer 4 at a flow rate of 0.5 mL/min and collect 0.5-mL fractions. The first greenish fractions correspond to some pigment–protein complex contaminants, including CP29, and the main green elution band at around 75  mM LiCLO4 corresponds to the CP47 complex. 4. The final elution band comes out at around 125 mM LiClO4, and it contains a mixture of CP47–D1D2Cyt b559 and D1D2Cyt b559 complexes. 5. The fractions of the main elution band that show a prominent room temperature absorption peak at around 675 nm and a small absorption band at around 620 nm (a vibronic band of Chl a) with similar absorbance to that at around 490 nm (the lowest energy carotenoid band) are collected and dialyzed overnight against buffer 4. This material will be used later to obtain purified CP47. 3.2.2. S-Sepharose, Strong Cation-Exchange Chromatography

1. Pass the material that does not bind to the TSK Toyopearl DEAE 650s column through a room-temperature, S-Sepharose column (1.0 × 10 cm), equilibrated with buffer 5.

3.2.3. Q-Sepharose, Strong Anion-Exchange Chromatography

1. Pass the material that does bind to the S-Sepharose column through a Q-Sepharose column (1.0 × 5  cm), equilibrated with buffer 5. 2. Dialyze the material that does not bind to the column overnight against buffer 6. 3. Load the dialyzed material onto a fresh Q-Sepharose column, equilibrated with buffer 6. 4. Wash the column with buffer 6 until the eluate becomes colorless. 5. Elute the green, bound material at a flow rate of 0.5 mL/min with a 10–70-mM MgSO4 linear gradient in buffer 6. 6. Collect 0.5-mL fractions and combine these with a prominent room-temperature absorption peak at 670  nm, a pronounced shoulder at 683 nm (see Note 4), and an absorption band at 620 nm that has a lower absorbance than the band at around 490 nm. Subsequently, dialyze the combined fraction pool twice for 1 h each against 1 L of buffer 6. 7. Concentrate the sample with 30-kDa cut-off Amicon Centricon tubes and keep frozen at −80°C (see Note 5). This is the purified CP43 complex.

3.3. Purification of CP47

1. Load the dialyzed, impure CP47 material from the first TSK Toyopearl DEAE 650s column onto another TSK Toyopearl DEAE 650s column (1.0 × 5.0 cm).

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2. Wash the column with buffer 4 until the eluate is colorless. 3. Apply a 0–175-mM LiClO4 linear gradient in buffer 4 and collect 0.5-mL fractions at a flow rate of 0.5 mL/min. 4. Take the room-temperature absorption spectrum of each fraction. Discard the fractions containing PSII reaction centers (those with a band at 542 nm). Pool those with a prominent peak at 674.8  nm and an absorption band at 620  nm that has an absorbance similar to that at around 490 nm. 5. Dialyze the pool against buffer 4, concentrate as convenient using 30-kDa cut-off Amicon Centricon tubes, and keep frozen at −80°C (see Note 5). This is the purified CP47. 3.4. Criteria to Assess the Quality of the Preparations

1. A rapid and simple method: room-temperature UV–Vis absorption spectra (Fig. 1). A high-quality preparation of CP43 should have a prominent peak at 670 nm and a distinct shoulder at 683 nm (see Note 4). The absorption intensity at around 620 nm due to Chl a is a little lower than that of the lowest energy carotenoid band at around 490 nm. A high-quality preparation of CP47 should have a maximum peak at 674.8 nm, and the absorbance intensity of the 620-nm band should be similar to that at around 490 nm.

Fig. 1. Room-temperature absorption spectra of isolated CP43 and CP47.

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2. Low-temperature absorption and fluorescence spectra. The low-temperature (4 K) absorption spectrum of CP43 shows a prominent peak at 669  nm, a shoulder at around 678  nm, and a small but distinct peak at 683 nm (i.e., the height of the 683-nm peak indicates the quality of the CP43 preparation). The low-temperature fluorescence spectrum of pure and native CP43 should have a single band at 685 nm. The low-temperature absorption spectrum of CP47 should have multiple bands at around 660, 670, 677, and 682 nm, with the maximum peak at 677 nm. The low-temperature fluorescence spectrum of pure and native CP47 should have a single band at 695 nm. 3.5. Additional Procedures to Analyze the Quality of the Preparations

1. Pigment analysis by HPLC. Pigments from CP43 or CP47 preparations can be extracted with 80% (v/v) cold aqueous acetone by sonicating the suspensions for 1  min. This releases pigments from the protein–detergent complexes, and the extracts are then centrifuged for 5 min in a microfuge. The white pellet, containing detergent and denatured protein, is discarded, and the pigment-containing supernatant is injected in a C-18, reversed-phase HPLC column (8). The chromatograms are monitored at 450 nm, and pure preparations show only two main peaks that correspond to Chl a and b-carotene. The appearance of Chl b and/or xanthophylls is a clear indication of contamination by other pigment–protein complexes. 2. Polypeptide composition. The polypeptide content of the CP43 or CP47 preparations can be analyzed by SDS-PAGE using 12% (w/v) acrylamide with 4 M urea. Coomassie Brilliant Blue stained gels should show a single band at 43 or 47 kDa for pure CP43 or CP47, respectively.

4. Notes 1. Good-quality PSII core complexes are a requisite for obtaining optimal results using the above-described procedures. The absorption spectrum of a good core complex should peak at around 674–675 nm, but cores with a peak maximum somewhat higher (up to 676 nm) are also acceptable for purification of CP43 and CP47 complexes. The exact peak maximum depends on the amount of contaminants that are often found in PSII core complex preparations. These contaminants are eliminated after the above-described, successive liquid chromatographic steps.

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2. Elimination of the extrinsic proteins from plant PSII membranes by treatment with 0.8 M Tris–HCl, pH 8.0, improves the yield of the PSII core complex extraction by the method described in Ref. (7). This treatment also increases somewhat the degree of disruption of the resulting core complex with LiCLO4 and DM. 3. The preparation of a buffer containing 4 M LiClO4 must be done at room temperature, and the buffer must be kept at that temperature until use. Lower temperatures result in the precipitation of LiClO4. After mixing one part of PSII core complex in buffer 1 (at 4°C) with one part of buffer 2 (at room temperature), the suspension is incubated at 4°C for 15 min with occasional hand mixing. All buffers containing DM should be prepared freshly before use. 4. When room-temperature absorption spectra are taken in the 350–750-nm region, it is helpful to examine expanded spectra in the 600–750-nm region. The CP43 shoulder at 683 nm is easier to see under these conditions (Fig. 1). If it cannot be seen, the CP43 preparation is of poor quality. 5. CP43 and CP47 complexes are stable at −80°C for a long time, and they can even be thawed and refrozen a few times without significant changes in their spectroscopic properties.

Acknowledgments This work was supported by the Ministry of Science and Innovation of Spain (Grants BFU2005-07422-C02-01 and AGL2008-00377) to R. P., and in USA by the Chemical Sciences, Geosciences, and Biosciences Division, Office of Basic Energy Sciences, US Department of Energy (M.S.). References 1. Kayami, N. and Shen, J. R. (2003) Crystal structure of oxygen-evolving photosystem II from Thermosynechococcus vulcanus at 3.7 Ǻ resolution. Proc. Natl Acad. Sci. USA 100, 98–103. 2. Loll, B., Kern, J., Zouni, A., and Biesiadka, J. (2005) Towards complete cofactors arrangement in the 3.0 Ǻ resolution structure of photosystem II. Nature 438, 1040–1044. 3. Guskov, A., Kem, J., Gabdulkhakov, A., Broser, M., Zouni, A., and Saenger W. (2009) Cyanobacterial photosystem II at 2.9-Ǻ resolution and the role of quinones, lipids, channels and chloride. Nat. Struct. Biol. 16(2), 334–342.

4. Bricker, T. M. and Frankel, L. K. (2002) The structure and function of CP47 and CP43 in photosystem II. Photosynth. Res. 72, 131–146. 5. Alfonso, M., Montoya, G., Cases, R., Rodríguez, R., and Picorel, R. (1994) Core antenna complexes, CP43 and CP47, of higher plant photosystem II. Spectral properties, pigment stoichiometry, and amino acid composition. Biochemistry 33, 10494–10500. 6. Dang, N. C., Zazubovich, W., Reppert, M., Neupane, B., Picorel, R., Seibert, M., and Jankowiak, R. (2008) The CP43 proximal antenna complex of higher plant photosystem II revisited: Modeling and hole burning study, I. J. Phys. Chem. B 112, 9921–9933.

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7. Ghanotakis, D. F., Demetriou, D. M., and Yocum, C. F. (1987) Isolation and characterization of an oxygen-evolving photosystem II reaction center core preparation and a 28 kDa Chl-a-binding protein. Biochim. Biophys. Acta 891, 15–21.

8. Larbi, A., Abadía, A., Morales, F., and Abadía, J. (2004) Fe resupply to Fe-deficient sugar beet plants leads to rapid changes in the violaxhantin cycle and other photosynthetic characteristics without significant de novo chlorophyll síntesis. Photosynth. Res. 79, 59–69.

Chapter 11 Preparation of Native and Recombinant Light-Harvesting Chlorophyll-a/b Complex Wolfgang Rühle and Harald Paulsen Abstract Procedures to isolate native light-harvesting chlorophyll-a/b complex (LHCIIb) and to reconstitute recombinant LHCIIb are described. Separation of trimeric from monomeric forms and free pigment by sucrose density-gradient ultracentrifugation can be applied to both native and reconstituted complexes. The preparations are characterized by their pigment composition, protein pattern, and spectral properties. Key words: Chlorophyll a/b-binding protein, Antenna protein, LHCII, Light-harvesting complex, Reconstitution in vitro

1. Introduction The apoprotein of the light-harvesting chlorophyll-a/b complex (LHCIIb) is the most abundant membrane protein in plants and one of the best-studied components of the photosynthetic apparatus. The latter partially owes to the fact that LHCIIb is more stable and easily isolated than most other pigment–protein complexes of the thylakoid membrane. The isolation procedure of Krupa et  al. (1) described here is easy and rapid. Although the resulting LHCIIb preparation has a low content of violaxanthin, a weakly bound carotenoid easily lost from native LHCIIb (2), the complexes are well suited for various spectroscopic studies (3, 4) and presumably also for crystallization (5, 6). Studies of structure–function relationship in LHCIIb are facilitated by the fact that the protein can easily be refolded in vitro. This allows mutation or altered pigment composition in recombinant versions of the complex. A technique to reconstitute

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LHCIIb from its apoprotein and pigments in  vitro has been devised by Plumley and Schmidt (7) and extended to bacterially expressed apoprotein (8). Subsequently, several slightly different procedures have been developed (9, 10). Other chlorophyll a/b complexes can be reconstituted with pigments using the same or a very similar protocol (11). In this chapter, we present a broadly applicable procedure leading to preparative amounts of trimeric recombinant LHCIIb.

2. Materials 2.1. Native LHCIIb

1. Pisum sativum (see Note 1). 2. Homogenization buffer: 50  mM Tricine–NaOH, pH 7.8, 0.4 M sorbitol. 3. Washing buffer: 5  mM EDTA–NaOH, pH 7.8, 50  mM sorbitol. 4. 5% (w/v) Triton X-100, stock solution (especially purified for membrane research) (Roche, Mannheim/Germany). 5. 1 M KCl. 6. 1 M MgCl2. 7. 0.5 M Sucrose. 8. Resuspension buffer: 50  mM Tricine–NaOH, pH 7.8, 100 mM sorbitol.

2.2. Recombinant LHCIIb

1. Bacterial strain expressing LHCIIb apoprotein Lhcb1/2 from pea (12, 13). 2. Solubilization buffer: 200 mM Tris–HCl, pH 9.0, 4% (w/v) Li-dodecylsulfate (LDS), 10  mM e-aminocapronic acid, 3 mM benzamidine, 25% (w/v) sucrose. 3. 1 M b-Mercaptoethanol. 4. 2 M KCl. 5. 0.1 M NiCl2. 6. 50 mM Tris–HCl, pH 7.5. 7. Octyl-b-d-glucopyranoside (OG) 10% (w/v) stock solution (Bachem, Heidelberg/Germany). 8. OG buffer: 1% (w/v) OG, 100 mM Tris–HCl, pH 9.0, 12.5% (w/v) sucrose. 9. TX buffer: 0.05% (w/v) Triton X-100, 0.01% (w/v) l-phosphatidyl-dl-glycerol dipalmitoyl (PG) (Avanti Polar Lipids, Alabaster/AL), 100 mM Tris–HCl, pH 7.5.

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10. Elution buffer: 0.05% (w/v) Triton X-100, 0.01% (w/v) PG, 10 mM Tris–HCl, pH 7.5, 300 mM imidazol. 11. Chelating Sepharose Fast Flow (GE Healthcare Bio-Sciences, Uppsala/Sweden). 2.3 Density Gradient

1. Gradient solution: 10 mM Tris–HCl, pH 7.0, 0.4 M sucrose, 0.06% b-d-dodecylmaltoside (Biomol, Hamburg/Germany). 2. Swing-out rotor (e.g., SW40, Beckman Coulter). 3. Ultracentrifuge tubes (e.g., for rotor SW40, Beckman Coulter).

2.4. Characterization of the Isolated LHCIIb

1. RP-18 HPLC column (Chromolith SpeedROD; Merck, Darmstadt/Germany). 2. Sodiumdodecylsulfate–polyacrylamide (SDS-PAGE) kit [12].

gel

electrophoresis

3. 50 mM HEPES, pH 7.5. 4. 10 mM phosphate buffer, pH 7.5. 5. 30-kDa size-exclusion membrane or G25 Sephadex gel filtration column.

3. Methods 3.1. Preparation of Native LHCIIb

Low-salt thylakoids are used as starting material for this procedure. Intrinsic proteins are solubilized by the non-ionic detergent Triton X-100, the LHCIIb is precipitated by K+ and Mg2+ salts, and then separated from other proteins by sedimentation through a sucrose layer. The resulting pellet consists mainly of trimeric LHCIIb and can further be separated from contaminating monomeric LHCIIb and minor antenna complexes by density-gradient ultracentrifugation (see Subheading 3.3).

3.1.1. Preparation of Low-Salt Thylakoids

All steps of thylakoid preparation are carried out at 4°C. 1. Homogenize about 300 g of young pea shoots (see Note 1) in a Waring blender with 1  l of ice-cold homogenization buffer. 2. Filter through four layers of cheesecloth and centrifuge for 5 min at 7,000 × g. 3. Wash the pellet once with 200 ml of cold washing buffer and centrifuge for 10 min at 10,000 × g. 4. Suspend the thylakoids in cold distilled water to a chlorophyll a + b concentration of 0.8 mg/ml (14) (see Note 2 for chlorophyll determination).

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3.1.2. Solubilization and Precipitation of LHCIIb

1. Add Triton X-100 from a 5% (w/v) stock solution to the ­low-salt thylakoids to obtain a final detergent concentration of 0.61% (w/v) (140 ml of 5% Triton per ml suspension; molar ratio of detergent/chlorophyll = 12/1). 2. Incubate the suspension at room temperature in the dark for 30 min with continuous stirring. 3. Centrifuge for 40 min at 30,000 × g at 4°C. 4. Discard the pellet and add KCl and MgCl2 from 1 M stock solutions in several portions to the supernatant to give final concentrations of 100 mM of KCl and 20 mM of MgCl2 (114 ml of KCl and 23 ml of MgCl2 per milliliter supernatant). 5. Gently stir the solution at room temperature for at least 10 min in the dark to precipitate the LHCIIb. 6. Prepare a cushion of 0.5  M sucrose in centrifuge tubes on which the suspension is carefully layered. The volume of the sucrose should be at least threefold the volume of the LHCIIb suspension. 7. Centrifuge at 10,000 × g for 10 min at 4°C (fixed-angle rotor is sufficient). 8. Resuspend the pellet in resuspension buffer and bring the solution to a chlorophyll concentration of 0.8 mg/ml. 9. Add Triton X-100 to give a molar detergent/chlorophyll ratio of 10/1 (116 ml of 5% (w/v) Triton per ml suspension). Stir for 10 min at room temperature. 10. Repeat precipitation with KCl and MgCl2 as above and centrifuge through a sucrose layer at 30,000 × g for 10 min. The chlorophyll a/b ratio should be below 1.4 after this step. Otherwise steps 8–10 should be repeated, with the exception that centrifugation is at 10,000 × g for 10 min. 11. Resuspend the pellet in resuspension buffer, and add KCl and MgCl2 as above but omit Triton. 12. Centrifuge through a sucrose layer for 10 min at 10,000 × g. 13. Resuspend the pellet in a small volume of distilled water and spin down for 3 min at 5,000 × g. 14. Repeat step 13 to remove any soluble detergent and ­remaining salt. 15. Adjust the suspension to a chlorophyll a + b concentration of 0.8 mg/ml and store the aliquots at –20°C or 16. Solubilize the native protein preparation for sucrose densitygradient ultracentrifugation of Subheading 3.3 with 1% dodecylmaltoside at a chlorophyll concentration of 1 mg/ml for 5 min on ice. Remove non-solubilized material by centrifugation at 15,000 × g for 2 min, and carefully layer the supernatant onto the gradient.

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3.2. Preparation of Recombinant LHCIIb

The procedure starts out from recombinant Lhcb1/2 protein carrying a C-terminal hexahistidyl (His) tag (13, 14). This protein is accumulated in the over-expressing bacteria as insoluble inclusion bodies. The pigments are isolated by acetone extraction of pea or spinach leaves. The procedure can be scaled up or down as needed. Recombinant LHCIIb, primarily formed as monomeric complexes, spontaneously oligomerizes into trimers when detergents are removed in the presence of phosphatidyl glycerol (13). This can conveniently be done by immobilizing the Histagged complexes on Ni2+-chelating affinity material (14).

3.2.1. Reconstitution of Recombinant Protein with Pigments

1. Dry an acetonic pigment extract (see Note 3) of pea or spinach leaves equivalent to 0.25 mg or 0.5 mg (see Note 4) of chlorophyll in a nitrogen stream and store aliquots for future use at –20°C. 2. Solubilize 250  mg of recombinant Lhcb1/2 apoprotein (inclusion bodies) in 300 ml of solubilization buffer and 30 ml of 1  M b-mercaptoethanol, and add water to make a final volume of 540 ml. 3. Incubate for 1 min in a boiling water bath. 4. Dissolve dried pigment extract corresponding to 0.25 mg of chlorophyll in 30 ml of ethanol. 5. Dissolve 0.25 mg of chlorophyll a in 30 ml of ethanol and add it to the pigment extract of step 4 (see Note 4). 6. Vortex the protein solution while adding the pigment solution of step 5 and vortex for at least another 30 s (see Note 5). Results are best when both protein and pigment solutions are at room temperature. 7. Add 75 ml of OG from stock solution to each vial and incubate for 10 min at room temperature to form mixed micelles of dodecylsulfate and octylglycoside. 8. Add 75 ml of 2 M KCl and place the solution on ice for 10 min to precipitate potassium dodecylsulfate. 9. Centrifuge at 4°C and 8,000 × g for 5 min and incubate the supernatant on ice for another 15 min. 10. Centrifuge at 4°C and 15,000 × g for 5 min. Aggregated pigments and rest of precipitated K-dodecylsulfate are removed by this step from the supernatant containing octylglycoside micelles with reconstituted complexes and free pigment.

3.2.2. Trimerization of Reconstituted LHCIIb on a Ni 2+-Chelating Column

1. Fill a 2-ml polypropylene chromatography column with Chelating Sepharose to obtain a sepharose bed of about 0.3– 0.5 ml, and wash thoroughly with water. 2. Add 0.5 ml of 0.1 M NiCl2 solution (the sepharose turns to a light green color).

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3. Add 1 ml of 50 mM Tris–HCl (pH 7.5) (the sepharose color changes to a light blue). 4. Equilibrate the column with 0.5 ml of OG buffer and transfer to 4°C. 5. Close the outlet of the column and mix the supernatant from the reconstitution experiment with the sepharose bed. Resuspend every 10 min for a total of 30 min to achieve complete binding of the protein to the Ni surface of the sepharose (see Note 6). 6. Wash the column with 3 × 0.5 ml of OG buffer. The brownish color of the eluate does not originate from pheophytin but from the reduction of some of the Ni2+ ions by mercaptoethanol. This does not interfere with the preparation. 7. Add 2 × 0.5 ml of TX buffer to change the detergent and provide the PG necessary for trimerization. 8. Finally, release the LHCIIb from the nickel–sepharose by the imidazole in 0.5 ml of elution buffer and collect all droplets with dark green color. The eluate contains a mixture of monomeric and trimeric LHCIIb which can be separated by sucrose density-gradient ultracentrifugation. 3.3. Separation of Monomeric and Trimeric LHCIIb

This procedure is suitable for both native and recombinant LHCIIb. 1. Prepare a sucrose density gradient by freezing the gradient solution in 12–14-ml ultracentrifuge tubes at –20°C and thawing it slowly at 4°C. 2. Remove the upper 500 ml of the gradient to provide a sharp step between the gradient and the sample. 3. Carefully load the gradient with solubilized native or reconstituted recombinant LHCIIb. No more than 500  mg of LHCIIb should be loaded; loading more will jeopardize the separation between monomeric complexes and unbound pigments; the separation of trimers is more robust. 4. Centrifuge at 200,000 × g for 16 h in a swing-out rotor. Figure 1 shows an example for the separation of monomeric from trimeric LHCIIb and free pigment in a preparation of native LHCIIb by sucrose density-gradient ultracentrifugation.

3.4. Characterization of the Isolated LHCIIb 3.4.1. Pigment Content

Table  1 is an example of yields and chlorophyll a/b ratios as a rough measure of purity after three precipitation steps. Chlorophyll concentrations are calculated from the absorbance of acetonic extracts (15). A more detailed analysis of the pigment composition in monomeric or trimeric LHCIIb is performed by extraction of the

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Fig. 1. Sucrose-density gradient separation pattern of a native LHCIIb preparation.

Table 1 Example of yield and chlorophyll a:b ratios as a rough measure of LHCIIb purity during successive precipitations Low-salt P1 = first P2 = second P3 = third thylakoids precipitate precipitate precipitate Total chlorophyll (mg)

  40.0

21.5

  7.6

0.9

Yield (%)

100.0

53.7

19.0

2.1

Chlorophyll a:b

   2.85

  2.33

  1.33

1.28

appropriate gradient band by 2-butanol and separation of pigments in an RP-18 HPLC column (8). Table  2 shows the pigment composition of the native and reconstituted trimeric LHCIIb bands. Lutein is used as a reference since two luteins are the most constant component of LHCIIb (16). The crystal structure of native LHCIIb reveals 14 chlorophylls (17, 18). Violaxanthin is only weakly bound and almost completely lost in this preparation (see Note 7).

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Table 2 Pigment stoichiometry of native and recombinant trimeric LHCIIb reconstituted with a chlorophyll a:b ratio of 3:1 and 7:1 Nx nLHCIIb

Vx

Lu Chlb

Chla

b-Car

0.8 ± 0.1 0.1 ± 0.1 2.0 6.4 ± 0.4 7.9 ± 0.7 nd

3:1 Reconstitution 0.8 ± 0.2 nd

2.0 7.0 ± 0.5 7.1 ± 0.6 nd

7:1 Reconstitution 0.9 ± 0.1 nd

2.0 6.4 ± 0.4 7.7 ± 0.4 nd

Data are in mol/2 mol lutein and represent means (±SD) of nine preparations nd not detectable

Fig. 2. Protein pattern at different steps of purification as revealed by a fully denaturing SDS-PAGE. Thyl  thylakoids; P1 first precipitation; P2 second precipitation. A Laemmli gel (17) (15% polyacrylamide) was stained with Coomassie.

3.4.2. Protein Pattern

The purification of native LHCIIb in Subheading 3.1 is demonstrated in Fig. 2 by a Coomassie-stained denaturing SDS-PAGE. Lane Thyl results from thylakoids and lane P1 from the first precipitation. After the second precipitation (P2), almost all other proteins except the apoproteins of LHCIIb and an unpigmented 65-kD component are removed. However, silver-stained gels show that there are still traces of proteins other than LHCIIb apoproteins, which shows the limitation of the procedure. Salt

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precipitation is suitable for preparing large amounts of LHCIIb free of other pigment-binding proteins within a short time but for further demands in purity, other principles of separation such as isoelectric focusing (3) are required. 3.4.3. Spectroscopy 3.4.3.1. Absorption Spectrum

The room-temperature absorption spectrum of LHCIIb has chlorophyll a maxima at 435 and 675 nm and chlorophyll b maxima at 471 and 650  nm as well as some shoulders in the blue region of the spectrum resulting from xanthophylls (see Fig. 3). For low-temperature absorption spectroscopy, the sucrosegradient bands were mixed with an 80% glycerol solution buffered by 50 mM HEPES (pH 7.5) to give a final glycerol content of 60% and subsequently measured in liquid nitrogen. Qy absorption forms of chlorophyll are clearly resolved at 649, 661, 670, and 675  nm. The derivative spectra reveal additional bands at 644, 654, 665, and 678 nm. Further absorption bands in native and reconstituted LHCIIb have been reported to appear at 4 K (4). The low-temperature absorption spectrum of the reconstituted protein complex depends partially on the chlorophyll a/b ratio in the pigment extract used for the reconstitution. At a chlorophyll a:b ratio of 3:1 in the pigment extract, some chlorophylls b occupy chlorophyll a-binding sites. Therefore, the chlorophyll a:b ratio is about 0.9, instead of around 1.3 in the native complex; the shortwave absorbing forms at 649, 654, and 661 nm are increased; and the two long-wavelength absorbing forms at 670 and 675 nm are weaker compared to the native complex (10). Upon reconstitution with a chlorophyll a:b ratio of 7:1, the absorption spectrum largely resembles that of the native complex.

Fig. 3. 77 K absorbance spectra of native trimeric LHCIIb (solid ) and its fourth derivative compared to reconstitutions with pigment extracts of 3:1 (dashed ) or 7:1 (dotted ) chlorophyll a:b ratios. Inset : identical fluorescence emission spectra of native and reconstituted trimeric LHCIIb at 77 K with chlorophyll b excitation at 470 nm.

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3.4.3.2. Fluorescence Emission

Low-temperature fluorescence spectroscopy was performed under the same conditions as absorption spectroscopy (inset in Fig.  3). Excitation at both 470  nm (chlorophyll b) and 430 or 410 nm (chlorophyll a) leads to an emission band at 680 nm (chlorophyll a) with no shoulder at 660 nm, the emission wavelength of chlorophyll b, indicating complete energy transfer from chlorophyll b to chlorophyll a. There is no difference between the native and reconstituted complexes. Significant formation of LHCIIb aggregates would be indicated by a fluorescence emission band at around 700 nm (19).

3.4.3.3. CD Spectrum

The UV region of CD spectra can provide information on the secondary protein structure; specifically, the bands between 200 and 240 nm present structural information on the amount of a helical secondary protein structure (10). There is almost no difference between the UV-CD signals of native and reconstituted LHCIIb. For this measurement, it is necessary to remove the sucrose and Tris from the gradient bands. This can be done either by repeated dilutions with a 10 mM phosphate buffer (pH 7.5) and subsequent ultrafiltration steps by centrifugations at 5,000 × g with a 30-kDa size-exclusion membrane, or by buffer exchange on a G25 Sephadex gel filtration column. The CD spectrum in the visible domain (see Fig. 4) of trimeric LHCIIb differs from that of the monomeric form by a significant shoulder at 642  nm (20) and a prominent minimum at 474 nm (6).

Fig. 4. CD spectra of monomeric reconstituted (dotted ), trimeric reconstituted (dashed ), and trimeric native LHCIIb (solid ). All spectra are divided by their absorbance at 675 nm for normalization. The two upper spectra are offset by 20 and 40 mdeg to separate them from each other.

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4. Notes 1. The pea plants should be 10–14 days old. Keep them in darkness for 12 h before starting the preparations to prevent starch accumulation. Avoid stems when harvesting. 2. Prepare an 80% acetonic extract of the thylakoids or protein complexes. Measure absorbance at 663.6, 646.6, and 750 nm. Subtract any absorbance at 750  nm from the 663.6 to 646.6 nm values. Calculate chlorophyll concentrations from Porra’s (15) equations considering dilution factors: chlorophyll a [mg/ml] = 12.25 × A663.6 − 2.55 × A646.6; chlorophyll b [mg/ml] = 20.31 × A646.6 − 4.91 × A663.6. 3. The preparation of a total pigment extract (8) is described briefly: Extract a pellet of thylakoids (prepared as described in Subheading 3.1.1) with acetone and centrifuge at 5,000 × g for 10 min. Partition the pigments into diethyl ether (0.25 volumes), freshly distilled over NaOH pellets, by adding an equivalent volume of 1.3 M NaCl. Repeat ether extractions, combining the ether fractions, as long as the lower acetone/ NaCl phase remains colored. Remove water from the ether fractions by freezing at –20°C and subsequent filtration. Evaporate the ether to dryness in vacuo. Solubilize the pigments in acetone, determine chlorophyll content (15), dry aliquots of 0.25 or 0.5  mg chlorophyll a + b in a nitrogen stream, and store at −20°C. Usually these preparations have a chlorophyll a:b ratio of about 3:1 (see Note 4). 4. For many applications, it is sufficient to use a chlorophyll a:b ratio of 3:1, which is the usual composition of thylakoids. In this case, you can replace the chlorophyll a addition in step 5 by an equal amount of total pigment extract. However, if the reconstitution is to match native LHCII with regard to the correct occupation of chlorophyll a- and chlorophyll b-binding sites, more chlorophyll a needs to be added to the reconstitution mixture. A chlorophyll a:b ratio of 7:1 avoids the occupation of some binding sites of chlorophyll a by chlorophyll b, which is often the case when starting out at a 3:1 ratio. Chlorophyll b-deficient thylakoids may also be an alternative source for a total pigment extract with increased chlorophyll a if you do not have access to a preparative HPLC. 5. It is crucial to avoid local supersaturation of the pigments before sequestering them in LDS micelles since this would lead to chlorophyll aggregation. Use 10-ml test tubes or vials large enough for vigorous vortexing without the need to close the lid. After vortexing, the samples may be transferred to 1.5-ml Eppendorf vials for the next steps.

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6. If the column is equipped with a flow control, the sample can also slowly be loaded to the packed column within 30 min. The washing and elution steps to be carried out are as described earlier. This will increase the binding capacity of the column and can lead to a higher LHCIIb concentration in the eluate. 7. Ruban et al. (2) published a procedure which yielded LHCIIb still containing substantial amounts of violaxanthin. They solubilize BBY particles or thylakoids with dodecylmaltoside and separate an “antenna band = A-band” by sucrose-gradient centrifugation. This A-band is then solubilized with low concentrations of dodecylmaltoside and separated with a second sucrose-gradient centrifugation. However, this procedure is only suitable for analytical purposes because yields are low.

Acknowledgments The authors thank Dr. Stephan Hobe and Dr. Volkmar Schmid for critically reading the manuscript. Work in the authors’ laboratory was funded by Deutsche Forschungsgemeinschaft (Pa324/5-3). References 1. Krupa, Z., Huner, N. P. A., Williams, J. P., Maissan, E., and James, D. (1987) Development at cold-hardening temperatures. Plant Physiol. 84, 19–24. 2. Ruban, A. V., Lee, P. J., Wentworth, M., Young, A. J., and Horton, P. (1999) Determination of the stoichiometry and strength of binding of xanthophylls to the photosystem II light ­harvesting complexes. J. Biol. Chem. 274, 10458–10465. 3. Bassi, R., Silvestri, M., Dainese, P., Moya, I., and Giacometti, G. M. (1991) Effects of ­non-ionic detergent on the spectral properties and  aggregation state of the light-harvesting ­chlorophyll a/b protein complex (LHCII). J. Photochem. Photobiol. B Biol. 9, 335–354. 4. Rogl, H., Schödel, R., Lokstein, H., Kühlbrandt, W., and Schubert, A. (2002) Assignment of spectral substructures to pigment-binding sites in higher plant light-harvesting complex LHC-II. Biochemistry 41, 2281–2287. 5. Kühlbrandt, W., Wang, D. N., and Fujiyoshi, Y. (1994) Atomic model of plant light-harvesting complex by electron crystallography. Nature 367, 614–621.

6. Hobe, S., Prytulla, S., Kühlbrandt, W., and Paulsen, H. (1994) Trimerization and crystallization of reconstituted light-harvesting chlorophyll a/b complex. EMBO J. 13, 3423–3429. 7. Plumley, F. G. and Schmidt, G. W. (1987) Reconstitution of chlorophyll a/b ­light-harvesting complexes: Xanthophylldependent assembly and energy transfer. Proc. Natl. Acad. Sci. U.S.A. 84, 146–150. 8. Paulsen, H., Rümler, U., and Rüdiger, W. (1990) Reconstitution of pigment-containing complexes from light-harvesting chlorophyll a/b-binding protein overexpressed in E. coli. Planta 181, 204–211. 9. Paulsen, H. and Schmid, V. H. R. (2002) Analysis and reconstitution of chlorophyll proteins, In Heme, Chlorophyll, and Related Molecules: Methods and Protocols (Smith, A. G. and Witty, M., eds.), Humana Press, Totowa, NJ, pp. 235–254. 10. Yang, C. H., Horn, R., and Paulsen, H. (2003) Light-harvesting chlorophyll a/b complex (LHCIIb) can be reconstituted in vitro from its completely unfolded apoprotein. Biochemistry 42, 4527–4533.

Preparation of Native and Recombinant Light-Harvesting Chlorophyll-a/b Complex 11. Schmid, V. H. R. (2008) Light-harvesting complexes of vascular plants. Cell. Mol. Life Sci. 65, 3619–3639. 12. Laemmli, U. K. (1970) Cleavage of structural proteins during the assembly of the head of bacteriophage T4. Nature 227, 680–685. 13. Kosemund, K., Geiger, I., and Paulsen, H. (2000) Insertion of light-harvesting chlorophyll a/b protein into the thylakoid – Topographical studies. Eur. J. Biochem. 267, 1138–1145. 14. Rogl, H., Kosemund, K., Kühlbrandt, W., and Collinson, I. (1998) Refolding of Escherichia coli produced membrane protein inclusion bodies immobilised by nickel chelating chromatography. FEBS Lett. 432, 21–26. 15. Porra, R. J., Thompson, W. A., and Kriedemann, P. E. (1989) Determination of accurate extinction coefficients and simultaneous equations for assaying chlorophylls a and b extracted with four different solvents: Verification of the concentration of chlorophyll standards by atomic absorption spectroscopy. Biochim. Biophys. Acta 975, 384–394. 16. Hobe, S., Niemeier, H., Bender, A., and Paulsen, H. (2000) Carotenoid binding sites

17.

18.

19.

20.

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in LHCIIb – Relative affinities towards major xanthophylls of higher plants. Eur. J. Biochem. 267, 616–624. Liu, Z., Yan, H., Wang, K., Kuang, T., Zhang, J., Gui, L., An, X., and Chang, W. (2004) Crystal structure of spinach major lightharvesting complex at 2.72 Å resolution. Nature 428, 287–292. Standfuss, J., Terwisscha van Scheltinga, A. C., Lamborghini, M., and Kühlbrandt, W. (2005) Mechanisms of photoprotection and non-photochemical quenching in pea lightharvesting complex at 2.5 Å resolution. EMBO J. 24, 919–928. Vasilev, S., Irrgang, K. D., Schrötter, T., Bergmann, A., Eichler, H. J., and Renger, G. (1997) Quenching of chlorophyll a fluorescence in the aggregates of LHCII: Steady state fluorescence and picosecond relaxation kinetics. Biochemistry 36, 7503–7512. Gülen, D., Knox, R., and Breton, J. (1986) Optical effects of sodium dodecyl sulfate treatment of the isolated light harvesting complex of higher plants. Photosynth. Res. 9, 13–20.

Chapter 12 Isolation and Characterization of Lamellar Aggregates of LHCII and LHCII-Lipid Macro-assemblies with Light-Inducible Structural Transitions Ilian Simidjiev, Zsuzsanna Várkonyi, Petar H. Lambrev, and Gyo˝  zo˝   Garab Abstract We describe the method of isolation of loosely stacked lamellar aggregates of LHCII that are capable of undergoing light-induced reversible structural changes, similar to those in granal thylakoid membranes (LHCII, the main chlorophyll a/b light-harvesting antenna complex of photosystem II). This unexpected structural flexibility of the antenna complexes depends largely on the lipid content that is retained during the isolation. As revealed by circular dichroism, in lipid–LHCII aggregates, the pigment–pigment interactions are very similar to those in the thylakoid membranes, while they differ significantly from those in solubilized trimers. The essence of the procedure is to adjust – for the plant material used – the proper conditions of detergent solubilization and purification that are mild enough for the associated lipids but provide sufficient purity. Microcrystals and most other LHCII preparations, which are more delipidated, are not capable of similar changes. The light-induced structural reorganizations can be enhanced by the addition of different thylakoid lipids, which – depending on the lipid species – also lead to the transformation of the lamellar structure. The preparation of different LHCII-lipid macro-assemblies is also described. Both in structurally flexible LHCII preparations and in thylakoids, the changes originate from a thermo-optic effect: fast local thermal transients, T-jumps, due to the dissipation of the (excess) excitation energy, which lead to elementary structural transitions in the close vicinity of the dissipating centers. This can occur because thylakoids and structurally flexible LHCII assemblies, but, e.g., not the microcrystals, exhibit a thermal instability below the denaturation temperature, and thus (local) heating leads to reorganizations without the loss of the molecular architecture of the constituents. We also list the main biochemical and biophysical techniques that can be used for testing the structural flexibility of LHCII, and discuss the potential physiological significance of the structural changes in light adaptation and photoprotection of plants. Key words: Chloroplasts, Dissipation, Excess excitation energy, Grana, LHCII, Light adaptation, Light-harvesting antenna, Lipid–protein interactions, Photoprotection of plants, Structural flexibility, Thermo-optic effect, Thylakoid lipids

Robert Carpentier (ed.), Photosynthesis Research Protocols, Methods in Molecular Biology, vol. 684, DOI 10.1007/978-1-60761-925-3_12, © Springer Science+Business Media, LLC 2011

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1. Introduction 1.1. Basic Observations

LHCII is the main chlorophyll a/b light-harvesting antenna complex of photosystem II (PSII). It is the most abundant ­pigment–protein complex in the biosphere. Its structure is known at near-atomic resolution (1). The primary function of this complex is to absorb light and transfer the excitation energy toward the reaction centers. By mediating the stacking of granal thylakoid membranes (2), LHCII is also known to stabilize the granal ultrastructure; its high self-aggregation capability explains the lateral segregation (sorting) of the two photosystems between the granum and stroma membranes (3). LHCII also participates in multilevel regulatory processes, thermal and light adaptation of plants (4, 5). LHCII, when isolated with mild detergent treatments (6, 7), readily assembles in loosely stacked lamellar aggregates, which – albeit less marked than in microcrystals – exhibit a significant degree of long-range chiral order of their chromophores (8) and form ordered 2D arrays (9). These lipid-enriched preparations, in contrast to more delipidated samples, such as microcrystals, are capable of undergoing light-induced reversible structural reorganizations (10, 11), which closely resemble the structural changes in the native thylakoid membranes (12) (Table 1). In LHCII, the rate of these changes is linearly proportional to the light intensity. In thylakoids, the changes also exhibit a similar, approximately linear light-intensity dependence above the saturation of photosynthesis. Hence, this reversible reaction “measures” the light intensity and by this means, the antenna system can react proportionally and reversibly to excess excitation.

1.2. Nature and Mechanism of the Structural Changes

As concerns the nature of the structural changes, a substantial degree of monomerization of LHCII has been shown to be involved both in vivo and in vitro. As for the physical mechanism, these reactions have been accounted for by a novel biological thermo-optic effect (13, 14). As indicated by chlorophyll fluorescence kinetic analyses, the structural changes also bring about alterations in the photophysical pathways, most notably in the regulated quenching of the singlet excited state of chlorophyll a molecules in the lightharvesting antenna (15–17). It seems likely that similar changes are responsible for the light regulation of LHCII phosphorylation at the substrate level (18). Monomerization may also be important in the accessibility of LHCII to proteases (19). In general, local structural rearrangements in the antenna may provide the system with a structural flexibility without perturbing the structural stability of the membranes. In granal thylakoids, this type of reorganizations of LHCII includes (1) the unstacking of membranes, (2) a lateral disorganization of the chiral macrodomains, and (3) monomerization of the LHCII trimers (20).

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Table 1 Comparison of microcrystalline (type IV) and delipidated disordered (type III) LHCII aggregates with loosely stacked lamellar aggregates (type II) of LHCII (cf. 7) and LHCII-containing thylakoid membranes

Feature

LHCII microcrystals, delipidated aggregates

LHCII lamellar aggregates

Thylakoids

References

Strong (weaker than in microcrystals)

Strong

(7, 8)

None Light-induced reversible DCD, or other structural changes

Yes, fully reversible up to 500 W/m2

Yes, fully reversible (7, 10, 17, 18, 24) up to 500 W/m2

Lipid content (rel. units)

60

330

500

(7)

Lipid enhancement of DCD

Not possible

Yes

No data

(11)

DF (reversible fluorescence quenching)

Weak

Strong

Non-photochemical (5, 15–17, 24) quenching

Light-induced reversible Mg2+release

Not detectable

2.5 per trimer

Yes, non-saturable (14) up to 500 W/m2

Thermal instability

High stability below 65°C

Breakdown of macrodomains: 40–50°C

Breakdown of macrodomains: 45–55°C

Light instability (>1,000 mE/m2 s, 15 min)

Very resistant at all temperatures up to 60°C

High instability (above 20–30°C)

High instability (13, 14, 20) (above 25–30°C)

Long-range order of chromophores (psi type CD)

Very strong in microcrystals; none in delipidated aggregates

(13, 14, 17)

500 W/m2 ≅ 2,700 mE/m2 s (of red photons)

1.3. LHCII-Lipid Macro-assemblies

The light-induced structural changes depend largely on the presence of lipids (7), and the addition of isolated thylakoid lipids enhances substantially the DCD (11). Freshly isolated LHCII (type II), which is prepared using the procedure described here, assembles in lamellar aggregates of loosely stacked sheets. This structure is retained after the addition of phosphatidylglycerol (PG) or sulfoquinovosyldiacylglycerol (SQDG). Upon the addition of digalactosyldiacylglycerol (DGDG), closed vesicles are

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formed, which stack to each other; upon the (gradual) addition of monogalactosyldiacylglycerol (MGDG), the loosely stacked lamellae are (gradually) transformed into onion-type structures or into stacked bundles of membranes (11). When disordered large aggregates (that contain less lipids than lamellar aggregates, but more than microcrystals) are to be restored by lipids, usually flat multilamellar arrays are obtained with all lipids, including MGDG (9); these, however, are not capable of undergoing light-induced reversible rearrangements (as measured by DCD of their so-called psi-type bands that originate from the long-range chiral order of the chromophores).

2. Materials All buffers should be kept in a refrigerator and used within a month. 2.1. Buffers

1. Buffer A: 50 mM Tricine–NaOH, pH 7.8, and 0.4 M sorbitol. 2. Buffer B: 20 mM Tricine–NaOH, pH 7.8, 5 mM EDTA, and 50 mM sorbitol. 3. Buffer C: 20 mM Tricine–NaOH, pH 7.8. 4. Buffer D: 0.5 M sucrose cushion equilibrated with 20 mM Tricine–NaOH, pH 7.8. 5. Buffer E: 50  mM Tricine–NaOH, pH 7.8, and 100  mM sorbitol.

2.2. Stock Solutions

1. 20% Triton-X 100 (Fluka) buffered with 20  mM Tricine– NaOH, pH 7.8. Store it at 4°C for up to 2 months in a dark brown bottle. 2. 2 M KCl. 3. 1 M MgCl2.

3. Methods 3.1. Isolation of LHCII

We use pea or spinach leaves with good turgor and dark green color, but other plants might be equally good. For instance, Krupa et al. (6) used rye; our method is based on their procedure, as described earlier (7). 1. Take 50–60 g of fresh pea leaves (or about 400 g of spinach), wash it with cold water and then in distilled water, and place in dark at 4°C for at least 2 h. Homogenize with a mixer in 250 ml of buffer A (see Note 1).

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2. Filter the homogenate through four layers of unused cheesecloth. Do not wash the cloth because any detergent may influence the structure of the isolated LHCII. 3. Centrifuge at 5,000 × g for 5 min. 4. Discard the supernatant and suspend the pellet in buffer B. Use medium-size brush for crude homogenization of the pellet, and then use Potter homogenizer. Be sure that all the material is finely homogenized, without visible big particles (see Note 2). 5. Centrifuge at 10,000 × g for 10 min. Discard the supernatant. 6. Resuspend the pellet in cold buffer C to obtain an absorbance of about 1 in a 0.2-mm cuvette at 680 nm (By this means, one can quickly adjust the chlorophyll content) (see Note 3). 7. Add Triton-X 100 (Fluka), from the solution, to obtain a final concentration of 0.7–0.9%. This is required for lamellar aggregates (type II). To obtain LHCII microcrystals (type IV – delipidated form), the final Triton concentration has to be 1.1–1.3% (see Note 4). 8. Stir the solution on ice continuously for at least 45 min, until the visible clouds disappear. Continue stirring up to 60 min if clouds are present (see Note 5). 9. Centrifuge at 30,000 × g for 40 min. The insoluble material will sediment, and the supernatant will predominantly contain LHCII (see Note 6). 10. Discard the pellet and add to the supernatant 20–25 mM of MgCl2 and 100–150  mM of KCl from the stock solutions. Stir gently to precipitate the crude LHCII. It takes about 5 min to see clouds appearing in the suspension. Then, continue to stir for another 10–15 min (see Notes 7 and 8). 11. The suspension is to be layered on buffer D. In order to wash out the detergent, the volume of this buffer has to exceed three times the volume of the LHCII suspension. Centrifuge at 10,000 × g for 15 min. 12. Resuspend the pellet in buffer E to obtain an absorbance of 0.8–1.0 in a 0.2-mm cuvette. 13. Add Triton-X 100 up to 0.2% final concentration, from 20% stock solution. Take the suspension in a screw-cap tube and shake it; clouds will appear (best seen in reflected light). Shake until the clouds partially disappear (see Note 9). 14. Precipitate as in step 10. Stir the suspension gently for about 15 min. 15. Lay the suspension on sucrose cushion (buffer D) and centrifuge at 30,000 × g for 40 min.

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16. For final purification, suspend the pellet in buffer E and add 20 mM of MgCl2 and 100 mM of KCl, from the stock to reach these final concentrations. Triton is omitted in this step. Lay the suspension on 0.5  M Tricine-buffered sucrose cushion (buffer D) and centrifuge at 10,000 × g for 10 min. The final purification could be omitted if the chlorophyll a/b ratio is between 1 and 1.3. Then, the solution is transferred to an Eppendorf tube, centrifuged at 3,000 × g for 5 min in an Eppendorf centrifuge, and resuspended in buffer C. This should be repeated three times. Finally, the LHCII is homogenized again in buffer E and the chlorophyll concentration and chlorophyll a/b are measured according to the procedure described by Arnon (21). 3.2. Preparation of LHCII–Lipid Macroassemblies

To obtain different thylakoid lipids, used for LHCII incorporation, total lipid extraction from thylakoid membranes was performed according to the procedure described by Bligh and Dyer (22), followed by separation of different lipid classes on thin layer chromatography (TLC) plates according to the procedure of Vigh et  al. (23). The lipids were stored in hexane containing 0.05% bromhydroxytoluene (BHT). The preparation of LHCII–lipid complexes was as follows: 1. Take the needed lipid concentration, transfer to a sharp-conical glass tube, and evaporate the hexane under nitrogen stream. 2. Add buffer C to the dry lipid, fill the tube with nitrogen or argon, and then close it with cap or Parafilm “M”. 3. Vortex vigorously until the lipids are completely dissolved. Usually SQDG, PG, and DGDG are dissolved upon 4–5 min of vortexing. During that time, place the tube on ice several times for about 30  s each time. Change vortexing and ice during the whole procedure of dissolving. Store the dissolved lipid mixture on ice. 4. To dissolve MGDG, add 0.05–0.08% Triton X-100 from the stock solution to buffer C. Then vortex vigorously for about 30 s, transfer the tube to ice for 30 s, and sonicate the suspension in a water bath sonicator for another 30 s. Repeat these steps till all lipid is dissolved, usually four to five times. 5. Add LHCII preparation in the desired amount to the lipid mixture, fill the tube with nitrogen or argon, close it, and vortex mildly for about 20 s. Store the mixture on ice and use it within 4 h.

3.3. Circular Dichroism Measurements

CD can be measured, e.g., in a Jobin-Yvon CD6 dichrograph (Fig. 1). It is perhaps the easiest and quickest test for the quality

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Fig. 1. Circular dichroism spectrum of loosely stacked lamellar aggregates of LHCII (type II) and microcrystalline LHCII aggregates (type IV).

of the preparation (7). The samples are usually placed at a ­distance of 5 cm from the photomultiplier. This can vary, but keeping the same geometry is important because of the involvement of differential scattering. Samples of 20 mg/ml of chlorophyll content can be measured in glass cuvettes with 1-cm optical path length. The presence of light-induced CD changes, DCD, is a clear indication of type II LHCII. This can be measured with a side illumination attachment. In our measurements, also shown in Fig. 2, the samples were illuminated at 90° with respect to the measuring beam. The actinic light from a 600-W tungsten halogen lamp passed through a heat filter (10 cm water) and 620-nm cut-off Corning 2-64 red filter, and was directed to the sample with a light guide. Stray light from the actinic beam was blocked with a broadband Corning 4-96 blue filter. Before CD measurements, LHCII samples were kept in dark, at room temperature for a few minutes. CD was continuously monitored at 495 nm. This is more convenient than in the red region, where fluorescence transients might contribute to the signal. (For monitoring CD in the red region, the two Corning filters are interchanged.) Since DCD is very sensitive to temperature, similar to DCD in thylakoid membranes (24), it is essential that the light-induced changes be recorded in the optimum temperature range, between 20 and 30°C. Note that DCD in some preparations exhibits

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Circular dichroism at 495 nm, E-4

−6.0 −6.5 −7.0 −7.5 −8.0

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Time (sec) Fig. 2. Light-induced CD changes in type II LHCII.

opposite sign. This is difficult to predict and not well understood. It seems that in all cases, the changes are between two states. This bistability may thus explain variations in the sign, which does not seem to affect the most commonly studied parameters, such as temperature and light-intensity dependencies, trimer-to-monomer transitions. 3.4. Lamellar Aggregates Mimicking the Native Thylakoid Membrane

In lamellar aggregates of LHCII as well as in the native thylakoid membranes, the pigments are organized in an ordered manner over distances of several hundred nanometers and as a result, these assemblies usually feature a strong psi-type CD signal (8, 12). The psi-type CD not only has a very large magnitude but is also extremely sensitive to the macro-arrangement of the chromophores (25, 26) and can be easily abolished, for example, by washing the thylakoid membrane suspension in a hypotonic and low-salt buffer, or decreasing the size of the LHCII aggregates (8). Under these conditions, the more structured CD bands, originating from excitonic interactions, become more clearly discernible. These bands provide a signature for the pigment organization at a lower structural level, since excitonic coupling can be formed within the individual pigment–protein complex or between neighboring complexes. Lamellar aggregates of LHCII having small or no psi-type CD are prepared as described in Subheading 3.1, except that at step 13 (second solubilization) Triton is added at a concentration of 0.5–0.7% and the final resuspension is in a buffer containing

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Circular dichroism (rel. u.)

1.0 0.5 0.0 −0.5 −1.0 −1.5 420 440 460 480 500 520

640 660 680 700

Wavelength (nm) Fig. 3. Circular dichroism spectra of lamellar disordered LHCII aggregates (solid line), thylakoid membranes in low-salt hypotonic buffer (dashed line), and LHCII trimers, solubilized in 0.1% n-dodecyl-b,d-maltoside (dotted line). The spectra are normalized to the negative maximum at 650 nm.

20 mM of Tricine (pH 7.8) without sorbitol (27). The CD spectra of these preparations have remarkable similarities to the spectra of unstacked thylakoid membranes (Fig.  3), showing that equivalent pigment–pigment interactions are formed in the LHCII in lamellar aggregates and in the native membranes (27). In contrast, after detergent solubilization of the thylakoid membranes or the lamellar aggregates, above the critical micelle concentrations, their respective spectra are virtually indistinguishable from the LHCII trimers. Thus, lamellar aggregates can serve as a rough model of the native membrane, and certainly serve as a better model of the state of LHCII in  vivo than solubilized trimers. These alterations in the CD, upon detergent solubilization of the trimers, and the accompanying changes in the absorbance and linear dichroism spectra, as well as in the fluorescence lifetimes, are probably caused by the losses of specific trimer–trimer contacts that are normally present in the thylakoid membranes and also in the lamellar aggregates (27). The native state of the trimers might also be protected by lipids, which could well explain the differences between the loosely stacked lamellar aggregates, retaining substantial amounts of lipids, and aggregates prepared from solubilized trimers, which are largely delipidated (7). In harmony with this hypothesis, Yang et al. (28) have found that LHCII proteoliposomes exhibit CD features similar to those of the excitonic CD of lamellar aggregates and thylakoid membranes.

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4. Notes 1. When using either pea, grown in the greenhouse, or spinach, from the local market or greenhouse, be sure that the plants are not treated with any kind of chemical agents (insecticides, pesticides, etc.). The leaves must be good-looking and fresh, with good turgor. This is really very important since all the following steps will depend almost solely on the plant material chosen. Homogenization of 50–60 g of pea needs about 150–200 ml of buffer A and for 400 g of spinach, use about 300–400 ml of buffer A. 2. When discarding the supernatant, make sure that the light upper part of the supernatant is also thrown away. Just keep the tube upside down for a minute until all unnecessary material drip out. 3. Be sure to homogenize the pellet until no particles are visible on the wall of Potter homogenizer. The chlorophyll concentration (i.e., the optical density) must be properly adjusted; otherwise there is a high risk that Mg ions cannot precipitate the LHCII. 4. The optimum concentration of Triton X-100 can vary from batch to batch. For example, the upper limit of 0.9% will be used when the material is fresh and good-looking spinach. The same is valid also for preparation of LHCII microcrystals; when preparing microcrystals, keep in mind that addition of higher than 1.6% Triton leads to inability to aggregate LHCII. 5. Clouds are visible for preparation of LHCII types I, II, and partially III. When LHCII type IV is prepared, no clouds are visible in a suspension and the color is dark green reddish. For type IV, if clouds are still present after initial addition of Triton X-100, add some more detergent and stir for a few minutes until clouds disappear. 6. When lamellar aggregates of LHCII are prepared, bigger pellet is obtained and for microcrystals of LHCII, very small or sometimes just a trace is obtained. 7. Take only the supernatant. Sometimes, especially with lower concentrations of Triton X-100, there is some loose pellet, which can spoil the experiment because it contains some unsolubilized thylakoid membranes and thus many additional proteins, and also larger amount of lipids. 8. Add Mg2+ and K+ in this order, quickly one after the other. For some unknown reason, when this order is changed, no aggregation is possible and the whole experiment might fail. 9. Addition of Triton X-100 should not exceed 0.2% final volume and shaking the tube with the detergent has to be short,

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2–3  min following by the addition of Mg2+ and K+ in this order. Continue shaking for another 10–15  min and stop when no more clouds are generated. Addition of the detergent has to be adjusted after measuring the optical density – higher detergent concentration for higher optical density.

Acknowledgments This work has been supported by grants from the Hungarian Fund for Basic Research (OTKA T30324, T34188 and T42696). References 1. Liu, Z. F., Yan, H. C., Wang, K. B., Kuang, T. Y., Zhang, J. P., Gui, L. L., An, X. M., and Chang, W. R. (2004) Crystal structure of spinach major light-harvesting complex at 2.72 Angstrom resolution. Nature 428, 287–292. 2. Arntzen, C. J. (1978) Dynamic structural features of chloroplast lamellae. Curr. Top. Bioenerg. 8, 111–160. 3. Garab, G., and Mustárdy, L. (1999) Role of LHCII-containing macrodomains in the structure, function and dynamics of grana. Aust. J. Plant Physiol. 26, 649–658. 4. Anderson, J. M., and Andersson, B. (1988) The dynamic photosynthetic membrane and regulation of solar energy conversion. Trends Biochem. Sci. 13, 351–355. 5. Horton, P., Ruban, A. V., and Walters, R. G. (1996) Regulation of light harvesting in green plants. Annu. Rev. Plant Physiol. Plant Mol. Biol. 47, 655–684. 6. Krupa, Z., Huner, N. P. A., Williams, J. P., Maissan, E., and James, D. R. (1987) Develop­ ment at cold hardening temperatures – the structure and composition of purified rye LHCII. Plant Physiol. 84, 19–24. 7. Simidjiev, I., Barzda, V., Mustárdy, L., and Garab, G. (1997) Isolation of lamellar aggregates of the light-harvesting chlorophyll a/b protein complex of photosystem II with longrange chiral order and structural flexibility. Anal. Biochem. 250, 169–175. 8. Barzda, V., Mustárdy, L., and Garab, G. (1994) Size dependency of circular dichroism in macroaggregates of photosynthetic pigment–protein complexes. Biochemistry 33, 10837–10841. 9. Simidjiev, I., Stoylova, S., Amenitsch, H., Jávorfi, T., Mustárdy, L., Laggner, P.,

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Holzenburg, A., and Garab, G. (2000) Selfassembly of large, ordered lamellae from nonbilayer lipids and integral membrane proteins in  vitro. Proc. Natl Acad. Sci. U. S. A. 97, 1473–1476. Barzda, V., Istokovics, A., Simidjiev, I., and Garab, G. (1996) Structural flexibility of chiral macroaggregates of light-harvesting chlorophyll a/b pigment–protein complexes. Light-induced reversible structural changes associated with energy dissipation. Biochemistry 35, 8991–8997. Simidjiev, I., Barzda, V., Mustárdy, L., and Garab, G. (1998) Role of thylakoid lipids in the structural flexibility of lamellar aggregates of the isolated light-harvesting chlorophyll a/b complex of photosystem II. Biochemistry 37, 4169–4173. Garab, G., Faludi-Dániel, Á., Sutherland, J. C., and Hind, G. (1988). Macroorganization of chlorophyll a/b light-harvesting complex in thylakoids and aggregates: Information from circular differential scattering. Biochemistry 27, 2425–2430. Cseh, Z., Rajagopal, S., Tsonev, T., Busheva, M., Papp, E., and Garab, G. (2000) Thermooptic effect in chloroplast thylakoid membranes. Thermal and light stability of pigment arrays with different levels of structural complexity. Biochemistry 39, 15250–15257. Garab, G., Cseh, Z., Kovács, L., Rajagopal, S., Várkonyi, Z., Wentworth, M., Mustárdy, L., Dér, A., Ruban, A. V., Papp, E., Holzenburg, A., and Horton, P. (2002) Light-induced trimer to monomer transition in the main light-harvesting antenna complex of plants: Thermo-optic mechanism. Biochemistry 41, 15121–15129.

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15. Barzda, V., Jennings, R. C., Zucchelli, G., and Garab, G. (1999) Kinetic analysis of the light-induced fluorescence quenching in lightharvesting chlorophyll a/b pigment–protein complex of photosystem II. Photochem. Photobiol. 70, 751–759. 16. Gruszecki, W. I., Grudzinski, W., Matula, M., Kernen, P., and Krupa, Z. (1999) Lightinduced excitation quenching and structural transition in light-harvesting complex II. Photosynth. Res. 59, 175–185. 17. Grudzinski, W., Krupa, Z., Garstka, M., Maksymiec, W., Swartz, T. E., Gruszecki, W. I. (2002) Conformational rearrangements in light-harvesting complex II accompanying light-induced chlorophyll a fluorescence quenching. Biochim. Biophys. Acta – Bioenergetics 1554, 108–117. 18. Zer, H., Vink, M., Keren, N., Dilly-Hartwig, H. G., Herrmann, R. G., Paulsen, H., Andersson, B., and Ohad, I. (1999) Regulation of thylakoid protein phosphorylation at the substrate level: Reversible light-induced conformational changes expose the phosphorylation site of the light-harvesting complex II. Proc. Natl Acad. Sci. U. S. A. 96, 8277–8282. 19. Yang, D. H., Paulsen, H., and Andersson, B. (2000) The N-terminal domain of the light-harvesting chlorophyll a/b-binding protein complex (LHCII) is essential for its acclimative proteolysis. FEBS Lett. 466, 385–388. 20. Dobrikova, A. G., Várkonyi, Z., Krumova, S. B., Kovács, L., Kostov, G. K., Todinova, S. J., Busheva, M. C., Taneva, S. G., and Garab, G. (2003) Structural rearrangments in chloroplast thylakoid membranes revealed by differential scanning calorimetry and circular dichroism spectroscopy. Thermo-optic effect. Biochemistry 42, 11272–11280.

21. Arnon, D. J. (1949) Copper enzymes in isolated chloroplasts: Polyphenoloxidase in Beta vulgaris. Plant.Physiol. 24, 1–15. 22. Bligh, E. G., and Dyer, W. J. (1959) A rapid method of total lipid extraction and purification. Can. J. Biochem. Physiol. 37, 911–917. 23. Vigh, L., Horváth, I., and Thompson, G. A. Jr. (1988) Recovery of Dunaliella salina cells following hydrogenation of lipids in specific membranes by a homogeneouus palladium catalyst. Biochem. Biophys. Acta – Biomembranes 937, 42–50. 24. Istokovics, A., Simidjiev, I., Lajkó, F., and Garab, G. (1997) Characterization of the light induced reversible changes in the chiral macroorganization of the chromophores in cloroplast thylakoid membranes. Temperature dependence and effect of inhibitors. Photosynth. Res. 54, 45–53. 25. Keller, D., and Bustamante, C. (1986) Theory of the interaction of light with large inhomogeneous molecular aggregates. II. Psi-type circular dichroism. J. Chem. Phys. 84, 2972–2979. 26. Garab, G., and van Amerongen, H. (2009) Linear dichroism and circular dicroism in photosynthesis research. Photosynth. Res. 101, 135–146. 27. Lambrev, P. H., Várkonyi, Z., Krumova, S., Kovács, L., Miloslavina, Y., Holzwarth, A. R., and Garab, G. (2007) Importance of trimer– trimer interactions for the native state of the plant light-harvesting complex II. Biochim. Biophys. Acta 1767, 847–853. 28. Yang, C., Boggasch, S., Haase, W., and Paulsen, H. (2006) Thermal stability of trimeric lightharvesting chlorophyll a/b complex (LHCIIb) in liposomes of thylakoid lipids. Biochim. Biophys. Acta 1757, 1642–1648.

Chapter 13 Protein Targeting Across and into Chloroplast Membranes Shari M. Lo and Steven M. Theg Abstract The protein complexes in the thylakoid membrane are composed of subunits derived from both the nuclear and chloroplast genomes. While less is known about the mechanisms of delivery of the plastidencoded subunits, the targeting mechanisms of the nuclear-encoded subunits have been more experimentally tractable. We have described in this chapter the methods used in our laboratory for investigations of the import of nuclear-encoded proteins across the chloroplast envelope membranes, and for their further delivery into or across the thylakoid membrane by one of the four distinct pathways. Key words: Protein transport, Chloroplast protein import, cpTat pathway, cpSec pathway, cpSRP pathway, Spontaneous insertion pathway

1. Introduction In chloroplasts, as in mitochondria, the major complexes responsible for electron transport and energy transduction contain polypeptide subunits that are encoded in both the organellar and nuclear genomes (1). The bigenomic origin of the photosynthetic machinery is especially pronounced, where the ratio of plastid- to nuclear-encoded subunits varies widely (2–4). Accordingly, the subunits encoded in the nucleus are translated on cytoplasmic 80S ribosomes and posttranslationally imported across the envelope membranes into the stroma. Most of them must further be integrated into the thylakoid membrane, and others are transported fully across the membrane to the lumen. Clearly, the assembly of the photosynthetic complexes is an elaborate process, requiring the subunits to cross as many as three membranes and to combine with their chloroplast-encoded partners.

Robert Carpentier (ed.), Photosynthesis Research Protocols, Methods in Molecular Biology, vol. 684, DOI 10.1007/978-1-60761-925-3_13, © Springer Science+Business Media, LLC 2011

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The targeting of nuclear-encoded proteins has been extensively studied using a small subset of transported substrates (5). These studies have shown the following: 1. With but few exceptions, all proteins appear to cross both envelope membranes through the so-called Toc and Tic machinery, separate translocation complexes located in the outer and inner envelope membranes, respectively, that act in concert to effect protein import into the stroma. 2. Proteins in the stroma that are destined for thylakoid membrane integration can follow one of the four routes, namely, the cpTat, cpSec, cpSRP, and the spontaneous integration pathways. 3. Proteins that reside in the thylakoid lumen can cross the membrane on the cpSec or cpTat pathways. The cpTat, cpSec, and cpSRP pathways all have counterparts in bacteria that mediate protein export from the cytoplasm, and in the latter two cases, in the endoplasmic reticulum controlling protein entry into the eukaryotic endomembrane system. The spontaneous integration pathway appears to be unique to thylakoids. These various protein targeting pathways are illustrated in Fig. 1.

Fig.  1. Targeting pathways in chloroplasts. Almost all nuclear-encoded proteins enter the plastid through the Toc/Tic machinery embedded in the envelope membranes. Some proteins are then integrated into the thylakoid membrane via the cpTat, cpSec, cpSRP, or spontaneous integration pathways; others are transported into the thylakoid lumen via the cpTat or cpSec pathways. The energy requirements for the different pathways are indicated.

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The study of the import of proteins into plastids and their distribution within is a prelude to investigations of the assembly of the photosynthetic complexes (6). This chapter describes the methods used in the authors’ laboratory to study each of the five protein targeting pathways found in chloroplasts.

2. Materials 2.1. Plant Growth

1. Seeds from Pisum sativa var. Little Marvel (Seedway LLC, Hall, NY). 2. Vermiculite.

2.2. In Vitro Transcription and Translation (see Note 1)

1. 2 mg of in vitro transcription plasmid encoding translocation substrate, fully linearized downstream of the coding sequence. In 50 ml restriction digestion reaction mixture. 2. OmniPur® phenol:chloroform:isoamyl alcohol, 25:24:1 (phenol:chloroform) in saturated TE buffer, pH 6.7, (EMD, Gibbstown, NJ). Store refrigerated at 2–8°C. 3. OmniPur® chloroform:isoamyl alcohol, 24:1 (chloroform:­ IAA), 3155 (EMD, Gibbstown, NJ). Store refrigerated at 2–8°C. 4. Pellet Paint® coprecipitant in 3  M sodium acetate, pH 5.2 (Novagen, Madison, WI). 5. RNase-free 3 M sodium acetate. 6. RNase-free 200 Proof (100%) ethyl alcohol. 7. RNase-free water. 8. RNA polymerase specific for in  vitro transcription plasmid (T7, T3, or SP6), along with 5× reaction buffer (P2075, P2083, or P1085) (Promega, Madison, WI). 9. RNase-free 1 mg/ml bovine serum albumin (BSA). 10. RNase-free 100 mM dithiothreitol (DTT) molecular grade. 11. RNase-free mixture of rNTPs (adenosine, uridine, cytidine, guanosine 5¢-triphosphate) at 2.5 mM concentration each in water, pH 7.5. 12. 5  mM m7G(5¢)ppp(5¢)G RNA Capping Analog in RNasefree water. 13. Recombinant RNasin® Ribonuclease Inhibitor (Promega, Madison, WI). The concentration varies, 40 U/ml is typical. 14. Wheat Germ Extract Kit (Promega, Madison, WI). Includes wheat germ extract, 1 M potassium acetate, 1 mM amino acid mixture minus methionine or leucine. 15. 1.0 mCi/ml l-[3,4,5-3H(N)]-leucine ([3H]-Leu) or 10.25 mCi/ ml l-[35S]-methionine ([35S]-Met) (Perkin Elmer, Boston, MA).

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16. 20 mM l-leucine and 20 mM l-methionine. 17. Temperature-controlled water bath. 2.3. Chloroplast Isolation

1. Grinding buffer (1× GB): 50  mM Tricine–KOH, pH 8.0, 330  mM sorbitol, 1  mM MgCl2, 1  mM MnCl2, 2  mM Na2EDTA, 0.1% BSA. 2. Import buffer 1 (1× IB1): 50  mM Tricine–KOH, pH 8.0, 330 mM sorbitol, 3 mM MgCl2. 3. Percoll (Sigma, St. Louis, MO). 4. Miracloth (Calbiochem/EMD, Gibbstown, NJ). 5. Isoascorbic acid. 6. Glutathione. 7. 80% acetone. 8. Waring blender, with sharpened blades.

2.4. Thylakoid Isolation and Stromal Extract Preparation

1. Isolated intact chloroplasts at 1 mg chlorophyll (Chl)/ml in 1× IB1. 2. Lysis buffer 1 (LB1): 10 mM MES–KOH, pH 6.5, 5 mM MgCl2. 3. Lysis buffer 2 (LB2): 10 mM HEPES–KOH, pH 8.0, 10 mM MgCl2. 4. 2× Import buffer 1 (2× IB1): 100 mM Tricine–KOH, pH 8.0, 660 mM sorbitol, 6 mM MgCl2 (see Note 2). 5. 2× Import buffer 2 (2× IB2): 100 mM HEPES–KOH, pH 8.0, 660 mM sorbitol, 20 mM MgCl2. 6. Stromal extract buffer (SEB): 90 mM HEPES–KOH, pH 8.0, 660 mM sorbitol, 10 mM MgCl2.

2.5. Chloroplast Protein Import Across the Envelope Membranes

1. Isolated intact chloroplasts at 1 mg Chl/ml in 1× IB1.

2.6. Thylakoid Protein Transport on the cpTat Pathway

1. Isolated thylakoids at 1 mg Chl/ml in 1× IB1.

2. 2× IB1. 3. 0.1 M ATP. 4. Adjustable (100 mE/m2 s) light source (see Notes 3 and 4).

2. 2× IB1. 3. [3H]-Leu, or [35S]-Met-labeled substrate: our laboratory has often used the precursor or intermediate forms of the 23 or 17 kDa subunits of the oxygen evolving complex, prOE23 or iOE23 and prOE17 or iOE17 (a.k.a. PsbP and PsbR, or OEE2 and OEE3). 4. Adjustable (100 mE/m2 s) light source.

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2.7. Thylakoid Protein Transport on the cpSec Pathway

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1. 4 mg Chl/ml Chloroplast lysate in 1× IB2. 2. 4× SE in 1× IB2. 3. [3H]-Leu, or [35S]-Met-labeled substrate: our laboratory commonly uses the precursor or intermediate forms of the 33 kDa subunit of the oxygen evolving complex, prOE33 or iOE33 (a.k.a. PsbO or OEE1). 4. 100 mM ATP in 1× IB1. Aliquots are stored in −20°C. Each aliquot is thawed before use and used only once. 5. Adjustable (150 mE/m2 s) light source.

2.8. Protein Integration into the Thylakoid Membrane via the cpSRP Pathway

1. 4 mg Chl/ml Chloroplast lysate in 1× IB2. 2. 4× SE in 1× IB2. 3. [3H]-Leu, or [35S]-Met-labeled substrate: our laboratory uses the precursor form of the light harvesting chlorophyll binding protein (prLHCP). 4. 100 mM ATP in 1× IB1. Aliquots are stored at −20°C. Each aliquot is thawed before use and used only once. 5. 100 mM GTP in 1× IB1. Aliquots are stored at −20°C. Each aliquot is thawed before use and used only once. 6. 6  mg/ml thermolysin with 150  mM CaCl2 and 150  mM CaCl2 alone in 1× IB1. Aliquots are stored at −80°C. Each aliquot is thawed before use and used only once. 7. 1× IB2 minus MgCl2: 50  mM HEPES–KOH, pH 8.0, 330 mM sorbitol. 8. 100 mM EDTA, pH 8.0. 9. Adjustable (50 mE/m2 s) light source.

2.9. Protein Integration into the Thylakoid Membrane via the Spontaneous Integration Pathway 2.10. 12.5% SDS-PAGE (for 9 gels)

1. Thylakoids at 1 mg Chl/ml in 1× IB2. 2. [3H]-Leu, or [35S]-Met-labeled substrate: we have used the precursor forms of PsbX or PsbW (prPsbX, prPsbW). 3. Adjustable (50 mE/m2 s) light source. 1. Resolving gel: 12.6 ml 1.5 M Tris–HCl, pH 8.8, 21.1 ml 30% acrylamide/bis, 0.51  ml 10% SDS, 16.2  ml H2O, 0.25  ml 10% ammonium persulfate (APS), 26  ml TEMED. Final volume: ~50 ml. 2. Stacking gel: 5.6  ml 0.5  M Tris–HCl, pH 6.8, 2.9  ml 30% acrylamide/bis, 0.225 ml 10% SDS, 13.7 ml H2O, 0.112 ml 10% APS, 22.5 ml TEMED. Final volume: ~23 ml. 3. Running buffer: 25 mM Tris–HCl, 192 mM glycine, 0.1% SDS.

2.11. 15.5% Tricine SDS-PAGE (per each gel) (7)

1. Acrylamide solution 1: 9.021  g acrylamide, 0.279  g bisacrylamide, bring to 20 ml with deionized distilled H2O.

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2. Acrylamide solution 2: 8.471  g acrylamide, 0.129  g bisacrylamide, bring to 20 ml with deionized distilled H2O. 3. 3x Gel buffer: 3 M Tris–HCl, pH 8.45, 0.3% SDS. 4. 15.5% Resolving gel: 1.5 ml acrylamide solution 1, 1.5 ml 3x gel buffer, 0.87  ml 70% glycerol, 0.63  ml H2O, 15  ml 10% APS, 1.5 ml TEMED. Final volume: ~4.5 ml. 5. 9.5% Spacer gel: 122 ml acrylamide solution 1, 200 ml 3x gel buffer, 272 ml H2O, 5 ml APS, 1 ml TEMED. Final volume: ~600 ml. 6. 3.5% Stacking gel: 80 ml acrylamide solution 2, 247 ml 3x gel buffer, 664 ml H2O, 8 ml APS, 1 ml TEMED. Final volume: ~1 ml. 7. Cathode buffer: 0.1 M Tricine–KOH, 0.1 M Tris–HCl, pH 8.25, 0.1% SDS. 8. Anode buffer: 0.2 M Tris–HCl, pH 8.9. 2.12. Protein Transport and Integration Quantification

1. Coomassie stain: 0.25% Coomassie brilliant blue, 40% methanol, 10% acetic acid. 2. 1× Destain solution: 7.5% acetic acid, 1% glycerol, 40% methanol. 3. 2,5-Diphenyloxazole (PPO): 1 mM in glacial acetic acid. 4. High-range Rainbow® molecular weight markers (Amersham/ GE Life Sciences, Piscataway, NJ). 5. 2× Sample buffer (2× SB): 125 mM Tris–HCl, pH 6.8, 4% SDS, 20% glycerol, 0.05 mg/ml bromophenol blue, 10% b-mercaptoethanol. 6. 100% Glacial acetic acid. 7. Fisherbrand® Pure Cellulose Chromatography Paper: thickness, 0.35 mm (Fisher Scientific, Pittsburgh, PA). 8. Mini-Protean® Gel Electrophoresis system (Biorad, Hercules, CA). 9. Slab gel dryer. 10. Storage phosphor screen and cassette. 11. X-ray film and X-ray film cassette. 12. Molecular Dynamic Storm® PhosphorImager Scanner and Image Quant program (GE Life Sciences, Piscataway, NJ). 13. Photo scanner with Adobe® Photoshop.

3. Methods 3.1. Plant Growth

1. Approximately 100 ml of pea seeds are soaked for a few hours in deionized water.

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2. Seeds are sown onto a wet bed of vermiculite in a flat plastic pan (35 × 20 × 6  cm) and covered by a shallow layer of wet vermiculite. 3. Plants are grown in a controlled environment chamber at 20°C in a 12/12 h light/dark cycle. 4. Seedling leaves are harvested at 9–14 days. 3.2. In Vitro Transcription and Translation of Substrates for use in the In Vitro Transport and Integration Assays

1. All steps are performed using RNase-free tips, tubes, and reagents. “RNase-free” will no longer be denoted for the remainder of this section. All centrifuge steps are performed in a tabletop centrifuge at room temperature and 15,000 × g (see Note 5). 2. Perform a phenol and chloroform extraction of the linearized plasmid DNA encoding the substrate in the preparation of in vitro transcription. Add 50 ml H2O to the microfuge tube (A) containing the linearized plasmid in 50  ml total restri­ ction enzyme digestion reaction. Add to this 100  ml phenol:chloroform. Mix by flicking the tube or pipetting up and down to fully homogenize the two phases (see Note 6). 3. Centrifuge the reaction in a tabletop centrifuge for 5  min. Carefully pipette the upper aqueous phase containing the plasmid DNA into a new microfuge tube (B) (see Note 7). 4. Add 50 ml H2O to the tube (B) containing the plasmid DNA. Add to this 100 ml chloroform:IAA. Mix by flicking the tube or pipetting up and down to fully homogenize the two phases. 5. Centrifuge the reaction in a tabletop centrifuge for 2  min. Carefully pipette the upper aqueous phase containing the plasmid DNA into a new microfuge tube (C). 6. Precipitate the plasmid DNA in tube (C) by adding 4 ml Pellet Paint® and 13  ml 3  M sodium acetate. Mix the reaction by pipetting up and down. Add 300 ml of 100% ethyl alcohol. Incubate the reaction at room temperature for 2  min. Centrifuge the reaction for 5 min. 7. A pink pellet should be present at the bottom of the tube, which is the precipitated plasmid DNA (see Note 8). Pipette and discard all the supernatant containing ethyl alcohol. Add 200 ml 70% ethyl alcohol. Mix the reaction gently to “wash” the pellet. Centrifuge the reaction for 5 min. 8. Pipette and discard the supernatant. Repeat step 7 with 100 ml 100% ethyl alcohol. Pipette and discard the supernatant. 9. Dry the precipitated DNA pellet completely by turning the tube upside down with the cap open. 10. Resuspend the dry DNA pellet in 11 ml H2O. Add to this the in vitro transcription mixture: 10 ml 5× transcription buffer,

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2.5 ml 1 mg/ml BSA, 5 ml 100 mM DTT, 10 ml 2.5 mM rNTPs minus rGTP, 3 ml 2.5 mM rGTP, 6 ml 5 mM RNA Capping Analog, 1 ml RNasin®, 1 ml appropriate RNA polymerase. Mix the reaction and incubate in a water bath at 37°C for 1 h. 11. Repeat the phenol:chloroform extraction (steps 2–9), starting with tube (A) containing 50 ml of the in vitro transcription mixture. 12. Resuspend the phenol:chloroform-extracted RNA pellet with 45 ml H2O, 5 ml 100 mM DTT, and 0.5 ml RNasin®. Incubate in a water bath at 50°C for 10 min. 13. Perform an in vitro translation using the Wheat Germ Extract Kit. For [3H]-Leu translations, dry down completely 150 ml [3H]-Leu in a clean microfuge tube under a gentle stream of nitrogen gas. To this tube, add 25 ml wheat germ extract, 4 ml 1 mM amino acid mixture minus leucine, 2.4 ml 1 M potassium acetate, 1 ml 100 mM DTT, 1 ml Rnasin®, 6.6 ml H2O, and 10 ml phenol:chloroform-extracted RNA. For [35S]-Met translations, add to a new tube 25 ml wheat germ extract, 4 ml 1  mM amino acid mixture minus methionine, 2.4  ml 1  M potassium acetate, 1 ml 100 mM DTT, 1 ml Rnasin®, 1.6 ml H2O, 5 ml [35S]-Met, and 10 ml phenol:chloroform-extracted RNA. Incubate at room temperature (25°C) for 2  h. After the incubation, add 15 ml of 20 mM nonradioactive l-leucine or l-methionine as appropriate (see Note 9). 3.3. Isolation of Intact Chloroplasts

All procedures should be carried out at 4°C with instruments prechilled on ice. 1. Mix in a 50-ml polycarbonate centrifuge tube 15 ml Percoll, 15 ml 2× GB, and 10 mg each of isoascorbic acid and glutathione. Centrifuge for 30 min at 37,000 × g (max) at 4°C, to form a continuous Percoll density gradient. Place tube on ice and use within a few hours. 2. Using a scissors cut the pea seedlings just below the leaves. Briefly grind the leaves in approximately 200 ml 1× GB in a Waring blender fitted with sharpened blades. Filter the slurry through a layer of Miracloth and divide the filtrate among four 50-ml centrifuge tubes. 3. Pellet the chloroplasts in the slurry by centrifuging the tubes in a swing-out rotor at 3,000 × g (max) for 5 min. 4. Resuspend the pellet in 2–4 ml 1× IB1 and carefully layer on top of the pre-formed Percoll gradient. Spin in a swing-out rotor at 8,000 × g (max) for 10 min with the brake off. Intact chloroplasts will collect in the lower of the two resulting ­chlorophyll-containing bands (Fig.  2). Aspirate the liquid above this band, and remove the chloroplasts to a single fresh

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Fig. 2. Separation of broken from intact chloroplasts in a linear Percoll density gradient.

50-ml tube using a Pasteur pipette. Wash the chloroplasts twice in 40 ml 1× IB1, pelleting at 1,500 × g (max) for 5 min. 5. Measure the Chl content of the thylakoids by mixing 10 ml thylakoids with 5 ml 80% acetone, filtering, and then reading the absorbance of the filtrate at 645, 663, and 720 nm in a spectrophotometer. Chl (in mg/ml) is calculated as 4.02 × (A663 − A720) + 10.14 × (A645 − A720). 6. Dilute the chloroplasts to 1 mg Chl/ml and keep in a covered ice bucket until use. 3.4. Thylakoid, Chloroplast Lysate, and Stromal Extract (SE) Preparation

All steps are performed with buffers and equipment precooled on ice or to 4°C. 1. Prepare thylakoids from 1 ml of 1 mg Chl/ml intact chloroplasts in 1× IB1. Pellet the chloroplast by centrifuging the chloroplasts at 4°C, 1,500 × g, for 5 min. Remove the supernatant. Resuspend the chloroplast pellet in 1 ml LB1. Incubate the resuspension on ice, in the dark for 5 min. Add 1 ml 2× IB1 and mix. Pellet the thylakoids by centrifugation at 4°C, 1,500 × g, for 5 min. Remove and discard the supernatant and resuspend the thylakoid pellet in 500 ml 1× IB1. Measure the chlorophyll concentration and adjust the final concentration of the thylakoid mixture to 1 mg Chl/ml using 1× IB1.

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2. Prepare 4 mg Chl/ml chloroplast lysate by pelleting 4 ml of 1 mg Chl/ml chloroplast by centrifugation at 4°C, 1,500 × g, for 5 min. Remove the supernatant. Resuspend the chloroplast pellet with 0.5  ml LB2. Incubate the resuspension on ice, in the dark for 5 min. Add 0.5 ml SEB. This results in 1 ml of 4 mg Chl/ml chloroplast lysate in 1× IB2. 3. Prepare 4  mg Chl/ml stromal extract (4× SE) by preparing 4  mg Chl/ml chloroplast lysate. Centrifuge the mixture at 4°C, 1,500 × g, for 10 min. Collect the supernatant, which is used at 4× SE (see Note 10). 3.5. Chloroplast Protein Import Assay

1. Add to a clear microfuge tube for each 60 ml reaction: 20 ml 2× IB1, 1.8 ml 0.1 M ATP (3 mM final), 2 ml radioactively labeled import substrate, other additions as dictated by the experiment, and water to bring the final volume to 40 ml (see Note 11). 2. Start the import reaction by adding 20 ml of chloroplasts at 1 mg Chl/ml and placing in front of the 100 mE/m2 s heatfiltered light source. Allow the reaction to proceed for 10 min (see Note 12). 3. Terminate the reaction by diluting the samples with 1 ml icecold 1× IB1 and placing on ice until all the reactions are completed. Samples are then pelleted in a microfuge at full speed for 3 min, and the supernatant aspirated (see Notes 13 and 14). 4. Depending on the requirements of the experiment, one can at this point include a 30-min incubation with thermolysin to digest remaining precursor that has not been imported. If this is to be included, resuspend the chloroplast pellet in 300 ml of ice-cold 1× IB1 minus MgCl2 with either 5 ml each of 6 mg/ ml thermolysin (0.1 mg/ml final concentration) and 150 mM CaCl2 or with 5  ml of 150  mM CaCl2 alone (2.5  mM final concentration). Incubate on ice in the dark for 30  min. Terminate the thermolysin digestion by adding 200 ml of icecold 1× IB1 minus MgCl2with 50 mM EDTA (20 mM final concentration). Mix. 5. Pellet the chloroplasts in a microfuge at full speed for 3 min. Aspirate the supernatant. 6. Resuspend the pellet in 40 ml 2× SB. Boil for 5 min in a water bath. 7. Dilute 2 ml of radioactively labeled precursor substrate used in the assays into 198 ml 2× SB. Boil for 5 min in a water bath. This is the standard for quantification (see Note 15). 8. Load 8 ml of the substrate standard generated in step 7 into the first 12.5% SDS-PAGE lane as a standard representing 20% of substrate used in the assay. Load 8 ml of each transport reaction in subsequent lanes.

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1. For each 60  ml transport reaction mix in a clear microfuge tube 2 ml radioactively labeled substrate, 20 ml 2× IB1, variables being tested, and water to bring the final volume to 40 ml. 2. Add 20 ml 1 mg/ml thylakoid in 1× IB1 to the reaction and place the microfuge tube in front of the heat-filtered light source adjusted to 100 mE/m2 s at room temperature to initiate transport. Allow transport to proceed for 10 min (see Note 12). 3. Terminate the transport reaction by centrifugation at 4°C, 1,500 × g, for 2 min. Aspirate the supernatant. Resuspend the thylakoid pellet completely in 40 ml of 2× SB. Boil the samples for 5 min in a water bath (see Note 16). 4. Dilute 2 ml of radioactively labeled substrate used in the assay into 198 ml 2× SB. Boil for 5 min in a water bath. This is the standard for quantification. 5. Subject samples to SDS-PAGE as described in Subheading 3.5, step 8. An example fluorograph is shown in Fig. 3.

3.7. Secretory Pathway (Sec) Transport Assay (10, 11)

1. For each 60  ml transport reaction mix in a clear microfuge tube 4  ml radioactively labeled substrate, 11.6  ml 2× IB2, 1.8 ml 100 mM ATP in 1× IB1, 25 ml 4× SE in 1× IB2, variables being tested, and water to bring the final volume to 50 ml. 2. Add 10 ml 4 mg/ml thylakoid in 1× IB2 to the reaction and place the microfuge tube in front of the heat-filtered light source adjusted to 150 mE/m2 s at room temperature to initiate transport. Allow transport to proceed for 10  min (see Note 12).

Fig. 3. iOE17 and iOE23 transport. The 50% denotes loading of 50% of the radiolabeled substrate used in the assay. Three lanes with identical transport assays are labeled with the appearance of the mature protein indicated. Samples were run on a 12.5% SDS-PAGE gel; a fluorograph is shown.

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Fig. 4. prOE33 transport. The 50% denotes loading of 50% of the radiolabeled prOE33 substrate used in the assay. Three lanes with identical transport assays are labeled with the appearance of the mature protein indicated. Samples were run on a 12.5% SDSPAGE gel; a fluorograph is shown.

3. Terminate the transport reaction by centrifugation at 4°C, 1,500 × g, for 2 min. Aspirate the supernatant. Resuspend the thylakoid pellet completely in 40 ml of 2× SB. Boil the samples for 5 min in a water bath (see Note 16). 4. Dilute 4 ml of radioactively labeled substrate used in the assay into 196 ml 2× SB. Boil for 5 min in a water bath. This is the standard for quantification. 5. Subject samples to SDS-PAGE as described in Subheading 3.5, step 8. An example fluorograph is shown in Fig. 4. 3.8. Signal Recognition Particle Integration Assay (12, 13)

1. For each 75  ml transport reaction mix in a clear microfuge tube 4 ml radioactively labeled substrate, 21 ml 2× IB2, 1.0 ml 100  mM ATP in 1× IB1, 1.0  ml 100  mM GTP in 1× IB1, 18.75 ml 4× SE in 1× IB2, variables being tested, and water to bring the final volume to 62.5 ml (see Note 17). 2. Add 12.5 ml of thylakoids at 4 mg Chl/ml in 1× IB2 to the reaction and place the microfuge tube in front of the heatfiltered light source adjusted to 50 mE/m2 s at room temperature to initiate integration. Allow transport to proceed for 10 min (see Note 12). 3. Terminate the integration reaction by centrifugation at 4°C, 6,000 × g, for 4 min. Aspirate the supernatant. Resuspend the thylakoid pellet completely in 300 ml of ice-cold 1× IB minus MgCl2 with either 5 ml of 6 mg/ml thermolysin and 150 mM CaCl2 or 5 ml of 150 mM CaCl2 alone. Incubate on ice in the dark for 30 min. 4. To terminate the thermolysin digestion, add 200  ml of icecold 1× IB2 minus MgCl2 with 50 mM EDTA. Mix. 5. Centrifuge at 4°C, 6,000 × g, for 4 min. Aspirate the supernatant. Resuspend the pellet with 500 ml of 1× IB2 minus MgCl2 with 5 mM EDTA.

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Fig. 5. prLHCP integration fluorography. The 50% denotes loading of 50% of the radiolabeled prLHCP substrate used in the assay. Three integration assays are labeled 1, 3, 5 with their corresponding thermolysin-treated aliquot labeled 2, 4, and 6, respectively. The appearance of the specific degradation product resulting from the proper integration of the substrate in the thylakoid membrane is indicated as LHCP-dp. Samples were run on a 12.5% SDS-PAGE gel; a fluorograph is shown.

6. Transfer the mixture into a fresh tube. 7. Centrifuge at 4°C, 6,000 × g, for 4 min. Aspirate the supernatant. Resuspend the pellet with 40  ml 2× SB with 50  mM EDTA. Boil the samples for 5 min in a water bath. 8. Dilute 4 ml of radioactively labeled substrate used in the assay into 196 ml 2× SB. Boil for 5 min in a water bath. This is the standard for quantification. 9. Subject samples to SDS-PAGE as described in Subheading 3.5, step 8. An example fluorograph is shown in Fig. 5. 3.9. Spontaneous Integration Assay (14, 15)

1. For each 60  ml transport reaction mix in a clear microfuge tube 2 ml radioactively labeled substrate, 20 ml 2× IB2, variables being tested, and water to bring the final volume to 40 ml. 2. Add 20 ml 1 mg/ml thylakoid in 1× IB2 to the reaction and place the microfuge tube in front of the heat-filtered light source adjusted to 100 mE/m2 s at room temperature to initiate transport. Allow transport to proceed for 10  min (see Note 12). 3. Terminate the transport reaction by centrifugation at 4°C, 15,000 × g, for 2 min. Aspirate the supernatant. Resuspend the thylakoid pellet completely in 40 ml of 2× SB. Heat the samples for 5 min in an almost-boiling water bath (see Note 18). 4. Dilute 2 ml of radioactively labeled substrate used in the assay into 198 ml 2× SB. Heat for 5 min in an almost-boiling water bath. This is the standard for quantification.

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Fig. 6. prPsbW integration fluorography. The 50% denotes loading of 50% of the radiolabeled prPsbW substrate used in the assay. Three lanes with identical integration assays are labeled with the appearance of the mature protein indicated. Samples were run on a 15.5% Tricine SDS-PAGE gel; a fluorograph is shown.

5. Load 8 ml of the substrate standard generated in step 4 into the first Tricine SDS-PAGE lane as a standard for 20% of substrates used in the assay. Load 8 ml of each transport reaction in subsequent lane. An example fluorograph is shown in Fig. 6. 3.10. 12.5% SDS-PAGE

We generally assess import, Tat and Sec transport, and signal recognition particle (SRP) integration into thylakoids using a 12.5% resolving gel in the Mini-Protean Tetra electrophoresis apparatus. 1. Using the multigel casting stand, prepare nine gels at a time. Prepare 50  ml of resolving gel solution as described in Subheading 2.10, item 1, adding APS and TEMED last. Fill the gels in the casting chamber, leaving room for a stacking gel. Overlay each gel with ~500 ml isopropanol. When they have polymerized, rinse the isopropanol off the gels with distilled water and blot dry the remaining water with a strip of chromatography paper. Pipette the stacking gel solution on top of the resolving gels. Insert combs, making sure to avoid air bubbles. When solidified, gels are removed from the casting chamber and stored in running buffer for up to a week at 4°C. 2. For use, remove the comb from the stacking gel and rinse the wells out with H2O. Load the gels into the Mini-Protean Tetra apparatus and add Running Buffer to the appropriate chambers in the apparatus.

3.11. 15.5% Tricine SDS-PAGE (7)

We generally assess Spontaneous integration into thylakoids using a 15.5% resolving gel in the Mini-Protean Tetra electrophoresis apparatus. 1. Prepare the resolving gel solution as described in Sub­ heading 2.11, item 4, adding APS and TEMED last. Fill the

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gels in the casting chamber, leaving room for a stacking gel. Overlay resolving gel with ~500  ml isopropanol. When the resolving gel is polymerized, rinse the isopropanol off the gel with distilled water and blot dry the remaining water with a strip of chromatography paper. Pipette the spacer gel solution ~2 mM thick, on top of the resolving gels. Overlay resolving gel with ~500 ml isopropanol. When the spacer gel is polymerized, rinse the isopropanol off the gel with distilled water and blot dry the remaining water with a strip of chromatography paper. Pipette the stacking gel solution on top of the spacer gel. Insert combs, making sure to avoid air bubbles. When polymerized, gels are removed from the casting chamber, wrapped in plastic wrap, and stored for up to a week at 4°C in an airtight container. 2. For use, remove the comb from the stacking gel and rinse the wells out with H2O. Load the gels into the Mini-Protean Tetra apparatus and add cathode and anode buffers to the appropriate chambers in the apparatus. 3.12. Protein Transport and Integration Quantification

1. Load 8 ml of each sample (20% of the reaction) onto either a precast 12.5% SDS-PAGE gel (for cpTat, cpSec, and cpSRP assays) or a precast 15.5% Tricine SDS-PAGE gel (for the spontaneous integration assay). 2. Load a lane with 10–20% of the radioactive substrate used in the assay as a standard. 3. Load a lane with prestained protein molecular weight marker, which can be useful to monitor the progress of the migration of samples in the gel. 4. Run the gel in a gel electrophoresis system such as MiniProtean Tetra® cell for about 1 h at 180 V (constant voltage) for 12.5% SDS-PAGE or about 2  h at 100  V and 4°C for 15.5% Tricine SDS-PAGE. 5. When the electrophoresis is completed, separate the plates, cut off the stacking gel, and cut a corner of the resolving gel in a distinctive manner. 6. Place the gel in a clean plastic container. The following solution incubations are carried out at room temperature on a gently rocking platform. For [35S]-labeled substrates, stain the gel in Coomassie R-250 solution for 15 min and destain two to three times for 10–15  min each with 1× destaining solution until distinct bands are present. For [3H]-labeled substrates, incubate the gel for 15–20  min in 100% glacial acetic acid, followed by 15–20 min in PPO solution, followed by two washes with deionized water, and two incubations in deionized water for 10 min each.

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7. Place a piece of chromatography paper just larger than the area of the gel onto the slab gel dryer and wet with deionized water. Place the wet gel on top of the chromatography paper, right side up. Cover the entire chromatography paper area with plastic wrap. Dry the gel for 1  h with the heat (set at 80°C) and vacuum on. 8. For [35S]-labeled substrates, place the gel dried onto chromatography paper into a storage phosphor cassette, making sure the gel faces the cleared phosphor screen directly. Close the cassette. Expose for 1 day at room temperature. For [3H]-labeled substrates, in a dark room: place the gel dried onto chromatography paper into an X-ray film cassette with an X-ray film on top, making sure the gel faces the film directly. Glow-in-the-dark paint may be used to mark the protein marker position on the dried gel before exposure to the X-ray film. Expose for 1 day at −80°C (see Note 19). 9. For [35S]-labeled substrates, scan the storage phosphor screen in a PhosphorImager. For [3H]-labeled substrates, develop the X-ray film and import into Adobe® Photoshop using a photo scanner with settings at Positive Film, Black and White Photo at 400 dpi. Convert the image to grayscale and invert the signals. 10. Quantitate the signals in the images using ImageQuant, setting the 10 or 20% substrate lane as a standard to compare the amount of transport from each assay (see Note 15).

4. Notes 1. While other laboratories have successfully utilized the TNT® Quick Coupled Transcription/Translation Systems (Promega, Madison, WI) to produce the substrates for their protein transport assays, we have found our method to produce the most efficient substrate in our lab. All assays and figures shown in this section used substrates produced using the in  vitro transcription and translation protocol presented here. Use of TNT® Quick Coupled Transcription/Translation substrates in the listed assays is also possible. TNT® Quick Coupled Transcription/Translation substrates are produced according to the manufacturer’s instructions. 2. All double-strength solutions, i.e., 2 ´ GB, 2 ´ IB1 and 2 ´ IB2, are diluted to 1× as needed, using deionized distilled water. The stock solutions are stored in −20°C for long-term storage (several months) and in 4°C as working stock. 3. It is important to ensure that the light source does not heat the samples. In our laboratory, we accomplish this by placing

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a solution of 1% CuSO4 in a flat-sided bottle between the lamp and the samples. 4. It is not difficult to inhibit the import reactions by having the light intensity too high. The optimal light intensity for each of the reactions described has not been determined, and we give values that yield successful results. Note that the light intensities used are considerably lower than that required for saturating rates of electron transport. 5. All phenol:chloroform steps are performed in a clean fume hood with inward air flow. Keep all instruments and equipment used as clean as possible. Practice RNase-free techniques to minimize the introduction of RNase into the reactions. 6. Take care to use the lower, organic phase of phenol:chloroform. The phenol:chloroform used is buffered to pH 6.7, as lower pH facilitates RNA stability. Organic waste generated is disposed as phenol:chloroform waste. 7. Take care not to mix the two phases in the reaction during transfer from the centrifuge or to the tube rack. Do not pick up any of the lower organic phase while collecting the upper aqueous phase. It is normal to leave some of the aqueous phase behind. 8. If no pellet forms, additional 100% ethyl alcohol may be added. Incubation on ice for 2 min or more may also facilitate formation of the pellet. 9. The incorporation of radioactive label may be quantitated. Normally, the minimum radioactive incorporation of a reaction that we would use in an assay is 100,000  cpm for [3H]-Leu and 50,000 cpm for [35S]-Met-labeled substrates. More or less RNA may be used in the translation mixture to achieve this target. 10. The preparation may be scaled up by using more chloroplast, lysis in LB2 at concentrations between 1 and 8 mg/ml chlorophyll and addition of equal volumes of SEB. Most commonly chloroplasts are lysed at 1  mg/ml chlorophyll but greater concentrations (up to 8  mg/ml chlorophyll) are required to produce more concentrated stromal extract. 11. In practice, we typically make up a master mix (or mixes) containing the stated volumes multiplied by n + 1, with n being the required number of samples. The master mixes are distributed 40 ml per tube before the chloroplasts are added to start the reactions. 12. In our laboratory, the 10-min time point for import, transport, or integration is chosen because this is at the end of the linear range for the reactions. This allows us to consider the extents of the reactions as rates. Longer incubation to see

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more import, transport or integration is possible, with published assays conducted for up to 30 min or longer. 13. This method is not suitable for exacting kinetics experiments as it does not cause an immediate cessation of the reactions (16). If an immediate halt is required, one can add, for instance, HgCl2 (17) or centrifuge the samples through silicone oil into acid (18). 14. Many laboratories include a step at this point to re-purify intact chloroplasts by centrifuging them through a 40% Percoll step gradient. This can be important if it is critical to lower the nonimported precursor concentration before running the samples in a gel. 15. In the quantitations it is necessary to account for any loss of radioactive amino acids that are present in the transit peptide and that are removed when the proteins are cleaved to their mature sizes after transport. 16. To remove all substrates bound to the thylakoids that may not be fully transported, optional thermolysin (19) treatment as detailed in Subheading 3.4, step 3 may be included. This treatment will verify that the evolution of the lower MW species in the transport assay is a truly “matured” or transported substrate. 17. One will need two reactions for each treatment, one with and one without thermolysin treatment. This is because the substrate used here is integrated into the thylakoid membrane, the correct integration of which produces a distinctive and specific degradation product (dp). 18. Some protein in the chloroplasts form aggregates upon boiling (for example, TatC and psbW). To prevent aggregation and allow denaturation of these proteins, the protein samples may be heated in a water bath at 70°C for 5–10 min. 19. To produce quantifiable X-ray film images, preflash the X-ray film with a single flash from an electronic photographic flash unit to introduce “absorbances between 0.1 and 0.2 above the background fog level of untreated film” (20). This will “increase sensitivity and give a fluorographic image that is proportional to the distribution of radioactivity in the sample” (20).

Acknowledgments This work is supported by US Department of Energy Grant DE-FG02-03ER15405.

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References 1. Aldridge, C., Cain, P., and Robinson, C. (2009) Protein transport in organelles: Protein transport into and across the thylakoid membrane. FEBS J 276, 1177–86. 2. Choquet, Y. and Vallon, O. (2000) Synthesis, assembly and degradation of thylakoid membrane proteins. Biochimie 82, 615–34. 3. Barber, J., Nield, J., Morris, E. P., Zheleva, D., and Hankamer, B. (1997) The structure, function, and dynamics of photosystem two. Physiol Plant 100, 817–27. 4. Jensen, P. E., Bassi, R., Boekema, E. J., Dekker, J. P., Jansson, S., Leister, D., Robinson, C., and Scheller, H. V. (2007) Structure, function and regulation of plant photosystem I. Biochim Biophys Acta 1767, 335–52. 5. Jarvis, P. (2008) Targeting of nucleus-encoded proteins to chloroplasts in plants. New Phytol 179, 257–85. 6. Hashimoto, A., Ettinger, W. F., Yamamoto, Y., and Theg, S. M. (1997) Assembly of newly imported oxygen-evolving complex subunits in isolated chloroplasts: Sites of assembly and mechanism of binding. Plant Cell 9, 441–52. 7. Schagger, H. (2006) Tricine-SDS-PAGE. Nat Protoc 1, 16–22. 8. Hynds, P. J., Robinson, D., and Robinson, C. (1998) The sec-independent twin-arginine translocation system can transport both tightly folded and malfolded proteins across the thylakoid membrane. J Biol Chem 273, 34868–74. 9. Cline, K. and McCaffery, M. (2007) Evidence for a dynamic and transient pathway through the TAT protein transport machinery. EMBO J 26, 3039–49. 10. Yuan, J. and Cline, K. (1994) Plastocyanin and the 33-kDa subunit of the oxygen-evolving complex are transported into thylakoids with similar requirements as predicted from pathway specificity. J Biol Chem 269, 18463–7. 11. 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

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nucleotide triphosphates. J Biol Chem 269, 3251–6. Cline, K. (1986) Import of proteins into chloroplasts: Membrane integration of a thylakoid precursor protein reconstituted in chloroplast lysates. J Biol Chem 261, 14804–10. 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, 32400–4. Kim, S. U., Robinson, D., and Robinson, C. (1996) An Arabidopsis thaliana cDNA encoding PS II-X, a 4.1 kDa component of photosystem II: A bipartite presequence mediates SecA/DELTA-pH-independent targeting into thylakoids. FEBS Lett 390, 175–8. Thompson, S. J., Kim, S. J., and Robinson, C. (1998) Sec-independent insertion of thylakoid membrane proteins – Analysis of insertion forces and identification of a loop intermediate involving the signal peptide. J Biol Chem 273, 18979–83. Leheny, E. A. and Theg, S. M. (1994) Apparent inhibition of chloroplast protein import by cold temperatures is due to energetic considerations not membrane fluidity. Plant Cell 6, 427–37. Reed, J. E., Cline, K., Stephens, L. C., Bacot, K. O., and Viitanen, P. V. (1990) Early events in the import/assembly pathway of an integral thylakoid protein. Eur J Biochem 194, 33–42. 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, 6730–6. Keegstra, K., Werner-Washburne, M., Cline, K., and Andrews, J. (1984) The chloroplast envelope: Is it homologous with the double membranes of mitochondria and gramnegative bacteria? J Cell Biochem 24, 55–68. Laskey, R. A. and Mills, A. D. (1975) Quantitative film detection of 3H and 14C in polyacrylamide gels by fluorography. Eur J Biochem 56, 335–41.

Chapter 14 Proteomic Analysis of Thylakoid Membranes Venkateswarlu Yadavalli, Sreedhar Nellaepalli, and Rajagopal Subramanyam Abstract Chlamydomonas is a model organism to study photosynthesis. Thylakoid membranes comprise several proteins belonging to photosystems I and II. In this chapter, we show the accurate proteomic measurements in thylakoid membranes. The chlorophyll-containing membrane protein complexes were precipitated using chloroform/methanol solution. These complexes were separated using two-dimensional gel electrophoresis, and the resolved spots were exercised from the gel matrix and digested with trypsin. These peptide fragments were separated by MALDI-TOF, and the isotopic masses were blasted to a MASCOT server to obtain the protein sequence. Matrix-assisted laser desorption/ionization-time of flight mass spectrometry (MALDI-TOF). The method discussed here would be a useful method for the separation and identification of thylakoid membrane proteins. Key words: 2D electrophoresis, Chlamydomonas, IEF, MALDI-TOF, Peptide fragments, Photo­ systems, SDS-PAGE, Thylakoid proteins, Trypsin digestion

1. Introduction The primary reactions of oxygenic photosynthesis occur at the thylakoid membrane and are catalyzed by several multi-molecular complexes including photosystem II (PSII), photosystem I (PSI), their associated light-harvesting complexes, the cytochrome b6f complex, and the ATP synthase. Each of these complexes consists of multiple subunits, pigments, and redox cofactors (1). Chlamydomonas reinhardtii is a valuable model system for determining the structure and function of polypeptides of the photosynthetic apparatus and the dynamic aspects of photosynthesis (2). We present here an easy analytical method for the separation and identification of thylakoid proteins by proteomics tools. The three major steps in proteome analysis are (1) the separation of complex protein mixtures, (2) the characterization of the separated Robert Carpentier (ed.), Photosynthesis Research Protocols, Methods in Molecular Biology, vol. 684, DOI 10.1007/978-1-60761-925-3_14, © Springer Science+Business Media, LLC 2011

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peptides, and (3) database searching. For the separation of complex protein mixtures, two-dimensional (2D) electrophoresis is currently the only technique that can reveal hundreds or even thousands of proteins at a time. Sample preparation is the important step in 2D electrophoresis, especially in the case of membrane proteins that are not soluble with common detergents. The use of b-dodecyl maltoside plays a key role in this method. Many protocols have been published for proteomic analysis of thylakoid membranes with 2D electrophoresis and mass spectrometry (MS) (3–11). Here, we show an advanced method of proteomic analysis of thylakoid membranes by 2D electrophoresis and Matrix-assisted laser desorption/ionization-time of flight mass spectrometry (MALDI-TOF).

2. Materials 1. Strain: Chlamydomonas reinhardtii strain CC125. 2. Medium (for 1.0 L): Cox Chlamydomonas medium (CC) (1×): Tris base (2.5 g), glacial acetic acid (1 mL), NH4NO3 (0.5 g), MgSO4 ∙ 7H2O (0.1 g), CaCl2 ∙ 2H2O (0.02 g), KCl (0.1  g), and Hutner’s trace elements solution (1  mL) prepared according to Ramesh and Webber (12). Store at 4°C. 3. Yeast extract should be added to the liquid CC medium (1×) at a final concentration of 0.1% and 50 mg/L KH2PO4 before autoclaving. 4. Cell resuspension buffer: 25  mM HEPES, pH 7.5, 0.3  M sucrose, 10 mM MgCl2, and 5 mM CaCl2. Store at 4°C. 5. Protease inhibitors: 1 mM benzamidine, 1 mM phenyl methyl sulfonyl fluoride (PMSF), 1  mM amino caproic acid, and 1 mM EDTA. Prepare 100 mM stocks and store at −20°C. 6. Thylakoid resuspension buffer I: 5  mM HEPES, pH 7.5, 0.3 M sucrose, and 10 mM EDTA. Store at 4°C. 7. Thylakoid resuspension buffer II: 5  mM Tris–HCl, pH 7.5, 0.2 M sorbitol, 5 mM CaCl2, and 10 mM MgCl2. Store at 4°C. 8. Sonicator (MSE). 9. Rehydration buffer: 7  M urea, 2  M thiourea, 2% 3-[(3-Cholamidopropyl) dimethylammonio]-1-propanesulfonate (CHAPS), 1% Biolyte pH 3-10NL (Bio-rad), 50 mM DTT, 0.01% Triton (v/v) X-100, 0.05% (w/v) b-dodecyl maltoside, and bromophenol blue (trace). Prepare fresh. 10. Protean IEF Cell (Bio-Rad). 11. Equilibration buffer I: 375 mM Tris–HCl, pH 8.8, 6 M urea, 2% (w/v) sodium dodecyl sulfate (SDS), 20% (v/v) glycerol, and 2% (w/v) DTT. Prepare fresh.

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12. Equilibration buffer II: 375  mM Tris–HCl, pH 8.8, 6  M urea, 2% (w/v) SDS, 20% (v/v) glycerol, and 2.5% (w/v) iodoacetamide. Prepare fresh. 13. Running buffer: 25 mM Tris–HCl, pH 8.3, 192 mM glycine, and 0.1% SDS. 14. Resolving gel concentrations (2D): 12% acrylamide and bisacrylamide, 375  mM Tris–HCl, pH 8.8, 0.1% SDS, 0.65  mL/L TEMED, and 3.5  mL/L of 10% ammonium persulfate. 15. Overlaying agarose: 0.5% agarose dissolved in 1× running buffer. Store at 4°C. 16. Protean II xi 2D cell (Bio-Rad). 17. Silver stain remover: 1:1 ratio of 30  mM K3[Fe(CN)6] and 100 mM Na2S2O3. Prepare fresh. 18. Wash solution: 50 mM NH4HCO3/acetonitrile in 1:1 (v/v). Prepare fresh. 19. Reduction buffer: 10  mM DTT in 50  mM NH4HCO3. Prepare fresh. 20. Alkylation buffer: 55 mM iodoacetamide in 50 mM NH4HCO3. Prepare fresh. 21. Trypsin (sigma) 10 ng/mL in 25 mM of NH4HCO3. 22. Speed vac (Labconco). 23. Zip-Tip c18 (Millipore). 24. Wetting solution for ZipTips: 100% acetonitrile. 25. Washing and equilibration solution for ZipTips: 0.1% trifluoroacetic acid in MilliQ water. 26. Elution solution for ZipTips: 0.1% trifluoroacetic acid/50% acetonitrile.

3. Methods 3.1. Growing of Chlamydomonas

1. A single colony from an agar plate was inoculated in 100 mL liquid medium (1X) according to the procedure described by Ramesh and Webber (12). 2. When the culture reached an OD of one at 730 nm, 5 mL of this culture was inoculated in 1.0  L of medium in a 2.5-L conical flask. 3. The Chlamydomonas was cultured on a shaker (115  rpm) under continuous light intensity of 35–40 mmol/m/s.

3.2. Preparation of Thylakoid Membranes

1. One liter of culture was harvested when the cells reached an OD of 1.0 at 730 nm by centrifuging at 4°C for 10 min at

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6,000 × g in a Beckman rotor JA 10 (wear gloves from this step onwards to avoid keratin contamination in later steps). 2. The pellets were suspended in 200 mL of cell resuspension buffer and centrifuged for 10 min at 9,000 × g in Sorvall GS3 rotor. 3. The resulting pellets were resuspended (2 × 108 cells/mL) in 50 mL of the same cell resuspension buffer containing protease inhibitors at a final concentration of 1 mM each. 4. The cells were broken using a sonicator by placing the cells on ice for four 20-s pulses with 3-min intervals. 5. The unbroken cells and starch granules were removed from the membrane suspension by centrifugation for 1.0  min at 1,000 × g in a SS 34 Sorvall rotor. 6. The above supernatant was centrifuged at 10,000 × g for 10 min in a SS 34 Sorvall rotor at 4°C. 7. The resulting pellet was resuspended in thylakoid resuspension buffer I and centrifuged at 20,000 g for 10 min in a SS 34 Sorvall rotor. 8. The pellet was resuspended in 2.0 mL of thylakoid resuspension buffer II. 3.3. Estimation of Chlorophyll Content

1. The chlorophyll concentration of thylakoids was determined by adding 10  mL aliquot of the thylakoid to 1  mL of 80% (v/v) acetone. 2. The aliquot was vortexed and then centrifuged for 2 min at room temperature in a microcentrifuge. 3. The supernatant was collected into a fresh Eppendorf tube without disturbing the pellet, to take the absorbance readings. 4. Chlorophyll concentration was determined by measuring absorption with a spectrophotometer at 645 and 663  nm using the following equation (13): Chlorophyll b (mg/mL) = (22.9 × A645)−(4.68 × A663) × dilution factor. Chlorophyll a (mg/mL) = (12.7 × A663)−(2.69 × A645) × ­dilution factor. Total chlorophyll (mg/mL) = C  hl a + Chl b = (20.21 × A645) + (8.02 × A663) × dilution factor.

3.4. Sample Preparation for 2D Electrophoresis

1. To precipitate the proteins, a 100-mL aliquot of thylakoid membrane solution having chlorophyll concentration of 1.0 mg/mL was taken into a sterilized Eppendorf tube. 2. A volume of 400 mL of methanol was added to the solution and vortexed thoroughly (modified method from Wessel and Fluegge (14) and Hippler et al. (15) (see Note 1).

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3. To this solution, 200 mL of water-saturated chloroform was added (see Note 2) and the solution was vortexed. 4. To the above fraction, 200 mL of deionized water was added, and the solution vortexed and centrifuged for 9,000 rpm in a microcentrifuge. 5. The protein precipitate was formed between the interphase. The upper and lower phases were carefully removed and discarded. 6. The protein precipitate was washed with 300 mL of methanol and then with another 300 mL of 95% methanol. 7. The pellet was air dried until methanol evaporated (see Note 3). 8. The pellet was solubilized with 400 mL of rehydration buffer by vortexing for 2 h (see Note 4). 9. The solubilized pellet solution was centrifuged at 12,000 rpm for 15 min at room temperature to avoid any unsolubilized proteins. The supernatant was saved in a fresh Eppendorf tube without disturbing the pellet. 10. Protein concentration from the above supernatant was determined according to the procedure described by Bradford (16). 3.5. Isoelectric Focusing (IEF)

1. To rehydrate the 17-cm immobilized pH gradient (IPG) strip, 300 mL of sample containing ~400 mg of protein was added to one side of rehydration tray placed on an even surface. 2. The IPG strip was placed with the gel side facing down on the sample from one end without forming any air bubbles. 3. The sample was allowed to stand for 10 min and then 3.0 mL of mineral oil was added on the backside of the strip and covered with a lid. 4. The sample was left with the mineral oil for at least 14 h to rehydrate the strip. Swelling of IPG strip may be observed after 14 h. 5. Two electrode wicks were placed on electrodes of the focusing tray by wetting with 8 mL of nano-pure water and placed in the PROTEAN IEF Cell system. 6. The strip was carefully removed from the rehydration tray by holding one side of the strip with a forceps, and the mineral oil was drained. 7. The IPG strip that has been rehydrated was loaded into the focusing tray by placing the gel side down and overlaying it with 3.0 mL of mineral oil and covering with the lid. 8. The following program is followed to run the isoelectric focusing at 20°C:

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Step 1: 250 V, 20 min, linear Step 2: 10,000 V, 2.30 h, linear Step 3: 10,000 V, 40,000 V-H, rapid. 9. After the isoelectric focusing is completed, the IPG strips can be run by SDS-PAGE immediately. Alternatively, IPG strips may be stored at −80°C for extended periods. 3.6. Equilibration of IPG Strip

1. After completion of isoelectric focusing, the IPG strip was carefully removed and the mineral oil drained. 2. The IPG strip was placed in the rehydration tray by facing the gel side up (see Note 5). 3. A volume of 6 mL of equilibration buffer I was added to the tray and IPG strip (after IEF run) was incubated on the rocker with slow motion for 15 min. 4. The equilibration buffer I was discarded by slanting one side and then 6 mL of equilibration buffer II was added and the tray incubated for 15 min on the rocker. 5. The equilibration buffer II (see Note 6) was discarded.

3.7. Preparation of Gels for SDS-PAGE

1. Glass plates were thoroughly cleaned with detergent (see Note 7). 2. The glass plates were assembled according to the instructions of the manufacturer and the resolving gel (12%) was poured allowing for a 1-cm gap for placing the IPG strip. 3. Water-saturated butanol was overlaid; this ensures that a flat top gel surface is achieved when the gel polymerizes. The gel was allowed to polymerize for at least 4 h before the run.

3.8. Second Dimension Running (2D Gel)

1. The top of the gels should be thoroughly washed with distilled water to remove all traces of butanol and unpolymerized acrylamide. 2. Agarose used for overlaying on the gel was melted in a microwave and maintained at 50°C in water bath. 3. The equilibrated IPG strip was immersed twice in 1× running buffer to ensure that excess equilibration solution has been removed. 4. The plastic backing was placed along a big glass plate with forceps and the strip was gently pushed down onto the top of resolving gel. 5. Overlay agarose was added to the well and the strip was slowly pushed into the well to ensure full contact between the IPG strip and the resolving gel (see Note 8). 6. After solidification of the agarose, the gel was mounted in the apparatus according to manufacturer’s instructions (Protein II xi 2D cell, Bio-Rad).

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7. In order to reduce the heat, a circulating water bath was connected to the Protean II xi (Bio-rad) 2D cell by setting the temperature to 15°C. 8. The gel box was filled with 1× running buffer and connected to the power supply. 9. The initial run was for 20 min at 80 V and then increased to 200 V. 10. A silver stain method was utilized to detect 2D spots (17) and gel was scanned using Amersham gel scanner. The gel spots were identified using PDQuest basic 8.0.1 imaging software (Alternatively, Coomassie stain can be done, in such case use 25 mM NH4HCO3 to destain in 3rd step of section 3.9). 3.9. In-Gel Digestion with Trypsin

1. The SDS gel was washed twice with water. For this study, we selected spots 1 and 2 only (Fig. 1) for in-gel digestion (see Note 9). 2. The spots were picked up manually with a clean scalpel, made into small 1-mm3 pieces, and transferred into sterile 0.5-mL vials. 3. To remove the silver stain, 30  mL of freshly prepared silver stain remover solution was added and vortexed for 30 min at room temperature. The procedure was repeated until clear gel pieces appeared. 4. The excess solution was removed and the gel particles were washed with wash solution for 15 min.

Fig.  1. Two-dimensional resolution of thylakoid membrane proteins from Chlamydo­ monas after chloroform/methanol precipitation. Horizontal gel dimension: isoelectric focusing on linear IPG gel strip between pH 3 (left border of the gel ) and pH 10 (right side of the gel ). Vertical gel dimension: SDS-PAGE. The SDS gel was silver stained.

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5. The remaining liquid was discarded and enough acetonitrile was added to cover the gel particles. 6. After adding acetonitrile, the gel pieces shrink and stick together; the excess acetonitrile was subsequently removed. 7. The gel particles were air dried. 8. Protein disulfide bridges were reduced by adding reduction buffer and incubating the gel for 45 min at 50°C. 9. After cooling to room temperature, excess liquid was removed and replaced quickly with the same volume of freshly prepared alkylation buffer. 10. The gel particles were incubated for 30 min at room temperature in the dark and the iodoacetamide was removed. 11. The gel particles were washed with 50  mM NH4HCO3/ acetonitrile in 1:1 (v/v) for 15 min, twice. 12. The gel particles were completely covered with acetonitrile. Extra acetonitrile was removed once the gel particles finished shrinking, and then the gel particles were allowed to air dry. 13. To this, enough freshly prepared trypsin enzyme solution (25 mM NH4HCO3 with 10 ng/mL trypsin) was added and the solution was incubated for overnight at 37°C. The trypsin solution can be added to cover the gel pieces. 14. The excess solution was carefully transferred to sterile 0.5-mL tubes and the gel pieces were extracted twice with 50% acetonitrile and 5% triflouroacetic acid. 15. These two extracts were combined with the earlier one (step 14) and concentrated with Speedvac. 3.10. ZIpTipc18 Purification (see Note 10)

1. The ZipTipc18 was wetted by depressing a pipette plunger to a dead stop, using the maximum volume setting of 10  mL, and aspirating the wetting solution into the tip. This procedure was repeated twice. 2. The ZipTipc18 was equilibrated with equilibration solution and dispensed to waste. This procedure was repeated two times. 3. To bind peptides and/or proteins to ZipTipc18, the pipette plunger was depressed to a dead stop into the sample. The sample was aspirated and dispensed using seven to ten cycles for maximum binding of complex mixtures. 4. The wash solution was aspirated into the tip and dispensed to waste (see Note 11). 5. The peptides and/or proteins were eluted with 4 mL of elution solution into a clean vial using a standard pipette tip (see Note 12). These peptides are pure and would be used for MALDI-TOF measurements.

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1. Aliquots of the sample measuring 2 mL were mixed with 1.5 mL of matrix (a saturated solution of a-cyano-4-hydroxycinnamic acid in 50% acetonitrile/0.1% trifluoroacetic acid) and a droplet of the mixture (about 1.5 mL) was spotted on the sample plate and allowed it to air dry for 5 min (18, 19) (see Notes 13–15). 2. Mass spectra were collected on Bruker’s autoflexIII mass spectrometer and analyzed with Flux analysis. 3. From the monoisotopic masses obtained, the amino acid sequences of matching tryptic fragments of Chlamydomonas thylakoid proteins were obtained using the MASCOT server.

3.12. Characterization of the Thylakoid Proteins

Figure 1 shows the 2D gel separation of thylakoid proteins. Spots 1 and 2 have been digested with trypsin and the obtained peptides were separated using MALDI-TOF mass spectrometry (Fig.  2). The monoisotopic masses were blasted using the MASCOT server. After searching the monoisotopic masses into MASCOT of spots 1 and 2 revealed the results as shown below: Spot: 1. Q93WL4_CHLRE Mass: 27420 Score: 45 Queries matched: 2 Light-harvesting chlorophyll a/b-binding protein LhcII – Chlamydomonas reinhardtii. Spot: 2. CB2_CHLRE Mass: 27003 Score: 100 Queries matched: 2 Chlorophyll a/b-binding protein of LHCII type I Chlamydomonas reinhardtii

Thus, any proteins from 2D gel spots could be prepared and analyzed using the same method of trypsin digestion and MALDITOF measurements, with appropriate BLAST searches.

4. Notes 1. Common problems with 2D PAGE are salts, polysaccharides, nucleic acids, and lipids. However, salt causes horizontal streaking in 2D electrophoresis. 2. Water-saturated chloroform can be prepared by mixing 4 mL of chloroform and 4 mL of MilliQ water in a 20 mL beaker. The mixture was stirred well for 10 min and allowed it to separate as two phases. Repeat the above step by adding another 4 mL of MilliQ water and than upper phase was removed by leaving a small layer. 3. The pellet should not be completely dry. Immediately add a suitable rehydration buffer. A completely dried pellet is difficult to dissolve in the rehydration buffer.

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Fig. 2. MALDI-TOF mass spectrum of the tryptic digests of thylakoid proteins measured in reflector mode. Upper panel corresponds to spot 1 and lower panel to spot 2 exercised from 2D gel electrophoresis (Fig. 1). The extracted monoisotopic peak list was submitted to the database search program of MASCOT. m/z is the mass-to-charge ratio.

4. Urea is sensitive to heat, so it should not be heated beyond 37°C to prevent carbomylation of proteins and the subsequent shift in isoelectric point.

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5. During equilibration steps, the gel side should face up, to facilitate good contact with buffer. Prepare the fresh equilibration buffers just before use. 6. Before completing this step, make sure that all the set-up for second dimension is ready with buffers and overlay agarose. 7. Glass plates should be thoroughly washed. It is the most common problem that causes point streaking. 8. Do not trap any air bubbles beneath the strip (gel side) while loading the IPG strips to the resolving gel to avoid vertical gaps in 2D pattern. 9. In gel digestion is the same for any spot of 2D gels. Also, measuring MALDI-TOF for any of the protein spots is similar. 10. ZipTipc18 purification is very important to avoid the salts and other contaminations. Thus, the peptides purified from this would give the accurate results in mass spectrometry. 11. A 5% methanol in 0.1% triflouroacetic acid/water wash can improve desalting efficiency. 12. Acetonitrile and methanol are volatile and evaporation can occur rapidly. If this occurs, add more eluent to recover the sample. Carefully aspirate and dispense the eluent through ZipTip at least three times without introducing air bubbles into the sample. Sample recovery can be improved by increasing the elution volume to 5 mL. 13. Spread the droplet and allow it to dry. It appears like a thin matrix film that is slightly yellowish in appearance. 14. All of the steps should be carried out using gloves. It is easy to contaminate the protein samples with keratin from the skin, and this can dominate the mass spectrum, making analysis of the fragmentation pattern very difficult. 15. In order to get promising results, high-grade chemicals should be used.

Acknowledgments RS is gratefully acknowledged for the financial support from DST NO.SR/SO/BB-34/2006, DST-FAST track NO SR/FT/ L-89/2006 and DB-RGYI, Government of India. VY and SN thank CSIR and UGC India for a fellowship in the form of JRF and SRF. We thank Mark Hunter, Department of Chemistry and Biochemistry, Arizona State University, USA, for critical reading of this chapter. We greatly acknowledge the School of Life Sciences, UOH-CREBB, for providing the facility of MALDITOF mass spectrometry at the University of Hyderabad.

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References 1. Rochaix, J. D. (2002) Chlamydomonas, a model system for studying the assembly and dynamics of photosynthetic complexes. FEBS Lett. 529, 34–38. 2. Elrad, E. D. and Grossman, A. R. (2004) A genome’s-eye view of the light-harvesting polypeptides of Chlamydomonas reinhardtii. Curr. Genet. 45, 61–75. 3. 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. 4. Julian, P. W. (2003) Thylakoid membrane proteomics. Photosynth. Res. 78, 265–277. 5. 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. 6. Ephritikhine, G., Ferro, M., and Rolland, N. (2004) Plant membrane proteomics. Plant Physiol. Biochem. 42, 943–962. 7. Jiang, L., He, L., and Fountoulakis, M. (2004) Comparison of protein precipitation methods for sample preparation prior to proteomic analysis. J. Chromatogr. A 1023, 317–320. 8. VanWijk, K. J. (2004) Plastid proteomics. Plant Physiol.Biochem. 42, 963–977. 9. Chen, X., Zhang, W., Xie, Y., Lu, W., and Zhang, R. (2007) Comparative proteomics of thylakoid membrane from a chlorophyll b-less rice mutant and its wild type. Plant Sci. 173, 397–407. 10. Kashino, Y., Harayama, T., Pakrasi, H., and Satoh, K. (2007) Preparation of membrane proteins for analysis by two-dimensional gel electrophoresis. J. Chromatogr. B 849, 282–292.

11. Mitra, S. K., Gantt, J. A., Ruby, J. F., Clouse, S. D., and Goshe, M. B. (2007) Membrane proteomic analysis of Arabidopsis thaliana using alternative solubilization techniques. J. Proteome Res. 6, 1933–1950. 12. Ramesh, V. M. and Webber A. N. (2004) Rapid isolation of photosystem I chlorophyllbinding protein from Chlamydomonas reinhardtii. Meth. Mol. Biol. 274, 19–28. 13. Arnon, D. I. (1949) Copper enzymes in isolated chloroplasts. polyphenoloxidase in Beta vulgaris. Plant Physiol. 24, 1–15. 14. Wessel, D. and Fluegge, U. I. (1984) A method for the quantitative recovery of protein in dilute solution in the presence of detergents and lipids. Anal. Biochem. 138, 141–143. 15. Hippler, M., Klein, J., Fink, A., Allinger, T., and Hoerth, P., (2001). Towards functional proteomics of membrane protein complexes: analysis of thylakoid membranes from Chlamydomonas reinhardtii. Plant J. 28(5), 595–606. 16. Bradford, M. M. (1976) A rapid and sensitive method for the quantitation of microgram quantities of protein using the principle of protein-dye binding. Anal. Biochem. 72, 248–254. 17. Chevallet, M., Luche, S., and Rabillound, T. (2006). Silver staining of proteins in polyacrylamide gels. Nat. protoc. 1, 1852–1857. 18. Shevchenko, A., Wilm, M., Vorm, O., and Mann, M. (1996) Mass spectrometric sequencing of proteins from silver stained polyacrylamide gels. Anal. Chem. 68, 850–858. 19. Subramanyam, R., Jolley, C., Brune, D.C., Fromme, P. and Webber, A.N., (2006) Characterization of a novel photosystem I-LHCI supercomplex isolated from Chlamydomonas reinhardtii under anaerobic (State II) conditions. FEBS Lett. 580, 233–238.

Chapter 15 Thylakoid Phosphoproteins: Identification of Phosphorylation Sites Anne Rokka, Eva-Mari Aro, and Alexander V. Vener Abstract Redox-dependent thylakoid protein phosphorylation regulates both the short- and long-term acclimation of the photosynthetic apparatus to changes in environmental conditions. The major thylakoid phosphoproteins belong to photosystem II (D1, D2, CP43, PsbH) and its light-harvesting antenna (Lhcb1, Lhcb2, CP29), but a number of minor phosphoproteins have also been identified. The detection methods traditionally include the radiolabeling techniques, electrophoretic separation of the phosphorylated and unphosphorylated forms of the protein, and the use of phosphoamino acid antibodies or phosphoprotein-specific dyes. The recent progress in mass spectrometry techniques and methods of proteomics allow for the successful identification and analyses of protein phosphorylation. In mass spectrometry approaches no exogenous tracer is needed and natural phosphorylation of proteins can be characterized with high sensitivity yielding the mapping of exact phosphorylation sites in the proteins as well. Various methods for the detection of thylakoid phosphoproteins, including the preparation of phosphopeptides for mass spectrometric analyses and techniques for phosphopeptide identification by electrospray ionization mass spectrometry (ESI-MS) are described. The experimental protocols for simultaneous identification of multiple phosphopeptides in complex peptide mixtures, enrichment of phosphopeptides by immobilized metal affinity chromatography (IMAC), and for their sequencing by tandem spectrometry are outlined. Key words: Light-harvesting protein of photosystem II, Reaction center proteins D1 and D2, CP43 antenna protein, PsbH protein, 12  kDa thylakoid phosphoprotein, Phospho-threonine (P-Thr) antibody, Mass spectrometry, 32P-ATP–labeling, IMAC

1. Introduction Several thylakoid proteins undergo reversible phosphorylation, the phenomenon, which is regulated by the redox state of thylakoid electron transfer components (1). Numerous thylakoid phosphoproteins have been reported, but the most vigorously studied ones are associated with photosystem II. Four of the Robert Carpentier (ed.), Photosynthesis Research Protocols, Methods in Molecular Biology, vol. 684, DOI 10.1007/978-1-60761-925-3_15, © Springer Science+Business Media, LLC 2011

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photosystem II core proteins, the reaction center D1 and D2 proteins, the chlorophyll a binding internal antenna protein CP43, and the small PsbH subunit are well-characterized phosphoproteins (2). Additionally, the minor antenna protein CP29 (Lhcb4) (3) and the two major LHCII proteins, Lhcb1 and Lhcb2 (2), are reversibly phosphorylated. Several novel thylakoid phosphoproteins have recently been reported, the thylakoid soluble phosphoprotein of 9 kDa (TSP9) being of particular interest in showing reversible association with the thylakoid membrane (4, 5). Although the exact physiological role of thylakoid phosphoproteins is still under discussion, they are likely to have various regulatory functions, including signaling for the regulation of gene expression, possibly both in the chloroplast and nuclear compartments. All so far best characterized thylakoid phosphoproteins become phosphorylated at threonine residues at or close to the N-terminus (6). From one up to three phosphorylation sites have been mapped in various thylakoid phosphoproteins (2, 4, 7, 8). Here, we describe the most commonly used methods to detect the phosphorylation of higher plant thylakoid proteins: labeling with 32P-ATP, phospho-specific dyes, immunoblotting with phospho-threonine (P-Thr) or protein-specific antibodies and mass spectrometric approaches. The mass spectrometry techniques allow also the mapping of exact phosphorylation sites in phosphoproteins.

2. Materials 2.1. Isolation of Thylakoids (Quick Protocol)

1. Homogenizer (Ultra Turrax T5FU) (IKA-Labortechnik Staufen, Germany). 2. Miracloth (Calbiochem). 3. NaF. 4. Buffer 1: 50 mM HEPES-NaOH, pH 7.4, 300 mM sucrose, 5 mM MgCl2, 1 mM Na-EDTA, 10 mM NaF, and 1% BSA. 5. Buffer 2: 10  mM HEPES-NaOH, pH 7.4, 5  mM sucrose, 5 mM MgCl2, and 10 mM NaF. 6. Buffer 3: 10 mM HEPES-NaOH, pH 7.4, 100 mM sucrose, 5 mM NaCl, 10 mM MgCl2, and 10 mM NaF. 7. Porra acetone: 80% acetone buffered with 25 mM HEPESKOH, pH 7.8.

2.2. SDS-PAGE

1. SDS-PAGE equipment. 2. Separating buffer: 1.5 M Tris–HCl, pH 8.8. 3. Stacking buffer: 0.5 M Tris–HCl, pH 6.8.

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4. 50% acrylamide/1.33% bis-acrylamide solution. 5. Ammonium persulphate: prepare fresh 10% solution in water before use. 6. N,N,N,N´-tetramethyl-ethylenediamine (TEMED, BioRad). 7. 20% SDS. 8. Laemmli solubilization buffer and electrophoresis buffer (9). 9. Prestained molecular weight marker (BioRad). 2.3. Western Blotting

1. P-Thr antibody (Zymed Laboratories Inc., San Francisco, CA and New England BioLabs). 2. AP-Goat Anti-Rabbit IgG (Zymed Laboratories Inc., San Francisco, CA). 3. BSA for blocking (Sigma). 4. PVDF membrane (polyvinylidene fluoride) (Millipore). 5. Whatman 3MM filter paper. 6. A semidry transfer system. 7. TBS: 20 mM Tris–HCl, pH 7.5, and 500 mM NaCl. 8. TTBS: TBS containing 0.05% Tween-20. 9. Chemiluminescence kit (New England BioLabs). 10. Fluorography cassettes or phosphoimager.

2.4. Phos-Tag Staining

1. Phos-tag Phosphoprotein Gel Stain (PerkinElmer). 2. Phos-tag dye-compatible gel imager (see Note 1). 3. Gel fixing solution: 50% methanol/10% acetic acid.

2.5. 32P-ATP Labeling and Isolation of Thylakoids

1. [g-32P] ATP. 2. Buffer 4: 50  mM Na2HPO4–NaH2PO4 (NaPB), pH 7.5, 0.3 M sucrose, and 5 mM MgCl2. 3. Osmotic shock medium: 10  mM NaPB, pH 7.5, 5  mM MgCl2, and 5 mM NaCl. 4. Buffer 5: 20 mM Tricine, pH 7.8, 5 mM MgCl2, and 5 mM NaCl. 5. Buffer 6: 50 mM Tricine, pH 7.8, 5 mM MgCl2, and 0.1 M sorbitol.

2.6. Mass Spectrometric Analyses of Phosphoproteins

1. Buffer 7: 330  mM sorbitol, 50  mM sodium phosphate, pH  7.8, 1  mM EDTA, 10  mM NaF, 0.15% BSA, 4  mM sodium ascorbate, and 7 mM l-cyctein. 2. Buffer 8: 10 mM sodium phosphate, pH 7.8, 5 mM MgCl2 and 10 mM NaF. 3. Buffer 9: 100  mM sorbitol, 50  mM sodium phosphate, pH 7.8, 5 mM MgCl2, and 10 mM NaF.

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4. Sequencing-grade modified trypsin (Promega, Madison, WI). 5. Acetyl chloride. 6. Anhydrous methanol (Aldrich). 7. Chelating Sepharose Fast Flow (Pharmacia). 8. FeCl3. 9. Acetonitrile (HPLC-grade). 10. ZipTipC18 (Millipore). 11. Triflouroacetic acid. 12. Formic acid.

3. Methods The methods described below outline four possible ways to determine thylakoid protein phosphorylation. ‘The 32P-ATP labeling, detection with P-Thr antibodies or phosphospesific dyes all are supplementary to each others, allow the detection of phosphoproteins and the estimation of their relative amounts. The mass spectrometric analyses provide information on exact phosphorylation sites in the proteins. Labeling of proteins with 32P orthophosphate is convenient for unicellular organisms but sometimes cumbersome for intact leaves and therefore the method is not described here. 3.1. Detection of Thylakoid Phosphoproteins by P-Thr Antibodies

This method has an advantage over labeling experiments in allowing the detection of endogenous level of thylakoid protein phosphorylation in  vivo under particular environmental conditions (Fig. 1). To avoid dephosphorylation during thylakoid isolation, all the isolation buffers should contain 10 mM NaF. When used to study the endogenous phosphorylation levels in vivo, the control experiments should include samples illuminated in far red (<700  nm) and red light (650 ± 10  nm) to completely dephosphorylate and phosphorylate the thylakoid proteins, respectively. Also a dilution series of thylakoids from leaves illuminated with red light should be included to reveal the linearity of the immunoresponse. Method is suitable also for studies on the regulation of thylakoid protein phosphorylation using intact chloroplasts, isolated thylakoid membranes or membrane subfractions. The intensity of the immunoresponse with different commercial antibodies differs between various phosphoproteins (Fig. 2) (see Note 2). The origin of immunoresponse with commercial P-Thr antibodies should always be specified (10, 11).

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Fig. 1. LHCII protein phosphorylation in differentially light-treated leaves demonstrated by immunoblotting with P-Thr antibodies and by [g-32P] ATP labeling. After dark adaptation, the leaves of mature pea plants were illuminated under different light conditions for 2 h before the isolation of thylakoid membranes. (a) LHCII protein phosphorylation in vivo detected by P-Thr antibody. (b) Autoradiogram of thylakoid membranes after in vitro phosphorylation with [g-32P] ATP (including 0.4 mM cold ATP). (c) P-Thr immunoblot following [g-32P] ATP labeling of thylakoid proteins, thus demonstrating the total pool of Thr-phosphorylatable LCHII proteins in thylakoid samples. Various light conditions for the treatment of intact pea leaves before thylakoid isolation are indicated as D darkness; LL 50 mmol photons/m2 s; HL 600 mmol photons/m2 s; and PSI and PSII light favoring PSI and PSII excitation, respectively (Figure has been published originally (22))

Fig. 2. Immunodetection of thylakoid phosphoproteins by two commercial antibodies. Lane 1, a polyclonal P-Thr antibody from New England BioLabs and lane 2, a polyclonal P-Thr antibody from Zymed Laboratories Inc. Thylakoids were isolated from spinach leaves, which had been illuminated for 30 min under 1,500 mmol photons/m2 s before thylakoid isolation. Thylakoid sample corresponding to 1 mg of chlorophyll was loaded in the well and the proteins were separated as described in Subheading 3.1.2. 3.1.1. Isolation of Thylakoid Membranes After In Vivo Phosphorylation (A Quick Protocol)

Leaves to be analyzed should be directly frozen in liquid nitrogen. All solutions are supplemented with 10  mM NaF to avoid dephosphorylation during thylakoid isolation and subfractionation. BSA (any high quality grade) and NaF are added before

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use. Avoid having thiol oxidants or reductants in the isolation buffers. All procedures should be carried out at 4oC under dim light. 1. Homogenize leaves (2 g) in 5 ml of a grinding buffer (buffer 1) for 5 s with a homogenizer/ultraturrax. 2. Filter the homogenate through Miracloth, and centrifuge at 1,500 × g for 4 min. 3. Resuspend the pellet in a lysis buffer (buffer 2) (4 ml), and centrifuge at 3,000 × g for 5 min. 4. Finally, resuspend the thylakoid pellet in a storage buffer (buffer 3) (50–100 ml). 5. Determined the chlorophyll concentration according to (12). 6. Freeze the thylakoids rapidly in liquid nitrogen and store at −70°C (see Note 3). 3.1.2. Separation of Thylakoid Proteins by SDS-PAGE and Transfer of Proteins to a PVDF Membrane

SDS-PAGE is run according to (9). 1. Prepare 0.75–1.0  mm thick polyacrylamide gels (160 ×  200 mm) using 15% acrylamide with 6 M urea in the separation gel. 2. Solubilize the thylakoid sample with Laemmli sample buffer for 5 min at 65°C. 3. Load the thylakoid sample corresponding to 1 mg chlorophyll in each well. 4. Run the gels at 10 mA/gel for 16 h (see Note 4). 5. Transfer the proteins to a PVDF membrane (Immobilon-P) according to manufacturer’s instructions (see Note 5).

3.1.3. Visualization of Proteins with P-Thr Antibody

1. Block the membrane with 5% bovine serum albumin prepared in TBS, for 1 h. 2. Incubate the membrane in primary antibody overnight (see Note 6). 3. Wash the membrane four times for 5 min with TTBS. 4. Incubate the membrane in the secondary antibody for 1–2 h. 5. Wash the membrane five times for 5 min with TTBS and two times for 5 min with TBS. 6. Detect phosphoproteins using a chemiluminescence kit according to manufacturer’s instructions (see Note 7). 7. For relative quantifications, scan the blots and compare intensities with controls included in the gel side by side with the samples.

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3.2. Immunodetection with Protein Specific Antibodies

3.2.1. Electrophoretic Separation of the Thylakoid Proteins and Visualization of D1 Protein Phosphorylation

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For some thylakoid proteins, it is also convenient to separate the phosphorylated and nonphosphorylated forms of the protein based on the slightly slower mobility of former during SDSPAGE, and then use a protein-specific antibody to detect both forms of the protein (10, 13, 14). For quantitative purposes, care should be taken to verify that both forms of the protein crossreact equally with the antibody. Successful separation of the phosphorylated and nonphosphorylated forms of the photosystem II D1 protein can be obtained using the following conditions. 1. Prepare and load gels as described in Subheading 3.1.2. 2. Run the gels at 30 mA/gel for 16 h (see Note 8). 3. Transfer the proteins to PVDF membrane according to manufacturer’s instructions. 4. Block the membrane with 6% fat-free milk, prepared in TBS, for 1 h. 5. Incubate the membrane in primary, D1 antibody overnight. 6. Continue as described under Subheading 3.1.3, step 3. 7. Quantify the immunoresponse using a laser densitometry and a software package for image quantification. Use appropriate standards and take care to work on the linear response range (see Note 9).

3.3. Detection of Thylakoid Phosphoproteins with Phospho-Specific Dye Phos-Tag

The benefit of this method compared to the detection of proteins with phospho-specific antibodies, is that proteins can be directly identified with mass spectrometry after in-gel digestion. Phos-tag phosphoprotein gel stain can detect close to 1 ng of phosphoprotein per band or a spot, depending on the phosphorylation state of the protein. The signal strength is linear over threefold of magnitude. Counterstaining with a variety of general total protein stains (like silver, Coomassie blue, or SYPRO Ruby) after Phostag staining is a useful way to estimate the phosphorylation state of a certain phosphoprotein. 1. Prepare, load, and run a gel as described in Subheading 3.1.2. 2. Fix the gel in fixing solution for 30 min. Use a rotary shaker to agitate the gel in the fixing solution (see Note 10). 3. Discard the fixer and repeat the fixation for 1 h to overnight. 4. Discard the fixer and incubate the gel in deionized water with gentle agitation for 10 min (see Note 11). 5. Repeat the step 4 twice. 6. Prepare Phos-tag Stain by diluting Phos-tag dye concentrate 1:100 with Stain Buffer (which are both in the kit). 25  ml ready Phos-tag Stain is adequate for one mini-gel and 170 ml for one 16 × 20 cm gel.

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7. Pour the staining solution into a clean plastic dish and place the gel into it. Protect the gel and staining solution from light all the times. Gently agitate the gel in the stain for 90 min at room temperature. 8. Dilute 60 ml of destain buffer (2× concentrate) with 60 ml of deionized water. 9. Discard the stain solution, and agitate the gel in 40  ml of destain buffer (1×) for 30 min. 10. Repeat the step 9 twice. 11. Discard the destain buffer, and agitate the gel in deionized water for 5 min. 12. Repeat the wash. 13. Image the gel using the available gel imaging system. 3.4. Labeling of Thylakoid Phosphoproteins with 32P-ATP

3.4.1. Isolation of Thylakoids for In Vitro Labeling with 32P-ATP

This method is convenient for intact chloroplasts and isolated thylakoid membranes. Because the method detects those phosphoproteins that are not already phosphorylated in the samples, it is important to isolate chloroplasts/thylakoids from plants kept under conditions that do not favor phosphorylation. If no labeling of phosphoprotein is detected for a particular protein known to be a phosphoprotein, either the entire pool of phosphoprotein is already phosphorylated in the sample under study or the kinase responsible for phosphorylation is inactive. When using [g-32P] ATP labeling, it is convenient to check the endogenous phosphorylation level of the protein under interest first with P-Thr antibodies (Fig. 2). 1. Homogenize leaves (20  g) in 40  ml of preparation buffer (buffer 4) three times for 3 s in a rotating-knife blender. 2. Filtrate the homogenate through four layers of nylon net (a pore size of 20 ml) or Miracloth. 3. Centrifuge the filtrate at 1,000 × g for 3 min. 4. Resuspend the pellet in the preparation buffer (buffer 4) (40 ml), and centrifuge at 2,400 × g for 5 min. 5. Resuspend the pellet in 40 ml of the osmotic shock medium, and homogenize three times with a Potter homogenizer. 6. Centrifuge at 2,400 × g for 5 min, and wash the pellet with a washing buffer (buffer 5). 7. Centrifuge at 2,400 × g for 5 min, and finally resuspend the thylakoid pellet in 3 ml of a phosphorylation buffer (buffer 6).

3.4.2. Labeling of Thylakoid Proteins with [g-32P] ATP and Visualization of the Phosphoproteins

1. Incubate thylakoids (0.3 mg chlorophyll/ml) in the presence of 0.1 mM [g-32P] ATP (0.02 mCi/mg chlorophyll) under a photon flux density of 300 mmol photons/m2 s at room temperature for 30 min (see Note 12).

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2. Spin down the thylakoids in an Eppendorf centrifuge with full speed for 1.5  min, and wash the excess label with cold phosphorylation buffer. 3. Centrifuge as above and resuspend the pellet in suitable buffer for further study. 4. Separate thylakoid proteins with SDS-PAGE as described under Subheading 3.1.2. 5. Dry the gel between two cellophane sheets, and expose to X-ray film at room temperature or use the phosphoimager. Take care to work on the linear response range. 3.5. Mass Spectrometric Analyses of Phosphoproteins

3.5.1. Preparation of the Surface-Exposed Peptides from Thylakoid Membranes

The methods described below outline preparation of the surfaceexposed peptides from thylakoid membranes, peptide esterification, enrichment of the phosphopeptides by immobilized metal affinity chromatography (IMAC), identification of phosphopeptides using electrospray ionization mass spectrometry (ESI-MS), and sequencing of the phosphorylated peptides by tandem mass spectrometry. 1. Homogenize 20  g of Arabidopsis or spinach leaves with a blender in 170 ml of ice-cold preparation buffer (buffer 7). 2. Filtrate the suspension through four layers of Miracloth and centrifuge for 3 min at 1,000 × g. 3. Resuspend the chloroplast pellet in 60 ml of the preparation buffer, and centrifuge for 5 min at 3,000 × g. 4. Resuspend the pellet in 50 ml of lysis buffer (buffer 8), and homogenize five times in a Potter grinder. 5. Centrifuge for 5 min at 7,500 × g. 6. Wash the pellet of thylakoid membranes twice with the wash buffer (buffer 9) and three times with 25  mM NH4HCO3 (pH 8.0) and 10 mM NaF (see Note 13). Centrifuge the thylakoids for 5 min at 6,000 × g after each resuspension. 7. Finally, resuspend the thylakoid pellet to 3  mg of chlorophyll/ml in 25  mM NH4HCO3 to the final volume about 2–3 ml, (see Note 14). 8. Incubate the thylakoid suspension with sequencing-grade modified trypsin (8 mg trypsin/mg chlorophyll) at 22oC for 3 h (see Note 15). 9. Freeze the sample in liquid nitrogen, thaw, and centrifuge the digestion products for 20 min at 14,000 × g. The supernatant containing released thylakoid peptides is used for mass spectrometric analyses directly or after enrichment of the phosphopeptides by IMAC. The released peptides could be methyl-esterified (19) for the blocking of peptides carboxylic groups. This will prevent unspecific binding of nonphosphorylated peptides to IMAC column.

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3.5.2. Esterification of Isolated Peptides

1. Lyophilize the peptides to dryness (see Note 16). 2. Add four times 20  ml of acetyl chloride to 0.5  ml of anhydrous methanol dropwise with stirring. The mixture is explosive. Therefore, the procedure should be carried out under the hood. 3. Add the reagent to lyophilized peptides (250 ml per 50 mg of chlorophyll used for trypsin digestion) and incubate at room temperature for 2 h with occasional vortexing. 4. Lyophilize the modified peptides to dryness. The methyl-esterification of peptides increases peptide mass of 14 Da per modification. This mass increment should be taken into account when samples are analyzed by mass spectrometry.

3.5.3. Enrichment of the Phosphopeptides by Immobilized Metal Affinity Chromatography

Enrichment of the phosphopeptides facilitates their subsequent identification and sequencing by mass spectrometry. Phospho­ peptides are affinity-enriched from the thylakoid peptides (prepared as described in Subheadings 3.5.1 and 3.5.2) by chromatography with immobilized Fe(III) or Ga(III) columns (see Note 17). 1. Pack columns with 50 ml of Chelating Sepharose Fast Flow beads. 2. Wash with 0.3  ml of water, followed by 0.3  ml 0.1% (v/v) acetic acid, and charge with 0.3 ml of 0.1 M FeCl3 or GaCl3. 3. Wash the unbound salts with 0.5 ml of 0.1% (v/v) acetic acid. 4. Mix the thylakoid peptides (0.2–0.3 ml) with an equal volume of 20% acetic acid and load onto the columns. 5. After washing the columns twice with 0.2 ml of 0.1% (v/v) acetic acid, elute the phosphopeptides with 300 ml of 20 mM of nonbuffered Na2HPO4. The phosphopeptides enriched by IMAC could be directly analyzed by LC-MS. However, for ESI-MS without prior HPLC, they first should be desalted on ZipTips. 1. Prewet ZipTipC18 with 50% acetonitrile in water and equilibrate with 0.1% triflouroacetic acid in water. 2. Load the peptides by pipetting of 10–20  ml of the peptide sample. 3. Then, wash the ZipTipC18 with 1% triflouroacetic acid in water. 4. Elute the desalted peptides by 10 ml of 50% acetonitrile in water.

3.5.4. Identification of Phosphopeptides Using ESI-MS

For selective identification of phosphopeptides in complex peptide mixtures, the precursor-ion scanning analyses by ESI-MS in negative ion mode is preferable (see Note 18). This type of mass spectrometric analysis could be performed using electrospray

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ionization triple quadrupole or hybrid (quadrupole-time-of-flight) mass spectrometers (see Note 19) equipped with either normal ion spray or nanoelectrospray ion source. The samples after desalting on ZipTips (see Subheading  3.5.3) should not be acidified with triflouroacetic acid or formic acid, which decreases negative ionization of the peptides. The peptides released from the surface of thylakoid membrane by trypsin (see Subheading 3.5.1) may be subjected to direct analysis being dissolved in water with 25 mM NH4HCO3. The nanoelectrospray capillaries are loaded with 2 ml of peptide solution and negative ionization mode full scan spectra are first recorded to determine the range of mass to charge ratios (m/z) of the major negative molecular ions present in the peptide mixture. Then, the precursor-ion scanning analyses with the detection of phosphoryl ions −79  m/z (PO3–) and −97  m/z (H2PO4–) are performed. The typical result of the precursor-ion scanning experiment for thylakoid peptides is shown in Fig.  3. The ­precursor-ion scanning experiment provides the information on the m/z for phosphorylated peptides present in the mixture. To determine the mass of the corresponding phosphopeptides, the full scan spectrum should be examined in the regions of m/z

Fig. 3. The precursor of −79 m/z ion (PO3-) spectrum for the mixture of the peptides released from the thylakoid membranes of Arabidopsis thaliana by trypsin. The spectrum is a result of 100 scans accumulated on a hybrid mass spectrometer API Q-STAR Pulsar i (Applied Biosystems, Foster City, CA) equipped with a nano-electrospray ion source (MDS Protana, Odense, Denmark). The positions of the signals produced by the major phosphopeptides present in the mixture are indicated by the names of corresponding proteins with the superscripts showing the negative charge state of corresponding ions. The sequences of the phosphopeptides are presented in the inset.

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with the signals in ­precursor-ion experiment. This allows the determination of the charge state of the corresponding phosphopeptide and calculation of its mass. The latter is used for the calculation of the m/z for the positive ions of corresponding phosphopeptides that are then subjected for tandem mass spectrometry as described in Subheading 3.5.5. 3.5.5. Sequencing of the Phosphorylated Peptides by Tandem Mass Spectrometry

The ESI-MS instruments used for the identification of the phosphorylated peptides (see Subheading 3.5.4) are also exploited for the fragmentation and sequencing of these peptides. To this end, the peptides should be ionized as positively charged, protonated ions. To increase the positive ionization, the peptide solutions (see Subheading 3.5.1 and 3.5.3) are acidified by the addition of formic acid to the final concentration 1–2% prior to the analyses. Complete sequencing of the phosphorylated peptides requires clean selection of their molecular ions for fragmentation (see Note 20), which is usually not feasible from the total nonfractionated mixture of thylakoid peptides (see Subheading 3.5.1). The mixtures of the phosphopeptides enriched by IMAC (see Subheading 3.5.3) are suitable for direct tandem mass spectrometry and sequencing. The nanoESI capillaries are loaded with 2 ml of the peptide mixture in 50% acetonitrile in water with 1% formic acid and positive ionization mode full scan spectra are first recorded. These spectra are analyzed for the presence of the ions with m/z ratios calculated from the results of the precursor ion analyses (see Subheading 3.5.4), according to the mass and possible ionization states of these phosphopeptides. The candidate ions are then selected and subjected to collision-induced dissociation. Inter­ pretation of the product ion spectra generally allows for the determination of the peptide sequence, including even complete de novo sequencing. The latter is usually feasible for the peptides not longer than 20 amino acids, including phosphopeptides (see Note 21).

4. Notes 1. There are available two different Phos-tag Phosphoprotein Gel Staining kits which are compatible with different excitation sources. The Phos-tag 540 dye can be excited with solid state YAG laser (532 nm), HeNe laser (543 nm), or xenon lamp with 540 nm excitation filter. Phos-tag 300/400 dye, which we used, can be used with UV transilluminator (excitation wave length 302 nm), blue LED laser (457 nm), argon laser (488 nm), or xenon lamp with 460 nm excitation filter. We used Geliance 1000 Imaging System (PerkinElmer).

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2. Commercial polyclonal P-Thr antibodies show very different cross-reaction with different thylakoid phosphoproteins. If you are interested in a specific phosphoprotein, it is worth checking of several available antibodies for best immunoresponse. With P-Thr antibodies, it is not possible to compare the phosphorylation intensities of different thylakoid phosphoproteins with each other. 3. Freezing and storage of thylakoids at −20°C should be avoided because of irreversible aggregation of proteins. 4. Do not run gels containing urea in cold room (4°C) because urea will precipitate. 5. Instead of using different anode and cathode buffers for protein transfer, they can be substituted with one continuous buffer system (48 mM Tris, 39 mM glycine, 0.0375% (w/v) SDS, and 20% methanol (v/v). 6. There are sometimes large variations in the immunoresponse between different antibody lots from the same manufacture. When you once find the most suitable antibody for your ­purposes, it is wise to order enough it for the entire set of experiments. 7. Nitrocellulose membranes are not compatible with the chemiluminescense kit, which has been mentioned in the Materials section. 8. Separation of the phosphorylated and nonphosphorylated forms of the D1 protein is obtained only if the proteins are allowed to migrate a long distance in the gel. It is good to use colored protein molecular weight marker to monitor the migration. 9. It is important to ensure that the used antibody will crossreact equally with both the phophorylated and nonphosphorylated protein forms. The antibody raised against a peptide corresponding to amino acids 234–242 of DE-loop of Synechocystis 6803 D1 shows similar reactivity to nonphosphorylated and phosphorylated D1 (15). 10. Polypropylene dishes are optimal for staining because the high-density plastic absorbs only a minimal amount of the dye. Clean and rinse the staining containers well before use to remove detergents which will interfere with staining. 11. It is important that the gel is completely immersed in water in order to remove all of the methanol and acetic acid which can interfere with Phos-tag stain. 12. To avoid dephosphorylation, add 10 mM NaF to phosphorylation mixture and to all solutions used after that step. But omit NaF in a case the phosphorylated thylakoids are used to study the dephosphorylation of phosphoproteins.

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13. The washes of thylakoid membranes with 25 mM ammonium bicarbonate (NH4HCO3) are important for two reasons. First, these washes lead to destacking of thylakoids, which increase the accessibility of trypsin to the surface-exposed regions of thylakoid membrane proteins. Second, the resulting supernatant of tryptic peptides in 25 mM NH4HCO3 is compatible with direct analyses by mass spectrometry. 14. The final resuspension of thylakoid membranes to a high chlorophyll concentration (about 3 mg of chlorophyll/ml) is important for good proteolytic digestion and for a high concentration of the resultant peptides for analyses. 15. It is important to make the proteolytic treatment at 22–25oC, but not at 37oC because a number of photosystem II phosphoproteins are rapidly dephosphotylated by the heat-shockactivated membrane protein phosphatase at 37oC (16). 16. The peptides should be lyophilized completely because any traces of water will decrease the efficiency of peptide esterification. 17. Using of IMAC with immobilized Fe(III) ions (17) as well as Ga(III) ions (18) is recommended because they could be complementary and neither of them is perfect. Specifically, different sets of thylakoid phosphopeptides could be enriched by either of these chromatographic techniques (7). The enrichment of the phosphopeptides could be increased by methylation of carboxylate groups in the peptides prior the IMAC (19); however, such a chemical modification leads also to a number of side reactions in the major thylakoid phosphopeptides. 18. The precursor-ion scanning technique allows for the selective determination of the mass to charge ratios (m/z) for phosphorylated peptides that produce the diagnostic phosphoryl ions −79 m/z (PO3–) and −97 m/z (H2PO4–) (4, 20, 21). In this type of mass spectrometric experiment, only the phosphorylated peptides present in the peptide mixture are detected because the detector is “blind” to all other, but the phosphoryl fragment ions and nonphosphorylated peptides do not produce signals. 19. In case of triple quadrupole mass spectrometer, the time required for a single precursor scan is shorter than that in a hybrid machine. This allows for the accumulation of a greater number of scans during the same experimental time if the ­triple quadrupole is used. However, hybrid mass spectrometers have higher sensitivity than triplequads. Moreover, both diagnostic phosphoryl ions −79  m/z (PO3–) and −97  m/z (H2PO4–) could be detected in the same experiment on a hybrid machine, while separate experiments should be ­performed for the ­detection of these ions by triple ­quadrupole.

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Detection of both ions is important because the intensity of the −79 and −97 m/z signals differs significantly for different thylakoid phosphopeptides (7). 20. The ions of individual phosphopeptides should be selected for fragmentation. The deconvolution of the phosphopeptide sequence is extremely hard if two or three different peptide ions penetrate through the selection window and fragmented simultaneously. Thus, the width of the selection window could be narrowed to 0.7 atomic unit. The narrowing of the selection increases the quality of the fragmentation spectra, but reduces the intensity of the signals. Accordingly, if the parent phosphopeptide ion is not in a close vicinity to the other peptide ions, the selection window could be broadened to 2 or 3 atomic unit width, which would increase the signal intensity due to the fragmentation of all isotopic forms of the phosphopeptide. 21. The fragmentation spectra of phosphopeptides allow the determination of the phosphorylated residues. A phosphoester bond between a phosphoryl group and a peptide is less stable than a peptide bond, which leads to a prominent neutral loss of phosphoric acid (H3PO4, 98 Da) from the positively charged peptide ions (7, 20, 21). The neutral loss of the phosphoric acid also occurs in the fragment ions containing phosphorylated residue. The series of y (C-terminal) and b (N-terminal) fragment ions without the neutral loss (originated from nonphosphorylated fragments of the peptide) together with the distinct ions that underwent the neutral loss (originated from phosphorylated peptide fragments) in each spectrum usually allow the identification of the phosphorylation sites.

Acknowledgments The work in authors laboratories is supported by the Academy of Finland, Nordic Energy Research (BioH2), Maj and Tor Nessling Foundation (E.-M.A.), The Swedish Research Council, and The Swedish Research Council for Environment, Agriculture and Spatial Planning (Formas) (A.V.V.). Figure 1 was reprinted with the permission of Blackwell Publishing Ltd. References 1. Vener, A. V. (2007) Environmentally modulated phosphorylation and dynamics of proteins in photosynthetic membranes. Biochim. Biophys. Acta 1767, 449–57. 2. Bennet, J. (1991) Protein phosphorylation in green plant chloroplasts. Annu. Rev. Plant Physiol. Plant Mol. Biol. 42, 281–311.

3. Bergantino, E., Dainese, P., Cerovic, Z., Sechi, S., and Bassi, R. (1995) A post-translational modification of the photosystem II subunit CP29 protects maize from cold stress. J. Biol. Chem. 270, 8474–81. 4. Carlberg, I., Hansson, M., Kieselbach, T., Schroder, W. P., Andersson, B., and Vener, A. V.

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6.

7.

8.

9. 10.

11.

12.

13.

Rokka, Aro, and Vener (2003) A novel plant protein undergoing light-induced phosphorylation and release from the photosynthetic thylakoid membranes. Proc. Natl. Acad. Sci. U.S.A. 100, 757–62. Fristedt, R., Carlberg, I., Zygadlo, A., Piippo, M., Nurmi, M., Aro, E.-M., Scheller, H. V., and Vener, A. V. (2009) Intrinsically unstructured phosphoproteins TSP9 regulates light harvesting in Arabidopsis thaliana. Biochemistry 48, 499–509. Rintamäki, E. and Aro, E.-M. (2001) Phosphorylation of photosystem II proteins, In Regulation of Photosynthesis (Aro, E.-M. and Andersson, B., eds.) Kluwer, Dordrecht, pp. 394–418. Vener, A. V., Harms, A., Sussman, M. R., and Vierstra, R. D. (2001) Mass spectrometric resolution of reversible protein phosphorylation in photosynthetic membranes of Arabidopsis thaliana. J. Biol. Chem. 276, 6959–66. Gómez, S. M., Nishio, J. N., Faull, K. F., and Whitelegge, J. P. (2002) The chloroplast grana proteome defined by intact mass measurements from liquid chromatography mass spectrometry. Mol. Cell Proteomics 1, 46–59. Laemmli, U. K. (1970) Cleavage of structural proteins during assembly of head of bacteriophage-T4. Nature 227, 680–5. Rintamäki, E., Salonen, M., Suoranta, U.-M., Carlberg, I., Andersson, B., and Aro, E.-M. (1997) Phosphorylation of light-harvesting complex II and photosystem II core proteins shows different irradiance-dependent regulation in vivo. Application of phosphothreonine antibodies to analysis of thylakoid phosphoproteins. J. Biol. Chem. 272, 30476–82. Bergo, E., Pursiheimo, S., Paakkarinen, V., Giacometti, G. M., Donella-Deana, A., Andreucci, F., Barbato, R., and Aro, E.-M. (2002) Rapid and highly specific monitoring of reversible thylakoid protein phosphorylation by polyclonal antibody to phosphothreonine-containing proteins. J. Plant. Phys. 159, 371–7. Porra, R. J., Thompson, W. A., and Kriedemann, P. E. (1989) Determination of accurate extinction coefficients and simultaneous-equations for assaying chlorophyll-a and chlorophyll-b extracted with 4 different solvents – Verification of the concentration of chlorophyll standards by atomic-absorption spectroscopy. Biochim. Biophys. Acta 975, 384–94. Callahan, F. E., Ghirardi, M. L., Sopory, S. K., Mehta, A. M., Edelman, M., and Mattoo, A. K.

14.

15.

16.

17.

18.

19.

20.

21.

22.

(1990) A novel metabolic form of the 32 kDa-D1 protein in grana-localized reaction center of photosystem II. J. Biol. Chem. 265, 15357–60. de Vitry, C., Diner, B. A., and Popot, J. L. (1991) Photosystem II particles from Chlamydomonas reinhardtii. Purification, molecular weight, small subunit composition, and protein phosphorylation. J. Biol. Chem. 266, 16614–21. Rintamäki, E., Salo, R., Lehtonen, E., and Aro, E.-M. (1995) Regulation of D1 protein degradation during photoinhibition of photosystem II in vivo: Phosphorylation of the D1 protein in various plant groups. Planta 195, 379–86. Rokka, A., Aro, E.-M., Herrmann, R. G., Andersson, B., and Vener, A. V. (2000) Dephosphorylation of photosystem II reaction center proteins in plant photosynthetic membranes as an immediate response to abrupt elevation of temperature. Plant Physiol. 123, 1525–36. Andersson, L. and Porath, J. (1986) Isolation of phosphoproteins by immobilized metal (Fe3+) affinity chromatography. Anal. Biochem. 154, 250–4. Posewitz, M. C. and Tempst, P. (1999) Immobilized gallium(III) affinity chromatography of phosphopeptides. Anal. Chem. 71, 2883–92. Ficarro, S. B., McCleland, M. L., Stukenberg, P. T., Burke, D. J., Ross, M. M., Shabanowitz, J., Hunt, D. F., and White, F. M. (2002) Phospho­ proteome analysis by mass spectrometry and its application to Saccharomyces cerevisiae. Nat. Biotechnol. 20, 301–5. Steen, H., Kuster, B., and Mann, M. (2001) Quadrupole time-of-flight versus triplequadrupole mass spectrometry for the determination of phosphopeptides by precursor ion scanning. J. Mass Spectrom. 36, 782–90. Shou, W., Verma, R., Annan, R. S., Huddleston, M. J., Chen, S. L., Carr, S. A., and Deshaies, R. J. (2002) Mapping phosphorylation sites in proteins by mass spectrometry. Methods Enzymol. 351, 279–96. Hou, C.-X., Pursiheimo, S., Rintamäki, E., and Aro, E.-M. (2002) Environmental and metabolic control of LHCII protein phosphorylation: revealing the mechanism for dual regulation of the LHCII kinase. Plant Cell Environ. 25, 1515-1525.

Chapter 16 Direct Detection of Free Radicals and Reactive Oxygen Species in Thylakoids Éva Hideg, Tamás Kálai, and Kálmán Hideg Abstract In plants, reactive oxygen species (ROS), also known as active oxygen species (AOS), are associated with normal, physiologic processes as well as with responses to adverse conditions. ROS are connected to stress in many ways: as primary elicitors, as products and propagators of oxidative damage, or as signal molecules initiating defense or adaptation. The photosynthetic electron transport is a major site of oxidative stress by visible or ultraviolet light, high or low temperature, pollutants or herbicides. ROS production can be presumed from detecting oxidatively damaged lipids, proteins, or pigments as well as from the alleviating effects of added antioxidants. On the contrary, measuring ROS by special sensor molecules provides more direct information. This chapter focuses on the application of spin trapping electron paramagnetic resonance (EPR) spectroscopy for detecting ROS: singlet oxygen and oxygen free radicals in thylakoid membrane preparations. Key words: EPR spectroscopy, Free radical, Leaf, Reactive oxygen species (ROS), Singlet oxygen, Spin trapping, Thylakoid

1. Introduction A free radical is defined as any species containing one or more unpaired electrons (1). Oxygen radicals are an important group among free radicals, although carbon- or sulfur-centered radicals and free radical oxides of nitrogen are also formed in living systems. Besides oxygen-centered radicals, the group of reactive oxygen species (ROS) also involves non-radical forms of oxygen, such as singlet oxygen or hydrogen peroxide. Although the collective terms “reactive species,” ROS, and “free radical” are frequently used as synonyms, the groups of chemical molecules they refer to are not identical (Fig.  1). All ROS are derivatives of molecular oxygen (Fig.  2). Superoxide is formed when one Robert Carpentier (ed.), Photosynthesis Research Protocols, Methods in Molecular Biology, vol. 684, DOI 10.1007/978-1-60761-925-3_16, © Springer Science+Business Media, LLC 2011

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Free radicals

ROS

RS

Oxygen radicals H2O2 RO

ROO ROOH

NO O2-

OH

CH3

O3 1O2

H

Fig. 1. Chemical classification of “reactive species.” Examples of oxygen free radicals: superoxide (O2•−), hydroxyl (•OH), peroxyl (•OOR), and alkoxyl (•OR) radicals. Examples of non-radical ROS: singlet oxygen (1O2), oxone (O3), hydrogen peroxide (H2O2), and other peroxides (ROOH). Free radicals that are not oxygen-centered are represented by sulfurcentered thiyl (RS•) and carbon-centered methyl (•CH3) radicals and the hydrogen atom (H•), which is the simplest free radical. Certain oxides of nitrogen such as NO• are also free radicals.

gy ner

E 2Ppx*

2Ppx

Singlet oxygen, 1 O2 (1Dg)

2Ppy*

2Ppy

Ele

ctro

2Psz Ground state,

sfer

tran

3O 2

n tr

ans

fe r

Superoxide radical, O2-

Peroxide ion, O22-

Fig. 2. Ground-state oxygen and its derivatives. Simplified illustration of electrons on molecular orbits.

electron is added to the ground-state oxygen molecule, addition of one more electron followed by protonation results in H2O2. Further reductions and protonations convert H2O2 into the neutral form, H2O. Singlet oxygen is also generated from molecular oxygen, but via energy transfer. In this reaction, the spin restriction of ground (triplet)-state oxygen is removed, so the oxidizing ability is very much increased (1–3).

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Plants, like any other living organism, constantly produce and extinguish ROS. Under natural conditions, there is usually no detectable pool of reactive species, because these react with other molecules shortly after being produced. Some of these reactions are harmful, leading to oxidative modification of membrane ­lipids, proteins, or DNA; while others, for example, the controlled cycling of electrons in the Asada–Halliwell cycle (4), the H2O2–dependent polymerization of phenols in lignin formation (1, 5), or cell wall loosening by ROS during growth (6, 7), are elements of ordinary plant metabolism. A complex system of enzymes and other ROS scavenging molecules keep the concentration of free radicals very low, contrary to continuous production (5, 8, 9). This balance may be tipped out under stress conditions, by increased radical production or by decreased antioxidant capacity. ROS production may increase as a result of imperfect electron transport, for example, in chloroplasts or in mitochondria, by electron leakage, or via energy transfer to oxygen. On the contrary, external sources (e.g., ozone, photosensitizers, or bipyridyl herbicides) may also induce free radical production in high concentrations at unexpected locations, compared to normal metabolic reactions. Scavenging these radicals may exhaust the antioxidant system, for example, by oxidizing the ascorbate and glutathione pools, if these cannot be restored (rereduced) fast enough. Also, adverse temperature or water conditions may decrease the activity of antioxidant enzymes, making them unable to process the increased load of free radicals. The photosynthetic electron transport is a major site of oxidative stress by visible or ultraviolet light, and pollutants or herbicides. High or low temperatures as well as adverse water conditions limit photosynthesis indirectly and may also trigger electron transport-related ROS (for reviews, see refs. 8–13). This chapter focuses on detecting radicals under conditions when the imbalance in the production and scavenging of reactive species allows specific external probes (ROS sensor molecules) to compete with the natural targets for ROS. Our examples are taken from research on singlet oxygen and free radical production in plants under stress by excess photosynthetically active radiation (photoinhibition). With minor adaptations, these techniques can be applied to study other conditions, when increased production of ROS is expected. Experimental work was partly supported by the Hungarian National Research Foundation (OTKA-NKTH K67597).

2. Materials 1. Grinding medium: 50 mM potassium phosphate, pH 7.2, 0.4 M sucrose, 15 mM NaCl, 5 mM MgCl2, and 5 mM ascorbate. 2. Resuspension buffer: 50 mM potassium phosphate, pH 7.2, 0.05 M sucrose, 15 mM NaCl, and 5 mM MgCl2.

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3. DMPO (5,5-dimethyl-pyrrolin N-oxide). 4. Tiron (1,2-dihydroxybenzene-3,5-disulfonate). 5. Clark-type oxygen electrode. 6. X-band spectrometers (e.g., Bruker ECS-106).

3. Methods 3.1. Thylakoid Membrane Isolation

1. Wash mature and healthy spinach leaves in tap water, dry on tissue or filter paper, and remove middle veins. 2. Chop the leaves in a blender with ice-cold grinding medium. Use approximately 120 mL of grinding medium per 200 g of leaf. Grind in repeated, short (10–15 s) intervals. 3. Filter the mixture through eight layers of cheesecloth and centrifuge at 7,000 g for 15 min. 4. Resuspend pellet in resuspension buffer to approximately one-third of the original volume. 5. Centrifuge at 300 g for 5 min in order to remove crude pellet and possible debris. 6. Centrifuge the supernatant again at 7,000 g for 15 min, and resuspended the pellet in the resuspension buffer to approximately 1,200–1,500 mg Chl/mL. 7. Carry out all centrifugation steps at 4°C, and keep the centrifuge tubes on ice while resuspending pellets. Store thylakoid preparations in 0.3–0.5 mL volumes at −80°C until use. 8. Once thawed, keep the samples on ice, in the dark, in resuspension buffer diluted to 60–200 mg Chl/mL until use. 9. Add ROS traps to diluted thylakoids at concentrations indicated below. 10. Once thawed, discarded surplus thylakoids or use them for experiments other than ROS detection. Hepes or Mes in the resuspension/reaction buffer may lower the amount of trapped ROS. Photosynthetic activity of the preparations should be checked, for example, by measuring steady-state oxygen evolution with a Clark-type oxygen electrode in the presence of an electron acceptor.

3.2. Spin Trapping EPR Spectroscopy

Electron paramagnetic resonance (EPR), also called electron spin resonance (ESR), spectroscopy detects the presence of unpaired electrons, providing an excellent technique for measuring free radicals. However, because free radicals are highly reactive and are usually present in small concentrations, their direct detection is practically impossible at room temperature, unless their radical

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Fig. 3. Representation of the principle of spin trapping indicating magnetic features and approximate lifetimes of the reagents and the product.

nature is stabilized by reacting with trapping molecules (14–17). The spin adducts produced in reactions between the free radical and the spin trap are also radicals but are reasonably stable for biological experiments (Fig. 3). Although singlet oxygen is not a free radical form of ROS, its reaction with specific traps may also yield EPR-detectable nitroxide radicals; therefore, 1O2 detection is also discussed here. Structures of spin traps applied in this study are shown in Figs. 4–6. These chemicals are commercially available from several companies. EPR spectra of spin adducts are usually measured at room temperature, using X-band spectrometers. In our experiments, a Bruker ECS-106 machine was used with the following settings: 1. EPR parameters: 9.45  GHz microwave frequency, 12  mW microwave power, 100 kHz modulation frequency, 0.2 mT modulation amplitude, 104–105 receiver gain, and 10.2  ms time constant. Optimal parameter values depend on the type of EPR spectrometer and cavity. 2. Thylakoid samples were measured in glass capillaries, at 15–20  ml volumes, although other sample holders may be more advantageous (see Note 1). Commercially available spin traps may need purification. For example, enolization of DMPO promotes the dimerization of this spin trap and after air oxidation, this adduct exhibits a six-line EPR spectrum. The concentration of dimers might be notable in old DMPO samples and even in stock DMPO solutions after several days. DMPO may also form DMPO-OH adducts in the presence of redox active metal ions (Fe2+/Fe3+, Cu+/Cu2+) and water. The chelation of these metal ions increases the electrophylic character of carbon of C=N bond, ultimately resulting in water addition (Fig. 7) (15). Ideally, spin traps alone, i.e., in the absence of ROS sources, should not show any EPR absorption. This should be checked by measuring their EPR spectra in the absence of thylakoids, in the same buffer and with the same instrument settings that will be used for the biological measurement. The following procedure removes most EPR active contaminants from DMPO: 1. Set up a semi-microscale distillation apparatus equipped with a magnet, a distilling flask, a three-way adapter, a thermometer, a condenser, a vacuum-adapter connected to vacuum

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a

R N

H

.

H N O

O

.

R

R= CH3

R= OH 2 mT *

* *

* *

*

DMPO sample addition withdrawal

b

2 mT 0

30

60 90 Photoinhibition (min)

120

*

**

DMPO sample addition withdrawal

** ** * * * *

**

*

2 mT

c 0

30

60 90 Photoinhibition (min)

120

Fig. 4. Spin trapping experiment using DMPO (50 mM) in isolated thylakoid membranes (100 mg/mL) exposed to 1,600 mmol/m2 s photosynthetically active radiation. (a) Structure of DMPO and the radical adduct formed upon reaction with a free radical represented by R•. Depending on the chemical nature of R•, radical adducts show EPR spectra characterized by different hyperline splitting. (b) and (c) Sampling strategy and typical EPR spectra. Single asterisks indicate the six lines belonging to a carbon-centered radical adduct, double asterisks identify the four lines of DMPO-OH.

line and nitrogen supply, and three receiving flasks with threelimbed multiple receiver. Be sure to dry the whole apparatus before use. A magnetic stirrer with heating, and liquid nitrogen or dry ice–acetone bath will also be needed. 2. Flush the distilling apparatus with N2 for 5 min, then place approximately 5  g of DMPO into the distilling flask and exchange  the N2 supply to vacuum line. Apply 0.4  mmHg ­vacuum and start to heat the distilling flask while stirring. Distil the DMPO at 73–75°C. Collect and discard about 0.5 mL as forerun.

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3. Change the receiver and collect 3–3.5 mL as main run, while chilling the receiving bulb with liquid nitrogen or a dry ice– acetone bath. 4. Exchange the vacuum line to the N2 supply and stop heating. 5. Use purified DMPO collected as the main run in spin trapping experiments. It is advisable to distribute the freshly distilled material into several small portions. Use a plastic-coated spatula to transfer frozen DMPO into cryovials. Store the DMPO under Ar and below −18°C. 6. For purifying smaller amounts of DMPO, a Kugelrohr distillation apparatus may also be used. 3.3. Applications of Spin Traps in Photosynthetic Membrane Preparations

3.3.1. Spin Trapping with DMPO

The following examples were taken from studies on ROS in the oxidative damage of isolated thylakoid membranes caused by excess photosynthetically active radiation (photoinhibition) (see Note 2). Hypothesis on the formation and further pathways of ROS in these samples are discussed elsewhere (18–23). It is important to note that results of ROS detecting experiments should always be interpreted as part of a complex study integrating as many aspects as possible, such as data on the stress-induced loss of physiologic function; oxidative damage of membrane proteins, lipids, or DNA; changes in antioxidant capacity; induction of repair processes; etc. 1. Keep isolated thylakoid membranes (see Note 3) at constant temperature in a temperature-controlled glass cuvette. 2. Realize photoinhibition by high-intensity photosynthetically active radiation (1,600  mmol/m2 s), for example, using a KL-1500 (DMP, Switzerland) lamp with optical fiber. 3. Use 50 mM DMPO as spin trap (24). Because the stability of DMPO adducts in this experimental system is too short for studying prolonged photoinhibition (22), do not keep the spin trap in the sample during the whole course of the experiment. 4. Measure nitroxide formation from DMPO in short, 5–10-min intervals: First, apply photoinhibition in the absence of the spin trap, then add DMPO and continue photoinhibition. Withdraw sample for EPR measurement 5 or 10  min after DMPO addition (Fig. 4b–c). Continuing this 5–10-min window of observation through the whole time course of the experiment allows comparing ROS production during the progression of photoinhibition. Typical examples of EPR spectra detected in the early and the late phase are shown in Fig. 4. The latter is a composite spectrum, composed of hydroxyl and carbon-centered radical adducts of DMPO, while the former signal is mainly due to the carboncentered radical adduct (18).

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3.3.2. Spin Trapping with Tiron

Although superoxide radicals also form characteristic adducts with DMPO, only experiments with this trap can give a straightforward answer for the involvement of superoxide radicals if these are the main radical species produced (25, 26). A relatively small concentration of superoxide (manifesting as a small amount of DMPO-OOH compared to other DMPO adducts) may not be noticed or may easily be detected as DMPO-OH (27, 28). A good alternative is to use Tiron, a spin trap for detecting superoxide radicals (29, 30) (see Note 4). Figure 5 illustrates trapping superoxide radicals with Tiron in illuminated thylakoid membranes. Isolated thylakoid membranes (100 mg Chl/mL) were exposed to 200 mmol/m2 s photosynthetically active radiation in the presence of 50 mM Tiron for 15 min. Without discussing mechanisms, we only note that only a small portion of superoxide is light stress related, and a similar spectrum was observed using higher light intensities (21). The addition of 10 mM of methyl-viologen markedly enhances superoxide production, as shown by the increase in the Tiron-OOH adduct (Fig. 5). a

O-

OH OH

HO3S

SO3H

. O2

. O SO3H

HO3S

0.5 mT

b

0.5 mT

c

0.5 mT

Fig. 5. (a) Structure of Tiron and its superoxide radical adduct, Tiron-OOH, EPR spectrum of Tiron-OOH. (b) and (c): Spin trapping experiments using Tiron (50 mM) in isolated thylakoid membranes (100 mg/mL). Samples were exposed to 200 mmol/m2 s photosynthetically active radiation for 15 min either in the absence (b) or in the presence (c) of 10 mM methyl-viologen.

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4. Notes 1. Because functioning plant samples are usually in the water-based preparation buffers, a critical step of EPR measurements is to lower the interference from the high microwave absorbance of water. This, on the one hand, calls for using small sample volumes, but, on the other hand, for using as much sample as possible to trap higher amounts of ROS. Optimal amounts may vary with the type and quality of the spectrometer, especially by its cavity. Signal: noise ratio may be improved by averaging several spectra if the stability of the spin adduct permits this. In the experiments illustrated here, nine spectra were averaged. Using an EPR flow cell allows the measurement of larger volumes with better signal: noise ratios than that obtained using capillaries or sample tubes, but the flow cell needs rigorous cleaning between changing samples in order to avoid artifacts. When using disposable sample tubes, comparative EPR quantification should be assured by keeping identical experimental conditions regarding both the samples (volume, concentration) and sample holders (uniform capillaries/tubes, the same central position in the cavity). 2. It is important to keep in mind that the added spin traps are in competition for ROS with potential sites of oxidative damage (the natural targets of ROS) and with any antioxidants present in the sample. In this way, failure to observe EPR signal from spin adducts does not necessarily mean that ROS were not produced in the sample. Similarly, when a free radical adduct is observed, the technique does not tell us whether the trapped ROS is a primary elicitor or a by-product of the studied stress reaction. Also, all spin trapping techniques underestimate real, in situ ROS concentration, because usually only a small fraction of the produced radicals is trapped. This can be somewhat improved by adding more spin trap, but it should be kept in mind that some spin traps inhibit photosynthetic reactions at higher concentrations. In our thylakoid membrane experiments, DMPO and Tiron were tolerated up to 80–100 mM concentrations, but other spin traps, such as pyridyl-nitrones, were found to be more toxic (22). Spin adducts (as well as spin traps themselves) may be reduced by plant metabolites into molecules without EPR activity. This can be checked by generating spin adducts chemically (i.e., without the biological sample) and comparing EPR signals in the absence and presence of untreated samples. Nitroxides, especially six-membered heterocyclic radicals, such as TempO, the singlet oxygen adduct of Temp

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a

1O

2

N

N

N

H

O

OH

.

Temp

diamagnetic

TempO

paramagnetic

TempOH

diamagnetic

TempO 2 mT

b 2 mT

c 2 mT

Fig. 6. (a) Structures of the sterically hindered amine Temp, its singlet oxygen adduct TempO and hydroxylamine TempOH, and the EPR spectrum of TempO. (b) and (c): Spin trapping experiments using Temp (10 mM) in isolated thylakoid membranes (100 mg/ mL). Samples were exposed to 1,600 mmol/m2 s photosynthetically active radiation for 15  min, then measured either directly (b) or after re-oxidizing ethylacetate-extracted TempOH into TempO (c).

(2,2,6,6-tetramethylpiperidine) (31), are very sensitive to reducing agents (32, 33), which convert the radical to a labile diamagnetic N-hydroxylamine (TempOH, Fig.  6a). The experiments illustrated in Fig. 6b and c were carried out similarly to the above ones with DMPO, but using 10 mM Temp. When Temp containing samples of photoinhibited thylakoids were measured directly, there was hardly any TempO signal (Fig. 6b). However, when the sample was extracted into an organic solvent, it was possible to restore the nitroxide radical (Fig.  6c). In order to achieve this, photoinhibited samples were mixed with ethylacetate (1:2 V/V) and allowed to separate into two phases for a few minutes. The upper, organic phase (containing the N-hydoxylamine but no photosynthetically active thylakoids) was removed and re-oxidized with a catalytic amount (10–30  mg) of PbO2 before EPR spectroscopy.

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Fig. 7. Possible decomposition pathways of DMPO.

3. Thylakoid preparations are good models for studying a number of photosynthetic reactions, but are less complex than leaves. Also, many in vivo stress response reactions occur on different time scales than the possibilities of thylakoid experiments. In this way, a direct extension of spin trapping experiments is not possible, because neither spin traps nor spin adducts endure long enough to report free radical production in leaves during environmental stress experiments lasting up to several days. Although clearly not a substitute for in  vivo monitoring, studies on leaf extracts may prove useful. In these studies, whole plants or detached leaves were exposed to stress conditions as long as necessary and then frozen in liquid N2. Frozen leaf powders were ground in a mortar with a phosphate buffer containing the ROS trap, then filtered and measured similarly to thylakoid samples. In these experiments, quick sample processing is essential, so only a small amount of leaves is used. The time between leaf harvesting and EPR measurement should not only be as short as possible, but also uniform for all samples. When evaluating results of these “post-in vivo” experiments, it is important to keep in mind that although samples were collected immediately after the cessation of stress, due to the time delay caused by sample processing, the trapped radicals are not primary products. In this way, the technique is not suitable for qualitative analysis of the ROS produced during the stress, but rather as a characterization of the antioxidant–pro-oxidant status of the stressed leaf (34, 35).

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4. Direct detection of reactive species with spin trapping is still a dynamically expanding field of biology. Although medical applications are dominant, the technique is gaining more and more recognition in plant studies. Extensive research is being carried out in several laboratories to develop improved spin traps with higher adduct stability and specificity. A few examples of spin traps that may prove more versatile than the traps listed in this chapter are pyridyl- (36), adamantyl-pyridyl(37), glycosylated (38), or phosphorylated (39) nitrones. An alternative to EPR spectroscopy is detecting ROS with double sensors, which utilize the principle of spin trapping, but their conversion to nitroxide radical can also be monitored as fluorescence quenching (40–43). An inverted mechanism has been utilized for glutathionyl radical detection with 4-((9-acridinecarbonyl)amino)-2,2,6,6-tetramethylpiperidine causing fluorescence increase in MPO-rich HL-60 cells, incubated in the presence of H2O2/phenol (44). Fluorescent sensors show great potential in in  vivo studies, for example, of leaves (45–47) or photosynthetic microorganisms (48), and the technique may also be extended to imaging (48, 49).

References 1. Halliwell, B. and Gutteridge, J.M. (2007) Free radicals in biology and medicine 4th ed. Oxford University Press Inc. NY. 2. Cadenas, E. (1989) Biochemistry of oxygen toxicity. Annu Rev Biochem 58, 79–110. 3. Elstner, E.F. (1987) Metabolism of activated oxygen species, in The biochemistry of plants (Davies, D.D., ed.) Vol 11, Academic Press, San Diego, pp. 253–315. 4. Asada, K. (1999) The water-water cycle in chloroplasts: Scavenging of active oxygens and dissipation of excess photons. Annu Rev Plant Physiol Plant Mol Biol 50, 601–639. 5. Asada, K. (2006) Production and scavenging of reactive oxygen species in chloroplasts and their functions. Plant Physiol 141, 391–396. 6. Schopfer, P. (2001) Hydroxyl radical-induced cell-wall loosening in vitro and in vivo: implications for the control of elongation growth. Plant J 28, 679–688. 7. Schopfer, P., Liszkay, A., Bechtold, M., Frahry, G., and Wagner, A. (2002) Evidence that hydroxyl radicals mediate auxin-induced extension growth. Planta 214, 821–828.

8. Smirnoff, N. (1995) Antioxidant systems and plant response to the environment, in Environment and plant metabolism (Smirnoff, N., ed.), Bios Scientific Publishers Ltd. Oxford, pp. 218–244. 9. Apel, K. and Hirt, H. (2004) Reactive oxygen species: Metabolism, oxidative stress, and signal transduction. Annu Rev Plant Biol 55, 373–399. 10. Foyer Ch. H., Lelandais M., and Kunert K.J. (1994) Photooxidative stress in plants. Physiol Plant 92, 696–717. 11. Veljovic´-Jovanovic´, S. (1998) Active oxygen species and photosynthesis: Mehler and ascorbate peroxidase reactions. Iugoslav Physiol Pharmacol Acta 34, 503–522. 12. Mittler, R., Vanderauwera S., Gollery, M., and Van Breusegem, F. (2004) Reactive oxygen gene network of plants. Trends in Plant Sci 9, 490–498. 13. Krieger-Liszkay, A. (2005) Singlet oxygen production in photosynthesis. J Exp Bot 56, 337–346. 14. Evans, C. A. (1979): Spin trapping. Aldrichim Acta 12, 23–29 15. Rosen, G.M., Britigain, B.E., Halpern, H. J., and Pou, S. (1999) Free radicals, Oxford University Press: Oxford.

Direct Detection of Free Radicals and Reactive Oxygen Species in Thylakoids 16. Janzen, E.G. (1980) Critical review of spin trapping in biological systems, in Free radicals in biology (Pryor, W.A., ed.), Vol IV., Academic Press, New York, pp. 115–154. 17. Floyd, R.A. (2009) Serendipitous findings while researching oxygen free radicals. Free Rad Biol Med 46, 1004–1013. 18. Hideg, É., Spetea, C., and Vass, I. (1994) Singlet oxygen and free radical production during acceptor and donor side induced photo­ inhibition. Studies with spin trapping EPR spectroscopy. Biochim Biophys Acta 1186, 143–152. 19. Hideg, É., Spetea, C., and Vass, I. (1994) Singlet oxygen production in thylakoid membranes during photoinhibition as detected by EPR spectroscopy. Photosynth Res 39, 191–199. 20. Hideg, É. and Vass, I. (1995) Singlet oxygen is not produced in photosystem I under photoinhibitory conditions. Photochem Photobiol 62, 949–952. 21. Hideg, É., Spetea, C., and Vass, I. (1995) Superoxide radicals are not the main promoters of acceptor side induced photoinhibitory damage in spinach thylakoids. Photosynth Res 46, 399–407. 22. Hideg, É., Takátsy, A., Sár, P.C., Vass, I., and Hideg, K. (1999) Utilizing new adamantyl spin traps in studying UV-B-induced oxidative damage of photosystem II. J Photochem Photobiol B: Biol 48, 174–179. 23. Hideg, É. (1997) Free radical production in photosynthesis under stress conditions, in Handbook of Photosynthesis (Pessarakli, M., ed.), Marcel Dekker, Newe York, pp. 911–930. 24. Janzen, E.G. and I-Ping Liu, J. (1973) Radical addition reactions of 5,5-dimethyl-1-pyrroline-1-oxide. ESR spin trapping with a cyclic nitrone. J Magn Reson 9, 510-512. 25. Hiramatsu M. and Kohno M. (1987) Determination of superoxide dismutase activity by electron spin resonance spectroscopy using the spin trap method. JEOL News 23A, 6–9. 26. Buettner, G.R. and Mason, R.P. (1990) Spintrapping methods for detecting superoxide and hydroxyl free radicals in vitro and in vivo. Meth Enzymol 186, 127–133. 27. Finkelstein, E., Rosen, G.M., and Rauckman, E.J. (1979) Spin trapping of superoxide. Mol Pharmacol 16 676–685. 28. Finkelstein, E., Rosen, G.M., and Rauckman, E.J. (1980) Spin trapping. Kinetics of the reaction of superoxide and hydroxyl radicals

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with nitrones. J Amer Chem Soc 102, 4994–4999. Greenstock, C.L. and Miller, R.W. (1975) The oxidation of Tiron by superoxide anion. Kinetics of the reaction in aqueous solutions and chloroplasts. Biochim Biophys Acta 396, 11–16 Miller, R.W. and MacDowall, F.D.H. (1975) The Tiron free radical as a sensitive indicator of chloroplastic photoautooxidation. Biochim Biophys Acta 387, 176–187 Lion, Y., Delmelle, M., and van de Vorst, A. (1976) New method of detecting singlet oxygen production. Nature 263, 442–443. Couet, W.R., Brash, R.C., and Tozer, T.N. (1985) Factors affecting nitroxide reduction in ascorbate solution and tissue homogenates. Magn Res Imaging 3, 83–87. Blinco, J.P., Hodgson, L.J., Morrow, J.B., Walker, J.R., Will, D.G., Coote, L.M., and Bottle, S.E. (2008) Experimental and theoretical studies of the redox potentials of cyclic nitroxides. J Org Chem 73, 6763–6771. Hideg, É. and Björn, L.O. (1996) Ultraweak light emission, free radicals and chilling sensitivity. Physiol Plant 98, 223–228. Hideg, É. and Vass, I. (1996) UV-B induced free radical production in plant leaves and isolated thylakoid membranes. Plant Sci 115, 251–260. Janzen, E.G., Dudley, R.L., and Shetty, R.V. (1979) Synthesis and electron spin resonance chemistry of nitronyl labels for spin trapping. a-phenyl N-[5-(5-methyl-2, 2-dialkyl-1,3-dioxanyl)] nitrones and a-(Nalkyl-pyridinium) N-(tert-butyl) nitrones. J Amer Chem Soc 101, 243–245. Sár, P.C., Hideg, É., Vass, I., and Hideg, K. (1998) Synthesis of a-aryl N-adamant-1-yl nitrones and using them for spin trapping of hydroxyl radicals. Bioorg Med Chem Lett 8, 379–384. Ouari, O., Chalier, F., Bonaly, R., Pucci, B., and Tordo, P. (1998) Synthesis and spin-trapping behaviour of glycosylated nitrones. J Chem Soc Perkin Trans 2, 2299–2307. Frejaville, C., Karoui, H., Tuccio, B., Le Moigne, F., Culcasi, M., Pietri, S., Lauricella, R., and Tordo, P. (1995) 5-(Diethoxyphosphoryl)-5methyl-1-pyrroline N-oxide: A new efficient phosphorylated nitrone for the in  vitro and in  vivo spin trapping of oxygen-centered radicals. J Med Chem 38, 258–265. Kálai, T., Hideg, É., Vass, I., and Hideg, K. (1998) Double (fluorescent and spin) sensors for detection of reactive oxygen species in the

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Hideg, Kálai, and Hideg thylakoid membrane. Free Rad Biol Med 24, 649–652. Hideg, É., Vass, I., Kálai, T., and Hideg, K. (2000) Singlet oxygen detection with sterically hindered amine derivatives in plants under light stress. Meth Enzymol 319, 77–85. Bilski, P., Hideg, K., Kálai, T., Bilska, M.A., and Chignell, C.F. (2003) Interaction of singlet molecular oxygen with double fluorescent and spin sensors. Free Rad Biol Med 34, 489–495. Likhtenshtein, G.I. (2008) Dual fluorophorenitroxide compounds, in Nitroxides (Likhtenshtein, G.I., Yamauchi, J., Nakatsuji, S., Smirnov, A.I. and Tamura, R. eds.) WileyVCH: Weinheim, pp. 256–264. Borisenko, G.G., Martin, I., Yhao, Q., Amoscato, A.A., Tzurina, Z.Z., and Kagan, V.E. (2004) Glutathione propagates oxidative stress triggered by myeloperoxidase in HL-60 cells. J Biol Chem 379, 23453–23462. Hideg, É., Kálai, T., Hideg, K., and Vass, I. (1998) Photoinhibition of photosynthesis in  vivo results in singlet oxygen production.

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Detection via nitroxide-induced fluorescence quenching in broad bean leaves. Biochemistry, 37, 11405–11411. Hideg, É., Barta, Cs., Kálai, T., Vass, I., Hideg, K., and Asada, K. (2002) Detection of singlet oxygen and superoxide with fluorescent sensors in leaves under stress by photoinhibition or UV radiation. Plant Cell Physiol 43, 1154–1164. Barta, Cs., Kálai, T., Hideg, K., Vass, I., and Hideg, É. (2004) Differences in the ROS generating efficacy of various ultraviolet wavelengths in detached spinach leaves. Funct Plant Biol 31, 23–28. Hideg, É., Kálai, T., Kós, P.B., Asada, K., and Hideg, K. (2006) Singlet oxygen in plants – its significance and possible detection with double (fluorescent and spin) indicator reagents. Photochem Photobiol 82, 1211–1218. Hideg, É. and Schreiber, U. (2007) Parallel assessment of ROS formation and photosynthesis in leaves by fluorescence imaging. Photosynth Res 92, 103–108.

Chapter 17 Assay of Photoinhibition and Heat Inhibition of Photosystem II in Higher Plants Nobuyoshi Nijo, Björn Lundin, Miho Yoshioka, Noriko Morita, and Yasusi Yamamoto Abstract When thylakoids of higher plant chloroplasts are exposed to excessive light or moderate heat stress, photosystem II reaction center-binding protein D1 is damaged. The photodamage of the D1 protein is caused by reactive oxygen species, mostly singlet oxygen, and also by endogenous cationic radicals generated by the photochemical reactions of photosystem II. Moreover, it was shown recently that the damage to the D1 protein by moderate heat stress is due to reactive oxygen species produced by lipid peroxidation near photosystem II. To maintain photosystem II activity, the oxidatively damaged D1 protein must be replaced by a newly synthesized copy, and thus degradation and removal of the photo- or heatdamaged D1 protein are essential for maintaining the viability of photosystem II. In this chapter, we describe the methods for assaying photoinhibition and heat inhibition of photosystem II in higher plant materials. Key words: Photosystem II, Light and heat stresses, D1 protein, Protein degradation and aggregation, Reactive oxygen species, Higher plants

1. Introduction Illumination of photosystem II (PSII) with excessive visible light induces reduction of the plastoquinone electron acceptors of PSII, QA and QB, and subsequent release of the fully reduced plastoquinones from their binding sites. These processes stimulate charge recombination between the reduced primary electron acceptor pheophytin and the oxidized primary electron donor P680+, thereby generating the triplet state of P680, which eventually reacts with oxygen to form singlet oxygen (1O2). The photodamage to the D1 protein under excessive illumination of

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healthy leaves, thylakoids, and PSII samples is caused mostly by 1 O2 produced in the over-excited PSII reaction center (1–3). This phenomenon is called acceptor-side photoinhibition of PSII (4, 5). The most prominent changes occurring to the D1 protein in the acceptor-side photoinhibition of PSII are (1) cleavage of the damaged D1 protein to N-terminal 23-kDa and C-terminal 10-kDa fragments and (2) formation of aggregates of the damaged D1 protein with the surrounding polypeptides (6, 7). Pioneering work by Greenberg et al. (8) demonstrated that the D1 protein is degraded in Spirodela oligorrhiza by strong illumination, where a 23.5-kDa breakdown fragment is produced. These results suggest that the primary cleavage of the D1 protein takes place at the stroma-exposed DE-loop of the D1 protein. In spite of subsequent efforts, however, the mechanisms underlying D1 cleavage have not been clarified in detail. Recently, several proteases, including a metalloprotease FtsH, were suggested to be involved in the proteolytic processes (9). Regarding the aggregation of the photodamaged D1 protein, three aggregation products of the D1 protein, namely D1/D2, D1/the a-subunit of cytochrome b559, and D1/CP43, have been identified (6, 7, 10). However, the precise cross-linking sites of these proteins remain to be determined. From the finding that the herbicide 3-(3,4-dichlorophenyl)1,1-dimethylurea (DCMU) inhibited the aggregation of the D1 protein, the structural change in the QB binding site of the D1 protein was suggested to be involved in this process (11). Since the steps related to the acceptor-side photoinhibition of PSII require oxygen, experiments should be carried out under aerobic conditions. It has been shown that the oxygen-evolving system of PSII is impaired when PSII is illuminated under high or low temperature conditions or under conditions where either Ca2+ or Cl− is depleted. In these circumstances, endogenous cationic radicals (ECR) are generated from oxidation of the primary electron donor P680, the secondary electron donor Tyr Z, and the antenna pigments. These radicals impair electron transport in PSII, inducing significant damage to the D1 protein in the lumen-exposed portion. This is called the donor-side photoinhibition of PSII (4–7). As the primary cause of the photodamage by the donorside photoinhibition is the generation of ECR, the reactions related to this process do not require oxygen (6, 7). It was suggested that cleavage of the D1 protein takes place at the lumenexposed AB-loop of the D1 protein, thereby producing 10-kDa N-terminal and 23-kDa C-terminal fragments (4, 5, 12). However, the cleavage of the D1 protein in the donor-side photoinhibition is usually not prominent, and more significant is the aggregation between the D1 protein and the antenna chlorophyll-binding protein CP43 (13).

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When the thylakoids were exposed to moderate heat stress such as 40°C for 30 min, the D1 protein was cleaved and cleavage products similar to those observed in the acceptor-side photoinhibition were detected (14). These results suggest that the site of damage in the D1 protein is the DE-loop of the D1 protein. Under moderate heat stress, lipid peroxidation near PSII was shown to be responsible for the production of reactive oxygen species and the subsequent damage to the D1 protein (15). The photodamaged D1 protein is degraded by the action of proteases in the chloroplasts. So far, several proteases involved in the degradation of the damaged D1 protein in the acceptor-side photoinhibition of PSII have been reported, while the protease(s) that recognizes the damaged D1 protein from the donor-side photoinhibition or the cross-linked products of the D1 protein remains to be identified (7, 9). Moreover, the proteases that degrade the primary cleavage products of the D1 protein are not known. Recently, aggregation of the photodamaged D1 protein and the surrounding polypeptides in PSII was shown to be an alternative pathway that the damaged D1 protein takes during light or heat stress (6, 7). In this chapter, we describe growth of plants (Arabidopsis thaliana), isolation of thylakoids, induction of photoinhibition and heat inhibition of PSII in vitro, measurement of PSII activity, detection of the cleavage and aggregation products of the photodamaged D1 protein by Western blot analysis, and assay of protease activity.

2. Materials 2.1. Growth of Arabidopsis thaliana

1. Miracloth.

2.1.1. Sterilization of Arabidopsis Seeds

3. Ethanol 70% (v/v).

2. Arabidopsis seeds. 4. SDS 0.5% (w/v). 5. Paper clip.

2.1.2. Germination of Arabidopsis Seeds

1. Sterilized Arabidopsis seeds. 2. Micropipette tip box containing 200-mL tips, sterilized. 3. Stock solution: (40 mL) 1 M KNO3, (12 mL) 1 M Ca(NO3)2, (8 mL) 1 M KH2PO4, (8 mL) 1 M MgSO4, and (8 mL) 1 M NH4Cl (total volume, 76 mL). 4. Micro solution: 46 mM H3BO3, 10 mM MnSO4, 0.77 mM ZnSO4, 0.32  mM CuSO4, 0.58  mM MoO3, and 0.25  mM NH4VO3 (total volume, 8 mL).

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5. Fe-EDTA solution: First, 1,000  mL solution containing 100  mM Na-EDTA, 330  mM KOH, and 90  mM FeSO4. Dissolve Na-EDTA and KOH together, and FeSO4 separately, and then mix these solutions. To oxidize Fe, air is bubbled in the Fe-EDTA solution for approximately 30 min. 6. Nutrient solution: 76 mL of stock solution, 8 mL of micro solution, 8 mL Fe-EDTA solution, and 8 mL H2O (total volume, 100 mL). 7. Agar solution: 125 mL H2O, 0.84 g agar, and 750 mL nutrient solution. 8. Spatulas. 9. Petri dish. 10. MilliQ water. 11. Glass pipettes. 12. Glass beakers. 2.1.3. Culture of Arabidopsis Seedlings

1. Arabidopsis seedlings with 4–6 leaves. 2. Containers to hold 0.5% nutrient solution. Any type of container can be used for the hydroponic culture system, but a shallow container 10–15 cm in height and capable of holding 20–40 L of water is recommended. 3. Air pump and plastic tube connected to the pump. 4. Plastic sheets. 5. Transparent plastic cups. 6. Sprayer. 7. Plant growth chamber: normal growth conditions are a photoperiod of 8  h light and 16  h darkness, light intensity 100 mmol photons/m2 s, and temperature 20°C.

2.2. Preparation of Thylakoids from Arabidopsis Leaves (Small-Scale Preparation)

1. Arabidopsis leaves, dark-adapted to avoid accumulation of starch. 2. Preparation medium: 50 mM Hepes-KOH, pH 7.8, 330 mM sorbitol, 10 mM KCl, 1 mM EDTA, 0.15% (w/v) BSA, 4 mM sodium ascorbate, and 7 mM l-cysteine. BSA, sodium ascorbate, and l-cysteine are added just before usage. 3. Washing medium: 50  mM Hepes-KOH, pH 7.8, 330  mM sorbitol, and 10 mM KCl. 4. Lysis medium: 5 mM MgCl2. 5. 2× resuspension buffer: 50  mM Hepes-KOH, pH 7.4, 660 mM sorbitol, 10 mM KCl, and 10 mM MgCl2. 6. 1× resuspension buffer: 25  mM Hepes-KOH, pH 7.4, 330 mM sorbitol, 5 mM KCl, and 5 mM MgCl2.

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7. Mortar or a mixer. 8. Centrifuge. 9. Brush or a glass homogenizer. 10. Acetone 80% (v/v). 11. Spectrophotometer. 2.3. Preparation of Thylakoids from Spinach or Pea Leaves (Large-Scale Preparation)

1. Spinach or pea leaves. 2. 5× Grinding medium (GM) stock solution: 0.25 M HepesKOH, pH 7.5, 1.65 M sorbitol, 5 mM MgCl2, 5 mM MnCl2, and 10 mM EDTA. 3. 1× GM: 50  mM Hepes-KOH, pH 7.5, 0.33  M sorbitol, 1  mM MgCl2, 1  mM MnCl2, 2  mM EDTA, 0.25% (w/v) BSA, and 5 mM Na-ascorbate. 4. ST buffer: 50  mM Tricine-KOH, pH 8.0, and 0.33  M sorbitol. 5. PBF-Percoll: 0.56 g polyethylene glycol # 4000, 0.185  g BSA, 0.185 g Ficoll, and 18.5 mL Percoll. 6. 0.5  M Na-ascorbate: 50  mM Hepes-KOH, pH 7.5, and 0.5 M Na-ascorbate. 7. Percoll gradient (20%): 1.5 mL 5× GM, 1.5 mL PBF-Percoll, 1.5  mg glutathione (reduced form), 75  mL of 0.5  M Na-ascorbate, and 4.5  mL distilled water (total volume, 7.5 mL). 8. Percoll gradient (50%): 3.6 mL 5× GM, 9.0 mL PBF-Percoll, 3.6 mg glutathione, 180 mL of 0.5 M Na-ascorbate, and distilled water (total volume, 18 mL). 9. Percoll gradient (80%): 2.0 mL 5× GM, 8.0 mL PBF-Percoll, 2.0 mg glutathione, and 100 mL of 0.5 M Na-ascorbate (total volume, 10 mL). 10. Blender. 11. Centrifuge and a swinging bucket rotor.

2.4. Preparation of Photosystem II-Enriched Membranes 2.5. Measurement of Photosystem II Activity

Spinach or pea leaves. The details of the materials and methods for preparation of these samples are presented elsewhere in the same volume (see Chapter 1 and Note 1). 1. Oxygen electrode (Hansatech, UK, or Rank Brothers, UK). 2. PAM (pulse-amplitude modulation) chlorophyll fluorometer (Walz, Germany). 3. Actinic light source. 4. Circulating water bath.

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2.6. SDS/UreaPolyacrylamide Gel Electrophoresis (SDS/ Urea-PAGE)

1. Acrylamide stock solution: 50% (w/v) acrylamide and 13.3% (w/v) bis-acrylamide (total volume, 500 mL). 2. Resolving gel solution: 0.6 M Tris–HCl, pH 8.8, 12.5% acrylamide (5 mL acrylamide stock solution), 6 M urea, and 0.1% (w/v) SDS (total volume, 20 mL). 3. Stacking gel solution: 0.125 M Tris–HCl, pH 6.8, 4.5% acrylamide (0.9 mL acrylamide stock solution), 6 M urea and 0.1% (w/v) SDS (total volume, 10 mL). 4. Electrophoresis buffer: 25  mM Tris, 0.192  M glycine, and 0.1% (w/v) SDS (pH adjustment not necessary) (total volume, 1 L). 5. Lysis buffer: 0.125 M Tris–HCl, pH 6.8, 6 M urea, 5% (w/v) SDS, 5% (w/v) b-mercaptoethanol, 5  mM EDTA, and 5% (w/v) sucrose (total volume, 30 mL). 6. Ammonium persulfate solution: 10% (w/v) ammonium persulfate. 7. TEMED (N,N,N¢,N¢-tetramethylene-ethylenediamine) solution. 8. Staining solution: 0.05% Coomassie brilliant blue R-250 (CBB), 50% methanol, and 7% acetic acid. 9. Destaining solution: 20% methanol and 7% acetic acid. 10. Molecular weight marker proteins: low molecular weight electrophoresis calibration kit (Pharmacia, Sweden). 11. A vertical electrophoresis apparatus and a power supply. 12. A shaker.

2.7. Western Blot Analysis

1. Transfer buffer: 25 mM Tris–HCl, pH 7.4, 0.192 M glycine, and 10% (v/v) methanol. 2. TBS: 20 mM Tris–HCl, pH 7.4, and 0.15 M NaCl. 3. TTBS: 20 mM Tris–HCl, pH 7.4, 0.15 M NaCl, and 0.05% (v/v) Tween 20. 4. Blocking buffer: 20 mM Tris–HCl, pH 7.4, 5% (v/v) skim milk, 0.02% (v/v) NaN3, 0.15  M NaCl, and 0.05% (v/v) Tween 20. 5. The first antibody solution: the first antibody (raised in a rabbit) diluted according to its titer, 20 mM Tris–HCl, pH 7.4, 1% (w/v) BSA, 0.01% (w/v) NaN3, 0.15 M NaCl, and 0.05% (v/v) Tween 20. 6. The second antibody solution: 20 mM Tris–HCl, pH 7.4, 1% (w/v) skim milk, 0.01% (w/v) the second antibody (horse radish peroxidase-coupled anti-rabbit goat antibody purchased from BioRad), 0.15 M NaCl, and 0.05% (v/v) Tween 20. 7. Probe-removing solution: 62.5  mM Tris–HCl, pH 6.7, 2% (w/v) SDS, and 100 mM b-mercaptoethanol.

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8. PVDF [poly(vinylidene difluoride)] membranes (Millipore, USA). 9. Electroblotting apparatus. 10. Power supply. 11. Shaker. 12. X-ray films and a film cassette. 13. Film developer. 14. An enhanced chemiluminescence (ECL) kit. 2.8. Protein Oxidation Analysis

1. OxyBlot protein oxidation detection kit (Chemicon, USA) containing the reagents A–E (see below). (a) 10× 2,4-dinitrophenylhydrazine (DNPH) solution. (b) 10× derivatization-control solution. (c) Neutralization solution. (d) Primary antibody: rabbit anti-DNP antibody. (e) Secondary antibody: goat anti-rabbit IgG (HRPconjugated). 2. 20% SDS. 3. 2× lysis buffer: 100  mM Tris, 2% (w/v) SDS, 20% (w/v) sucrose, 0.06% bromophenol blue (BPB), and 100 mM DTT. Add distilled water to 100 mL. 4. 10× phosphate-buffered saline (PBS), 26 g NaH2PO4 H2O, 115  g Na2HPO4, and 85  g NaCl. Add distilled water to 1,000 mL. 5. 1× PBS-T: 100 mL 10× PBS and 0.5 mL Tween20. Add distilled water to 1,000 mL. 6. NuPAGE antioxidant (Invitrogen, USA). 7. PVDF membranes (Millipore, USA). 8. Electrophoresis apparatus. 9. Electroblotting apparatus. 10. Blocking/dilution buffer: 10  g BSA. Add 1× PBS-T to 1,000 mL.

2.9. Activity Gel (Gelatin/SDS– Polyacrylamide Gel Electrophoresis)

1. Gelatin/SDS–polyacrylamide gel: 0.2% (w/v) gelatin in the resolving gel of the SDS–polyacrylamide gel (see Note 2). 2. Thylakoids or other protease-containing fractions. 3. Renaturing buffer: 50 mM Tris–HCl, pH 8.0, 5 mM MgCl2, and 1% (w/v) Triton-X100. 4. Incubation buffer: 50  mM Tris–HCl, pH 8.0, and 5  mM MgCl2.

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5. Staining solution (see Subheading 2.6.). 6. Destaining solution (see Subheading 2.6.). 7. Lysis buffer (see Subheading 2.6.). 8. Electrophoresis apparatus. 9. Knife or razor blade. 10. Glass homogenizer (Teflon pestle and a glass container).

3. Methods 3.1. Growth of Arabidopsis thaliana Seedlings 3.1.1. Sterilization of Arabidopsis Seeds

1. Place 100–200 seeds in a Miracloth (5 × 5 cm), make a package, and seal with a paper clip. 2. Soak the seeds in 70% ethanol for 15 min and then in 0.5% SDS (w/v) for 7 min. 3. Wash the seeds 4–5 times in distilled water until there is no foam. 4. Seal 200-mL micropipette tips by melting the tip briefly with a gas burner, and then dip them in ice-cold water to cool rapidly. 5. Fill a whole micropipette tip box with sealed tips and sterilize it on the week of use (it is important to have 1–2 cm space between the tips and the box cover). After autoclaving, wrap the box with layers of parafilm to keep it sterile.

3.1.2. Germination of Arabidopsis Seeds

1. Sterilize the Arabidopsis seeds (if already sterilized, soak in 70% ethanol and wash in distilled water). 2. Warm the sterilized agar solution until it becomes liquid and fill each sealed tip with the agar to make a small dome. 3. Unwrap the Miracloth containing the sterilized seeds on a sterilized Petri dish. 4. Pick up the seeds using a spatula and place one seed at a time on the dome of a tip. If it is difficult to take up seeds with the spatula, dip the spatula briefly in milliQ water before picking up the seeds. 5. Wrap the box with parafilm and place it at 4°C for ~48 h and then at 18–22°C under white-light illumination (100– 120 mmol photons/m2 s).

3.1.3. Culture of Arabidopsis Seedlings

1. Two weeks after germination of the seeds, remove the parafilm from the box, open the lid for a few seconds to ventilate the seeds, and then return the box to the growth chamber (see Note 3). 2. When 4–6 leaves have developed, transfer the seedlings to the hydroponic culture system. Fill the container with 0.5% (v/v) nutrient solution.

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3. Cut the tips containing the seedling between half and twothirds of the length from the top and place them in a plastic sheet with perforations in line for insertion of the tips. 4. The seedlings need to be covered with a transparent plastic cup or some other transparent cover for about 2 weeks. Spray water daily over the seedlings to make the transfer to the hydroponic system less stressful. 5. Approximately 6–8  weeks after transfer to the hydroponic system, the leaves are ready to be harvested (16). 3.2. Preparation of Thylakoids from Arabidopsis Leaves (Small-Scale Preparation)

1. Place the leaves in cold distilled water for 30 min. 2. Grind the leaves with a mortar or a mixer in the preparation medium and filter the mixture through four layers of cheese cloth (pore size 20 mm). 3. Centrifuge the filtrate at 1,000g for 3 min. 4. Discard the supernatant and, with a brush, gently resuspend the pellet in the washing medium to a volume 0.4 times that of the initial preparation medium. 5. Centrifuge the suspension at 1,000g for 3 min. 6. Discard the supernatant and resuspend the pellets in the washing medium using either a brush or a glass homogenizer. 7. Determine the chlorophyll concentration in an 80% acetone extract of the suspension by measuring absorbance at 663 and 645 nm with a spectrophotometer. 8. Resuspend the thylakoids in the lysis medium to give a chlorophyll concentration of 0.2  mg/mL, and homogenize the samples using either a brush or a glass homogenizer. Keep the sample on ice for 5 min and add an equal volume of 2× resuspension buffer. 9. Centrifuge at 5,000g for 5 min. 10. Discard the supernatant and using a brush gently resuspend the pellet in 1× resuspension medium. 11. Centrifuge at 5,000g for 5 min. 12. Discard the supernatant and using a brush gently resuspend the pellets in 1× resuspension medium to give a chlorophyll concentration between 0.5 and 2 mg/mL. 13. Measure the chlorophyll concentration.

3.3. Preparation of Thylakoids from Spinach or Pea Leaves (Large-Scale Preparation)

1. Wash the leaves with tap water and cool them on ice. 2. Cut the leaves into squares (2 × 2 cm) with a knife. 3. Homogenize the leaves with a pre-cooled blender. Add 500 mL of GM per 100 g of leaves. 4. Filter the homogenate through four layers of cheesecloth.

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5. Centrifuge the filtrate at 3,000g for 90 min. 6. Suspend the precipitates in a small volume of GM. 7. Load the suspension on a Percoll gradient (20–50–80%) and centrifuge in a swinging bucket rotor at 5,500g for 10 min. 8. The intact chloroplasts form a green band at the interface between the 50 and 80% Percoll. Collect the band with a pipette. 9. Dilute the intact chloroplast fraction with five volumes of ST. 10. Centrifuge at 1,000g for 90 s to obtain the intact chloroplasts as a precipitate. 3.4. Photoinhibitory Illumination

1. Photoinhibition of PSII by visible light can be induced by a conventional light source, such as a slide projector with a 1,000-W tungsten lamp. Heat cut-off filters should be placed between the lamp and the samples. 2. For incubation at a specific temperature, the samples are placed in transparent plastic sample tubes (1  mL volume), which are then placed in a reaction vessel connected to a temperature-controlled circulating water bath. We constructed the reaction vessel from transparent acrylic plates. 3. The sample tubes are illuminated by light from the side. The light intensity is measured with a quantum sensor QSPAR (Hansatech, UK).

3.5. Heat Treatment

1. Thylakoid samples in black sample tubes are placed in a temperature-controlled water bath or a heating block. 2. After heat treatment, the samples are cooled on ice to stop the heat stress.

3.6. Measurement of Photosystem II Activity

1. Measure oxygen evolution using an oxygen electrode. Consult the manufacturer’s manual for details. 2. Measure optimum quantum yield Fv/Fm of chlorophyll fluorescence using a PAM. Consult the manufacturer’s manual for details.

3.7. SDS/Urea– Polyacrylamide Gel Electrophoresis

The method is based on that of Laemmli (17). A 12.5% acrylamide gel is used here but a gradient gel (e.g., from 10 to 20%) can also be used. To improve resolution, 6 M urea is included both in the stacking and resolving gels, and 0.6 M Tris (18) is used in the resolving gel. Samples are solubilized in the lysis buffer containing 6 M urea. For the electrophoresis conditions (see Note 4), the gel is stained with CBB. Alternatively, the proteins are electroblotted onto a PVDF membrane and stained with CBB.

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3.8. Assay of the Degradation Fragments and Aggregates of the D1 Protein Generated by the Acceptor-Side Photoinhibition of PSII with Western Blot Analysis and Fluorography (Fig. 1)

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1. Separate the proteins in the samples using SDS/urea–PAGE. 2. The proteins in an SDS/urea–PAGE gel are transferred to a PVDF membrane. The power supply is set to 200  mA (constant current) and run for 1 h at 4°C. 3. We use a wet-type blotting apparatus since proteins are evenly transferred to the PVDF membranes. 4. Wash the PVDF membrane in TBS for 5 min on a shaker. A small plastic container can be used for the incubation. 5. Incubate the membrane in blocking solution for 1  h with shaking. 6. Wash the membrane twice with TTBS. 7. Incubate the membrane in a solution containing the first antibody for 1 h with shaking. 8. Briefly wash the membrane twice with TTBS, then wash once with TTBS for 15 min and twice for 5 min. Incubate the membrane in the solution containing the second antibody for 1 h. 9. Rinse the membrane with TTBS. 10. Transfer the membrane to another plastic container and add the ECL detection solution. The ECL detection solution is prepared by mixing equal volumes of two reagents (denoted solutions 1 and 2). Use 4 mL of the solution for one 10 × 10 cm membrane. Incubate for 1 min. 11. Place the PVDF membrane in a plastic sack and seal it with a sealer.

Fig. 1. The effects of heat stress on the D1 protein of spinach thylakoids. (a) Proteins blotted on a PVDF membrane after SDS/urea–PAGE and stained with Coomassie brilliant blue. (b) A fluorogram of the Western blot analysis with the antibody against the DE-loop of the D1 protein. Spinach thylakoids were incubated at 40°C for 30 min at different pHs specified at the top of the lanes, where either Mes-KOH (indicated by *1) or Tricine-KOH (indicated by *2) was used as a buffer. The symbols (+) and (−) indicate heated and non-heated, respectively. The arrows on the right-hand side of the fluorogram indicate the bands representing the D1 protein, the aggregates and degradation products of the D1 protein. The protein markers are in the left-hand lane of the PVDF membrane.

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12. Under safe light in a dark room, lay the sack with the membrane in an X-ray film cassette and place a section of X-ray film over the sack. 13. After exposure of the film to the PVDF membrane for an appropriate period, develop the film with an X-ray film developer. 14. Reuse of the membrane requires removing the antibodies; incubate the membrane in 30–50 mL of the probe-removing solution in a tightly covered plastic container for 1 h at 50°C with shaking. 15. Transfer the membrane to another container and rinse the membrane several times with TTBS. 16. Incubate the PVDF membrane with TTBS for several minutes with shaking. Repeat this step until the smell of b-mercaptoethanol is not detectable. 17. The membrane is ready for probing with another antibody, beginning with step 4 above (see Note 5). If the membrane is not to be used immediately, store it in a blocking solution in a refrigerator. 3.9. Protein Oxidation Analysis (Fig. 2)

1. Transfer 5 mL of a sample into a 0.5-mL Eppendorf tube (see Note 6).

Fig. 2. A fluorogram showing the oxidized proteins in spinach thylakoids under moderate heat stress. An OxyBlot protein oxidation detection kit (Chemicon, USA) was used for the assay. The thylakoids were incubated at either normal (20°C) or heat-stressed (40°C) conditions for 30 min in the presence (+) and absence (−) of oxygen. Many proteins were found to be already oxidized before the moderate heat stress, but after the heat stress, several proteins were further oxidized, which are indicated by arrows. The increase in protein oxidation was not observed in anaerobic conditions.

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2. Add 5 mL of 20% SDS to the samples and incubate at 20°C for 20 min. 3. Derivatize the samples by adding 10 mL of 1× DNPH solution. To the aliquot of the negative control, add 10 mL of 1× derivatization-control solution instead of the DNPH solution. 4. Incubate both tubes at 20°C for 15 min. 5. Add 7.5 mL of neutralization solution. 6. Denature the samples by adding 27.5 mL of 2× lysis buffer. 7. Separate the protein sample by SDS/urea–PAGE (see Note 7). 8. Western blot analysis (see Notes 8 and 9). 3.10. Assay of Protease(s) by Gelatin/ Polyacrylamide Gel Electrophoresis (Fig. 3)

1. Prepare a gelatin/SDS/urea–polyacrylamide gel. 2. Subject the thylakoids or other protease-containing fractions to gelatin/SDS/urea–PAGE at 4°C. 3. The SDS is removed from the gel by incubating it in renaturation buffer at 25°C for 30 min. Transfer the gel to incubation buffer at 37°C for 16 h.

Fig.  3. Gelatin/acrylamide gel electrophoretogram showing the protease activity of spinach thylakoids. (a) The thylakoids were treated with 2 M KSCN to solubilize a metalloprotease, and after centrifugation, the supernatant was assayed for protease activity with gelatin/acrylamide gel electrophoresis. To the protease-containing fraction, various cofactors or EDTA, an inhibitor of metalloproteases, was added, or the protease was inactivated by heating before the electrophoresis: −, control (no addition), +Zn (+0.5 mM ZnCl2), +ATP (+2 mM ATP and 5 mM MgCl2), +Zn, ATP (+0.5 mM ZnCl2, 2 mM ATP and 5 mM MgCl2), +EDTA (+1 mM EDTA), Heated (the fraction was heated at 80°C for 30 min to inactivate the protease). The arrow indicates the protease. (b) As a control, trypsin was loaded on the gelatin/SDS-PAGE gel. The proteases were detected as white bands against a blue background by CBB staining. The arrow indicates the locations of trypsin. The relative molecular masses of marker proteins are shown on the right-hand side of the fluorogram. In (b), trypsin was overloaded, and broad transparent bands were detected at a higher molecular mass range in addition to the band of trypsin.

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4. Stain the gel with CBB. Proteases are detected as transparent bands in the gel where the gelatin has been digested by protease activity. 5. To extract the proteases from the gel, the gel is not stained with CBB and is homogenized with a glass homogenizer. Proteases are extracted from the homogenate (see Note 11).

4. Notes 1. When preparing the membrane samples, spinach, pea, or Arabidopsis leaves are usually placed in a dark cold room overnight to remove starch in the chloroplasts. It should be noted that all the sample preparation steps are carried out at 4°C in the dark under green safe light, because unnecessary exposure of the samples to light may affect the results of photoinhibition and heat-inhibition studies. After preparation, the samples are immediately frozen in liquid nitrogen and stored at −80°C. During storage, the samples are kept in sample tubes shielded from light by wrapping with aluminum foil. 2. For the conventional SDS–PAGE, urea is omitted from the gels and the lysis buffer described in Subheading 2.6. 3. Work in a clean area. 4. A mini gel is used routinely for the electrophoresis. The running condition is 3–5 mA (constant current) for 12 h (overnight) or 20–25 mA for 2 h. 5. To identify the proteins involved in the protein aggregation, the PVDF membranes are reused for immunoblotting with several specific antibodies. 6. Avoid a high chlorophyll concentration (>1 mg chlorophyll/ mL). 7. Add 0.5 mL of NuPAGE antioxidant to the electrophoresis buffer (running buffer). 8. Dilute the primary antibody stock 1:150 with blocking/ dilution buffer. 9. Dilute the secondary antibody stock 1:300 with blocking/ dilution buffer. 10. To separate SDS-resistant proteases, the step for removing SDS is omitted. The SDS-resistant proteases are detected as white bands at the top of the separation gel, indicating the digestion of gelatin in a blue background, as was shown previously by Sokolenko et al. (19).

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11. The extracted protease activity can be assessed by several methods, including a sensitive fluorescence assay (e.g., EnzChek Protease Assay Kits are available from Molecular Probes Inc., USA). References 1. Macpherson, A. N., Telfer, A., Barber, J., and Truscott, T. G. (1993) Direct detection of singlet oxygen from isolated Photosystem II reaction centres. Biochim Biophys Acta 1143, 301–309. 2. Hideg, É, Spetea, C., and Vass, I. (1994) Singlet oxygen production in thylakoid membranes during photoinhibition as detected by EPR spectroscopy. Photosynth Res 39, 191–199. 3. Mishra, N. P., Francke, C., van Gorkom, H. J., and Ghanotakis, D. F. (1994) Destructive role of singlet oxygen during aerobic illumination of the photosystem II core complex. Biochim Biophys Acta 1186, 81–90. 4. Barber, J. and Andersson, B. (1992) Too much of a good thing: light can be bad for photosynthesis. Trends Biochem Sci 17, 61–66. 5. Aro, E.-M., Virgin, I., and Andersson, B. (1993) Photoinhibition of photosystem II. Inactivation, protein damage and turnover. Biochim Biophys Acta 1143, 113–134. 6. Yamamoto, Y. (2001) Quality control of photosystem II. Plant Cell Physiol 42, 121–128. 7. Yamamoto, Y., Aminaka, R., Yoshioka, M., Khatoon, M., Komayama, K., Takenaka, D., Yamashita, A., Nijo, N., Inagawa, K., Morita, N., Sasaki, T., and Yamamoto, Y. (2008) Quality control of photosystem II: impact of light and heat stresses. Photosynth Res 98, 589–608. 8. Greenberg, B. M., Gaba, V., Mattoo, A. K., and Edelman, M. (1987) Identification of a primary in  vivo degradation product of the rapidly-turning-over 32 kd protein of photosystem II. EMBO J 6, 2865–2869. 9. Andersson, B. and Aro, E.-M. (1997) Proteolytic activities and proteases of plant chloroplasts. Physiol Plant 100, 780–793. 10. Ishikawa, Y., Nakatani, E., Henmi, T., Ferjani, A., Harada, Y., Tamura, N., and Yamamoto, Y. (1999) Turnover of the aggregates and cross-linked products of the D1 protein generated by acceptor-side photoinhibition of photosystem II. Biochim Biophys Acta 1413, 147–158. 11. Yamamoto, Y. and Akasaka, T. (1995) Degradation of antenna chlorophyll-binding

12.

13.

14.

15.

16.

17. 18.

19.

protein CP43 during photoinhibition of photosystem II. Biochemistry 34, 9038–9045. De Las Rivas, J., Andersson, B., and Barber, J. (1992) Two sites of primary degradation of the D1-protein induced by acceptor or donor side photo-inhibition in photosystem II core complexes. FEBS Lett 301, 246–252. Henmi, T., Yamasaki, H., Sakuma, S., Tomokawa, Y., Tamura, N., Shen, J.-R., and Yamamoto, Y. (2003) Dynamic interaction between the D1 protein, CP43 and OEC33 at the lumenal side of photosystem II in spinach chloroplasts: evidence from light-induced cross-linking of the proteins in the donor-side photoinhibition. Plant Cell Physiol 44, 451–456. Yoshioka, M., Uchida, S., Mori, H., Komayama, K., Ohira, S., Morita, N., Nakanishi, T., and Yamamoto, Y. (2006) Quality control of photosystem II: cleavage of reaction center D1 protein in spinach thylakoids by FtsH protease under moderate heat stress. J Biol Chem 281, 21660–21669. Yamashita, A., Nijo, N., Pospíšil, P., Morita, N., Takenaka, D., Aminaka, R., Yamamoto, Y., and Yamamoto, Y. (2008) Quality control of photosystem II: reactive oxygen species are responsible for the damage to photosystem II under moderate heat stress. J Biol Chem 283, 28380–28391. Norén, H., Svensson, P., and Andersson, B. (2004) A convenient and versatile hydroponic cultivation system for Arabidopsis thaliana. Physiol Plant 121, 343–348. Laemmli, U. K. (1970) Cleavage of structural proteins during the assembly of the head of bacteriophage T4. Nature 227, 680–685. Ikeuchi, M. and Inoue, Y. (1988) A new 4.8kDa polypeptide intrinsic to the PSII reaction center, as revealed by modified SDS-PAGE with improved resolution of low-molecularweight proteins. Plant Cell Physiol 29, 1233–1239. Sokolenko, A., Altschmied, L., and Herrmann, R. G. (1997) Sodium dodecyl sulfate-stable proteases in chloroplasts. Plant Physiol 115, 827–832.

Chapter 18 Photosystem II Reconstitution into Proteoliposomes and Methodologies for Structure–Function Characterization David Joly, Sridharan Govindachary, and Mário Fragata Abstract This chapter discusses the photosystem II (PSII) reconstitution into proteoliposomes. In the first part of the chapter, protocols are outlined for the preparation of lipid bilayer vesicles (liposomes) constituted of individual thylakoid lipids or their mixtures, for the preparation of PSII particles, and for the incorporation of the PSII particles into the liposomes. In the second part of the chapter, methodologies are described for the structure–function characterization of the PSII–lipid complexes (proteoliposomes). This includes the sodium dodecylsulfate-polyacrylamide gel electrophoresis determination of the PSII proteins, the measurement of oxygen-evolving activity of PSII in the proteoliposomes, the study of structural changes of the PSII proteins upon their incorporation into the lipid bilayers by Fourier transform infrared (FT-IR) spectroscopy, and the characterization of the PSII activity by fluorescence induction. Key words: Chloroplast lipids, Infrared spectroscopy, Lipid bilayer vesicles, Lipid phase transitions, Liposomes, Oxygen evolution, Photosystem II, Proteoliposomes, Thylakoid membrane, Fluorescence induction

1. Introduction The chloroplasts of higher plants and algae have three membranes (outer, inner, and thylakoid) constituted of similar glyco- and phospholipids (1). That is, monogalactosyl diacylglycerol (MGDG; nonionic lipid), digalactosyl diacylglycerol (DGDG; nonionic lipid), sulfoquinovosyl diacylglycerol (SQDG; anionic lipid), phosphatidylglycerol (PG; anionic lipid), phosphatidylinositol (PI; nonionic lipid), and phosphatidylcholine (2–4) (PC; zwitterionic lipid between pH 2 and 10 (see Note 1). Their average compositions in the outer, inner, and thylakoid membranes, as reported in (5, 6), are displayed in Table 1. The table shows that the inner and thylakoid membranes have about the same lipid content. On the Robert Carpentier (ed.), Photosynthesis Research Protocols, Methods in Molecular Biology, vol. 684, DOI 10.1007/978-1-60761-925-3_18, © Springer Science+Business Media, LLC 2011

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Table 1 Average lipid compositions (in mol%) in the outer, inner, and thylakoid membranes of the chloroplast of higher plants Chloroplast lipids

Chloroplast membrane

MGDG

DGDG

SQDG

PG

PC

PI

Refs.

Outer

17

39

6

10

32

5

(5)

Inner

49

30

5

8

6

1

(5)

Thylakoid

52

26

6.5

9.5

4.5

1.5

(5)

Thylakoid

54

24

9.6

7.4

4.8

(6)

DGDG digalactosyl diacylglycerol, MGDG monogalactosyl diacylglycerol, PC phosphatidylcholine, PG phosphatidylglycerol, PI phosphatidylinositol, SQDG sulfoquinovosyl diacylglycerol

contrary, the outer membrane has relative PC and MGDG compositions that differ considerably from the lipid content observed in the two other membranes. That is, a quite larger amount of PC in relation to MGDG. A major question in the study of structure–function relations of the chloroplast lipids in photosynthesis is how to examine the functional dissimilarities of the three chloroplast membranes, i.e., the import of cytosolically synthesized proteins through the outer and inner membranes (7) or the primary processes of photosynthesis in the thylakoids (8), in the context of the similarities and dissimilarities of lipid structure and composition referred to above. Several methods are available for the study of the role of chloroplast lipids (see Note 2). Hereunder, the reconstitution of purified photosystem II particles into proteoliposomes composed of chloroplast lipids is described. This methodology is useful in (1) the study of the thylakoid lipids effect on oxygen evolution and electron transfer and (2) the characterization of the lipid– protein interactions that are at the origin of protein folding or misfolding. In the first part of this chapter, protocols are given for the incorporation of photosystem II (PSII) complex into lipid bilayer vesicles (proteoliposomes) constituted of individual thylakoid lipids or their mixtures. The second part of the chapter contains methodologies for the characterization of PSII function in the proteoliposomes by measuring oxygen evolution with the Clark electrode, and the study of structural changes of the PSII proteins upon their incorporation into the lipid bilayers by Fourier transform infrared (FT-IR) spectroscopy.

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2. Materials In many instances it is necessary to purify the salts and lipids used in the preparation of proteoliposomes, as well as to analyze the chemical structure of the fatty acid chains of the glyco- and phospholipids. The procedures for the purification of salts are given in Note 3, those for purification of lipids in Note 4, and the fatty acid analysis is discussed in Note 5. 2.1. Preparation of Lipid Bilayer Vesicles

1. Amicon cell equipment. 2. Buffer C: 20 mM MES–NaOH, pH 6.5, 400 mM sucrose, 15 mM NaCl, and 5 mM MgCl2. 3. Chromatography columns. 4. Gas chromatograph (e.g., Varian, model 3700, equipped with a Shimadzu integrator, model C-R3A). 5. HPLC equipment. 6. Lipids: (a) DGDG (digalactosyl diacylglycerol). (b) MGDG (monogalactosyl diacylglycerol). (c) SQDG (sulfoquinovosyl diacylglycerol). (d) PC (phosphatidylcholine). (e) PG (phosphatidylglycerol). (f) PI (phosphatidylinositol). 7. Millipore equipment. 8. Sepharose-4B gel (2.5 × 60 cm) (Pharmacia Fine Chemicals). 9. Sonicator (e.g., Heat Systems-Ultrasonics, Plainwiew, LI). 10. Sonifier cell disrupter (Model W-225R, Heat SystemsUltrasonics, Plainwiew, LI).

2.2. Preparation of Photosystem II Particles

1. Barley seeds (or other adequate plant material). 2. Buffer A: 50 mM Tricine–NaOH, pH 7.8, 400 mM sorbitol, 10 mM NaCl, and 5 mM MgCl2. 3. Buffer B: 50 mM Tricine–NaOH, pH 7.8, 10 mM NaCl, and 5 mM MgCl2. 4. Buffer C: 20 mM MES–NaOH, pH 6.5, 400 mM sucrose, 15 mM NaCl, and 5 mM MgCl2. 5. Buffer D (PSII storage buffer): 20 mM MES–NaOH, pH 6.5, 400 mM sucrose, 15 mM NaCl, 5 mM MgCl2, and 30% (v/v) glycerol. 6. Cheesecloth tissue. 7. Triton X-l00.

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8. 80% (v/v) Acetone. 9. Vermiculite and polythene trays (or other adequate material). 2.3. Photosystem II Reconstitution into Proteoliposomes 2.4. SDSPolyacrylamide Gel Electrophoresis

1. Buffer C: 20 mM MES–NaOH, pH 6.5, 400 mM sucrose, 15 mM NaCl, and 5 mM MgCl2. 1. Acrylamide stock solution: 45% (w/v) acrylamide, 1.2% bisacrylamide. 2. SDS 10% (w/v). 3. Ammonium persulfate 10% (w/v). 4. TEMED. 5. Urea. 6. Isopropanol. 7. 4X Stacking gel buffer: 0.0541  M Tris–0.0267  M H2SO4, pH 6.1. 8. Stacking gel: 5% acrylamide (1.1 mL acrylamide stock solution), 6  M Urea, 0.1% SDS (100  mL SDS stock solution), 2.5 mL 4X Stacking gel buffer (total volume, 10 mL). At last: 0.05% ammonium persulfate (100  mL stock solution) and 0.05% TEMED (5 mL). 9. Resolving gel buffer: 0.4244  M Tris–0.0308  M HCl, pH 9.18. 10. Resolving gel: 15% acrylamide (6.67  mL acrylamide stock solution), 6 M Urea, 0.1% SDS (200 mL SDS stock solution), 4  mL resolving gel buffer (total volume, 20  mL). At last: 0.05% ammonium persulfate (200  mL stock solution) and 0.04% (v/v) TEMED (8 mL). 11. Sample buffer: 4X Stacking gel buffer (250  mL), glycerol (250 mL), 2-mercaptoethanol (25 mL), SDS (200 mL SDS stock solution), urea (720 mg), bromophenol blue (a few grains). 12. 5X Lower reservoir buffer: 0.4244 M Tris–0.0308 M HCl, pH 9.18. 13. 20X Upper reservoir buffer: 0.04  M Tris–0.04  M borate, pH 8.64, SDS 2% (w/v). 14. Staining solution: 10% (v/v) acetic acid, 50% (v/v) methanol, 0.125% (w/v) Coomassie Brilliant Blue R-250. 15. Destaining solution: 10% (v/v) acetic acid, 30% (v/v) methanol. 16. A kit of protein molecular mass markers.

2.5. Measurement of Oxygen Evolution

1. Argon or nitrogen cylinder. 2. Buffer C: 20 mM MES–NaOH, pH 6.5, 400 mM sucrose, 15 mM NaCl, and 5 mM MgCl2.

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3. 2,5-Dichloro-p-benzoquinone (DCBQ). 4. Dithionite. 5. Clark-type electrode equipment (e.g., Hansatech D.W. Oxygen Electrode Unit, King’s Lynn, Norfolk, UK). 2.6. Structural Study of PSII and PSII–Lipid with FT-IR Spectroscopy

1. Argon or nitrogen cylinder. 2. Buffer C: 20 mM MES–NaOH, pH 6.5, 400 mM sucrose, 15 mM NaCl, and 5 mM MgCl2. 3. FT-IR spectrophotometer. 4. Software program for the mathematical treatment of the infrared data; the use of one of the Spectra-Calc programs from Galactic Industries Corporation (Salem, NH) is recommended, but any appropriate mathematical software is acceptable. 5. Infrared spectroscopy optical windows (BaF2 or ZnSe).

2.7. Characterization of the PSII Activity by Fluorescence Induction

1. Hansatech plant efficiency analyzer (PEA) fluorometer. 2. Leaf clip. 3. Plastic cylindrical vial and sample holder. 4. Buffers B and C.

3. Methods The first part of the methods (Preparation of PSII Proteoliposomes) outlines the protocols for (1) the preparation of lipid bilayer vesicles, (2) the extraction of photosystem II particles, and (3) the photosystem II reconstitution into proteoliposomes. The second part of the methods (Characterization of PSII Proteoliposomes) describes procedures for (1) the SDS-PAGE, (2) the measurement of oxygen evolution in PSII, (3) the structural study of the proteoliposomes by FT-IR spectroscopy, and (4) the characterization of the PSII activity using fluorescence induction. 3.1. Preparation of PSII Proteoliposomes

The protocols described in Subheadings  3.1.1–3.1.3 are wellknown, reliable procedures for (1) the preparation of lipid bilayer vesicles (liposomes) of about 250 Å (25  nm) diameter, (2) the preparation of photosystem II (PSII) particles, and (3) the reconstitution of the PSII particles into the lipid bilayer vesicles (proteoliposomes), respectively. It is noted, however, that several other procedures for performing the same purpose are available in the literature, as well as a vast array of other methods designed for fundamental and applied research such as the many present day therapeutical purposes. Note 6 guides the reader to some of those methodologies.

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3.1.1. Preparation of Lipid Bilayer Vesicles

Before attempting to form lipid bilayer vesicles (liposomes) in aqueous media, one might consider any available information on the phase behavior of the lipids, a question that is crucial for the success of any preparation (this is discussed in Note 7). In short, one shall inquire first which lipids are bilayer-forming or nonbilayerforming; second, one shall address the question of the phase transition temperatures of the lipids. The latter point is particularly important since in most cases it is not possible to obtain a liposome preparation at temperatures below the gel-to-liquid crystal phase transition temperature of the lipid. For example, lipid vesicles never form in aqueous mixtures of dipalmitoyl phosphatidylcholine (DPPC) with water at room temperature (see Note 7). In fact, the temperature of most laboratories is well below the gel-to-liquid crystal transition of DPPC, which is 41°C (14), thereby indicating the need for heating the water–DPPC mixture to temperatures above the 41°C limit. The method described hereunder is a standard procedure used for the formation of lipid bilayer vesicles constituted of either one lipid class or binary and ternary mixtures of different lipid species. The following steps are those described in (4, 15, 21, 22): 1. 20 to 150 mg of the purified lipids (see Note 4) are first solubilized in diethyl ether (or another appropriate organic solvent), and then dried under a current of nitrogen or argon to avoid oxidation of the lipids. Note, however, that the solubilization step can be skipped if the preparation of the bilayer vesicles is made with only one purified lipid species. 2. Step 1 is followed immediately by dispersion of the dried lipids in 10  mL of buffer C to give a lipid concentration of 10 mg/mL in the final solution. 3. The suspension obtained in step 2 is subjected to sonication in a test tube with a titanium sonicator probe for 15 min at 160 W output in a sonifier cell disrupter, having nitrogen or argon bubbling into the solution (see Note 8). 4. The mixture obtained in step 3 is centrifuged at 100,000 × g for 1 h and the supernatant is concentrated to 1 or 2 mL in an Amicon cell (or a similar equipment). 5. Step 4 is followed by fractionation of the lipid dispersion in a Sepharose-4B column and elution with the above-described buffer. Then, 5-mL fractions are collected. A typical elution diagram is given in Fig. 1. 6. Only the homogeneous fractions constituted of unilamellar vesicles (class III vesicles; see Fig. 1) are retained for the preparation of proteoliposomes.

3.1.2. Preparation of Photosystem II Particles

Before proceeding further into the description of the methods used here, it is useful to emphasize that

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Fig. 1. Typical elution diagram of phosphatidylcholine vesicles prepared by ultrasonic dispersion in a buffer containing 20 mM MES (2-[N-morpholino]ethanesulfonic acid)– NaOH (pH 6.5), 400 mM sucrose, 15 mM NaCl, and 5 mM MgCl2 followed by ultracentrifugation (100,000 × g; 1 h) and gel filtration in a Sepharose-4B column. The volume of each fraction is 5 mL. I, class I liposomes constituted of large multilayered vesicles; II, class II liposomes constituted of mixtures of class I and class III; III, class III liposomes constituted of small single-layered vesicles of about 20–25 nm diameter. Reproduced from (58) with permission from Humana Press.

1. The following protocols are designed for the extraction of PSII particles from barley (Hordeum vulgare) seedlings; these protocols may need to be changed if other biological materials are used to obtain the PSII particles. 2. All the steps described in this section are performed under green dim light at temperatures in the 0–4°C range. 3. All glassware, buffers, and other materials must be precooled in a refrigerator before use and maintained in containers filled with ice during the extraction procedures. 4. All biological materials (chloroplasts, thylakoid membranes, and PSII particles) must always be kept in closed dark containers filled with ice, unless one chooses to work in a cold room under green dim light. 3.1.2.1. Cultivation of Barley Seedlings

Soaking the seeds in water: 1. 200  g of barley seeds are first washed thoroughly and then spread inside polythene trays; water is added to the trays till the seeds are immersed.

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2. Cover the seeds with a layer of white tissue paper. 3. Leave the trays in darkness for 24 h at room temperature. Planting the seeds on vermiculite: All the steps below should take place in a growth chamber, if possible. 1. Add vermiculite to polythene trays to a height of 2–3 cm. 2. Spread the barley seeds (presoaked in water) on the surface of the vermiculite and sprinkle the seeds with water; the water should be spread over the whole area of the polythene tray. 3. Spread vermiculite over the seeds (see step 2) to a height of 1 cm in order to cover all the seeds. 4. Let the seeds germinate and grow for about 6–8 days at 23 ± 2°C and a light intensity of 200 ± 20 mmol/m2s. 5. After germination, sprinkle the seeds every day with about 200 mL water until the seedlings harvest. 3.1.2.2. Isolation of Chloroplasts and Thylakoid Membranes

The thylakoid membranes are obtained from chloroplasts of 6- to 8-day-old-barley (H. vulgare) seedlings according to the procedure of Berthold et al. (23) with modifications described in (24, 25). 1. Barley leaves are cut to 1 cm length and introduced in a precooled homogenizer jar to full volume. The barley leaves fragments are then homogenized (10–15 s) at 0°C in the precooled buffer A. The homogenization should take place in the dark. Cover the homogenizer jar with a black cloth during operation. 2. Filter the homogenized slurry through eight layers of cheesecloth tissue, and then centrifuge the filtrate at 1,000 × g for 5 min at 4°C. 3. The pellet obtained in step 2 (chloroplasts) is dispersed in buffer A to a total volume of 200–400 mL using a brush and a vortex mixer; then, the dispersion is centrifuged at 1,000 × g for 5 min at 4°C. 4. The chloroplast pellet is suspended in buffer B using brush and vortex mixer; this is followed by a centrifugation at 1,000 × g for 5 min at 4°C. 5. The pellet obtained in step 4 (thylakoid membranes) is suspended in a minimal volume of precooled buffer B, using brush, vortex mixer and a teflon homogenizer; this step might be followed by a centrifugation at 1,000 × g for 5 min at 4°C. 6. The pellet of thylakoid membranes obtained in step 5 is suspended in buffer C to give a final chlorophyll (Chl) concentration of 2 mg/mL and stored at about –80°C until use. 7. The Chl concentration is estimated using 80% (v/v) acetone solutions according to the method of Arnon (26). (A useful procedure is described in Note 9.).

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1. First, the thylakoid membranes kept at –80°C (see Subheading “Isolation of Chloroplasts and Thylakoid Membranes”) must be brought to 0°C in a container filled with ice. 2. The chlorophyll concentration in the thylakoid samples might be estimated according to the procedure in the Subheading “Isolation of Chloroplasts and Thylakoid Membranes.” 3. The oxygen-evolving activity of the thylakoid membranes might be assayed prior to the isolation of the PSII particles. 4. Using a vortex, mix 1 mL of the thylakoid membrane suspension (2 mg Chl/mL) with 0.5 mL of buffer C. 5. With very gentle shaking, add dropwise 0.5 mL of 8% Triton X-l00 to the mixture obtained in step 4 to give a final Chl concentration of 1 mg/mL; the final Triton X-100:Chl ratio should be 20:1. Very importantly, during the addition of Triton X-100 solution, the use of a vortex or any other mechanical shaking device should be avoided; this step must be done slowly, but as fast as possible, and it shall be noted finally that the use of gloves is mandatory to avoid any skin contact with Triton X-100 solution. 6. The suspension obtained in step 5 is incubated in the dark for 20 min at 4°C and then centrifuged at 1,000 × g for 3 min at 4°C to precipitate the unbroken thylakoids. 7. The supernatant of step 6 is transferred to one or several prechilled centrifuge tubes and centrifuged immediately at 29,000 × g for 30 min at 4°C. 8. The supernatants of each centrifuge tube of step 7 (containing Triton X-100) are discarded, and 1  mL of buffer C is added to the pellet of each of the tubes, followed by dispersion of the pellets using a brush. 9. Add 10–15  mL of buffer C to each one of the centrifuge tubes in step 8 and centrifuge them immediately at 29,000 × g for 30 min at 4°C; in this step, the residual Triton X-100 shall be removed. 10. The pellets of all tubes used in step 9 (PSII particles) are pooled together upon suspension in small volumes of buffer D to give a final volume of 1 mL. 11. Immediately after step 10, or as soon as possible, it is useful to determine the characteristics of the PSII particles just ­prepared by measuring (1) their chlorophyll content, (2) the oxygen-evolving activity, and (3) the polypeptide composition with SDS-polyacrylamide gel electrophoresis. It is worth noting that the electrophoresis test is particularly important to check whether the PSII particles are contaminated with photosystem I proteins. 12. Finally, the PSII particles are stored at –80°C until use.

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3.1.3. Photosystem II Reconstitution into Proteoliposomes

The steps outlined here for the preparation of PSII–lipid complexes have been described in (4, 24, 25). It is important to note that all the following steps were performed in green dim light and at temperatures of ~4°C; one may work either in a cold room in green dim light or maintain all buffers and other materials in dark containers filled with ice. To prepare the PSII–lipid membranes, follow these steps: 1. To wash the PSII particles obtained as described above (see Subheading  3.1.2), the solubilized particles are first centrifuged at 29,000 × g for 30 min at 4°C and then suspended in buffer C; this step is repeated once using again buffer C as the suspension medium. 2. The washed PSII particles obtained in step 1 are gently mixed in a vortex with the lipid vesicles (see Subheading 3.1.1); the lipid:chlorophyll ratio is maintained at 20:1 (w/w) throughout the work (4). 3. If one needs to study the effect of ions on the activity of the PSII–lipid complex, the salts can be added at this stage. 4. Then, the protein–lipid mixtures are incubated in darkness at 0°C for 20 min. 5. Immediately after the incubation step, an aliquot of the preparation is used for oxygen evolution determination according to the procedure outlined below (see Subheading 3.2.2). 6. Concomitantly, washed PSII samples with no lipid added is handled in exactly the same way and their oxygen evolution activity is measured.

3.2. Characterization of PSII Proteoliposomes

The protocols described here include first standard procedures of SDS-PAGE and measurement of oxygen evolution with a Clark-type electrode (see Subheadings 3.2.1 and 3.2.2). Next, Subheading 3.2.3 outlines the essentials of the FT-IR spectroscopy methodology, which has been successfully used in a large number of studies. For example, the examination of structural changes in the photosynthetic membrane is induced by a variety of effects such as UV-B radiation (27) and heat stress (28). The examples discussed hereunder are limited to the analysis of the “spectral window” between 1,700 and 1,480/cm (Figs.  2–4) where most bio-lipids have no significant infrared absorption. Procedures are described for the preparation of samples for FT-IR spectroscopy, the measurement of infrared spectra, and the analysis of infrared data. Finally, the Subheading 3.2.4 describes the methodology for the characterization of the PSII activity using fluorescence induction.

3.2.1. Sodium Dodecyl Sulfate-Polyacrylamide Gel Electrophoresis

The polypeptide composition of the PSII particles is analyzed by SDS-PAGE in order to determine whether they are contaminated with photosystem I (PSI) proteins (see list of PSI and PSII polypeptide subunits in (8, 30).

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.8 1. PSII-DGDG 2. PSII 3. DGDG

Absorbance

.6

.4

.2

1 I

Y8a,b

II

Y19a

2 3

0 1800

1700

1600

1500

Wavenumber, cm-1 Fig. 2. Infrared spectra of photosystem II (PSII), digalactosyl diacylglycerol (DGDG) and the PSII–DGDG complex between 1,800 and 1,480/cm. I amide I region (1,700–1,630/ cm), II amide II region (1,580–1,520/cm), Y8a,b and Y19a regions of the 8a and 8b (1,630–1,580/cm) splitting components of the fundamental mode no. 8 and the high frequency splitting component 19a (1,520–1,500/cm) of the phenol ring according to Wilson (39). Note that in the “spectroscopic window ” from 1,700 to 1,480/cm, DGDG has only a negligible absorbance. Reproduced from (58) with permission from Humana Press.

First, it is remarked that the protocols outlined hereunder (preparation of the gels for SDS-PAGE as well as many other experimental details) are taken from the procedures described by Chua (29), which should, therefore, be consulted before proceeding further in this section. In addition, it is specially important to note the following: 1. All steps described below are performed at room temperature. 2. SDS and TEMED might be stored at room temperature. 3. Always prepare fresh solutions of urea and ammonium persulfate. 4. While handling acrylamide (monomeric product), use a mouth-and-nose mask and a pair of gloves; this precaution is not necessary upon polymerization of the acrylamide. The standard proteins and PSII polypeptides are resolved on a linear gradient gel with the separating gel containing 15% acrylamide and 6 M urea and the stacking gel having 5% acrylamide

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Amide I-II region .2

Absorbance

.15

.1

H2O

.05 D2O 0 1800

1700

1600

1500

Wavenumber, cm-1 Fig. 3. Infrared absorbance of H2O and D2O between 1,800 and 1,480/cm, that is, in the spectral regions of the amide I, amide II, and the splitting components of the fundamental modes no. 8 and no. 19 of the phenol ring (cf. Fig. 2). Reproduced from (58) with permission from Humana Press.

and 6 M urea. Before loading into the gel slots, the PSII samples are first solubilized with SDS. This is followed by a centrifugation at 13,600 × g for 5 min to remove the unsolubilized membranes. Upon electrophoresis, the gels are stained for 3 h in the staining solution and then in the destaining solution. To estimate the molecular mass of the proteins, a plot of log molecular mass vs. the migrated distance (in centimeters) is used for the unknown proteins and standard protein markers. The molecular masses of the markers are 15, 25, 30, 35, 50, 75, 105, 160, and 250 kDa. The PSII polypeptide subunits usually detected in the electrophoresis gel are the 16-, 23-, and 33-kDa extrinsic proteins, the S protein, the a-cyt b559, the proximal antenna proteins (CP43 and CP47), and the D1 and D2 proteins in the reaction center (34 and 32 kDa, respectively) (8). 3.2.2. Measurement of Oxygen Evolution

Measurement of electron transport through PSII estimated as oxygen evolution in the PSII particles and the PS II–lipid complex is performed at 25°C with a Clark-type electrode (see Note 10). The reaction medium (total volume: 2  mL) contains the following: 1. 1,960 mL of the measurement buffer C 2. 20 mL of 350 mM DCBQ as exogenous electron acceptor at the QA site (8)

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3 ABSu

2

Absorbance

FSD

1 2nd-D

0 4th-D 1800

1700

1600

1500

Wavenumber, cm-1 Fig.  4. Resolution enhancement of a untreated infrared spectrum of photosystem II (ABSu) using the Fourier self deconvolution (FSD), second derivative (2nd-D) and fourth (4th-D) derivative methods. The calculations were performed with the “Mathematical tool” of the Spectra-Calc program (see text). Reproduced from (58) with permission from Humana Press.

3. 20 mL samples of PSII particles or PSII–lipid complex (final concentration: 12.5 mg Chl/mL) The PSII and PSII–lipid preparations are then illuminated with white light at saturating light intensity. Note that usually the universal unit used for expressing oxygen-evolving activity is mmol O2/mg Chl/h. For the measurement of the pH effect on oxygen evolution, one may choose to adjust the pH of the buffer media in the pH 3–10 range with HCl or NaOH. This practice is useful if one wants to avoid changes of chemical composition of the reaction media as is the case when one uses three different buffer systems to cover the pH 3–10 range. 3.2.3. Structural Study of PSII and PSII–Lipid with FT-IR Spectroscopy

FT-IR spectroscopy is widely used in structural studies of proteins (31). Among those studies, a large number was dedicated to the examination of structure–function relations in the photosynthetic membrane (32–35). Although the useful infrared range spans from about 4,000 to 600/cm, most works are performed between

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1,700 and 1,480/cm, i.e., the amide I and amide II regions. The reason for this is that these spectral regions contain a set of vibrational that which are related to several periodic structures found in many proteins. In addition, the amide I and amide II regions are very sensitive to conformational changes in the proteins secondary structures such as those caused by ultraviolet light (27) or thermal effects (28, 34, 36). Table 2 collects the wavenumbers and vibrational modes of the major bands between ~1,780 and 1,500/cm in the infrared spectra of photosystem II and also gives information on the protein periodic structures that were deduced from the spectral data. The data in Table 2 are from (27, 31, 35–39) and literature cited therein. In brief: 1. The region from ~1,780 to 1,700/cm is characterized by vibrational modes originating in the lipids and chlorophylls. 2. The region from 1,700 to 1,600/cm (amide I) is associated with an in-plane C=O stretching vibration (~80%) weakly coupled with some contributions from CN stretching and CCN deformation. 3. The region from 1,580 to 1,520/cm (amide II) is assigned to an in-plane NH bending strongly coupled with CN stretching. 4. The spectral bands at about 1,608, 1,590, and 1,508/cm originate in the vibrational modes of the phenol ring, e.g., in tyrosine. Hereunder, the question of the “spectral window” between 1,700 and 1,480/cm in lipid–protein studies is discussed first, and then the methods are outlined (1) for the preparation of samples for FT-IR spectroscopy, (2) for the measurement of infrared spectra, and (3) for the analysis of infrared data. 3.2.3.1. “Spectral Window” Between 1,700 and 1,480/ cm in Infrared Spectra of Bio-Lipids

It is interesting to note that in the study of lipid–protein interactions one takes advantage of a very useful “spectral window” between 1,700 and 1,480/cm where most lipids do not absorb infrared radiation (35, 40). This is clearly seen in Fig. 2 for the interaction between PSII and the galactolipid DGDG (digalactosyl diacylglycerol). That is, from about 1,700 to 1,480/cm the DGDG absorbance is negligible comparatively to the large absorbance of infrared light by the PSII and PSII–DGDG complexes. This means that within the 1,700–1,480/cm range any lipid-induced change of the infrared spectra of the PSII–lipid complex is not affected by the lipid absorbance, which is observed either at wavenumbers greater than 1,700/cm or smaller than 1,480/cm.

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Table 2 Wavenumbers and vibrational modes of the major spectral bands between 1,760 and 1,500/cm in the infrared spectrum of photosystem II with indication of protein periodic structures in the amide I and amide II regionsa Spectral region

Wavenumber (per cm)

Vibrational modes and protein periodic structuresb

Amides Ic, IId

1,627 1,572, 1,564 1,550, 1,531

daNH3+ or dNH2 naCOO− dsNH3+

Tyrosine

1,620–1,580 ~1,608 ~1,590 1,520–1,500

n8a ringe n8b ringe n19a ring (~1,508/cm) + deformatione

Chl + lipids

1,760–1,692 ~1,739 ~1,737 ~1,693 ~1,662

nCO (sn1 or sn2 ester CO) of lipids nCO (ester CO) of chlorophylls nCO (free keto CO) of chlorophylls Bound keto CO of chlorophylls

Vibrational modes

Protein periodic structures Amide I

Amide II

1,696–1,620 1,696–1,665 1,658–1,654 1,648–1,641 1,640–1,620 1,580–1,520 ~1,545

Turns (e.g., b-turns), antiparallel b-sheet (~1,693/cm) a-helix + some random coil structures Random structures, loops b-sheet (~1,636/cm), extended chain (b-strand: ~1,626/cm) Some a-helical conformations

Data from Arrondo et al. (31), Gabashvili et al. (35), Ségui et al. (27), and literature cited therein Chl chlorophyll, d bending, da asymmetric bending, ds symmetric bending, n stretching, na asymmetric stretching c Amide I: In-plane C=O stretching vibrations weakly coupled with CN stretching and CCN deformation d Amide II: In-plane NH bending strongly coupled with CN stretching e Wilson et al. (39) a

b

3.2.3.2. Preparation of Samples for FT-IR Spectroscopy

Infrared investigations of protein–lipid interaction are carried out in deuterium oxide (D2O) to overcome the strong overlapping effect of the H2O absorption in the amide I and amide II regions (see Fig. 3). The protocols given below, which are based on those described in (41), outline (1) a modified ­procedure for preparing liposomes and proteoliposomes in D2O and (2) the steps for preparation of samples for layering on BaF2 or ZnSe plates.

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Preparation of Liposomes and Proteoliposomes in D2O

1. The lipid bilayer vesicles (liposomes) are prepared according to the protocols described in Subheading  3.1.1 except for replacement of the water solvent with D2O. 2. For preparing the PSII–lipid membranes (proteoliposomes), the PSII particles obtained in Subheading  3.1.2 are first washed in D2O by centrifugation at 28,700 × g for 7 min at 4°C, and then complexed with the liposomes obtained in step (a) (see Subheading 3.1.3).

Preparation of Samples for Layering on BaF2 or ZnSe Plates

1. Aliquots of the PSII or PSII–lipid samples (~1.5 mg Chl/mL) obtained as described in step 1 are centrifuged at 28,700 × g for 7 min at 4°C. 2. The pellet is suspended in 3 mL D2O and centrifuged a second time (28,700 × g; 7 min; 4°C). 3. The final pellet is suspended again in a D2O volume sufficient to make a solution with a chlorophyll concentration of 0.333 mg/mL. 4. An aliquot of the solution is immediately layered on 25 mm diameter BaF2 or ZnSe plates, which are at once stored in darkness on filter paper placed on ice in a controlled temperature chamber (~0°C). 5. The samples are then dehydrated for about 90 min under a current of nitrogen. In the above conditions any 1H/2H exchange is minimized, thereby assuring that the results are not influenced significantly by the presence of D2O molecules in the PSII samples.

3.2.3.3. Measurement of Infrared Spectra

The infrared absorbance measurements are made in a Nicolet FT-IR spectrophotometer, model 420, or another appropriate instrument. In the Nicolet FT-IR instrument, 100 interferograms are in general collected and co-added using its Omnic software facility. The infrared spectra are obtained upon subtraction of the spectrum of the BaF2 or ZnSe plates used for deposition of the samples. The spectral resolution is between 1 and 2/cm.

3.2.3.4. Analysis of Infrared Data

The FT-IR spectra are processed using the GRAMS/386 SpectraCalc program, or another appropriate software. Hereunder, information is provided (1) for the baseline correction, (2) for the mathematical analysis of FT-IR spectra, (3) for the determination of the contents in protein periodic structures, and (4) for the calculation of difference spectra. 1. Baseline correction. Prior to data processing, or before any mathematical treatment of the FT-IR spectra, it is important to perform a baseline correction of the original spectra between about 4,000 and 600/cm using the “baseline correction tool” of the Spectra-Calc program.

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2. Mathematical analysis of FT-IR spectra. Identification of the band maxima frequencies (or wavenumbers) is first done on inspection of the FT-IR absorbance spectra. However, the amide I and amide II envelope is usually rather featureless (cf. PSII and PSII–DGDG spectra in Fig. 2), thereby rendering in a first approximation the analysis of the infrared spectra difficult or not worthy. To cope with this disadvantage one turns to mathematical methods of “resolution enhancement.” For example, the “mathematical tool” of the Spectra-Calc program offers the possibility of performing various types of resolution enhancement: (1) the Fourier self deconvolution (FSD), (2) the second and fourth derivatives, and (3) curve-fitting analyses using Gaussian, Lorentzian, and Voigt functions. (a) Fourier self deconvolution (FSD). In this method the effective spectral bandwidth is synthetically narrowed. As apodization filters, FSD uses either the Bessel or the Boxcar function, although in most instances the Bessel function is preferred. Upon application of this method it becomes easier to identify the principal bands under a much larger multiband envelope with overlapping individual band features. This is seen in Fig. 4, which displays a typical “untreated” absorbance spectrum of PSII together with its FSD spectrum. (b) Second and fourth derivatives. The “Differentiate tool” of the Spectra-Calc program provides the first and second derivatives using the Savitsky-Golay convolution method (Analytical Chemistry, 1964, vol. 36, page 1627 et seq). To obtain the fourth derivative, one uses the second derivative spectrum as the starting data for calculation. Now, it is important to note for good interpretation of the data that a “band maximum in the original spectrum” (and the FSD spectrum) is displayed as a “band minimum in the second derivative spectrum” and as a “band maximum in the fourth derivative spectrum.” This is clearly seen in the amide I-II spectrum of PSII displayed in Fig. 4. As an example, one may wish to verify in Fig. 4 that the a-helix band maximum (Table 2) seen in the original (“untreated”) spectrum at approximately 1,655.7/cm is displayed at 1,657.7/cm in the FSD, second derivative and fourth derivative spectra, that is, a displacement of about 2/cm to a higher wavenumber. In most instances, the 1,657.7/cm wavenumber, or near, is taken as a better estimation of a-helix band maxima. (c) Curve-fitting analyses. The “curve-fitting tool” of the Spectra-Calc program automatically calculates the best fit of Gaussian, Lorentzian, or Voigt bands, which make up

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a complex set of overlapping peaks. The program works first on a guess at the number of overlapping bands present within the multiband envelope, their peak positions, their peak widths, and the peak types (e.g., Gaussian, Lorentzian, or their mixtures). Then, it iterates to find a combination of band heights, positions, and widths, which best fit the experimental data. It is here that the data obtained with the FSD, second derivative and fourth derivative calculations (see above), are extremely important. In fact, all calculations suggested in this subsection (b) should be performed sequentially and followed by comparisons with data available in the literature. The final result of the “curve-fitting analysis” is a set of synthetized spectral bands that represent the several periodic substructures that make up the molecular or supramolecular structure of the protein or protein complexes under study (Table 2). 3.2.3.5. Determination of the Contents in Protein Periodic Structures

The procedure is quite simple. First, the total area of the amide I-II envelope surface from about 1,700 to 1,500/cm is determined using the “integration tool” of the Spectra-Cal program. Second, the individual areas of the different spectral bands obtained in the “curve-fitting analysis” described above are as well determined with the “integration tool.” Third, the percentages of a-helix, b-sheet, b-strands, or any of the other structures are found by computing the ratio of the individual band areas in relation to the total area of the amide I-II surface. Examples of this type of calculations are seen in Figs. 2 and 3 of (40). For identifying a given protein periodic structure (a-helix, b-sheet, b-strand, etc.) with a specific spectral band, use the information in Table 2 or data given elsewhere in the literature.

3.2.3.6. Calculation of Difference Spectra

To obtain the difference spectra, i.e., the [PSII–lipid]-minus[PSII] spectra, the methods developed in (35, 42, 43) are applied. First, the areas under the spectral envelope in the 1,700–1,500/ cm region, i.e., amides I and II, for the PSII and the PSII–lipid spectra are determined using the “integration tool” of the SpectraCalc program. Second, the PSII and the PSII–lipid areas are normalized according to the expression An = kAo, where Ao is the integral of the observed surface under the band envelope calculated with the Spectra-Calc Program, k is a scale factor that normalizes Ao to 100, and An is the normalized area, i.e., 100. Thirdly, the [PSII–lipid] − [PSII] subtraction of the “normalized spectra” is performed.

3.2.4. Characterization of the PSII Activity by Fluorescence Induction

Measurement of chlorophyll (Chl) a fluorescence is widely used to probe photosynthetic efficiency because it is noninvasive, easy, fast, and reliable. Kautsky and Hirsh (44) first found

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a relationship between the fluorescence and photosynthetic activity during illumination of dark-adapted leaves. It was later found that fluorescence increased with the photochemical reduction of the primary quinone acceptor of PSII, QA (45). Excitation of the PSII reaction center (RC) leads to a charge separation with the formation of the reversible primary radical pair P680+Pheo−. The primary radical pair is stabilized by QA and the secondary donor tyrosine Z (YZ), thus forming YZ+P680PheoQA−. Therefore, the efficient trapping of excitation by PSII is directly dependant on the capacity of QA to accept electrons from Pheo−. As a consequence, a high trapping efficiency when QA is oxidized (open RC) leads to an efficient dissipation of absorbed light energy as photochemistry. On the opposite, a low trapping efficiency when QA is reduced (close RC) increases the dissipation of absorbed light energy as fluorescence (46, 47). Improvement of time resolution of fluorometers in the last decades allowed the measurement of fluorescence kinetics at microsecond time-scale with commercial instruments. Upon continuous illumination of dark-adapted leaves with high light intensity, Chl a fluorescence rises from its minimal level F0 (or O) to its maximal lever Fm (or P) in three distinct phases delimited by two clear peaks, J and I (48, 49). This O-J-I-P fluorescence rise is also called fluorescence induction (FI). The O-J rise reflects the fast reduction of QA within the first ms of illumination (49) while J-I probably represents the accumulation of RC in QA−QB2− (50). I-P is related with the removal of nonphotochemical quenching of Chl excited states by oxidized plastoquinone (PQ) molecules of the thylakoid membrane pool during their progressive accumulation as PQH2 following PSII activity (51, 52). In summary, FI kinetics is strongly related with the reduction of quinone acceptors by PSII (see Note 11 and Table 3).

Table 3 Approximative half-times and origin of fluorescence induction phases Phase

Half-time (ms)

Origin

O-J

0.15–0.70

Reduction of QA; accumulation of PSII in QA−QB and QA−QB−

J-I

5–15

Accumulation of PSII in QA−QB2− and QA−QBH2

I-P

40–120

Accumulation of PQH2

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However, the reduction of the PSII acceptor side is dependent on the input of electrons from the donor side of PSII. Consequently, FI is also highly sensitive to changes in the activity of the oxygenevolving complex (54, 55). Thus, FI is an interesting tool to study changes in structure and function of the whole PSII. It can be used with leaves or preparation of chloroplasts, thylakoids, or isolated PSII, but the clear I-peak is lost upon breaking the chloroplast membrane (56, 57). 3.2.4.1. Measuring Fluorescence Induction in Leaves

Fluorescence induction can be measured in leaves prior to isolation of PSII. It is a fast way to assess the integrity of the photosynthetic apparatus in leaves. 1. The duration of the measurement should be determined. It should be long enough to reach Fm, which is within few hundreds of ms in healthy leaves at saturating light intensity. In most cases, 1 or 2 s is enough. 2. The intensity of excitation light should be set to get the highest fluorescence signal, through avoiding saturation of the detector with out-of-range signals. The chosen intensity should allow variation of fluorescence level between samples and will vary with plant species and growth conditions. With the PEA, 40–50% of the maximal intensity is needed that corresponds to a photon flux density of 1,600– 2,000 mmol/m2s. 3. Place the leaf on the leaf clip. The adaxial side of the leaf should be upward (in the direction of the probe). The leaf should be dark-adapted for at least 5 min by closing the leaf clip before measurement (see Note 12). 4. Start the measurement protocol. Healthy mature leaves should have an Fv/Fm ratio higher than 0.78 (the highest is the better). Upload the data on a computer and verify if the FI curve shows a typical O-J-I-P pattern (Fig. 5, trace 1). At least five to ten repetitions with different leaves are needed to get a representative average of a group. It should be noted that the shape of FI can be affected by the age of leaves and the position where fluorescence was measured on leaves.

3.2.4.2. Measuring Fluorescence Induction in PSII Preparations

1. Measurements of FI with liquid photosynthetic samples are done with cylindrical plastic vials supplied by Hansatech. They can hold 4 mL of solution. 2. Put the vial in the sample holder; put Buffer C and then PSII preparation in the vial. Total volume should be 4  mL and chlorophyll concentration should be 25  mg/mL or less. Photosynthetic sample should be homogeneously mixed in the buffer (see Note 13).

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Fig. 5. Typical fluorescence induction traces measured with a leaf (1), thylakoid membranes (2), and PSII membranes (3). Data acquired before 40 ms are not shown because they are artifacts ascribed to the response time of the instrument.

3. Cover the sample holder with the fluorescence head. Darkadaptation period should be the same for all measurements (minimum 1 min). 4. Measure the fluorescence induction curve with maximal excitation light intensity. Fluorescence induction can also be measured in thylakoid membranes. In that case, Buffer C should be replaced by Buffer B, and final Chl concentration can be increased to 50 mg/mL. 3.2.4.3. Interpretation of FI Measurements

The maximal quantum yield of photochemistry measured by the Fv/Fm ratio (Fv = Fm − F0) is the most commonly used fluorescence parameter. It is a good indication of the leaf health and of the PSII activity/integrity. To analyze further the state of PSII, one can plot the fluorescence kinetics on a logarithmic time-scale. As a result, the three phases will become clear if measurement was done with leaves (Fig. 5, trace 1). For isolated thylakoids and PSII, FI will look like a biphasic O-J-P curve (Fig. 5, traces 2 and 3) even though the J-I phase is still present (but is minor for PSII). By comparing two different measurements carried out with the same photosynthetic material, changes in the shape of FI traces can indicate which part of PSII was affected by a treatment. Table 4 shows typical changes that can be observed and their cause.

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Table 4 Basic interpretations of typical changes in fluorescence induction traces Symptoms

Diagnostic

Decrease in J-I-P amplitude

PSII cannot reduce efficiently its PQ acceptors. Donor side is probably slowed

O-J rise is aborted, fluorescence drops after J and J-I-P is strongly diminished

The donor side of PSII is strongly impaired and cannot supply efficiently electrons

O-J amplitude is increased

Electron transfer from QA− to QB may be slowed or a nonreducible molecule may occupy the QB pocket

F0 is decreased

There may be a quencher of Chl excited states in antenna/Chl concentration may be decreased

F0 is increased

A fluorescence quencher was removed (PQ pool may be partly reduced prior to measurement; this would also lead to an increased J-level) or excitation transfer in PSII antennae may be perturbated

I-P rise is slowed and/or diminished

Accumulation of PQH2 is slowed or impaired

4. Notes 1. Phosphatidylcholine in the thylakoid membrane. It has been clearly demonstrated that phosphatidylcholine is one of the components of the outer membrane of the chloroplast (2). However, its presence in the thylakoids is still a matter of controversy (3). Notwithstanding this conjecture, it is interesting to remark that the oxygen-evolving activity of purified PSII particles is enhanced upon their incorporation into liposomes constituted of phosphatidylcholine (4). 2. Methodologies for structure–function studies of chloroplast lipids. Most methodologies available for the study of the chloroplast lipids role in structure–function relations are discussed in (9), which is a repository of data and ideas. So to say, a “bible” for anyone investigating structure–function of the chloroplast lipids. Additions to this knowledge were presented at the 12th International Congress on

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Photosynthesis (10). In short, the major methods used ­presently are the ­following: (1) isolated proteins reconstituted into proteoliposomes, (2) membranes with lipids inactivated by chemical modification or by immunological methods, (3) membranes selectively depleted of lipids by chemical treatment, and (4) mutants with genetically induced changes in lipid composition. Method (1) gives direct information on the lipid–protein interactions and protein folding and misfolding; but, this method cannot always assure whether those interactions take place in vivo, thereby indicating the need for the application of other methods. Methods (2) and (3) are very useful to study the outer surfaces of the membranes; however, they may present serious difficulties partly related to the large dimension of the proteins used to interact with the much smaller surface of the polar head of the chloroplast lipids. Method (4) should facilitate, in principle, the interpretation of the data; however, it is not clear in some cases whether a mutant that is deficient in, e.g., DGDG and displays a concomitant loss of photosynthetic activity, does perform this new function as the result of mutation-induced defects in the “thylakoid membrane” or in the “protein import apparatus” located in the outer membrane of the chloroplast. These considerations indicate the need for the application of combinations of the above-described methods in the study of structure– function relations of chloroplast lipids. 3. Salts purification. Commercially available inorganic salts may be contaminated with organic impurities that usually give rise to changes in the surfaces properties of native biological membranes and artificial lipid bilayer vesicles (liposomes). This important matter is discussed, for example, in (11) (cf. their Fig. 4b). The membrane properties affected by the presence of organic impurities in inorganic salts are, among other possible interferences, the polarity of the lipid–water interface and the evolution of interfacial reaction kinetics (11), and the lipid–protein interactions. The following procedure for purification of the salts is recommended. First, the salts are washed successively with three different organic solvents (methanol, benzene, and chloroform) to remove the organic impurities. Second, the purified salts are dried at 150–160°C for 24 h to remove any traces of adsorbed solvent. Third, the degree of purification of the salts obtained by this procedure is checked by surface tension experiments. The surface tension data of the purified salts dissolved in water (demineralized; distilled) might be in the range of values published in the literature. See, for instance, CRC (ed.) (1980–1981) Handbook of Chemistry and Physics, 61st ed., CRC Press, Boca Raton, p. F-43.

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4. Lipids purification. The routine purification procedure used in (12, 13) takes place in a HPLC (Water Associates, Milford, MA) equipped with two 510 pumps, an automatic gradient controller (model G80), a Rheodyne injector (model 7126) (Cotati, CA), with a 1-mL loop and a programmable ultraviolet-visible detector set at 205  nm. The lipid separation is done on a silica cation-exchange column operated in isocratic mode. All solvents (HPLC grade) used in the mobile phase are filtered through a membrane of 0.2-mm pore size (Millipore, Bedford, MA), and degassed under vacuum prior to their introduction into the HPLC system. The mobile phase is a n-hexane:isopropyl alcohol:water mixture, 70:30:2 (v:v:v). The flow rate is 10 mL/min at 2,400 PSI. The samples are dried in a stream of nitrogen, then dissolved in the elution mixture to give a final concentration of 5  mg/mL, followed by injection via the Rheodyne rotatory injector. The purified lipids are thereupon dissolved in chloroform and kept in a deep-freezer until use. 5. Fatty acid chains analyses. Analyses of the fatty acid chains of the lipids (12, 13) are done by gas chromatography of methyl esters formed from methanolysis of the lipid (0.1 mg) with 1  mL of BF3/MeOH 14% wt/vol (Pierce Chemical Co.). The mixture is heated 15 min at 100°C followed by cooling for approximately 10 min; then n-hexane and bidistilled water are added to separate the fatty acid methyl esters, which are dried by treatment with anhydrous Na2SO4 followed by a current of nitrogen. The residue is dissolved in 50  mL of n-hexane. The fatty acid analysis described in (12, 13) is carried out in a Varian gas chromatograph (model 3700), equipped with a Shimadzu integrator (model C-R3A), using nitrogen as the carrier gas. 6. Other methods for preparation and use of liposomes. The evolution of the liposome methodology has been in constant progress since its early days (see, e.g., (14) and literature cited therein). In fact, from their initial purpose as useful models of biological molecules (15, 16), the liposomes became an indispensable ingredient in the study of lipid–protein interactions (proteoliposomes), and a powerful tool as drug transport vehicles in vivo for therapeutical uses such as the delivery of anticancer agents to malignant tumors (17). In which concerns the purpose of this chapter, one notes that papers are regularly published on (1) the formation of small size liposomes of about ~250 Å (0.025  mm) diameter for new biochemical and physical–chemical studies as the lipid–porphyrin interactions described in (17) and (2) the preparation of large unilamellar vesicles (LUV or “giant liposomes”) of sizes ranging from 10 to 500  mm diameter using such sophisticated methods and equipment as those discussed in (18).

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7. Phase behavior of the thylakoid lipids. The phase behavior of a lipid species in its mixtures with other lipids and water is usually well described by the phase diagram of the lipid. It is noted, in this respect, that a notable account of available phase diagrams and other related questions is given in the excellent review by Marsh (19). Among the many other works that were published on these matters, those by Williams (2) and Garab et  al. (20) are especially interesting since they are directly related to the chloroplast lipids. For practical purposes, there are two main reasons that render useful the consultation of the phase diagram, if available, of the lipid that one intends to use in the preparation of liposomes or proteoliposomes. The first reason is to identify whether the lipid in question is a “bilayerforming” or a “nonbilayer-forming” species; in the latter case, one has to use binary or ternary mixtures of lipids. The second reason is to identify the gel-to-liquid crystal transition temperature of the lipid in order to know whether the formation of liposomes shall take place at room temperature (14, 19). In fact, it is important to emphasize that the preparation of lipid vesicles should always be performed above the phase transition temperature of the lipid species being used. This precaution is usually unnecessary with native lipids, which have in general transition temperatures several degrees below 0°C, but is essential if the vesicles are prepared with a synthetic lipid such as DPPC (dipalmitoyl phosphatidylcholine) whose transition temperature is around 41°C (14), that is, well above the room temperature of most laboratories. Detailed information on the phase behavior and hydration of most phospholipids, e.g., phosaphatidylcholine and phosphatidylglycerol, is seen in Section II of (19), whereas Section III.6 (19) gives information on the glycolipids referred to in the Introduction, i.e., digalactosyl diacylglycerol, monogalactosyl diacylglycerol, and sulfoquinovosyl diacylglycerol. Furthermore, in (14, 19) one finds interesting discussions on the basic physical–chemical relationship between the cis and trans configurations of the carbon atoms in the fatty acid chains and the phase transition temperatures. 8. Sonication in a capped tube. It is worth noting that the sonication step can be also performed, whenever possible, in a special capped tube that can be obtained from Heat SystemsUltrasonics (Plainwiew, LI). The use of the capped tube is interesting as it avoids contamination of the lipid preparations with the titanium particles from most sonicator probes. 9. Estimation of chlorophyll concentration. The chlorophyll concentrations of chloroplasts, thylakoid membranes, or PSII particles are estimated using 80% acetone solutions according to the procedure of Arnon (26). In short, an aliquot of volume Vs

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of the sample under study is added to a given volume of 80% acetone, Va. The mixture is first vortexed and then centrifuged for 5 min in a IEC clinical centrifuge (or another similar model). The supernatant is carefully transferred to test tubes paying particular attention not disturb the centrifugation pellet. Then, the absorbance (A) of the solutions is measured in a spectrophotometer at 645 and 663 nm. Finally, the chlorophyll concentration in mg/mL, [Chl], is calculated using the expression [Ch1]=[(20.2 × A645 + 9.02 × A663) × Va] / Vs where A645 and A663 are the absorbances at 645 nm and 663 nm, respectively. 10. The oxygen electrode. In general, the Hansatech Company (http://www.hansatech-instruments.com) furnishes with their equipment a detailed theoretical and experimental documentation containing useful information for the successful ­measurement of oxygen evolution in various photosynthetic materials. In addition, a copy of the book The Use of the Oxygen Electrode and Fluorescence Probes in Simple Measurements of Photosynthesis by Prof. David A. Walker can be obtained from Hansatech. 11. Origin of O-J-I-P phases. Interpretation of FI is still a matter of debate. In addition to recent papers cited here, readers are invited to consult a review by Lazar (53) for an overview of the various interpretations in the literature. 12. Dark-adaptation of leaves prior to measurement of fluorescence induction. If leaves were light adapted before measurement, plants should be dark adapted for at least 1  h to allow the leaves to reach an homogenous steady state of dark adaptation. If fluorescence induction measurements have to be compared between various treatments, care must be taken to always use the same period of dark adaptation since significant changes in traces happen especially during the first 30 min of dark adaptation. 13. Efficient mixing of liquid samples. The sample holder can be put on a magnetic plate and a small magnetic stirrer can be inserted to the vial to facilitate the mixing of the photosynthetic sample with the buffer. However, the sample should not be stirred during fluorescence induction measurements, since the excitation beam penetrates only in the first centimeter of the cylindrical vial.

Acknowledgments This work was supported in different occasions by grants from the NSERC Canada, the Université du Québec à Trois-Rivières, the Département de chimie-biologie, and the Groupe de recherche en biologie végétale (GRBV).

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infrared spectroscopy. Biochemistry 33, 11650–11655. 37. Krimm, S. and Bandekar, J. (1986) Vibrational spectroscopy and conformation of peptides, polypeptides, and proteins. Adv. Protein Chem. 38, 181–364. 38. Surewicz, W. K. and Mantsch, H. H. (1988) New insight into protein secondary structure from resolution-enhanced infrared spectra. Biochim. Biophys. Acta 952, 115–130. 39. Wilson Jr., E. B., Decius, J. C., and Cross, P. C. (1955) The Theory of Infrared and Raman Vibrational Spectra, McGraw-Hill, New York. 40. Menikh, A. and Fragata, M. (1994) Protein secondary structure of the photosystem II complex studied by FT-IR spectroscopy. Changes in the amide I region mediated by digalactosyl diacylglycerol and divalent cations. J. Mol. Struct. 319, 101–107. 41. Fragata, M., Nénonéné, E. K., Maire, V., and Gabashvili, I. S. (1997) Structure of the phosphatidylglycerol-photosystem II complex studied by FT-IR spectroscopy. Mg(II) effect on the polar head group of phosphatidylglycerol. J. Mol. Struct. 405, 151–158. 42. Heimburg, T. and Marsh, D. (1993) Investigation of secondary and tertiary structural changes of cytochrome c with anionic lipids using amide hydrogen exchange measurements: a FTIR study. Biophys. J. 65, 2408–2417. 43. Lee, D. C., Haris, P. I., Chapman, D., and Mitchell, R. C. (1990) Determination of protein secondary structure using factor analysis of infrared spectra. Biochemistry 29, 9185–9193. 44. Kautsky, H. and Hirsch, A. (1931) Neue versuche zur kohlensäureassimilation. Naturwissenschaften 48, 964. 45. Duysens, L. N. M. and Sweers, H. E. (1963) Mechanism of two photochemical reactions in algae as studied by means of fluorescence, in Studies on Microalgae and Photosynthetic Bacteria (Japanese Society of Plant Physiologists, Ed.), University of Tokyo Press, Tokyo, pp 353–372,. 46. Butler, W. L. (1978) Energy distribution in the photochemical apparatus of photosynthesis. Annu. Rev. Plant Physiol. 29, 345–378. 47. Lavergne, J. and Trissl, H. W. (1995) Theory of fluorescence induction in photosystem II: derivation of analytical expressions in a model including exciton-radical-pair equilibrium and restricted energy transfer between photosynthetic units. Biophys. J. 68, 2474–2492. 48. Neubauer, C. and Schreiber, U. (1987) The polyphasic rise of chlorophyll fluorescence upon onset of strong continuous illumination:

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Chapter 19 Physical and Chemical Immobilization Methods of Photosynthetic Materials Lise Barthelmebs, Robert Carpentier, and Régis Rouillon Abstract Isolated photosynthetic materials have a relatively short active life time that limits their effective use. To circumvent this limitation, various immobilization techniques have been designed to improve their stability both under storage and working conditions. The immobilization methods are identified either as chemical or physical procedures depending on whether covalent bonds are established or not. In this chapter, two immobilization methods frequently used are described: a physical one based on the entrapment of photosynthetic materials in photo-crosslinkable poly(vinylalcohol) polymer bearing styrylpyridinium groups (PVA-SbQ) and a chemical one where the photosynthetic materials are immobilized by coreticulation in an albumin-glutaraldehyde cross-linked matrix (BSA-Glu). Different immobilization procedures in relation with various photosynthetic materials are mentioned. Key words: Immobilization, Stabilization, Photosynthetic, Polyvinylalcohol, Glutaraldehyde, Bovine serum albumin, Chloroplast, Photosystem I, Photosystem II, Thylakoid membrane, Cyanobacteria

1. Introduction In thylakoid membranes, light energy is transduced into chemical energy. At this level, light is absorbed by the photosynthetic ­pigments imbedded in chlorophyll-protein complexes and its energy is transferred to the reaction centers of photosystem I and II, where a charge separation takes place. During this process, water molecules are cleaved and negatively charged species are formed. These properties confer to thylakoid membranes a great potential for various biotechnological applications. However, the isolated photosynthetic materials have a relatively short active life time, which limits their effective use. In order to improve the

Robert Carpentier (ed.), Photosynthesis Research Protocols, Methods in Molecular Biology, vol. 684, DOI 10.1007/978-1-60761-925-3_19, © Springer Science+Business Media, LLC 2011

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Table 1 Main physical and chemical methods used to stabilize the photosynthetic materials Physical

Adsorption

Gel entrapment

Chemical

Reticulation Coreticulation Glutaraldehyde Different proteins

DEAE cellulose Filter paper disk Glass microfiber filter Alumina filter disk Polysaccharide gel   Agar   Alginate   Carrageenan   Agarose Protein gel   Gelatin Synthetic gel   Polyacrylamide   Polyurethane   Photo-crosslinkable resin   Poly(vinylalcohol)   Poly(vinylalcohol) bearing styrylpyridinium groups   Silica        

Glutaraldehyde Gelatin Collagen Bovine serum albumine

stability of photosynthetic materials both under storage and working conditions, a variety of immobilization techniques have been designed (Table  1). They are identified either as chemical or ­physical procedures depending on whether covalent bonds are established or not (1). Physical methods involve the adsorption of the photosynthetic material on supports or the inclusion in a gel. The immobilization by adsorption is a simple and economic technique, which does not damage the activity of the biological material, However, the interaction forces are weak, hence desorption of the biocatalyst may be observed. The immobilization by inclusion is preferentially applied to the immobilization of photosynthetic material. In this technique, the biocatalyst is entrapped into a three-dimensional network of the gel that could be natural or synthetic. Covalent techniques, widely used for enzyme immobilization, are not suitable for photosynthetic material because of the denaturing effect of the binding agent and the consequent loss of

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activity. Nevertheless, as glutaraldehyde has been shown to preserve the Hill reaction activity in chloroplasts, this cross-linker is used for the immobilization of photosynthetic material. The denaturing effect of glutaraldehyde can be reduced by the addition of proteins during the polymerization. In this chapter, two immobilization methods frequently used are described: a physical one based on the entrapment of photosynthetic materials in photo-crosslinkable poly(vinylalcohol) polymer bearing styrylpyridinium groups (PVA-SbQ) (2–7) and a chemical one where the photosynthetic materials are immobilized by coreticulation in an albumin-glutaraldehyde cross-linked matrix (BSA-Glu) (6, 8–15). Different immobilization procedures are mentioned. In the PVA-SbQ method (Fig. 1), the biological material is stabilized without chemical bonding but is confined into a network established by the polymerized PVA-SbQ. In the BSA-Glu method, the glutaraldehyde builds a network of covalent bondings with the free NH2 groups of photosynthetic membrane proteins and albumin. The role of albumin is mainly to protect the photosynthetic material against too many bondings so that an adequate biological function is retained.

CH2–CH

—CH2 –CH–CH2 –CH O

O

l

OH m

CH2–CH O

n

C=O

CH

CH3

DP = 2l + m +n

CH

l = DP

CH

SbQ content

100 m = DP(100 – DS)/100

n = DP (DS – 2. SbQ content)/100 N+ CH3

X-

or

-(Ch2)2SO3 - : betaine form

Fig. 1. Structure of PVA-SbQ.

DP : Degree of polymerization DS : Degree of saponification

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In comparison to other immobilization methods, the use of PVA-SbQ or BSA-Glu allows a better preservation of the photosynthetic activity. After storage of 90 days at 4°C in the dark and in dry state, a mixture of chloroplasts and thylakoid membranes entrapped in PVA-SbQ polymers retains 20% of the initial activity (measurements of oxygen production). The same mixture stored at −18°C shows 20% activity after a 427-day storage (2). After immobilization in BSA-Glu, thylakoid membranes maintained in the dark at 4°C remain stable about 200  h. After 1,000-h storage, the photosynthetic activity was near 40% of initial activity (8). Various immobilized materials were often used in phytotoxicity tests and also as biological receptor in biosensors (16), in order to detect chemicals which interact with the process of photosynthesis. Despite the largely differing mode of immobilization, the procedures led to strikingly similar detection capabilities for the herbicides tested (6).

2. Materials 2.1. Immobilization Procedures in PVA-SbQ

1. Immobilization buffer: 25  mM MOPS-NaOH, pH 7.9, containing 300 mM mannitol, 2 mM ethylenediamine tetraacetic acid (EDTA), 1% (w/v) bovine serum albumin (BSA) for mixtures of chloroplasts and thylakoid membranes or straight chloroplasts or thylakoid membranes (see Note 1); or 20  mM MES-NaOH, pH 6.2, for photosystem II submembrane fractions; or 20 mM sodium phosphate solution, pH 7.0 containing 0.15 mM NaCl and 1 mM MgCl2, for cyanobacteria. 2. 11% (w/v) poly(vinylalcohol) bearing styrylpyridinium groups solution (PVA-SbQ) (see Notes 2 and 3). 3. Petri dishes. 4. Microtiter plates.

2.2. Immobilization Procedure in BSA-Glu 2.2.1. Preparation of Samples in Petri Dishes or Test Tubes

1. Immobilization buffer: 50 mM phosphate buffer, pH 7.1, for thylakoid membranes; or 20  mM Mes-NaOH, pH 6.5, for photosystem II submembrane fractions; or 20 mM TricineNaOH, pH 7.8 containing 10 mM KCl, 10 mM NaCl, and 5 mM MgCl2, for photosystem I submembrane fractions. 2. 20% (w/v) BSA solution. 3. 1.5% (v/v) glutaraldehyde solution. 4. Petri dishes. 5. Test tubes.

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1. Immobilization buffer: 15  mM MES-NaOH, pH 6.5, ­containing 0.5 M mannitol, 0.1 M NaCl, 5 mM MgCl2, and 0.05 mM chloramphenicol. 2. 10% (w/v) BSA solution in immobilization buffer. 3. 10% (v/v) glutaraldehyde solution. 4. Screen-printed electrodes.

2.2.3. Preparation of Samples in Microtiter Plates Wells

1. Immobilization buffer: 25 mM MOPS-NaOH, pH 7.9 containing 300 mM mannitol, 2 mM ethylenediamine tetraacetic acid (EDTA), and 1% (w/v) bovine serum albumin. 2. 20% (w/v) BSA solution. 3. 1.5% (v/v) glutaraldehyde solution. 4. Microtiter plates.

3. Methods 3.1. Immobilization Procedures in PVA-SbQ 3.1.1. Preparation of a Thin Film (Fig. 2)

This procedure was used to immobilize spinach mixtures of chloroplasts and thylakoid membranes (2), straight chloroplasts (2), spinach thylakoid membranes (3), spinach photosystem II submembrane fractions (4) ,or whole cells of cyanobacterium Synechococcus sp. PCC 7942 (5). 1. Weigh 3 g of PVA-SbQ solution. 2. Add 500 mL of photosynthetic material diluted in the immobilization buffer to a chlorophyll concentration of 3 mg/mL for mixtures of chloroplasts and thylakoid membranes (2), straight chloroplasts (2), thylakoid membranes (3), or photosystem II submembrane fractions (4), and 0.8  mg/mL for whole cells of cyanobacteria (5) (see Notes 4 and 5). 3. Homogenize with a spatula (see Note 6), and centrifuge the mixture for 5 min at 200 × g (see Note 7). 4. Spread in a mold to obtain a thin film (diameter: 3 cm. Mixture weigh: 0.8 g), then homogenize the thin film on a mechanical shaker for 5 min at room temperature in the dark. 5. Polymerize under UV light (235 and 365 nm) for 2 min (see Note 8) and dry for 24 h at 4°C in the dark. 6. Store the thin film at 4°C or −18°C in a dry state in the dark in Petri dishes. 7. Cut samples in the thin film with a hollow punch (see Note 9).

3.1.2. Preparation of Samples in Petri Dishes

This procedure was used to immobilize spinach thylakoid membranes (6). 1. Weigh 3 g of PVA-SbQ solution.

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mixture of PVA-SbQ (3 g) + biological material* (500 ml)

thin film diameter: 3 cm mixture weight: 0.8 g

polymerization: 2 min. UV: 235 and 365 nm

drying for 24h at 4°C in the dark

storage at 4°C or -18°C in a dry state and in the dark

sample** to detect photosynthetic activity

0.126 cm2

*Chlorophyll concentration: 3 mg/mL (spinach material) or 0.8 mg/mL **Chlorophyll content: 6mg (spinach material) or 1.6mg (cyanobacteria)

Fig. 2. Preparation of a thin film with photosynthetic materials immobilized in PVA-SbQ.

2. Add 175  mL of thylakoid membranes preparation (initial ­concentration of 2 mg chlorophyll/mL) to 375 mL of immobilization buffer. 3. Mix the PVA-SbQ solution and the photosynthetic preparation. 4. Homogenize with a spatula (see Note 6), and centrifuge the mixture for 5 min at 200 × g (see Note 7). 5. Deposit the immobilization mixture (60 mg samples) in Petri dishes (see Note 10). 6. Polymerize under UV light (235 and 365 nm) for 2 min (see Note 8), and dry for 24 h at 4°C in the dark. 7. Store the samples at 4°C or −18°C in a dry state in the dark in Petri dishes. 3.1.3. Preparation of Samples in Microtiter Plates Wells

This procedure was used to immobilize spinach thylakoid membranes (7). 1. Weigh 1.5 g of PVA-SbQ solution. 2. Dilute the PVA-SbQ solution (1:4) with immobilization buffer. 3. Mix with 250 mL of thylakoid membranes preparation (initial concentration of 2 mg chlorophyll/mL).

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4. Homogenize with a spatula (see Note 6), and centrifuge the mixture for 5 min at 200 × g (see Note 7). 5. Place 150 mL of this mixture in a microtiter plate well, and dry the plate during 24 h at 4°C in the dark until 30 mL of solution per well is left. 6. Polymerize the resulting mixture under UV light (235 and 365 nm) for 2 min (see Note 8), and incubate at −18°C for 2 days. 7. Store the microtiter plate at 4°C or −18°C in a dry state in the dark (see Note 11). 3.2. Immobilization Procedure in BSA-Glu 3.2.1. Preparation of Samples in Petri Dishes or Test Tubes

This procedure was used to immobilize spinach thylakoid membranes (6, 8, 9), spinach photosystem II submembrane fractions (10), or spinach photosystem I submembrane fractions (11). 1. Mix 1.65 mL of immobilization buffer specific for the preparation of photosynthetic material used with 1.25 mL of 20% BSA and 1 mL of 1.5% glutaraldehyde (see Note 12). 2. Incubate for 2 min at room temperature. 3. Add 0.6 mL of photosynthetic material diluted to 3.3 mg chlorophyll/mL (8, 11) or 2 mg chlorophyll/mL (6, 9, 10) (see Note 13) and mix for 3–4 s using a vortex mixer. 4. Distribute the preparation into 80 mL samples in appropriate vessels, such as Petri dishes or test tubes, to obtain the desired shape and volume of immobilized material (see Note 14). 5. Store for 2 h at −20°C. 6. Thaw the immobilized preparations at 4°C in the dark for at least 2 h before use (see Note 15). 7. Cover the sample with the resuspension buffer to prevent them from drying. 8. Wash with distilled water before use (see Note 16).

3.2.2. Immobilization on the Surface of Screen-Printed Electrodes

This procedure was used to immobilize photosystem II submembrane fractions prepared from the thermophilic cyanobacterium Synechococcus elongatus (12). 1. Mix 970  mL of 10% BSA in immobilization buffer with 970 mL of photosystem II submembrane fractions diluted to 200 mg chlorophyll/mL. 2. Add 60 mL of 10% glutaraldehyde and mix for 3–4 s using a pipette tip. 3. Place 5  mL on the surface of the working electrode (see Note 17), and leave the electrode in the dark at 4°C until the solidification of the mixture. 4. Use the electrode or store it at −20°C before use.

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3.2.3. Preparation of Samples in Microtiter Plates Wells

This procedure was used to immobilize spinach thylakoid membranes (7) 1. Mix 1.25 mL of 20% BSA with 120 mL of 1.5% glutaraldehyde and 1.65 mL of immobilization buffer. 2. Add 200 mL of thylakoid preparation (initial concentration of 2  mg chlorophyll/mL), and mix for 3–4  s using a vortex mixer. 3. Place 50  mL of this mixture in a microtiter plate well, and store at −18°C for 2 days. 4. Stored at 4°C before use (see Note 11).

3.3. Determination of Chlorophyll Concentration

1. Add 10  mL of photosynthetic membranes preparation to 5 mL acetone 80% in a conical tube, and mix carefully using a vortex mixer (see Note 18). 2. Centrifuge in a bench-top centrifuge for a few minutes to remove precipitated proteins. 3. Verify the exact volume (5  mL) and adjust if necessary to compensate the evaporation. 4. Measure the absorbance at 663 and 645 nm. 5. Taking the dilution of the membranes in the acetone solution into account, the chlorophyll concentration (mg/mL) in the  membrane preparation is calculated from the following equation: 0.5 (22.22 (A645) + 9.05 (A663)), in which A645 and A663 represent the absorbance values at the respective wavelengths.

4. Notes 1. A 1% BSA concentration in the immmobilization buffer increases the stability of the thylakoid membranes (2). 2. The PVA-SbQ polymers are gifts from Toyo Gosei Kogyo Co, LTD Japan. 3. Different PVA-SbQ polymers with different degrees of polymerization can be used. With thylakoid membranes, the residual photosynthetic activity and the storage stability increase with the degree of polymerization (2). 4. Chlorophyll concentration was determined according to Arnon (17) or Porra et al. (18). 5. Other initial chlorophyll concentrations were also used: 1 or 2 mg/mL. 6. The PVA-SbQ solutions are viscous and cannot be homogenized with a vortex mixer.

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7. This operation allows eliminating bubbles. The ­centrifugation speed must be low to avoid sedimentation. 8. An UV lamp to read chromatographic plates is sufficient. 9. This method allows the evaluation the chlorophyll concentration, e.g., a 0.126 cm2 sample contains 6 mg of chlorophyll (spinach material) with an initial concentration of 3 mg chlorophyll/mL or 1.6 mg (cyanobacteria) with an initial concentration of 0.8 mg chlorophyll/mL . 10. The immobilized samples used for testing contain 20  mg chlorophyll. 11. It is possible to measure the activity of photosynthetic thylakoid membranes immobilized in microtiter plate wells using colorimetric methods. 12. Several immobilization buffers were tested for each type of photosynthetic membranes and the buffer used here provides the best retention of native photosynthetic activity and storage stability. 13. The chlorophyll concentration in the immobilization medium is important to obtain the concentration/albumin-glutaraldehyde ratio that provides the best preservation of the photosynthetic membrane integrity together with optimal immobilization. 14. The final immobilized material presents a soft sponge-like green tissue structure. It is important to choose the appropriate vessel for immobilization since it can be used to mold the immobilized material to any shape or volume required. 15. Best activities are obtained after at least 12 h. 16. For some applications, the immobilized material may be crushed into small particles using a mortar and pestle (10). 17. This volume is used for a big electrode printed in PVC. With a smaller ceramic electrode, 0.5 or 1  mL are placed on the surface of the working electrode. 18. The method of Arnon (17) was initially used but is now modified according to Porra et al. (18). The sample of photosynthetic membranes should be added to the acetone while mixing. This is necessary to minimize the amount of chlorophyll that remains bound to the precipitated proteins.

Acknowledgments Part of this work was supported by the European Community within the EC contract QLK3-CT-2002-01629.

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References 1. Papageorgiou, G.C. (1987) Immobilized photosynthetic microorganisms. Photosynthetica 21, 367–383. 2. Rouillon, R., Mestres, J.J., and Marty, J.L. (1995) Entrapment of chloroplasts and thylakoids in polyvinylalcohol-SbQ. Optimization of membrane preparation and storage conditions. Anal. Chim. Acta 311, 437–442. 3. Rouillon, R., Sole, M., Carpentier, R., and Marty, J.L. (1995) Immobilization of thylakoids in polyvinylalcohol for the detection of herbicides. Sensor. Actuat. B 26–27, 477–479. 4. Rouillon, R., Boucher, N., Gingras, Y., and Carpentier, R. (2000) Potential for the use of photosystem II submembrane fractions immobilised in poly(vinylalcohol) to detect heavy metals in solution or in sewage sludges. J. Chem. Technol. Biot. 75, 1003–1007. 5. Rouillon, R., Tocabens, M., and Carpentier, R. (1999) A photoelectrochemical cell for detecting pollutant-induced effects on the activity of immobilized cyanobacterium Synechococcus sp. PCC 7942. Enzyme Microb. Technol. 25, 230–235. 6. Laberge, D., Rouillon, R., and Carpentier, R. (2000) Comparative study of thylakoid membranes sensitivity for herbicide detection after physical or chemical immobilization. Enzyme Microb. Technol. 26, 332–336. 7. Piletskaya, E.V., Piletsky, S.A., Sergeyeva, T.A., El’skaya, A.V., Sozinov, A.A., Marty, J.L., and Rouillon, R. (1999) Thylakoid membranesbased test-system for detecting of trace quantities of the photosynthesis-inhibiting herbicides in drinking water. Anal. Chim. Acta 391, 1–7. 8. Loranger, C. and Carpentier, R. (1994) A fast bioassay for phytotoxicity measurements using immobilized photosynthetic membranes. Biotechnol. Bioeng. 44, 178–183. 9. Laberge, D., Chartrand, J., Rouillon, R., and Carpentier, R. (1999) In vitro phytotoxicity screening test using immobilized spinach thylakoids. Environ. Toxicol. Chem. 18, 2852–2858. 10. Carpentier, R. and Lemieux, S. (1987) Immobilization of a photosystem II submem-

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brane fraction in a glutaraldehyde crosslinked matrix. Appl. Biochem. Biotechnol. 15, 107–117. Bonenfant, D. and Carpentier, R. (1990) Stabilization of the structure and functions of a photosystem I submembrane fraction by immobilization in an albumin glutaraldehyde matrix. Appl. Biochem. Biotechnol. 26, 59–71. Koblizek, M., Maly, J., Masojidek, J., Komenda, J., Kucera, T., Giardi, M.T., Mattoo, A.K., and Pilloton, R. (2002) A biosensor for the detection of triazine and phenylurea herbicides designed using photosystem II coupled to a screen-printed electrode. Biotechnol. Bioeng. 78, 110–116. Euzet, P., Giardi, M.-T., and Rouillon, R. (2005) A crosslinked matrix of thylakoids coupled to the the fluorescence transducer in order to detect herbicides. Anal. Chim. Acta 539, 263–269. Breton, P., Euzet, P., Piletsky S.A., Giardi, M.-T., and Rouillon, R. (2006) Integration of photosynthetic biosensor with molecularly imprinted polymer-based solid phase extraction cartridge. Anal. Chim. Acta 569, 50–57. Touloupakis, E., Giannoudi, L., Piletsky, S.A., Guzzella, L., Pozzoni, F., and Giardi, M.-T. (2005) A multi-biosensor based on immobilized Photosystem II on screenprinted electrodes for the detection of herbicides in river water. Biosens. Bioelectron. 20, 1984–1992. Campas, M., Carpentier, R., and Rouillon, R. (2008) Plant tissue-and photosynthesis-based biosensors. Biotechnol. Adv. 26, 370–378. Arnon, D.I. (1949) Copper enzymes in isolated chloroplasts . Polyphenoloxydase in B. vulgaris. Plant Physiol. 24, 1–15. Porra, R.J., Thompson, W.A., and Kriedemann, P.E. (1989) Determination of accurate extinction coefficients and simultaneous equations for assaying chlorophylls a and b extracted with four different solvents: verification of the concentration of chlorophyll standards by atomic absorption spectroscopy. Biochim. Biophys. Acta 975, 384–394.

Chapter 20 Identifying Chloroplast Biogenesis and Signalling Mutants in Arabidopsis thaliana Verónica Albrecht, Gonzalo M. Estavillo, Abby J. Cuttriss, and Barry J. Pogson Abstract The chloroplast is the largest and arguably the most complex of the three energy organelles in the plant cell. The biogenesis of the chloroplast requires a combination of thousands of proteins encoded by the chloroplastic and nuclear genomes. Chloroplast function is also subject to modifications to enable responses to changes in environmental and developmental stimuli. As a consequence, interorganelle signalling and coordination between the chloroplast and nucleus is critical for the biogenesis and function of the chloroplast. Coordination and signalling during biogenesis is referred to as biogenic control and during the function as operational control (1). In this article, we report on two different mutant screens as examples of strategies for identifying mutations that affect biogenic and operational control signalling pathways and processes. We also describe strategies for the analysis and genotyping of the mutants. Key words: Photoprotection, Arabidopsis, Ascorbate peroxidase, Luciferase, Chloroplast biogenesis, Mapping, Positional cloning, Oxidative stress

1. Introduction The survival of plants and consequently, most of life on earth, is ultimately dependent on the chloroplast. Upon illumination germinating seedlings start to undergo a series of morphological and physiological changes among which the greening and thus the formation of chloroplast is the most dramatic and required for subsequent photosynthetic reactions. Light energy is converted to chemical energy through the process of photosynthesis. However, the exposure of a plant to light exceeding that which can be utilised in photochemistry leads to inactivation of photosynthetic functions (photoinhibition) and oxidative stress due to

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the production of reactive oxygen species (ROS), such as H2O2, superoxide (O2−), hydroxyl radicals, and singlet oxygen (1O2) (2). ROS are extremely detrimental to plant survival as they can induce the oxidation of lipids, proteins, and enzymes necessary for the proper functioning of the chloroplast and cell as a whole (3). It is therefore of great interest to understand the processes a plant utilises to assemble its chloroplast to limit damage during biogenesis, and how it protects itself from excess light during operation in changing environments, including the imposition of excess light and/or drought stress. In order to investigate both regulatory pathways of chloroplast biogenesis in seedlings as well as of pathways involved in the induction of the oxidative stress indicator, ASCORBATE PEROXIDASE2 (APX2), we have designed two different screens to identify potential chloroplast biogenesis or photoprotective mutants, respectively. The power of a genetic screen is to identify novel genes; the weakness is that an inadequately designed screen will identify unrelated genes. Thus, in each instance, consideration needs to be given to the biology of the process being investigated. The potential complication of a mutant screen for chloroplast ­biogenesis is distinguishing between mutants that impair function, e.g. components of the photosynthetic apparatus and ­mutations required for biogenesis. Consequently, a screen for chloroplast biogenesis mutants was based on identifying mutants with pale green cotyledons and relatively wild type true leaves. This strategy targets genes specific to biogenesis of the chloroplast from its progenitors, the proplastid and etioplast during seedling development. Screens for mutations that perturb photoprotection include, but are not limited to the altered induction of nonphotochemical quenching (NPQ) (4), increased chlorophyll fluorescence (5), aberrant carotenoid accumulation (6), and delayed greening (7). With respect to identifying genes and proteins required for retrograde signalling from the chloroplast to the nucleus in response to excess light and drought, the strategy was to focus on a direct assay, namely, changes in the transcription of a nuclear-encoded gene that is induced by oxidative stress in the chloroplast. Under normal growth conditions, plants have evolved several mechanisms to protect themselves against the adverse effect of ROS formed during cellular metabolism (8), including photoprotective compounds, such as carotenoid. Another line of defence consists of enzymes, such as SOD (superoxide dismutase) and APX (ascorbate peroxidase), that can dismutate O2− radicals and convert H2O2 to water, respectively (reviewed in (2)). Some members of the ascorbate peroxidase gene family, APX1 and APX2, are known to be transcriptionally regulated. They are induced by high light, increased H2O2 levels and are regulated by the redox state of the PQ pool (9). The screen for aberrant APX2 expression was

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performed by monitoring altered luminescence of the reporter gene, luciferase, under the control of the APX2 promoter, APX2::LUCIFERASE. This has the advantage of consecutive screening for three classes of mutants: plants constitutively expressing APX2; plants with an altered APX2 response after exposure to high light; and plants in which APX2 exhibited an increased expression 24 h after exposure to 1 h of high light.

2. Materials 1. Plants: (a) For chloroplast biogenesis mutant screen: EMSmutagenized seeds for the screening and wild type Arabidopsis thaliana ecotype Landsberg erecta (Ler) for backcrossing and Columbia (Col) for mapping. (b) For signalling mutants screen: as in (a), but here the mutagenised seeds containing the luciferase reporter gene APX2 promoter construct are in Col ecotype. Thus, for creating the mapping population transgenic Ler harbouring the APX2 reporter gene fusion are needed. Otherwise, a different trait in the mutant is required for screening the mapping population, such as variegation (alx13) or altered leaf shape (alx8) (10, 11). 2. Seeds sterilising solution: 25% (v/v) bleach, 0.05% (v/v) Tween 20. 3. 70% ethanol (v/v). 4. MS agar 1% (w/v) plates without sucrose. 5. Ethyl methanolsulfonate (EMS). 6. MS media: 1× MS salts (4.3  g/L), sucrose 0.2% (w/v), vitamins 1×, 1% (w/v) phytagar (12). 7. d-Luciferin firefly, potassium salt (synth.). Make 0.1  M aliquots and store in –80°C. 8. Plates: Opaque 96-well cell culture plates; black is preferable, white can be used. 9. Luminescence counter such as TopCount NXT microplate scintillation and luminescence counter and/or cooled CCD camera and dark chamber or equivalent. 10. Agarose 1000. 11. Molecular markers: such as the Arabidopsis MapPairs microsatellite marker set from Research Genetics, or designing markers around the Cereon polymorphism database (http://www. arabidopsis.org/Cereon/) and ordering primer sets individually.

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12. Standard PCR reagents for mapping. 13. Shorty buffer: 0.2 M Tris–HCl, pH9, 0.4 M LiCl, 25  mM EDTA, 1% (w/v) SDS. 14. Eppendorf tubes.

3. Methods (Fig. 1) 3.1. Mutagenesis 3.1.1. Generate or Purchase Mutant Populations

3.2. Mutant Screens

If you wish to saturate the genome with mutations, you need to consider both the number of mutations per seed and the total size of the population. Other works have used seeds ranging between 5,000 and 125,000 (M1) seeds for EMS-mutagenised populations (13). Assuming about 100 mutations per seed, then 5,000 is typically sufficient. M1 populations can also be purchased or generated “in house” as described (14) (see Note 1). Ecotype selection is important if you anticipate a positional cloning approach to gene identification, as molecular markers are based on sequence differences between ecotypes. Previously, Col and Ler ecotypes were preferable because both have been sequenced, giving rise to a plethora of molecular markers available for this combination (15), however, with the sequencing of more than a dozen Arabidopsis ecotypes at the time of writing and up to 1,001 planned (see http://1001genomes.org), this is now less of an issue (16). A crucial step in identifying desired mutants is the design of the initial screen. Avoid multiple variables and labour intensive screens. Design the screen to be semi-automated or visual.

Fig. 1. Strategy for screening and identifying mutants.

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The screens described here utilise both ways. For the biogenic screen, a visual screen was applied on an EMS-mutagenised population of Ler looking for pale cotyledon mutants on MS media plates. For the operational control screen, a semi-automated phenotypic screen was used for the EMS-mutagenised population of an APX2-luciferase reporter line using the level of light emission due to enzyme activity of phenotyping. 3.2.1. Mutant Screens 3.2.1.1. Chloroplast Biogenic Control Mutants

1. Sterilise seeds: 3 min in the sterilising solution, 5 min in 70% (v/v) ethanol, wash four times in sterile water, and plate on MS plates containing 1% (w/v) agar without sucrose. (many chloroplast mutants can overcome their pale phenotype on sucrose) (see Note 2). 2. The identification of the chloroplast biogenesis mutants can be visually performed. Transfer 7-day-old seedlings exhibiting pale cotyledons on a new MS plate and check if newly appearing true leaves are pale, pale green, or wild type green. The mutants need to be transferred to a new plate to avoid being overgrown by the better growing wild type-like seedlings. 3. Grow selected plants and harvest the seeds from self-fertilisation for checking in the next generation.

3.2.1.2. Chloroplast Operational Control Mutants

1. Sterilise seeds: 3 min in the sterilising solution, 5 min in 70% (v/v) ethanol, wash four times in sterile water, and plate on MS plates containing 1% (w/v) agar without sucrose. 2. When emerging roots are about 5 mm long (approx. 4 days), carefully transfer individual seedlings to 96-well plate containing 200  ml of 0.8% (w/v) MS agar with 50–100 mM luciferin per well using a small spatula (see Note 2). Let the plantlets adapt for up to 1 week so luciferin is evenly taken up throughout the plants (see Note 3). In each 96-well plate also include one non-transgenic plant and one non-mutagenised APX2-luciferase transgenic plant as controls. 3. The activity of the enzyme luciferase is measured in the signalling mutants after 1 week comparing the luminescence generated by luciferase activity using the TopCount micro­ plate scintillation and luminescence counter (see Note 4). Measure at 0  h, after 1  h high-light stress and 24  h after return to moderate light. Gain of function mutations could either have increased constitutive (basal) luminescence or a higher (or more rapid) level of expression under high-light stress due to altered induction of the APX2-luciferase transgene. Loss of function mutations will either reflect intragenic lesions in the APX2-luciferase transgene or, more interestingly, second-site mutations that reduced expression of the transgene.

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4. Normalisation and background measurement of the plate and the seedlings must be performed. For this, the TopCount plate reader will first need to be normalised with a normalisation plate provided by the manufacturer, correcting background and cross-well emission. Depending on the concentration of luciferin, determine how long to count each well (between 5 and 20  s) (see Note 5). Count each plate twice and use average values of each well. Determine an arbitrary cut-off value based on the control luminescence. We chose a cut-off value of a greater than twofold induction or repression of APX2 as compared to non-mutagenised control plants. 5. Grow selected plants and harvest the seeds from self-fertilisation for checking in the next generation. 3.2.1.3. Screen for Different Type of APX2 Mutants

Different type of mutants with altered expression of APX2 could be screened. For example, highly induced APX2 phenotypes could be detected by illuminating plates with high light (1,000 mmol/m2 s) for 1 h and recount each plate using the same parameters. This screen will identify the loss of function mutants or mutants with extremely high induction of APX2. Alternatively, mutants with constitutively higher APX2 expression could be screened by transferring plates back to the growth cabinet (80 mmol/m2 s) and recounting luminescence after 15 h.

3.2.2. Re-screening Putative Mutants

Seeds from the self-putative mutant plants were sterilised and plated out as described previously. The screen has to be repeated for several seedlings of the same mother plant in the same way as the mutants have been isolated.

3.3. Allelism

If more than one line displays a similar phenotype, check for allelism. Perform reciprocal crosses (17) between homozygous mutants and analyse the F1 progeny for the mutant phenotype. Likewise, test for allelism against similar known phenotypes (see Note 6).

3.4. Establish Isogenic Line

Backcross mutants to the parental ecotype and self the resulting F1 progeny. A total of four backcrosses will remove about 95% of the superfluous mutations, leaving a near isogenic line. Saturation mutagenesis generally implies that there will be numerous mutations per line and each backcross will theoretically exclude half of these spurious mutations in the resulting progeny, which will be homozygous for the altered luminescence phenotype. A segregation analysis during these backcrosses can be informative in the case of a complicated phenotype to determine if the phenotype is in fact pleiotropic, or due to other mutations in different genes (e.g. dominant, recessive, or semi-dominant).

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3.5. Create Mapping Population

Cross the homozygous mutant with alternative ecotype, preferably Ler (assuming mutant is in a Col background) (17). Note that it would be best to use Ler plants harbouring the APX2 reporter gene fusion for the signalling mutants. Self-fertilise the resulting F1 progeny to produce a F2 mapping population for use in positional cloning. F2 plants are selected for mapping based on their phenotype, namely, those that are homozygous for the mutant phenotype and thus homozygous at that locus. The selection of homozygous F2 mutant lines requires a clear and defined mutant phenotype; otherwise, the selection of mutant plants as well as the subsequent mapping will be difficult. Aim for about 150 mutant plants in the first round of coarse mapping (15). In case of unclear mutant phenotype or dominant mutations, the selected plants need to be selfed and analysed in the F3 generation for being homozygous for the selected phenotype.

3.6. Positional Cloning

Having identified mutants, the next step is to find the disrupted gene responsible for the altered phenotype. Currently, several approaches are possible: map-based cloning using known single nucleotide polymorphisms (SNPs) and cleavage amplified polymorphisms (CAPs) markers between Ler and Col ecotype, insertion and deletion (indel) arrays with 70-mer oligonucleotide elements representing indels between the two ecotypes (18), or the newly arising technologies associated with deep-sequencing or re-sequencing of the mutant genome (19). Here, we describe the original method of map-based cloning.

3.6.1. Markers

Select markers that are evenly spread across the genome. A good start is ordering a kit (see Subheading 2, step 11) Alternatively, use the modified set of markers listed below, which was compiled by Lukowitz and colleagues (20) (see Note 7 and Table 1). In Arabidopsis, the most convenient and commonly used molecular marker types are based on microsatellites and SNPs. Microsatellite markers rely on differences between the two ecotypes in short repetitive sequences (also called SSLPs or simple sequence length polymorphisms (21)), which can be differentiated by PCR amplification of the microsatellite followed by size fractionation on agarose or acrylamide gels. SNPs can be utilised for molecular marker design in a number of different ways. If the base change between ecotypes results in the loss or gain of a restriction site, then differences can be utilised by way of a CAPs marker (22). If a restriction site can be created around a SNP by primer design, then the marker is called a derived-cleaved amplified polymorphism (dCAP) (23, 24). Both of these marker types require PCR amplification of the SNP region, followed by a restriction digest to differentiate between different ecotypes and are visualised on an agarose gel. There are other rapid strategies that have been developed by a number of companies and researchers,

III

(MDC16)

(MFE16)

(F18B3)

(T17J13)

  43

  70

  86

(T7F6)

  73

  20

(F13B15)

  50

(F28P22)

113

(T26I20)

(F14J16)

  81

  30

(F14J22)

  72

(T18C20)

(F14M2)

~50

  11

(T22C5)

  39

II

(F21M12)

  10

I

BAC

cM

Chr.

nga 6

ciw 4

ciw 11

nga 162

nga 168

F13B15

ciw 3

ciw 2

Nga 111

nga 280

ciw 1

F14M2

ciw 12

F21M12

Marker

TGGATTTCTTCCTCTCTTCAC

GTTCATTAAACTTGCGTGTGT

CCCCGAGTTGAGGTATT

CATGCAATTTGCATCTGAGG

TCGTCTACTGCACTGCCG

CCGCATCCTTTGCGTTTA

GAAACTCAATGAAATCCACTT

CCCAAAAGTTAATTATACTGT

CTCCAGTTGGAAGCTAAAGGG

CTGATCTCACGGACAATAGTGC

ACATTTTCTCAATCCTTACTC

GGCAAGAAATTGCGTTGTG

AGGTTTTATTGCTTTTCACA

GGCTTTCTCGAAATCTGTCC

Forward primer

Table 1 Microsatellite markers modified after Lukowitz and colleagues (20)

ATGGAGAAGCTTACACTGATC

TACGGTCAGATTGAGTGATTC

GAAGAAATTCCTAAAGCATTC

CTCTGTCACTCTTTTCCTCTGG

GAGGACATGTATAGGAGCCTCG

GCATTGAGGTCCAGAGGAAA

TGAACTTGTTGTGAGCTTTGA

CCGGGTTAATAATAAATGT

TGTTTTTTAGGACAAATGGCG

GGCTCCATAAAAAGTGCACC

GAGAGCTTCTTTATTTGTGAT

CCCGGTCGGAGCTATCTT

CTTTCAAAAGCACATCACA

TTACTTTTTGCCTCTTGTCATTG

Reverse primer

143/123

190/~215

179/~230

107/89

151/135

314/339

230/~200

105/~90

128/162

105/85

159/~135

332/308

128/~115

200/~160

Product (Col/Ler )

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V

IV

(MQJ16)

(MIK22)

(MFO20)

(MAE1)

  71

  88

115

(T9A14)

104

  42

(FCA2)

  70

(MED24)

(T6G15)

  47

  10

(T15B16)

  10

MAE1

ciw 9

PHYC

ciw 8

MED24

nga 1107

FCA2

ciw 6

ciw 5

TTATTTTTCCACAAGTTCTTTTCTT

CAGACGTATCAAATGACAAATG

CTCAGAGAATTCCCAGAAAAATCT

TAGTGAAACCTTTCTCAGAT

TGCCTCCTTGGGAAAGTG

GCGAAAAAACAAAAAAATCCA

AGCTGCATCTGGARCRACGG

CTCGTAGTGCACTTTCATCA

GGTTAAAAATTAGGGTTACGA

ACTTGGGTCAGAGGACAAAC

GACTACTGCTCAAACTATTCTTCGG

AAACTCGAGAGTTTTGTCTAGATC

TTATGTTTTCTTCAATCAGTT

GGCCCAAGCACACCTACA

CGACGAATCGACAGAATTAGG

AAGCTCCCTCGGTCTACCAT

CACATGGTTAGGGAAACAATA

AGATTTACGTGGAAGCAAT

205/163

165/~145

207/222

100/~135

447/408

150/140

240/204

162/~148

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such as using high resolution melt during real time PCR, to detect the SNPs (25). An advantage of PCR-based markers is that they identify the genotype of both chromosomes. For a particular marker locus, one can visualise a single “Col-size” fragment, a “Ler-size” fragment, or in the case of a heterozygote, both fragments. In the case that the original mutant was identified in a Col background and crossed with a Ler wild type plant, and assuming the mutation is recessive, both chromosomes in 100% of F2 plants with the mutant phenotype, will be Col at the point of the mutation. Near the mutated gene, both chromosomes will have predominantly Col sequence and hence that locus is “linked”. Further away from the mutation, more recombination events will have taken place, introducing increasingly more Ler sequence. At positions that are “unlinked” to the mutation, segregation will be random and the chromosomes will average 50/50 Col/Ler sequence. This information can be used to determine linkage between markers and the mutant locus, estimate distances between linked markers (see Note 8 and Subheading 3.6.2) and hence determine the position of a particular mutation. This is the basis for positional cloning. 3.6.2. Pool Samples and Run Markers

In the first instance, test about three molecular markers per chromosome on DNA extracted from the 150 segregating F2 plants. Pooling of samples (preferably in groups of 5) for a bulked segregant analysis works well with a large population and considerably reduces cost and time. However, it is not advisable for small pool sizes (see Note 9). Analyse results by scoring individual plants as CC, CL, or LL for each of the molecular markers where C = Col and L = Ler (see Note 8). Determine the recombination frequency as the number of chromosomes with an L ecotype divided by the total number of chromosomes tested (i.e. two times the number of plants) and multiplied by 100. A recombination frequency of 50%, which is the maximum that is statistically possible, indicates unlinked markers. Conversely, the lower the frequency the closer the marker is to the mutant locus. A recombination frequency of 1 indicates one chromosomal crossover between the mutant locus and the marker’s locus per hundred F2 plants, and this can be used to approximately calculate genetic distance (centimorgans, cM). While genetic distances is not directly equivalent to physical distance due to variable recombination frequencies across chromosomes, a distance of 1 cM is equivalent to about 250 kb in Arabidopsis. DNA extraction: 1. Grind frozen leaf material and add 400 ml of Shorty buffer. 2. Denature material at 65°C for 10 min. 3. Centrifuge samples for 10  min at high speed (room temperature).

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4. Transfer supernatant into a new eppendorf tube. 5. Precipitate DNA using 350  ml isopropanol, mix well, and centrifuge for 5 min at high speed. 6. Wash pellet with 500 ml 70% (v/v) ethanol. 7. Resuspend dried pellet in 400 ml TE buffer. The isolated DNA can be used for crude and fine mapping. It should be stored at 4°C. 3.6.3. Fine Mapping

The initial mapping round and linkage analysis will give some idea of the position of the mutation. Screen more seeds from the F2 population for plants that are homozygous for the mutation. Collect both leaf tissue and seeds from each plant. The objective of fine mapping is to identify plant(s) with a crossover very close to the mutant locus, i.e. <0.1–0.01  cM (i.e. a 1/1,000 to 1/10,000 event) (Fig. 2). Thus, depending on the frequency of the recombination events on the chromosome you need a population in the order of 1,000 to up to 5,000 to be confident of identifying such plants.

3.6.4. Quick Screen to Identify Recombinant Plants

You will need to screen the entire population with two flanking markers that are easy to score. This will identify a few hundred recombinants with crossovers (breakpoints) between the markers and mutant locus. Calculate the genetic distance (cM) to determine the approximate location of the mutant locus. Plants that are LL or CL for the marker indicate a recombination event between the marker and the mutant locus, which will be CC. The remaining plants without breakpoints between the flanking markers, i.e. CC, are uninformative and can be discarded. The recombinant plants with breakpoints will help narrow down the region of the mutation and can therefore be used for further fine mapping analysis. The next step in gene identification depends on the resolution that could be achieved by fine mapping and the identification of candidate genes. Note if the mutant phenotype is too labour-intensive to score, it may be easier to identify recombinants and then test for zygosity of your mutant of interest.

3.6.5. Candidate Approach

To identify candidate genes in the region identified by fine mapping, check sequence databases (see Note 10) for genes that might account for the phenotype. Amplify candidate genes (standard PCR) from your various mutant alleles and look for polymorphisms, such as deletions, that are resolvable on a standard gel or sequence the entire gene to identify single base pair mutations common to EMS mutagenesis. To check for aberrant transcripts, for instance due to mutations in the intron/exon splice site, amplify the transcript by RT-PCR and run the products on a standard gel. To identify altered transcript levels, use real time

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1. Use mutagenised Columbia seed stock

1. Identify mutants that alter a biological process. 2. Cross mutant (Columbia (C) ecotype) with wild type Landsberg erecta (L) ecotype to introduce the Ler alleles of the genetic markers into the mutant line. 3. Self F1 generation. 4. Select homozygous mutants from F2 progeny. 5. Determine linkage to an initial set of genetic markers distributed across the five chromosomes. 6. Increase size of mapping population. 7. Undertake fine mapping on increased mapping population with more markers spanning the region. 8. Initiate gene identification strategies.

Mutant (mutmut ) 2.

mutmut

x

CC

MUTMUT Phenotype LL

3.

Genotype

Initial Round of Mapping F 2 plant #14,mutmut

mutMUT

F1

4.

CL

#14

a

LL

b

CC

c

CL

5.

marker a : unlinked to mut locus #14

F2 generation

C L

6. marker c : linked

#14 C L

7. F2 plant #14 a

b d g h e f c

b c Initial round

8.

F2 plant #33 b CC CC

g h

F2 plant #1024 b

CL CC

c

g h

c

CC CL

The recombinations in plants #33 and #1024 between the markers g and h, respectively, demonstrate that the mutation is located between those two markers g.

Fine mapping

Search for candidate genes between g and h and initiate gene identification and complementation strategies

Fig. 2. Mapping strategy.

PCR or Northern blot analysis. If fine mapping does not reduce the interval sufficiently and/or it is impossible to pick candidates, narrow down the remaining region by complementing mutant plants with vectors containing one or more candidate genes (10).

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Additionally, another option is to use microarrays to probe for differential expression of all genes. It is also worth considering sequencing the whole region, either by Sanger or deep sequencing technologies (19). 3.6.6. Complementation and Alternative Approaches

If a mutation has been identified, the mutation has to be confirmed to be responsible for the mutant phenotype. Several approaches can be used. An assay has to be performed by complementing the mutant phenotype with a wild type allele either as cDNA or genomic (10, 26, 27). Another approach to confirm and recreate the mutant phenotype is to generate knockdown plants for the candidate genes by RNA silencing (28) or search the Arabidopsis database for T-DNA insertion mutants (www.arabidopsis.org). This latter strategy is quite important if you only have a single allele as many researchers and some journals, for example The plant cell, require multiple alleles for the validation of critical phenotypes.

4. Notes 1. M1 populations can also be purchased from Lehle Seeds, (www.arabidopsis.com, Round Rock, Texas, USA). Tagged mutant populations, such as T-DNA insertion lines, are convenient. A stable tag can easily be identified and the interrupted gene identified by TAIL-PCR (29). Untagged populations, such as chemical, require time consuming positional-cloning approaches to gene identification. However, chemical mutagenesis can produce a range of subtle mutation types, such as missense mutations, promoter mutations, knockdowns, etc, which is advantageous in the case of a lethal knockout. It is also advisable to check if the observed phenotype is listed in TAIR (www.arabidopsis.org). You can search the database by keywords, and sometimes representative photos are provided. 2. Seeds could initially be plated out on higher agar concentration (i.e. 1.5% w/v) MS plates. This stops roots from growing deep and facilitates the transfer of the seedlings to 96-well plates. 3. Depending on the expression level of the gene of interest, it is advisable to determine the ideal concentration of luciferin to be used as this affects the luminescence signal. 4. An alternative approach is to use a cooled CCD camera (−60 to −80°C) in a dark box with image analysis software to visualise the induction of luciferase in individual plants in situ without transferring to the microtitre plate. This strategy has been successfully applied by a number of researchers (30, 31).

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However, the quality of the camera (in particular, a very low dark noise) and dark box are critical as the level of luminescence is very low and may require exposure times of 5–20 min. 5. The effect of high light may be very transient and thus limiting luciferase expression. Therefore, do not count too many plates at once. Also, results from the primary screen can be variable due to different plant sizes, altered luciferin uptake, etc. Thus, it is essential to re-screen candidate lines. It would be advisable to assay luciferase activity per gram fresh weight. Also check to determine if APX2 mRNA is also correspondingly altered to determine that the mutation is extragenic to APX2:LUCIFERASE: that is, the lesion is not in the actual APX2:LUCIFERASE transgene. 6. Allelic mutant plants exhibiting different mutations in the same gene can confirm if the disrupted gene is responsible for the mutant phenotype. This is particularly important for pleiotropic phenotypes, especially in EMS mutants that can have up to 100 mutations per plant. 7. All markers anneal at 55°C. For exact conditions, see (20). The PCR products are analysed on a 3.0% (w/v) agarose (for SNPs with less than 5 bp difference, a high resolution agarose can be used at 3.5 to up to 4% (w/v), such as Agarose-1000). The gel has to be poured hot to reduce “smiling” and increase resolution. High percentage gels will have to be run significantly longer than a normal 1% (w/v) agarose gel. To differentiate markers with less than 10 bp difference, try to keep marker products short (100 bp or less) and optimise reactions carefully to reduce primer dimer interference. An alternative method is to look for mutations by high resolution melting (HRM) of PCR amplicons by real time PCR as described by Chateigner-Boutin (32). Although this method may require some optimization and be more expensive than conventional methods, it has the advantage of being faster and sensitive. 8. The genetic map distance (D) can be determined in terms of map units or cM. For many purposes, recombination frequency is a good approximation of map distance. Recom­ bination frequency (r) is calculated as number of Ler plants for a marker (where LL equals 2 and LC equals 1) divided by the double total number of plants and expressed as a percentage. 9. If you use significantly less plants in the initial mapping round, a pooled approach may be statistically misleading. Therefore, it is preferable for samples to be kept separate. 10. The Arabidopsis Information Resource (TAIR, http://www. arabidopsis.org/), MIPS A. thaliana database (MATDB,

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http://mips.gsf.de/proj/thal/db/index.html) and the National Center for Biotechnology Information (NCBI, http://www. ncbi.nlm.nih.gov/) are all good resources for annotated sequence data.

Acknowledgments We would like to thank Dr. Phil Mullineaux (John Innes Institute, Norwich, UK) for kindly providing us with EMS-mutagenised APX2-luciferase seed. We also thank Jan Bart Rossel (Australian National University) for input into the first version of this manuscript. References 1. Pogson, B.J., Woo, N.S., and Forster, B. (2008) Small ID: plastid signalling to the nucleus and beyond. Trends Plant Sci 13(11), 602–609. 2. Niyogi, K.K. (1999) Photoprotection revisited: genetic and molecular approaches. Annual Rev Plant Physiol Plant Mol Biol 50, 333–359. 3. Foyer, C.H. and Noctor, G. (1999) Leaves in the dark see the light. Science 284(5414), 599–601. 4. Niyogi, K.K., Grossman, A.R., and Bjorkman, O. (1998) Arabidopsis mutants define a central role for the xanthophyll cycle in the regulation of photosynthetic energy conversion. Plant Cell 10(7), 1121–1134. 5. Meurer, J., Meierhoff, K., and Westhoff, P. (1996) Isolation of high-chlorophyll-fluorescence mutants of Arabidopsis thaliana and their characterisation by spectroscopy, immunoblotting and northern hybridisation. Planta 198(3), 385–396. 6. Pogson, B., McDonald, K.A., Truong, M., Britton, G., and DellaPenna, D. (1996) Arabidopsis carotenoid mutants demonstrate that lutein is not essential for photosynthesis in higher plants. Plant Cell 8(9), 1627–1639. 7. Park, H., Kreunen, S.S., Cuttriss, A.J., DellaPenna, D., and Pogson, B.J. (2002) Identification of the carotenoid isomerase provides insight into carotenoid biosynthesis, prolamellar body formation, and photomorphogenesis. Plant Cell 14(2), 321–332. 8. Asada, K. (1999) The water-water cycle in chloroplasts: scavenging of active oxygens and dissipation of excess photons. Annu Rev Plant Physiol Plant Mol Biol 50, 601–639.

9. Karpinski, S., Reynolds, H., Karpinska, B., Wingsle, G., Creissen, G., and Mullineaux, P. (1999) Systemic signaling and acclimation in response to excess excitation energy in Arabidopsis. Science 284(5414), 654–657. 10. Wilson, P.B., Estavillo, G.M., Field, K.J., Pornsiriwong, W., Carroll, A.J., Howell, K.A., Woo, N.S., Lake, J.A., Smith, S.M., Harvey, M.A. et  al. (2009) The nucleotidase/phosphatase SAL1 is a negative regulator of drought tolerance in Arabidopsis. Plant J. 58(2), 299–317. 11. Woo, N., Badger, M., and Pogson, B. (2008) A rapid, non-invasive procedure for quantitative assessment of drought survival using chlorophyll fluorescence. Plant Methods 4(1), 27. 12. Murashige, T. and Skoog, F. (1962) A revised medium for rapid growth and bioassays with tobacco tissue culture. Physiol Plant 15, 473–497. 13. Lightner, J. and Caspar, T. (1998) Seed mutagenesis of Arabidopsis. Methods Mol Biol 82, 91–103. 14. Leyser, O. (2000) Mutagenesis. Methods Mol Biol 141, 133–144. 15. Jander, G., Norris, S.R., Rounsley, S.D., Bush, D.F., and Levin, I.M. (2002) Last RL: Arabidopsis map-based cloning in the postgenome era. Plant Physiol 129(2), 440–450. 16. Ossowski, S., Schneeberger, K., Clark, R.M., Lanz, C., Warthmann, N., Weigel, D. (2008) Sequencing of natural strains of Arabidopsis thaliana with short reads. Genome Res 18(12), 2024–2033. 17. Koornneef, M., Alonso-Blanco, C., and Stam, P. (1998) Genetic analysis. Methods Mol Biol 82, 105–117.

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18. Salathia, N., Lee, H.N., Sangster, T.A., Morneau, K., Landry, C.R., Schellenberg, K., Behere, A.S., Gunderson, K.L., Cavalieri, D., Jander, G. et al. (2007) Indel arrays: an affordable alternative for genotyping. Plant J 51(4), 727–737. 19. Lister, R., Gregory, B.D., and Ecker, J.R. (2009) Next is now: new technologies for sequencing of genomes, transcriptomes, and beyond. Curr Opin Plant Biol 12(2), 107–118. 20. Lukowitz, W., Gillmor, C.S., and Scheible, W.R. (2000) Positional cloning in Arabidopsis. Why it feels good to have a genome initiative working for you. Plant Physiol 123(3), 795–805. 21. Bell, C.J. and Ecker, J.R. (1994) Assignment of 30 microsatellite loci to the linkage map of Arabidopsis. Genomics 19(1), 137–144. 22. Jarvis, P., Lister, C., Szabo, V., and Dean, C. (1994) Integration of CAPS markers into the RFLP map generated using recombinant inbred lines of Arabidopsis thaliana. Plant Mol Biol. 24(4), 685–687. 23. Michaels, S.D. and Amasino, R.M. (1998) A robust method for detecting single-nucleotide changes as polymorphic markers by PCR. Plant J. 14(3), 381–385. 24. Neff, M.M., Neff, J.D., Chory, J., and Pepper, A.E. (1998) dCAPS, a simple technique for the genetic analysis of single nucleotide polymorphisms: experimental applications in Arabidopsis thaliana genetics. Plant J 14(3), 387–392. 25. Herrmann, M.G., Durtschi, J.D., Bromley, L.K., Wittwer, C.T., and Voelkerding, K.V. (2006) Amplicon DNA melting analysis for mutation scanning and genotyping: crossplatform comparison of instruments and dyes. Clin Chem 52(3), 494–503. 26. Albrecht, V., Ingenfeld, A., and Apel, K. (2006) Characterization of the snowy cotyledon

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Chapter 21 Expression of Genes in Cyanobacteria: Adaptation of Endogenous Plasmids as Platforms for High-Level Gene Expression in Synechococcus sp. PCC 7002 Yu Xu, Richard M. Alvey, Patrick O. Byrne, Joel E. Graham, Gaozhong Shen, and Donald A. Bryant Abstract Synechococcus sp. PCC 7002 is an ideal model cyanobacterium for functional genomics and biotechnological applications through metabolic engineering. A gene expression system that takes advantage of its multiple, endogenous plasmids has been constructed in this cyanobacterium. The method involves the integration of foreign DNA cassettes with selectable markers into neutral sites that can be located on any of the several endogenous plasmids of this organism. We have exploited the natural transformability and powerful homologous recombination capacity of this organism by using linear DNA fragments for transformation. This approach overcomes barriers that have made the introduction and expression of foreign genes problematic in the past. Foremost among these is the natural restriction endonuclease barrier that can cleave transforming circular plasmid DNAs before they can be replicated in the cell. We describe herein the general methodology for expressing foreign and homologous genes in Synechococcus sp. PCC 7002, a comparison of several commonly used promoters, and provide examples of how this approach has successfully been used in complementation analyses and overproduction of proteins with affinity tags. Key words: Synechococcus, Gene expression, Affinity-tag, Homologous recombination, Cyanobacteria, Transformation, Plasmid

1. Introduction Synechococcus sp. PCC 7002 is a unicellular, euryhaline cyanobacterium that possesses several characteristics that make it well suited for fundamental biological research (1–10) as well as biotechnological and industrial applications. These properties include, but are not limited to, its fast growth rate; tolerance of high light

Robert Carpentier (ed.), Photosynthesis Research Protocols, Methods in Molecular Biology, vol. 684, DOI 10.1007/978-1-60761-925-3_21, © Springer Science+Business Media, LLC 2011

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intensities; natural transformability; facultative photoheterotrophy and heterotrophy; and fully sequenced genome (11–15) (see Note 1). Recently, significant advances obtained from genome sequencing, proteomics, transcriptomics, metabolomics, functional genomics, and the creation of new strains and insights from mutagenesis and genetic modification provide a solid basis for further utilization of this strain in metabolic engineering. Synechococcus sp. PCC 7002 naturally harbors six endogenous plasmids, denoted pAQ1 (4809 bp), pAQ3 (16,103 bp), pAQ4 (31,972  bp), pAQ5 (38,515  bp), pAQ6 (124,030  bp), and pAQ7 (186,459  bp) (16); (see National Center for Biotechnology Information, http://www.ncbi.nlm.nih.gov/). Although the exact copy numbers of these plasmids per cell vary somewhat with growth conditions, the largest two plasmids always appear to have the same copy number as the chromosome (copy number per cell in exponential phase, chromosome: pAQ1: pAQ3: pAQ4: pAQ5: pAQ6: pAQ7 = 6:~50:27:15:10:6:6; T. Li and D. A. Bryant, unpublished results). So long as the introduced DNA does not interfere with the expression of any essential gene(s), we have consistently observed that expression cassettes can specifically be targeted by homologous recombination into any of these six plasmids or the chromosome with little or no apparent impact on cell viability. The smallest of the plasmids, pAQ1, has the highest copy number and appears to be indispensable (Fig. 1a). Attempts to cure strains of this plasmid have failed (17) (D. A. Bryant, unpublished results). However, a strain of Synechococcus sp. PCC 7002 that had spontaneously been cured of pAQ4 was isolated. This result demonstrates that pAQ4 carries no genes that are essential for the growth of Synechococcus sp. PCC 7002 under standard laboratory growth conditions and could indicate that wild-type copies of this plasmid would be completely displaced under ­antibiotic selection conditions (T. Li, J. Marquardt, J. Zhao, and D. A. Bryant, unpublished results). Although pAQ1 has been the primary target for gene overexpression in our laboratory, we have found that the second smallest plasmid, pAQ3 (16,103 bp), is also useful for gene expression applications (Fig.  1b) (see Note 2). In order to streamline the routine introduction and expression of foreign genes and homologous genes into the plasmids of this cyanobacterium, we have developed gene expression cassettes that include: an exogenous (usually cyanobacterial) promoter, multicloning sites, a deca-histidine (His10–) affinity tag, a transcription terminator, and an antibiotic resistance cartridge. These DNA elements are linked in tandem between two flanking DNA regions derived from the appropriate plasmid target in order to target the overexpression cassette to specific neutral sites in these two plasmids (Figs. 1 and 2).

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Fig. 1. pAQ1, pAQ3, and a generic expression vector that can be used for the introduction of genes into Synechococcus sp. PCC 7002. Regions of homology, and thus targets for homologous recombination, are indicated as “Flank A” and “Flank B.” Open reading frames are indicated with gray arrows and appropriate labels. (a) Endogenous, wild-type plasmid pAQ1 from Synechococcus sp. PCC 7002. (b) Endogenous, wild-type plasmid pAQ3 from Synechococcus sp. PCC 7002. (c) Generic representation of an expression vector that can be made to insert expression constructs into the genome of Synechococcus sp. PCC 7002. The backbone of this vector, the region from the SphI and NsiI restriction sites that includes the AmpR (ampicillin resistance cassette) and Rep (pMB1) (origin of replication) are from pGEM-7zf.

2. Materials 2.1. Manipulation of DNA

1. Equipment and supplies for performing agarose gel electrophoresis of DNA (gel-casting system, power supply, agarose, ethidium bromide, TAE buffer, 5× loading buffer, etc.).

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Fig. 2. Schematic showing the construction and use of the pAQ1 expression system to express a gene under the control of the cpcBA promoter of Synechocystis sp. PCC 6803 in Synechococcus sp. PCC 7002. Relevant restriction enzyme recognition sequences are indicated. Briefly, a generic gene, Gene X, is amplified by PCR to include an NdeI restriction site at the start codon of the gene of interest and a BamHI site after the stop codon. The resulting product is then digested and cloned into pAQ1Ex-PcpcBA to form pAQ1Ex-PcpcBA-GeneX, which is then either digested with SphI or NsiI or used as template in a PCR reaction using primers pAQ1flankBF and pAQ1flankAR. This linear DNA fragment is used to transform wild type cells of Synechococcus sp. PCC 7002. After homologous recombination and spectinomycin selection, the foreign DNA will be recombined into endogenous pAQ1 plasmid to form the new plasmid, from which GeneX will be expressed.

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2. Equipment and supplies for performing PCR (PCR machine, aerosol-barrier tips, sterile water, etc.). 3. DNA gel extraction kit. 4. Plasmid extraction/purification kit. 5. High-Fidelity DNA Polymerase (e.g., Phusion™) (see Note 3). 6. 2¢-Deoxyribonucleoside 5¢-triphosphate (dNTP) mixture for PCR. 7. Custom-designed primers. 8. Appropriate restriction enzymes and incubation buffers. 9. T4 DNA ligase and ligation buffer. 10. Plasmids containing an appropriate antibiotic resistance marker. 11. Supplies for ethanol precipitation of DNA. 12. Standard cloning vector (e.g., pGEM-7Zf(+/−) vectors) (Promega, Madison WI). 13. Transformation-competent Escherichia coli (e.g., DH5a or DH10B). 14. Media for the growth of E. coli, (e.g., Luria-Bertani (LB) medium containing appropriate antibiotics). 2.2. Transformation of Synechococcus sp. PCC 7002

The protocol for the transformation of Synechococcus sp. PCC 7002 has been described in detail previously (18). The materials listed here are largely a duplication of that information. 1. Equipment for the cultivation of Synechococcus sp. PCC 7002 in liquid medium and on solid medium (an illuminated incubator or aquarium bath at 38°C, sterile gas delivery/bubbling device, etc.). 2. Liquid A+ medium (Medium A containing 1.0 g NaNO3/L) (26) prepared with or without glycerol (see Note 4 and ref. (19)). 3. Plates with solid A+ medium prepared with 1.5% (w/v) BactoAgar (Becton Dickinson, Sparks, MD) and with or without glycerol (see Note 5). 4. Appropriate antibiotics (see Note 6). 5. Benchtop centrifuge (Eppendorf 5417C).

2.3. Materials for Expression Analysis (Optional)

1. Polyacrylamide gel electrophoresis reagents (polyacrylamide, sodium dodecyl sulfate, Coomassie Brilliant Blue, etc.) (20). 2. Gel electrophoresis equipment. 3. French press, sonicator, or Bugbuster™ reagent from Novagen (EMD Chemicals Inc., Madison, WI).

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4. Immunoblotting equipment (e.g., Semi-dry Transfer Cell, BioRad, Hercules, CA) and supplies (nitrocellulose membranes, etc.). 5. Protein concentration determination kit (such as Coomassie Plus Protein Assay Reagent (Pierce, Rockford, IL)). 6. Antitag antibodies (e.g., His6 Antibody from Rockland Immunochemical (Gilbertsville, PA)). 7. Nickel-nitrilotriacetic acid (Ni-NTA) affinity resin. 8. Chromatography column. 9. RT-PCR kit (e.g., Roche OneStep™ RT-PCR kit (Roche Diagnostics, Indianapolis, IN). 10. RNA Isolation kit (e.g., Roche Diagnostics, Indianapolis, IN).

3. Methods 3.1. Creating a Site-Specific Targeting Vector for Gene Expression

3.1.1. PCR Amplification and Cloning of Integrating Flank Sequences

Our expression cassettes for Synechococcus sp. PCC 7002 comprise several components, including a heterologous (usually cyanobacterial) promoter, multicloning sites, a deca-histidine (His10–) affinity tag, a transcription terminator, and an antibiotic resistance cartridge all linked in tandem between two flanking DNA regions derived from the appropriate plasmid target (Fig. 1c). These flanking regions are required for efficient integration of the expression cassette into the target DNA of Synechococcus sp. PCC 7002 through double homologous recombination (see Notes 7 and 8). If it is anticipated that a site will be used repeatedly, it is very convenient to have these flanking sequences cloned into a standard E. coli plasmid, such as the pGEM®-7Zf(+/−) vectors using standard molecular biology techniques. 1. Amplify (see Note 3) flank A and flank B by PCR using custom-designed primers with appropriate restriction sites for cloning and downstream integration of antibiotic resistance cassette and gene of interest (see Note 9 for an example). 2. Confirm the success of the PCR reaction by electrophoresis of the DNA on an agarose gel. 3. If the reaction was successful, precipitate the DNA from the reaction mixture with 70% (v/v) cold ethanol to remove the PCR reaction buffer. 4. Digest the PCR product with the appropriate restriction enzymes according to the manufacturer’s protocol. 5. Purify the DNA digestion product by electrophoresis on a 0.8% (w/v) agarose gel; excise the appropriate region of the gel containing the DNA and purify the DNA using a DNA gel extraction/purification kit.

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6. Set up a ligation reaction using one of the inserts above and a similarly prepared E. coli cloning vector. Allow the mixture to incubate for several hours or overnight at 16°C to room temperature. 7. Transform the ligation product into transformation-competent cells of E. coli strains DH5a or DH10B. 8. After incubation at 37°C for 1 h in 1.0 mL of E. coli growth medium SOC, the cells should be plated on solid LB media containing appropriate antibiotic(s) and incubated at 37°C overnight. 9. Confirm successful ligation by picking a few (10–20) colonies and by growing them individually overnight in 3.0  mL of ­liquid LB medium with antibiotic(s). Isolate plasmids using a plasmid DNA isolation kit, and confirm the presence of the correct insert by either restriction digestion or DNA sequencing (or both). 10. Repeat the process for the second flank and ligate it into the previously verified plasmid to produce the first insert. 3.1.2. Cloning of Additional Components of Expression Cassette

Along with the appropriate flanking sequences, the transformation cassette must also include an antibiotic resistance marker gene (see Note 6) and the gene of interest to be heterologously expressed (see Notes 9–13). These can be subcloned from another vector or amplified by PCR and cloned into the newly constructed vector described above, which contains the flanking sequences for site-directed double homologous recombination (see Notes 9 and 16 and Fig. 1c for example).

3.2. Preparations for Transformation into Synechococcus sp. PCC 7002

In order to ensure high transformation efficiency and appropriate site-specific integration of the gene of interest, linear fragments of DNA should be used to transform Synechococcus sp. PCC 7002. This can most conveniently be accomplished by amplifying the construct to be introduced in the cyanobacterial cells by PCR by using the outermost pair of primers that were also used to amplify the two flanking sequences (Fig. 2).

3.2.1. PCR of Cassette from Vector to Create Transformation Fragment 3.2.2. Preparation of Cells of Synechococcus sp. PCC 7002 for Transformation

3.3. Transformation of Synechococcus sp. PCC 7002 3.3.1. Simplified Transformation Protocol

Liquid cultures are grown in A+ medium. Prepare an exponentially growing culture (OD730  nm = 1.0–1.5) of Synechococcus sp. PCC 7002 at 38°C in liquid medium A+ bubbled slowly for about 1 to 3 days with sterile air supplemented with 1% (v/v) CO2. This protocol is a slightly modified and streamlined version of the protocol described previously (18). 1. Mix prepared linearized transformation cassette DNA (1–5 mg or 5–10 mL of a typical PCR amplification reaction) and 2 mL of the liquid culture in a fresh, sterile bubbling tube and incubate under saturating illumination (about 250  mE/m2s) at

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38°C for 5 h to overnight while gently bubbling with sterile air supplemented with 1% (v/v) CO2. A negative control, in which no DNA is added to a 2-mL aliquot of cells, should also be prepared. 2. After incubation, the cells should be aliquoted into a sterile 1.5-mL microcentrifuge tube (generally enough will have evaporated after overnight incubation so that this will be the complete contents of the tube) and concentrate the cells by centrifugation for 1 min at 1,500 × g (e.g., ~4,000 rpm in the benchtop centrifuge). Remove most of the cleared medium with a micropipetor, and then spread the remaining cell suspension over a solid A+ plate containing the appropriate antibiotic(s). The plate(s) should then be incubated under moderate illumination (about 150 mE/m2s) at 30°C for about 7–10 days, or until colonies are visible. 3. Transfer several colonies onto fresh plates containing solidified A+ medium with the appropriate antibiotic(s), and incubate again under the same conditions as in step 2 above. Colonies can be transferred using sterile toothpicks or by resuspension in 1–2 mL of sterile A+ media. Use of a stereo dissecting microscope may aid in the detection and recovery of colonies. Increased incubation time will allow colonies to become larger, and they will correspondingly be easier to manipulate. Colonies can be repeatedly streaked on fresh media until a homozygous isolate is obtained (two to three restreakings from a single colony is usually sufficient unless the wild-type gene product is strongly selected for under the growth conditions employed (see Note 14). 4. For each clonal isolate, streak one colony in a small area (about 1 cm2) on a fresh plate and incubate until a thick patch of cells appears. This patch of cells can be used to verify the segregation of the wild-type and mutant alleles (see Notes 14 and 15), and the cell patch can also serve as a source of cells to inoculate liquid cultures for further study or for long-term storage. Strains may be frozen and stored as described in Note 16. After having successfully isolated strains resistant to the newly introduced antibiotic, it may additionally be necessary to confirm the presence and expression of the introduced gene (see Notes 9 and 17–20).

4. Notes 1. Genome database sites: (A) Kazusa DNA Research Institute, Cyanobase: http://genome.kazusa.or.jp/cyanobase/ , (B) Joint Genome Institute, DOE: http://img.jgi.doe.gov/cgi-bin/pub/

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main.cgi , (C) National Center for Biotechnology Information: http://www.ncbi.nlm.nih.gov/genomes/lproks.cgi 2. In some cases, it may be useful to express two or more genes separately from distinct locations in the genome. We have had success targeting additional constructions to pAQ3 (Fig. 1b), and homologous recombination has been observed for all other plasmids. Similar neutral sites could also be developed for any of the remaining plasmids or in the chromosome of Synechococcus sp. PCC 7002. We have successfully used genes encoding proteins conferring antibiotic resistance to the following: streptomycin/spectinomycin, kanamycin, gentamicin, chloramphenicol, erythromycin, and ampicillin as suitable selection agents (see Note 6). 3. A typical 50-mL PCR reaction using Phusion™ DNA polymerase and genomic DNA from Synechococcus sp. PCC 7002 as template contains: 10  mL 5× HF (High Fidelity) buffer, 1 mL of 10 mM dNTP, 2.5 mL of each 10 mM primer, 100 ng genomic DNA, 0.5 mL Phusion™ DNA polymerase, and sterile water to produce a final volume of 50 mL. The reaction conditions are: one cycle of initial denaturation at 98°C for 30  s, 35 cycles of denaturation at 98°C for 10  s, 20  s of annealing at a temperature +3°C of the lower Tm primer, 20 s per kb of extension at 72°C, and one cycle of final extension at 72°C for 10 min. 4. Because medium A+ contains no organic carbon source, Synechococcus sp. PCC 7002 may exhibit slow growth when genes affecting the photosynthetic apparatus are expressed. Improved growth may be obtained by providing 10  mM glycerol, which allows mixotrophic (or photoheterotrophic or heterotrophic) growth, as an additional carbon and energy source under such conditions. However, because the wild-type strain of Synechococcus sp. PCC 7002 is sensitive to glycerol, a glycerol-tolerant variant must first be isolated. A simplified method for obtaining such a strain is to grow a WT culture in 25 mL of medium A+ containing 1 mM glycerol for 2–3 days to a relatively high turbidity for a starter culture. This liquid culture should be subcultured in medium A+ containing 10  mM glycerol and again grown to early stationary phase (OD730 nm of ~2.0–2.5). During the first growth period, the glycerol-sensitive cells will not grow or will die, but glyceroltolerant cells will nevertheless survive and eventually grow out. Once the OD730 nm has recovered to a value greater than 1.0, the cells should be plated on solid A+ medium containing 10  mM glycerol. Selection of a fast growing colony will provide a glycerol-tolerant strain that can then be used in transformation experiments as before, and the resulting transformants will be much better adapted for the growth on A+ media containing glycerol.

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5. Synechococcus sp. PCC 7002 seems to have a strong preference for the growth on BactoAgar (As described on the bottle from Becton, Dickinson, Sparks, MD, “A solidifying agent in which extraneous matter, pigmented portions and salts have been reduced to a minimum.”). Use of other solidifying agents (generic agar, or a gellan gum-based product, such as Phytagel or Gelrite) may be acceptable for some applications, but plating efficiencies will be severely reduced. 6. The useable concentrations of antibiotics for Synechococcus sp. PCC 7002 are: gentamicin: 50  mg/mL; streptomycin: 100 mg/mL; spectinomycin: 50 mg/mL; kanamycin: 100 mg/ mL; ampicillin: 5–50 mg/mL; chloramphenicol: 10 mg/mL; and erythromycin: 20 mg/mL (18). 7. Although flanking sequences as short as 100 bp can be sufficient to target an insertion to an appropriate site by homologous recombination, longer flanking regions (0.5–1.0 kb) are more efficient. There are some practical limitations because the use of longer sequences may complicate and/or reduce the efficiency of other manipulations. The endogenous plasmids of Synechococcus sp. PCC 7002 are good targets for integration and gene expression due to their relatively high copy number per cell, which may aid in maximizing homologous recombination as well as promoting higher transcript levels. The design of the flanking regions can be performed such that a significant region of a gene, operon or the plasmid target is removed; alternatively, they may be designed to preserve most of the plasmid sequence (see Fig.  1a and b). It may be advantageous in some cases to eliminate significant portions of the plasmid in order to avoid unforeseen restrictions on plasmid size and reduce possible affects on copy number that now these large plasmids may have. The best design and configuration conditions should be considered on a case-by-case basis and then optimized empirically (see Note 9). 8. If integration into an untested site is desired, it may be beneficial to test the selected site by ligating the flanking sequences to an antibiotic resistance cassette and transforming this construction into Synechococcus sp. PCC 7002 to determine if there are any unanticipated effects on viability. This test can be easily be accomplished by using PCR for the amplification of all three fragments, along with appropriate restriction endonuclease digestions to create “sticky ends,” followed by ligation using T4 DNA ligase to create a linear DNA molecule as follows: left flank-antibiotic resistance cassette-right flank. The products of this reaction can be directly transformed into Synechococcus sp. PCC 7002 using the outlined procedures (see Subheading 3.3.1).

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9. The example below outlines the construction of the expression vector pAQ1Ex-PcpcBA, which is the cassette that we have found to produce the highest gene expression levels obtained so far. The methods described can easily be adapted for other specific purposes, however. Two regions of pAQ1, a 512-bp region of pAQ1 from nucleotide 4196 to 4707 and a 377-bp region of pAQ1 from nucleotide 2635 to 3011, were amplified by PCR (using primers pAQ1flankAF, pAQ1flankAR, pAQ1flankBF, and pAQ1flankBR, respectively; see Table 1). These two fragments were digested with SalI and XbaI restriction endonucleases, respectively. The resultant products were ligated to the aadA gene from the plasmid pSRA81 (18) that had been amplified by PCR by using primers aadAF and aadAR (Table 1). The aadA amplicon, which encodes an enzyme conferring streptomycin and spectinomycin resistance, was digested with both SalI and XbaI restriction endonucleases. The resulting ligation reaction was used to transform Synechococcus sp. PCC 7002 wild-type cells (see Subheading 3.3) in order to determine if they would be viable with the new construct, which deletes 1184 bp of pAQ1 from bases 3012 and 4195. The cells were viable and the resulting region was then subcloned after PCR amplification (using primers pAQ1flankBF and pAQ1flankAR; Table  1) from the cyanobacterial DNA into the NsiI and SphI sites of the multicloning site of pGEM-7zf to form pGEM-AQSp. The gene insertion site from pET-16b was then amplified by PCR using primers pet16bF and pet16bR (Table 1), digested with NcoI and ligated to the PCR-amplified cpcBA (phycocyanin) operon promoter from Synechocystis sp. PCC 6803 (nucleotides 728,063 to 727,469 using primers cpcBproF and cpcBproR; Table 1), which had also been digested with NcoI. The resulting ligation product was purified by agarose gel electrophoresis, digested with EcoRI and XbaI, and ligated into similarly digested pGEM-AQSp to form pAQ1ExPcpcBA. Alternative promoter regions, which contain EcoRI and NcoI restriction sites introduced during PCR amplification, can be cloned into pAQ1Ex-PcpcBA by replacing the cpcBA promoter cassette, thereby producing constructions that allow for different transcription rates (see Note 10). In order to avoid recombination problems that might lead to plasmid integration into the chromosome, overexpression constructions have been introduced into strains in which there were no regions of sequence identity between the ­chromosome (or other plasmids) and the expression plasmid. Operationally, for foreign genes, the promoter with the ­highest measured activity to date is the phycocyanin (cpcBA) ­promoter from Synechocystis sp. PCC 6803, and this promoter has been the first choice for the majority of our studies

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Table 1 DNA oligonucleotides sequences (added restriction digestion sites are underlined) Primer name

Sequence (5¢ to 3¢)

pAQ1flankAR

TTTGGCGCATGCGGGGTTTTCTCGTGTTTAGGC

pAQ1flankAF

AAGGCAGTCGACCTTTCTCTTATGCACAGATGGG

pAQ1flankBF

AACAGGATGCATCTCTCACCAAAGATTCACCTG

pAQ1flankBR

TCACGGTCTAGAGCGAATTCGCCTCCTGAATAAATCTATTTATAC

pAQ1F

ATTAGCATGCCGCTCTCACCAAAGATTCACCTG

pAQ1R

ATTAATGCATGAGTGGGGGAAAACGAACC

cpcBproF

AGGAATTCGTTATAAAATAAACTTAACAAATCTAT

cpcBproR

CGTCGACCATGGAATTAATCTCCTACTTGACTTTATG

PkanF

GAACACGTAGAATTCCAGTCCGCAGAAAC

PkanR

CATCGAGCTCAACCATGGGAAACGATCC

AADAF

GGAGGCTCTAGACGAACGCAGCGGTGGTAAC

AADAR

TTTTTGGTCGACGTCGAGCGAATTGTTAGACATTA

pet16bR

TCCTCCTCTAGACAAAAAACCCCTCAAGACCCGT

pet16bF

TTTAAGAAGGAGATATACCATGG

eyfpF

GAATTCTCGCATATGGTGAGCAAGGGCGAGGA

eyfpR

GTCGCGGGATCCTTACTTGTACAGCTCGTCCATG

fdcpecpR

CAAATGCCATGGATTGTTTCTCCTGTTAACGA

fdcpecpF

CACTAAGAATTCCGGTGACACAGCATTATACTTA

6psbAF

GTTTACGAATTCAAAAAACGACAATTACAAGAAAGTA

6psbAR

TCGTGGCCATGGGGTTATAATTCCTTATGTAT

7942crtGF

CTTAATTCGAGGTCATATGAACCTGCT

7942crtGR

ACAGTTTACTGCTGGGATCCCCTAGG

(see Note 10). Transcriptional profiling suggests the psaAB promoter may produce a similar level of transcription under standard growth conditions (M. Ludwig and D. A. Bryant, unpublished results). 10. The yellow fluorescent protein (YFP) variant of the green florescent protein (GFP) has been an excellent reporter for studying the parameters of this system. The YFP gene from plasmid pEYFP (Clontech, Mountain View, CA) was amplified using primers eyfpF and eyfpR (Table  1) and digested with NdeI and BamHI restriction endonucleases. The fragment

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was then cloned into the similarly digested pAQ1Ex-PcpcBA plasmid to yield pAQ1Ex-PcpcBA::YFP. Characterization of YFP expression proceeded using the following protocol: (a) Grow a culture (1L, A+ medium containing 50 mg spectinomycin/mL) of the Synechococcus sp. PCC 7002 strain harboring pAQ1Ex-PcpcBA::YFP to late exponential phase. (b) Harvest cells by centrifugation at 5,000 × g (Sorvall SLC4000 rotor). (c) Resuspend pelleted cells in 40  mL lysis/binding buffer (50 mM Na2HPO4, 300 mM NaCl, 10 mM imidazole) and transfer to a 50 mL conical tube. (d) Centrifuge suspension again at 5,000 × g in a Piramon Technologies F13-14x50cy rotor, and resuspend the resulting pellet in 20 mL lysis/binding buffer. (e) Disrupt cells by three passages through a chilled French pressure cell operated at 138 MPa. (f) Clarify the extract by ultracentrifugation at 90,000 × g in a Beckman 70Ti rotor for 35 min. (g) To the cleared lysate, add a 1.0 mL aliquot of Qiagen Ni-NTA resin that has been washed and resuspended in lysis/binding buffer. Incubate the mixture with constant mixing for 1 h at 4°C. (h) Apply the slurry to a column and elute the protein using treatments with lysis/binding buffer containing imidazole at 30 mM, 50 mM, 100 mM, 250 mM, and 500 mM concentrations by using wash volumes of 4 mL, 0.5 mL, 0.75 mL, and 0.5 mL, respectively. (i) Analyze YFP content by electrophoresis on a 10% (w/v) polyacrylamide gel in the presence of sodium dodecylsulfate (20) (see Fig. 3a). (Note: As a preliminary measure, the YFP content of each sample can be examined by visually assessing the fluorescence of the aliquots by using UV excitation light.). In order to determine the relative strength of expression that occurs using our expression system, several different variants of the pAQ1Ex expression cassette were constructed. The Synechocystis sp. PCC 6803 cpcBA promoter was removed by digestion of pAQ1Ex-PcpcBA::YFP with NcoI and EcoRI ­allowing alternative promoters to be cloned in its place using the same restriction enzymes. The amount of YFP florescence per OD730  nm was determined using SLM-Aminco 8100C ­fluorometer modernized for computerized data acquisition by On-Line Instrument Systems (Bogart, GA). The following promoters were tested: the cpcBA promoter from Synechocystis sp. PCC 6803; the cpeC promoter from Fremyella

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Fig. 3. Use of YFP as a reporter for gene expression in Synechococcus sp. PCC 7002: (a) YFP, expressed from its gene under the control of the cpcBA promoter from Synechocystis sp. PCC 6803, was isolated from whole-cell extracts by nickel chelation chromatography. Samples eluted with increasing concentrations of imidazole were exposed to UV light to reveal the presence of the YFP and were analyzed using SDS-PAGE. (b) Results of a promoter assay using a series of expression cassettes controlling YFP expression using several different promoters. Abbreviations are as follows: WT wild-type cells with no expression plasmid, cpcB strain transformed with the Synechocystis sp. PCC 6803 cpcBA promoter controlling YFP expression, cpeC strain transformed with the Fremyella diplosiphon cpeC promoter controlling YFP expression, Kan strain transformed with the promoter from aphII (kanamycin resistance cassette) controlling YFP expression, psbA2 strain transformed with the Synechocystis sp. PCC 6803 psbA2 promoter controlling YFP expression.

diplosiphon using primers fdcpecpF and fdcpecpR (Table 1); the ­promoter for the aphll gene in the kanamycin resistance cassette of pRL161 using primers PkanF and PkanR (28); and the psbA2 promoter from Synechocystis sp. PCC 6803 using

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primers 6psbAF and 6psbAR (Table  1). The results of this study are shown in Fig. 3b; they may provide some assistance in choosing an appropriate promoter for particular expression studies. 11. Additional considerations for the construction of an expression platform are the choice of a promoter and the inclusion of an insertion site that may or may not include the ability to add sequence tags to a protein of interest. If the gene being introduced into Synechococcus sp. PCC 7002 originates from another cyanobacterium, its native promoter can often be used to drive the expression. The choice of a promoter will largely be based on the specific application for the protein being expressed, and it may be very useful to express the gene from a noncognate promoter. We have characterized a few promoters for use with this system (see Note 10). Our general strategy has been to avoid using promoters from Synechococcus sp. PCC 7002 in favor of those from other closely related cyanobacteria, especially Synechocystis sp. PCC 6803, so as to reduce the possibility of recombination with the duplicated regions of the chromosome or plasmid. However, this is obviously not a problem if the gene(s) is introduced into a strain in which the promoter (and/or gene sequence) in question has been previously deleted. We are currently developing a system that uses an inducible promoter that can be externally controlled by manipulations of the growth medium or the growth conditions. Examples of tightly regulated promoters that could be used are nirA (regulated by nitrogen source) (21) or sbtA (regulated by CO2 availability) (22). Although these have not yet been fully implemented for Synechococcus sp. PCC 7002, the lacI-lacO repression system functions effectively in Synechococcus elongatus sp. PCC 7942 (23) and a copper-regulated promoter has been used successfully with Nostoc sp. PCC 7120 (24, 25). Unfortunately, in neither of these examples is transcription completely attenuated under nonpermissive conditions. Depending on how well characterized a selected promoter might be, it may be wise to be conservative when deciding how much of the promoter region to include, as expression may be significantly enhanced by upstream “up elements.” Similarly, it may also be important to consider downstream cis-elements involved in translation of a gene after transcription. It may often be best to include the entire 5¢-untranslated region up to the first codon, along with a chosen promoter to achieve optimal expression. 12. Foreign DNA inserts of up to 14 kbp have successfully been introduced into pAQ1 using this system. No upper limit on fragment size has yet been determined.

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13. A final point for consideration in the construction of a gene expression system is the configuration of the site for insertion of the gene of interest. This should also include the consideration of any expression tag(s) that would allow for the detection or purification (or both) of the protein of interest. Depending on the downstream application, deca-histidine, strep- or flag-tags can greatly facilitate the detection (see Note 19) and purification of the protein of interest (see Note 10). Expression tags can often be incorporated into the PCR primers used for the amplification of a coding sequence of interest, but a more convenient option may be to integrate a gene insertion/expression site from a commercially available E. coli expression vector into the Synechococcus sp. PCC 7002 expression vector. If one utilizes the multicloning site of pET-16b or similar vector, then proteins can be expressed as His10-tagged variants if the forward primer for PCR is designed to include an NdeI site. Alternatively, untagged versions of the protein of interest can be produced if NcoI is selected as the N-terminal restriction enzyme. Another alternative is to introduce genes together with their native promoters. In this case, the pAQ1Ex-PcpcBA expression vector can be digested with EcoRI and BamHI to remove the promoter from the construct, and the DNA fragment of interest can be cloned in its place using these same enzymes. 14. Depending on the desired use of the created strain, confirmation of complete segregation may not be required because the created strain represents a gain of function for the cells. This differs from the situations in which genes are being insertionally inactivated, for which it is desirable (although not always achievable) to have complete segregation of alleles. Confirmation of complete segregation should be conducted using PCR before any mutant strain is cultivated in the absence of antibiotics in order to insure that the strain does not revert to wild-type copies of the plasmid target in question (e.g., pAQ1). 15. It is generally possible to perform PCR amplification of a region of interest from the genome of Synechococcus sp. PCC 7002 by using the template DNA available from a few cells scraped from a plate. The actual number of cells required to provide the DNA template for successful PCR amplification can be quite small. An amount of cells that is just barely visible to the eye, recovered on the end of a pipet tip, is usually sufficient. Alternatively, cells from a plate can be resuspended in sterile water (~20 mL, in a 1.5-mL microcentrifuge tube) at density at which the blue-green color is just barely visible. After boiling and centrifugation, the resulting solution can be

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used in place of purified DNA as a template for PCR. Experience indicates that these methods work best for smaller amplicons (<3 kb). 16. Freezing and storage of wild type and mutant strains of Synechococcus sp. PCC 7002 can be performed as previously described (29): harvest cells from a late-exponential phase culture by centrifugation, and resuspend the cells in fresh A+ medium containing 10 mM glycerol and 5% (v/v) methanol at one-tenth of the original culture volume. The cell suspension should be aliquoted into 1-mL samples in cryovials and frozen in a 5100 Cryo 1°C Freezing Container (“Mr. Frosty,” Nalgene #5100-0001, Rochester, NY) that allows for very slow freezing of the cells (around 1°C per minute). Cells may be revived by rapidly melting the contents of the cryovial in water at 35°C. The cells should be harvested by centrifugation, and resuspended in fresh A+ medium containing 10 mM glycerol. The culture should be incubated in the dark at 38°C for a day prior to incubation at low to moderate light intensity. 17. One method of determining if active protein products are being produced is to perform a biochemical assay for the gain of function conferred by the new protein. An example of  this is provided by our introduction of a gene with sequence ­similarity to crtG into Synechococcus sp. PCC 7002. In cyanobacteria, carotenoids function in energy transfer and photoprotection, and also play important roles structurally in biogenesis and stabilization of membranes and photosynthetic complexes (30). The crtG gene encodes a 2,2¢-b-ionone ring hydroxylase, and the encoded protein was anticipated to play a role in carotenoid biosynthesis in S. elongatus sp. PCC 7942 (31). No gene with sequence similarity to crtG is present in Synechococcus sp. PCC 7002. The introduction of this gene under the control of the aphII promoter into Synechococcus sp. PCC 7002, which synthesizes several carotenoids, including b-carotene, echinenone, cryptoxanthin, zeaxanthin, myxoxanthophyll, and newly identified synechoxanthin (32), caused no phenotype apparent to the unaided eye. However, it did alter the carotenoid content of the cells in the expected manner and led to the production of two new carotenoids, caloxanthin, and nostoxanthin, which are not normally synthesized (Fig. 4). Methods for pigment extraction and HPLC analysis have been described (32, 33). Pigments were extracted from washed cell pellets of Synechococcus sp. PCC 7002 strains by sonication in acetone:methanol (7:2, v/v) or methanol (100%). Extracts were routinely centrifuged (>12,000 × g) to remove debris and filtered through 0.2 mm Teflon (polytetrafluoroethylene) syringe filters.

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wild-type B

Absorbance (491nm)

S X E

M1

Z

B

+ CrtG N

S

C X E

M2 10

20

30

40

50

Elution Time (min) Fig. 4. Comparison of HPLC chromatograms of the carotenoids extracts from the cells of wild type and a transformant of Synechococcus sp. PCC 7002 that harbors pAQ1Ex-Pkan-crtG. The letters identifying specific carotenoids are defined as follows: S synechoxanthin, M1 myxol-2¢-fucoside, Z zeaxanthin, X cryptoxanthin, E echinenone, B b-carotene, M2 an unidentified myxoxanthophyll derivative, mostly likely 2-hydroxy-myxol-2¢-fucoside, N nostoxanthin, C caloxanthin.

Pigments were separated by HPLC (Agilent Model 1100) equipped with a diode array detector (model G1315B) and controlled with Agilent ChemStation software (Agilent Technologies, Palo Alto, CA) on a 25 cm by 4.6 mm analytical Discovery 5 mm C18 column (Supelco, Bellefonte, PA). 18. The expression of genes encoding proteins with readily discernable phenotypes makes it trivial to confirm protein synthesis (see Notes 9 and 17) and Figs. 3 and 4 concerning YFP and CrtG). However, if confirming the expression of the protein of interest is not trivial, and if the coding sequence for the gene of interest includes a hexa- or deca-His-tag or some other common epitope tag for which commercial antibodies are available, a simple way to check for transformants expressing the protein of interest is immunoblotting with antibodies directed against the introduced peptide tag. Wild-type cells or cells from a control transformation (e.g., the empty vector) should be used as a negative control for these experiments. (a) Harvest cells from a liquid culture (20–30 mL) grown photoautotrophically until late exponential phase are by centrifugation at 10,000 × g at 4°C in a Sorval SLC-1500 rotor. (b) Resuspend the cell pellet in 0.5 mL Bugbuster™ Master Mix cell lysis buffer. (c) Incubate the suspension on a shaking platform for 15 min at room temperature.

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(d) Prepare a cleared cell lysate by centrifugation at 14,000 × g. (e) Examine the preparation using polyacrylamide gel electrophoresis in the presence of sodium dodecylsulfate (SDS-PAGE) (20) and immunoblotting (27). If little is known about the potential solubility of the protein of interest, both the supernatant and pellet fractions from this centrifugation step should be used. Alternatively, larger scale cultures of these transformants can be grown, and the protein of interest can be isolated using Ni-NTA affinity chromatography to determine if the protein of interest is being produced (see example in Note 9). 19. If the expression of the introduced gene cannot be determined by immunoblotting, it may be instructive to confirm that the gene of interest has actually been successfully integrated at its desired target site. PCR is the most simple and effective way to verify the presence of the expression construct in transformant cell lines. As long as the amplicons from the transformant and wild-type cells are of sufficiently different sizes to be easily distinguishable after gel electrophoresis, primers used to produce the flanking sequences can be used to generate a product, whose size is indicative of successful transformation. Untransformed, wild-type cells should also be used as a negative control for these experiments. If PCR was used to produce the original transforming fragment, the gene of interest contained within the product amplified from the cyanobacterial cells overexpressing the gene of interest can be sequenced to determine if any unintended sequence errors were introduced during the transformation process. 20. If protein production cannot be confirmed according to the procedure in Note 18 above, but the gene is found to be present in the strain (see Note 19), reverse transcriptase-PCR (RT-PCR) experiments may be conducted to determine if the gene of interest is actually being transcribed. This can be done by using total RNA isolated from transformant cells as template(s) for the amplification. Untransformed WT cells, or cells transformed with an empty vector, can be used as the negative control for these experiments. If the desired transcripts are not detectable, additional isolates of the transformed Synechococcus sp. PCC 7002 should be assayed for expression because an expression-silencing mutation may have occurred in the particular isolate assayed. If none of the isolated transformants can be shown to express the introduced gene, alternative promoters may need to be considered (see Note 10) along with the possibility that the expression of the introduced gene may be strongly selected against because of deleterious effects conferred by its product.

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Acknowledgments This research has been partly supported by grant MCB-0519743 from the National Science Foundation as well as by subcontract 00001041 from Princeton University to D.A.B. The primary award for this subcontract is Multi-University Research Initiative (MURI) grant FA-9550-05-1-0365 from the U. S. Air Force Office of Scientific Research to Dr. G. Charles Dismukes, Department of Chemistry, Princeton University. References 1. Bryant, D. A., de Lorimier, R., Guglielmi, G., and Stevens, S. E. Jr. (1990) Structural and compositional analyses of the phycobilisomes of Synechococcus sp. PCC 7002. Analyses of the wild-type strain and a phycocyanin-less mutant constructed by interposon mutagenesis. Arch. Microbiol. 153, 550–560. 2. Zhou, J., Gasparich, G. E., Stirewalt, V. L., de Lorimier, R., and Bryant, D. A. (1992) The cpcE and cpcF genes of Synechococcus sp. PCC 7002: Construction and phenotypic characterization of interposon mutants. J. Biol. Chem. 267, 16138–16145. 3. Shen, G. and Bryant, D. A. (1995) Characterization of a Synechococcus sp. strain PCC 7002 mutant lacking Photosystem I. Protein assembly and energy distribution in the absence of the photosystem I reaction center core complex. Photosynth. Res. 44, 51–53. 4. Sakamoto, T. and Bryant, D. A. (1999) A novel nitrate/nitrite permease in the marine cyanobacterium Synechococcus sp. strain PCC 7002. J. Bacteriol. 181, 7363–7372. 5. Shen, G., Saunée, N. A., Williams, S. R., Gallo, E. F., Schluchter, W. M., and Bryant, D. A. (2006) Identification and characterization of a new class of bilin lyase: The cpcT gene encodes a bilin lyase responsible for attachment of phycocyanobilin to Cys-153 on the b-subunit of phycocyanin in Synechococcus sp. PCC 7002. J. Biol. Chem. 281, 17768–17778. 6. Maresca, J. A., Graham, J. E., Wu, M., Eisen, J. A., and Bryant, D. A. (2007) Identification of a fourth family of lycopene cyclases in photosynthetic bacteria. Proc. Natl. Acad. Sci. U.S.A. 104, 11784–11789. 7. Shen, G., Schluchter, W. M., and Bryant, D. A. (2008) Biogenesis of phycobiliproteins. I. cpcS-I and cpcU mutants of the cyanobacterium Synechococcus sp. PCC 7002 identify a

8.

9.

10.

11.

12.

13.

14.

heterodimeric phycocyanobilin lyase specific for b-phycocyanin and allophycocyanin subunits. J. Biol. Chem. 283, 7503–7512. Shen, G., Leonard, H. S., Schluchter, W. M., and Bryant, D. A. (2008) CpcM post-translationally methylates asparagine-71/72 of phycobiliprotein beta subunits in Synechococcus sp. PCC 7002 and Synechocystis sp. PCC 6803. J. Bacteriol. 190, 4808–4817. Jin, Z., Heinnickel, M., Krebs, C., Shen, G., Golbeck, J. H., and Bryant, D. A. (2008) Biogenesis of iron-sulfur clusters in photosystem I: Holo-NfuA from the cyanobacterium Synechococcus sp. PCC 7002 rapidly and efficiently transfers [4Fe-4S] clusters to apo-PsaC in vitro. J. Biol. Chem. 283, 28426–28435. Graham, J. E. and Bryant, D. A. (2009) The biosynthetic pathway for the synthesis of the myxol-2¢-fucoside in the cyanobacterium Synechococcus sp. strain PCC 7002. J. Bacteriol. 191, 3292–3300. Buzby, J. S., Porter, R. D., and Stevens, S. E., Jr. (1983) Plasmid transformation in Agmenellum quadruplicatum PR-6: Construction of biphasic  plasmids and characterization of their ­transformation properties. J. Bacteriol. 154, 1446–1450. Kimura, A., Hamada, T., Morita, E. H., and Hayashi, H. (2002) A high temperature-sensitive mutant of Synechococcus sp. PCC 7002 with modifications in the endogenous plasmid, pAQ1. Plant Cell Physiol. 43, 217–223. Akiyama, H., Kana, S., Hirano, M., and Miyasaka, H. (1998) Nucleotide sequences of plasmid pAQ1 of marine cyanobacterium Synechococcus sp. PCC 7002. DNA Res. 5, 127–129. Chen, X. and Widger, W. R. (1993) Physical genome map of the unicellular cyanobacterium Synechococcus sp. strain PCC 7002. J. Bacteriol. 175, 5106–5116.

Expression of Genes in Cyanobacteria: Adaptation of Endogenous Plasmids 15. Essich, E., Stevens, S. E., Jr., and Porter, R. D. (1990) Chromosomal transformation in the cyanobacterium Agmenellum quadruplicatum. J. Bacteriol. 172, 1916–1922. 16. Roberts, T. M. and Koths, K. E. (1976) The blue-green alga Agmenellum quadruplicatum contains covalently closed DNA circles. Cell 9, 551–557. 17. Porter, R. D. (1986) Transformation in cyanobacteria. Crit. Rev. Microbiol. 13, 111–132. 18. Frigaard, N. U., Sakuragi, Y., and Bryant, D. A. (2004) Gene inactivation in the cyanobacterium Synechococcus sp. PCC 7002 and the green sulfur bacterium Chlorobium tepidum using in vitro-made DNA constructs and natural transformation. Methods Mol. Biol. 274, 325–340. 19. Lambert, D. H. and Stevens, S. E., Jr. (1986) Photoheterotrophic growth of Agmenellum quadruplicatum PR-6. J. Bacteriol. 165, 654–656. 20. Laemmli, U. K. (1970) Cleavage of structural proteins during the assembly of the head of bacteriophage T4. Nature 227, 680–685. 21. Qi, Q., Hao, M., Ng, W. O., Slater, S. C., Baszis, S. R., Weiss, J. D., and Valentin, H. E. (2005) Application of the Synechococcus nirA promoter to establish an inducible expression system for engineering the Synechocystis tocopherol pathway. Appl. Environ. Microbiol. 71, 5678–5684. 22. Wang, H. L., Postier, B. L., and Burnap, R. L. (2004) Alterations in global patterns of gene expression in Synechocystis sp. PCC 6803 in response to inorganic carbon limitation and the inactivation of ndhR, a LysR family regulator. J. Biol. Chem. 279, 5739–5751. 23. Clerico, E. M., Ditty, J. L., and Golden, S. S. (2007) Specialized techniques for sitedirected mutagenesis in cyanobacteria. Methods Mol. Biol. 362, 155–171. 24. Ghassemian, M., Wong, B., Ferreira, F., Markley, J. L., and Straus, N. A. (1994) Cloning, sequencing and transcriptional studies of the genes for cytochrome c-553 and plastocyanin from Anabaena sp. PCC 7120. Microbiology 140, 1151–1159.

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25. Buikema, W. J. and Haselkorn, R. (2001) Expression of the Anabaena hetR gene from a copper-regulated promoter leads to heterocyst differentiation under repressing conditions. Proc. Natl. Acad. Sci. U.S.A. 98, 2729–2734 26. Stevens, S. E., Jr., Patterson, C. O. P., and Myers, J. (1973) The production of hydrogen peroxide by blue-green algae: A survey. J. Phycol. 9, 427–430 27. Shen, G., Zhao, J., Reimer, S. K., Antonkine, M. L., Cai, Q., Weiland, S. M., Golbeck, J. H., and Bryant, D. A. (2002) Assembly of Photosystem I: I. Inactivation of the rubA gene encoding a membrane-associated rubredoxin in the cyanobacterium Synechococcus sp. PCC 7002 causes a loss of photosystem I activity. J. Biol. Chem. 277, 20343–20354 28. Elhai, J. and Wolk, C. P. (1988) A versatile class of positive-selection vectors base on the nonviability of palindrome-containing plasmids that allows cloning into long polylinkers. Gene 68, 119–138. 29. Brand, J. J. (2003) Cryopreservation of cyanobacteria. http://www-cyanosite.bio. purdue.edu/protocols/cryo.html 30. Bryant, D. A. and Frigaard, N.-U. (2006) Prokaryotic photosynthesis and phototrophy illuminated. Trends Microbiol. 14, 488–496. 31. Iwai, M., Maoka, T., Ikeuchi, M., and Takaichi, S. (2008) 2,2’-beta-hydroxylase (CrtG) is involved in carotenogenesis of both nostoxanthin and 2-hydroxymyxol 2’-fucoside in Thermosynechococcus elongatus strain BP-1. Plant Cell Physiol. 49, 1678–1687. 32. Graham, J. E., Lecomte J. T. J., and Bryant, D.  A. (2008) Synechoxanthin, an aromatic C40 xanthophyll that is a major carotenoid in the cyanobacterium Synechococcus sp. PCC 7002. J. Nat. Prod. 71, 1647–1650. 33. Graham, J. E. and Bryant, D. A. (2008) The biosynthetic pathway for synechoxanthin, an aromatic carotenoid synthesized by the euryhaline, unicellular cyanobacterium Synechococcus sp. strain PCC 7002. J. Bacteriol. 190, 7966–7974.

Chapter 22 Construction of Gene Interruptions and Gene Deletions in the Cyanobacterium Synechocystis sp. Strain PCC 6803 Julian J. Eaton-Rye Abstract A series of protocols are presented for the storage, growth, transformation, and characterization of wild type and mutant strains of Synechocystis sp. strain PCC 6803. These protocols include the isolation of genomic DNA and the strategies required for the construction of specific gene interruptions or deletions in this organism. This cyanobacterium has been used widely as a model for photosynthesis research, and the sequence of its genome is available at CyanoBase (http://genome.kazusa.or.jp/cyanobase/). The details provided in this chapter do not assume any previous experience in working with cyanobacteria and are intended to enable new investigators to take advantage of a wide range of gene modification and mutation mapping techniques that have been adapted for use in this system. Key words: Cell culture, CyanoBase, DNA isolation, Growth curves, Mutagenesis, Oxygen evolution, Synechocystis

1. Introduction The cyanobacterium Synechocystis sp. strain PCC 6803 (hereafter Synechocystis PCC 6803) is a naturally transformable ­Gram-negative prokaryote that performs oxygenic photosynthesis (1, 2). Transformation is achieved by the uptake of DNA by the type IV pili and incorporation into the host genome by homologous double recombination (3, 4). A glucose-tolerant strain has been developed that is able to grow photoheterotrophically in the presence of photosystem (PS) II-specific herbicides, such as atrazine or diuron, thereby facilitating the study of PSII through oligonucleotidedirected mutagenesis (4, 5). The widespread use of Synechocystis PCC 6803 as a model system led to its selection as the first photoautotrophic organism to have its genome sequenced,

Robert Carpentier (ed.), Photosynthesis Research Protocols, Methods in Molecular Biology, vol. 684, DOI 10.1007/978-1-60761-925-3_22, © Springer Science+Business Media, LLC 2011

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and this information is accessible through CyanoBase at: http:// genome.kazusa.or.jp/cyanobase/ (6). The substrain selected for genomic sequencing is glucose-tolerant, but is noncompetent for transformation with exogenous DNA. A discussion of the different substrains together with an overview of the genomic structures and functional genomics of Synechocystis PCC 6803 is provided in ref. 7. The objective of the present chapter is to describe the basic protocols for growing this organism, constructing gene interruptions or deletions, and confirming the genomic and basic phenotype of the resulting mutants. The details provided assume no previous experience working with cyanobacteria, but they are intended to provide a series of protocols that may be used for the successful application of more advanced mutagenesis studies with this model organism.

2. Materials 2.1. Cell Culture

1. Growth flasks (see Fig. 1). 2. Cole Palmer Masterflex tubing (CP-96400-25).

Fig. 1. A modified Erlenmeyer flask used to culture Synechocystis PCC 6803 cells based on an original design described by John G. K. Williams in ref. 4. The 0.2 mm Millipore filter is connected to a glass capillary tube with an outer diameter of 6.0 mm and an inner diameter of 1.7 mm that stops 10 mm from the bottom of the flask. The connection is made with Cole Palmer Masterflex tubing. The sidearm is made with glass tubing that has a 20 mm outer diameter, and it is covered with a stainless steel culture-tube cap. This whole unit is autoclaved before use.

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Table 1 Composition of stock solutions for BG-11 liquid medium (1) Stock

Ingredients

Amount

100× BG-11

NaNO3 MgSO4 ∙ 7H2O CaCl2 ∙ 2H2O Citric acid 0.25 M NaEDTA, pH 8.0 Trace minerals Deionized water

149.60 g/L 7.49 g/L 3.60 g/L 0.60 g/L 1.12 mL/L 100.00 mL/L up to 1 L

Trace minerals

H3BO3 MnCl2 ∙ 4H2O ZnSO4 ∙ 7H2O Na2MoO4 ∙ 2H2O CuSO4 ∙ 5H2O Co(NO3)2 ∙ 6H2O Deionized water

2.860 g/L 1.810 g/L 0.222 g/L 0.390 g/L 0.079 g/L 0.0494 g/L Up to 1 L

1,000×

Ammonium iron(III) Citrate, browna

0.60 g/100 mL

1,000×

Na2CO3

2.00 g/100 mL

1,000×

K2HPO4

3.05 g/100 mL

Also known as ferric ammonium citrate

a

3. Millipore 0.2 mm filters (Millex FG50: MISLFG-050-10). 4. Aquarium pump(s). 5. BG-11 stock solutions as shown in Table 1. 6. Glucose, herbicide, and antibiotic stock solutions as shown in Table 2. 7. Antibiotic-resistance cassettes as shown in Table 3. These can be obtained from the author upon request. 8. Sodium thiosulfate. 9. 10 mM TES-NaOH (pH 8.2). 10. Bacteriological agar. 11. Disposable 1 mL sterile pipets. 12. Parafilm (see Note 1). 13. Laminar-flow hood. 14. Growth cabinet (see Note 2).

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Table 2 Additions for BG-11 liquid medium Solution

Stock

Final concentration

Notes

Glucose

1 M

5 mM

Autoclave; store at 4°C

20 mM

20 mM

Make in methanol; store at –20°C

20 mM

20 mM

Make in ethanol; store at –20°C

Chloramphenicol

50 mg/mL

25 mg/mL

Make in ethanol; store at –20°C

Erythromycin

25 mg/mL

25 mg/mL

Make in ethanol; store at –20°C

Kanamycin

50 mg/mL

25 mg/mL

Filter-sterilize; store at –20 or 4°C

Spectinomycin

50 mg/mL

25 mg/mL

Filter-sterilize; store at –20 or 4°C

Atrazine Diuron

a

b

2-chloro-4-ethylamino-6-isopropylamino-s-triazine 3,4-dichloro-1,1-dimethyl urea (DCMU)

a

b

Table 3 Antibiotic-resistance markers

2.2. The Use of Cyanobase 2.3. Cloning Sequences Obtained from Cyanobase 2.4. Transformation of Synechocystis PCC 6803

Resistance marker

Plasmid from which cassette was derived

Accession or reference

Chloramphenicol

pBR325

L08855

Erythromycin

pE194

J01755

Kanamycin

pUC4K (Tn 903)

X06404

Spectinomycin

pHP45Ω

K02163

Software packages for primer design and restriction mapping. pUC or pBR derivatives as cloning vectors: pT7Blue-T (Novagen, Madison, WI, USA), pUC19 (New England BioLabs, Beverly, MA, USA), pBluescript (Stratagene, La Jolla, CA, USA), and the pGEM-T (Promega, Madison, WI, USA) vector systems. 1. Sterile test tubes with culture-tube caps. 2. Gloves. 3. 70% Ethanol. 4. Scissors and small forceps. 5. Whatman Nuclepore polycarbonate membrane (Whatman catalog number: WHA111107). 6. Inoculation loop for streaking plates.

circles

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1. 300-mL centrifuge bottles. 2. Sodium iodide. 3. Tris buffer 1:50 mM Tris–HCl, pH 8.0, 5 mM EDTA, and 50 mM NaCl. 4. Lysozyme. 5. N-Lauryl sarcosine. 6. 45-mL screw-capped phenol-resistant polypropylene centrifuge tubes. 7. Tris–HCl, pH > 7.8, equilibrated phenol. 8. Chloroform (containing 1/25 isoamylalcohol). 9. 3 M Sodium acetate, pH 5.0. 10. Absolute ethanol. 11. 70% Ethanol. 12. TE buffer: 10 mM Tris–HCl, pH 8.0, and 1 mM EDTA. 13. Speedvac (Savant, Holbrook, NY, USA) or equivalent instrument.

2.6. Basic Phenotypic Characterization

1. Clark-type oxygen electrode with FLS1 (or equivalent) light source by Hansatech (King’s Lyn, Norfolk, UK). 2. Melles Griot OG 590 sharp cutoff red glass filter. 3. Neutral density filter set. 4. BG-11: 25 mM HEPES (pH 7.5). 5. 1 M Sodium bicarbonate (NaHCO3–). 6. 100 mM Methyl viologen. 7. Ascorbic acid. 8. 2,3,5,6-Tetramethyl-p-phenylenediamine (TMPD). 9. 250 mM Potassium ferricyanide. 10. 50 mM 2,5-dimethyl-p-benzoquinone. 11. 3-(3,4-dichlorophenyl)-1,1-dimethylurea (Diuron).

3. Methods 3.1. Cell Culture of Synechocystis PCC 6803

The wild type can be obtained directly from the Pasteur Culture Collection (see Note 3) and the glucose-tolerant strain, utilized in PSII studies, is available on request from the author. The cells are maintained on BG-11 plates at 30°C (see Note 4). 1. The stock solutions required to make BG-11 medium are given in Table 1. 2. To obtain 1 L of BG 11, 987 mL of deionized H2O is combined with 10 mL of 100× BG 11 and 1 mL each of 1,000×

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ammonium iron (III) citrate brown, 1,000× Na2CO3, and 1,000× K2HPO4; however, for plates also add 15.0 g/L agar, 10 mL of TES-NaOH buffer, and 3 g/L sodium thiosulphate (see Note 5). 3. After autoclaving, allow the medium to cool to 55°C before adding glucose, herbicides, or antibiotics if these are required. 4. The plates should be poured and dried in a laminar-flow hood and cyanobacterial cells should be spread using standard microbiological sterile techniques (see Note 6). 5. The plates should be placed in a growth chamber under constant illumination at 15–25 mE/m2s (see Note 2). 6. To avoid condensation the plates should be placed right-side up and to minimize contamination, they can be wrapped with parafilm (see Note 1). For successful liquid cultures modified Erlenmeyer flasks are recommended as shown in Fig.  1. Modified 200 and 500  mL flasks are usually the most useful but 2 L flasks work well when large cultures are required. 1. Perform inoculation and sampling of flasks in a laminar-flow hood taking all possible care to maintain sterile conditions. 2. Scrape cells from a BG-11 plate using a disposable 1 mL pipet and introduce into the culture flask so that several clumps of cells are clearly visible. 3. Liquid cultures are bubbled using an aquarium pump to pass air through a Millipore 0.2 mM filter; however, before turning on the pump cultures should be left to stand for at least 2 h (an overnight wait is also suitable). This procedure facilitates the growth of the cultures. 4. Cell growth is monitored by measuring the turbidity or optical density (OD) of the culture at 730 nm. 5. Liquid cultures should not be grown beyond an OD730 nm of 2.0 if the culture is to be diluted for further use. To avoid inaccuracy when determining the OD730 nm of a culture, it should be diluted to an OD730 nm < 0.4 to prevent inaccuracies resulting from light scattering (NB. an OD730 nm of 0.25 = 1 × 108 cells/ mL) (4). 6. When glucose, herbicides, or antibiotics are present in liquid media and plates, they should be added at the final concentrations supplied in Table 2. The antibiotic-resistance cassettes for resistance to the antibiotics in Table 2 are derived from the plasmids given in Table 3. 7. The wild type and mutant cultures can be stored at −80°C in sterile BG-11 containing 15% glycerol.

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3.2.1. Obtaining Sequence from CyanoBase

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The initial step in the construction of a gene interruption or deletion is to obtain the sequence of the target gene from CyanoBase for the design of PCR primers and the construction of a restriction site map of the intended PCR product. 1. In an Internet browser enter: http://genome.kazusa.or.jp/ cyanobase/. 2. In CyanoBase select the Synechocystis PCC 6803 genome by clicking on “Synechocystis sp. PCC 6803” in the Species column. In the next window that opens, type the name of the gene you wish to interrupt or delete into the box and press Search. If you do not know the name of your gene, perform a BLAST search (see Note 7). 3. CyanoBase will respond by providing the unique identifier for this gene in the genome and by clicking on this number a detailed summary of the information available on this gene will be displayed. The nomenclature used to identify specific genes is explained in the notes (see Note 8). 4. To ensure that sufficient sequence is obtained for efficient homologous recombination, select 1 kb upstream and downstream of the gene by entering the values into the dialog boxes and press Retrieve DNA sequence (see Note 9). 5. Save the nucleotide sequence obtained in a format that will enable the sequence to be entered into a standard package that will give a restriction map. 6. Exit CyanoBase.

3.3. Cloning Sequence Obtained from CyanoBase: The Construction of Plasmids for the Introduction of Mutations into Synechocystis PCC 6803 Cells 3.3.1. Designing PCR Primers

In this section, the design of PCR primers and the selection of a suitable vector for cloning the sequence selected from CyanoBase is explained. Primers can be designed for the sequence obtained from CyanoBase by entering it into a program for primer design or alternatively primers can be designed by eye. Typically, primers should be ~ 20 nucleotides long and the 3¢ end should contain two or three G or C bases. All standard criteria for designing primers apply, such as avoiding sequences that can pair within a primer, primer dimers, and maintaining equal G/C content in both primers of a pair. If the objective is to inactivate the target gene, the restriction map of the sequence should be examined to identify a suitable site within the open-reading frame (ORF) that can be utilized for cloning in an antibiotic-resistance cassette for gene interruption and subsequent selection of the cyanobacterial transformant (see Note 10). Flanking primers should be designed with a view to keeping the final product less than 2.5  kb to facilitate cloning. The primers may be designed across restriction sites if this will

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simplify cloning or restriction sites can be introduced, although in the latter case longer primers may be necessary. If the objective is to delete the target gene, primers should be designed to PCR the upstream and downstream regions on either side of the ORF. Both products can then be separately cloned into suitable vectors (see Subheading  3.3.2) and an antibioticresistance cassette then cloned at either the 3¢ end of the upstream product or the 5¢ end of the downstream product. Standard cloning techniques can then be used to join the fragments utilizing the antibiotic selection. The resulting construct, therefore, has the upstream and downstream sequences flanking the antibioticresistance marker and when introduced into Synechocystis PCC 6803 cells, this construct will replace the target ORF with the antibiotic-resistance cassette as a result of double homologous recombination. As before, to facilitate the cloning steps, suitable restriction sites can be incorporated into the primer design, but as noted above, primers may need to be extended (e.g., up to ~ 30 nucleotides) to facilitate annealing since the introduced substitution(s) will create a mismatch with the genomic DNA template. 3.3.2. Vector Selection

Synechocystis PCC 6803 can be easily transformed with pT7Blue-T, pUC19, pBluescript, and the pGEM-T vector systems. If the required restriction site for cloning the antibiotic-resistance marker in the ORF is also present in the polylinker of the plasmid, this can be initially removed by cutting and blunting the site, followed by religation of the plasmid, before cloning in the PCR insert. It is also important to ascertain if any of the flanking DNA that will be used for recombination contains any sequence for neighboring genes and an appropriate PCR enzyme, with or without proofreading, should be chosen. In the simplest cases, the use of T/A cloning into pGEM-T or pGEM-T Easy is recommended.

3.4. Transformation of Synechocystis PCC 6803

Once the construct to interrupt or delete the gene of interest has been assembled the corresponding interruption mutant or deletion mutant, respectively, is created by the transformation of the plasmid DNA into Synechocystis PCC 6803 using the protocol outlined in this section.

3.4.1. Cyanobacterial Transformation

For this procedure, ensure that all work surfaces and equipment are sterile (see Note 11). 1. Start a fresh cyanobacterial culture in BG-11, adding glucose and antibiotics as appropriate. Grow for 2–3 days, but do not let the OD730 nm increase above 0.5 as the transformation efficiency declines in late log phase. 2. In a laminar-flow hood, transfer the cells from the flask to 50  mL sterile Falcon tubes and spin down cells at approx. 2,760 × g for 5 min at room temperature.

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3. Discard the supernatant and resuspend the pellet in 2  mL BG-11 medium and measure the OD730 nm (see Note 12). 4. Each transformation will be performed using a 0.5 mL volume of the cells at an OD730 nm of 2.5. Dilute sufficient cells to an OD730 nm of 2.5 using BG-11 medium (see Note 13). 5. Aliquot 0.5  mL volumes of cells (at an OD730  nm = 2.5) into sterile test tubes with culture-tube caps. 6. Add 2–10 mg of DNA in 2–10 mL of TE buffer to each tube and shake. However, leave one tube without the addition of DNA as a control. 7. Place tubes in a rack in a growth chamber at 30°C for 6 h in the light and shake the tubes once at 3 h. 8. Place the sterile Whatman Nuclepore polycarbonate membrane circles (referred to hereafter as membranes) onto BG-11 plates that do not contain antibiotics in a laminar-flow hood and plate  200  mL of the cells onto the membranes. Allow membranes to dry before transferring plates to the growth room (see Note 14). 9. Grow for 12 h and then transfer the membranes to a BG-11 plate that contains the appropriate antibiotic at half the concentration listed in Table 2. After 3 days, transfer the membranes onto BG-11 plates containing the appropriate antibiotic at the concentration given in Table 2 (see Note 14). 10. Colonies should appear after 1 or 2 weeks. Pick colonies and streak a single colony on a BG-11 plate containing the appropriate additions (e.g., antibiotics) as shown in Table  2 (see Notes 14 and 15). 11. To ensure segregation of the transformants restreaks should be performed at approximately weekly intervals for 3 or 4 weeks. PSII-specific mutants are normally fully segregated after this time, but other mutations may require additional restreaking for several months. It should be noted that a failure to segregate implies that an essential gene has been targeted for disruption. 3.5. Verification of Mutant Genotype

Because Synechocystis PCC 6803 cells contain approx. ten copies of their genome, it is essential to confirm that all copies contain the mutation. The most rapid method for establishing the complete segregation of gene interruptions or deletions is by colony PCR. However, this method may not always work and PCR from isolated genomic DNA may be preferred (see Note 16). Alternatively, a Southern blot may be performed on isolated genomic DNA. In this section, the methods for colony PCR screening and isolating genomic DNA are presented (see Note 17). For Southern blotting, the large-scale genomic DNA isolation procedure is recommended.

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3.5.1. Colony PCR

1. Make up a 50 mL PCR reaction mix in a microfuge tube, but do not add the polymerase enzyme at this step. 2. Using an inoculation loop, add a cyanobacterial colony to the PCR reaction mix and disperse with low-speed vortexing. 3. Collect the sample with a very short (< 2 s) spin in a microfuge. 4. Add the polymerase enzyme and gently mix the sample with the pipet tip. Run the PCR reaction. 5. After the PCR reaction, spin for 30 s at maximum speed in a microfuge (typically approx. 13,600 × g). Next, transfer the supernatant to a clean microfuge tube and analyze the products on an agarose gel.

3.5.2. Rapid Small-Scale Genomic DNA Isolation from Plate Cultures

1. Scrape all the cells from a 3-week-old plate, resuspend in 200 mL of saturated NaI (2 g/mL) in a microfuge tube, and incubate at 37°C for 10 min. 2. Add 1 mL distilled deionized (dd) H2O to dilute the NaI and centrifuge at 13,600 × g for 10 min. 3. Discard the supernatant and resuspend the pellet by pipeting with a 1 mL pipet in 400 mL of Tris buffer 1. 4. Add 100 mL of a 50 mg/mL lysozyme solution and incubate for 10 min at 37°C followed by the addition of 100 mL of a 10% N-lauryl sarcosine solution. 5. Gently mix the tube by inversion and incubate at 37°C for 10 min. 6. Add 600 mL of Tris–HCl (pH > 7.8) equilibrated phenol and mix gently on a rotating wheel for 20 min. 7. Centrifuge at 13,600 × g for 10 min, and transfer the upper aqueous phase to a new tube using a 1 mL pipet with the end removed to avoid shearing the DNA. 8. To this new tube, add 600  mL of chloroform solution and gently mix on a rotating wheel for 20 min. 9. Transfer the upper aqueous phase, as in step 7, to a clean tube and ethanol precipitate the DNA by adding a one-tenth volume of sodium acetate solution and two and a half volumes of absolute ethanol. Place the tube at –20°C for 3 h, and then collect the DNA by centrifugation at 13,600 × g for 20 min at 4°C. 10. Discard the supernatant and wash the pellet in chilled 70% ethanol (−20°C) to remove salt. Centrifuge at 13,600 × g for 5 min and aspirate off the supernatant. Dry the pellet at room temperature and then resuspend in 50 mL of TE buffer and store at –20°C.

3.5.3. Large-Scale Genomic DNA Extraction from Liquid Culture

1. Weigh a 300-mL centrifuge bottle, add 75–250  mL of cell culture, and centrifuge at 7,000 × g for 5  min. Remove the

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supernatant and weigh the bottle again. Determine the weight of the cell paste (see Note 18). 2. Resuspend the cell paste in saturated NaI: for 1 g of paste use 2 mL of saturated NaI (2 g NaI per mL H2O) and incubate at 37°C for 20 min. 3. Fill up the centrifuge bottle with water to dilute the NaI. 4. Centrifuge for 10 min at 2,760 × g at 4°C, and then pour off the supernatant. 5. Resuspend the pellet in 8 mL of a solution of Tris buffer 1 per gram of cells. Resuspend completely by passing through an 18-gage needle. 6. Per gram of cells, add 1.25 mL of a 50 mg/mL solution of lysozyme and mix gently. Incubate at 37°C for 20 min (see Note 19). 7. Add 1 mL of 10% N-lauryl sarcosine (per gram cells), invert tube gently to mix, and then incubate at 37°C for 20 min. 8. Put lysate into a 40 mL phenol-resistant screw-capped plastic tube, add an equal volume of Tris–HCl (pH > 7.8) equilibrated phenol, and place parafilm around the lid to prevent leakage (see Note 20). 9. Mix gently on a wheel for 60  min and then centrifuge at 2,760 × g for 10 min at 4°C. 10. Use a pipet with a 5 mL tip (to prevent shearing of the DNA) to remove the aqueous (upper) phase, which should be pink, and place in a clean phenol-resistant tube. 11. Add an equal volume of chloroform solution to the aqueous phase, put parafilm around the lid and mix gently on a wheel for 45 min. 12. Centrifuge at 2,760 × g for 10 min at 4°C and transfer the top aqueous phase to another clean phenol-resistant centrifuge tube. For high quality DNA, it is recommended to repeat steps 8–12 before proceeding to step 13. 13. Ethanol precipitate the DNA by adding a one tenth volume of sodium acetate and two and a half volumes of absolute ethanol. Shake gently and place tube at −20°C overnight. 14. Centrifuge at 23,000 × g for 15 min at 4°C and then aspirate off the ethanol. 15. Wash the pellet with chilled 70% ethanol (−20°C), centrifuge at 2,760 × g for 5  min at 4°C and aspirate off the ethanol. 16. Put parafilm over the top of the tube and puncture several times with a pin. Dry in a Speedvac. 17. Resuspend the pellet in 300  mL of TE buffer and store at –20°C in a microfuge tube (see Note 21).

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3.6. Basic Phenotypic Characterization: Measurements of Growth Curves and Oxygen Evolution in Whole Cells 3.6.1. Photoautotrophic Growth Curves

Although the appropriate phenotypic characterization will depend on the specific mutations that have been introduced, growth curves and oxygen evolution assays provide a simple assessment of photosynthetic performance in mutant strains. Both of these measurements are described in this section. 1. For each strain, grow a starter culture to an OD730 nm of ~ 0.8. As appropriate these cultures may contain glucose and antibiotics (see Note 22). 2. Working in a laminar-flow hood pour 40  mL of the starter culture into a 50 mL sterile Falcon tube and spin-down the cells at 2,760 × g for 5 min at room temperature. 3. Ensuring that sterile conditions are maintained, decant the supernatant and resuspend the pellet using a vortex mixer in 5 mL of BG-11. Next, top-up the Falcon tube with 40 mL of BG-11 medium and spin-down the cells as in step 2. 4. Repeat the wash in step 3 two further times to remove any traces of glucose. 5. After the final wash, resuspend the pellet in 5 mL of BG-11 using a vortex mixer and then, taking care to maintain sterile conditions, determine the OD730 nm of the cells (typically a 1 in 100 dilution is required to obtain an accurate reading). 6. Prepare culture flasks by pouring in 150 mL of BG-11 and adding the appropriate antibiotics. Next, add the cells so that the starting OD730 nm in the flask is approx. 0.05. Measure the exact OD730 nm and record this as the zero time point. 7. Attach the flasks to an aquarium pump and ensure that all cultures are bubbling evenly. Measure the OD730 nm of the cultures every 12 or 24 h for 7 days (see Note 23).

3.6.2. Oxygen Evolution

The overall rate of photosynthesis, the rate of photosynthetic electron transport through the thylakoid membrane, as well as the activities of both photosystems can be easily assayed using a Clark-type oxygen electrode and an FLS1 (or equivalent) light source. Saturating actinic light is required for these measurements, and this can be experimentally determined by measuring the activity as a function of light intensity with the aid of neutral density filters. It is recommended that the actinic light be passed through a Melles Griot OG 590 sharp cutoff red glass filter as this reduces photoinactivation of the samples. Activity measurements should be performed at 30°C in BG-11solution. The chlorophyll a concentration should be between 10 and 20 mg/mL and the chlorophyll a should be extracted with methanol. The chlorophyll a determinations can be performed according to MacKinney (8). The overall rate of photosynthesis can be measure by adding 10 mM NaHCO3− from the fresh 1 M stock solution. To measure

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PSII activity, the electron acceptor should be 1.0 mM K3Fe(CN)6 with 0.2  mM 2,5-dimethyl-p-benzoquinone added to mediate between PSII and the nonpenetrating K3Fe(CN)6 (see Note 24). Whole chain electron transport can be assayed in isolated thylakoid membranes (see Note 25) by adding 2  mM methyl viologen from the methyl viologen stock solution. This reaction will measure oxygen uptake and the reader is referred to ref. 9 for a complete discussion of the relationship between oxygen uptake and electron transport. A PSI partial reaction can be measured in the presence of 1  mM ascorbic acid and 1  mM 2,3,5,6-tetramethyl-p-phenylenediamine as electron donor (see Note 24). For this reaction, 2 mM methyl viologen should be present as the electron acceptor and 20  mM diuron should be present to block PSII activity. 3.7. Extending the Mutagenesis System: Information on Additional Methods to Expand the Utility of This System

The flexibility of Synechocystis PCC 6803 as a model organism for the study of gene knockouts is not restricted to the antibioticresistance markers identified in Table 3. By using combinations of the antibiotic-resistance cassettes multiple genes can be inactivated or deleted in a single strain (10–12). Additionally, markerless deletions can be obtained in a two-step process using the sacB gene isolated from Bacillus subtilis. This gene encodes levansucrase which catalyzes the breakdown of sucrose to glucose and fructose derivatives (13). Expression of this gene has been shown to be lethal to Anabaena PCC 7120 and Synechocystis PCC 6803 (14, 15). In the first transformation sacB, in tandom with a selectable marker, is introduced into the cyanobacterium with flanking sequences so as to replace the target gene. The resulting strain is both resistant to the antibiotic but sensitive to growth on 5% sucrose. In the second step, the strain is transformed with a construct that contains only the flanking sequences upstream and downstream of the inserted sacB and antibiotic-resistance cassette. The markerless deletion mutant obtained is able to grow on sucrose, but is sensitive to the antibiotic. Synechocystis PCC 6803 has also been widely used for protein structure/function studies through the application of oligonucleotidedirected mutagenesis. For these experiments, the target gene must first be replaced by an antibiotic-resistance cassette and full segregation demonstrated so as to ensure that all copies of the gene have been deleted. To introduce mutations, the target gene, together with an antibiotic-resistance cassette and flanking upstream and downstream sequences, is cloned into a vector, such as pGEM-T, that can be used for the generation of ssDNA. The resulting construct is transformed into Escherichia coli strain CJ236. This strain is dUTPase and uracil-N-glycosylase deficient (dut−ung−) and therefore allows the stable incorporation of dU rather than dT into the single-stranded template. After the template has been used for in vitro mutagenesis reactions, it is transformed

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into a dut+ung+ strain of E. coli, thus selecting against the parental strand. This procedure positively selects for the form of the plasmid containing the mutated gene, and this can then be reintroduced into the Synechocystis PCC 6803 strain, in which the target gene was deleted, thus creating the desired mutant. Details of this procedure can be found in (16–18) and examples of their specific use with Synechocystis PCC 6803 may be found in (19, 20) (see Note 26). Further details of a range of mutagenesis-based strategies to study protein function in Synechocystis PCC 6803 can be found in ref. 5.

4. Notes 1. Parafilm may restrict gas exchange and limit the growth of the cyanobacterium. A commercially available Saran foodwrapping product can also be used. 2. Lighting can be provided by fluorescent tubes or metal halide bulbs. 3. Strain can be requested from the Pasteur Culture Collection by email at [email protected] and the Website is http://www. pasteur.fr. 4. Synechocystis PCC 6803 is a mesophilic bacterium and is able to grow optimally between 30 and 34°C. 5. When cooled to room temperature a brown precipitate is formed and the bottle should be shaken before use. When 10 mL of 1 M TES-NaOH (pH 8.2) is added the amount of deionized water should be adjusted accordingly. The organism is basophilic and grows well between pH 8.0 and pH 10.0. 6. BG-11 plates can be stored at 4°C; however, when antibiotics are present storage periods should not exceed 4 weeks. 7. In CyanoBase, you can perform a protein or nucleotide BLAST search by pasting in your query sequence, and this will identify the corresponding gene(s) in Synechocystis PCC 6803. To perform the BLAST search scroll to the Tools at the bottom of the window that opens when you initially type: http://genome.kazusa.or.jp/cyanobase/. 8. The nomenclature used to identify genes in CyanoBase begins with three letters followed by four numerals. The initial letter stands for the species and the second letter indicates gene length, where “l” indicates longer than 100 codons and “s” less than 100 codons. The third letter indicates the direction of transcription on the circular genome map (“r” for right and “l” for left). The number corresponds to the sequential order of the putative ORFs.

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9. When requesting nucleotide sequence care must be taken with respect to the direction of the gene given under the “Genome context” information. If the direction is complement, then the nucleotide sequence will initiate with the larger number and terminate with a smaller number. 10. Ideally, a single unique restriction site is selected for the introduction of an antibiotic-resistance cassette, but if this is not possible, a combination that results in an internal deletion of the gene may also be used providing that at least 0.4  kb is available for recombination at both ends. 11. Wash the laminar flow down with 70% ethanol, ethanol and flame your scissors and forceps, wear gloves when handling the filters (even rub ethanol over your gloves first). The Falcon tubes and test tubes used must also be sterile. 12. To accurately determine the OD730 nm, dilute an aliquot so the reading of the diluted sample is < 0.4. 13. For high transformation efficiencies, the OD730 nm of the cells to be transformed should be between 2.0 and 3.0. 14. When using the glucose tolerant strain to study mutations at the level of photosynthetic electron transport 5 mM glucose should be present in the BG-11 plates during the first 12 h. Subsequently, if specifically studying mutations in PSII proteins, 5 mM glucose and 20 mM atrazine or diuron should be present in the plates to remove any selection pressure for the formation of PSII-specific revertants. 15. It is recommended that each BG-11 plate be divided into four quarters and four colonies picked with one streaked out in each quadrant. These can then undergo subsequent restreaks so that four replicates of each transformation are brought through the segregation steps. 16. The procedure in Subheading  3.5.1 for colony PCR may be enhanced by initially resuspending the colony in 200  mL TE containing 1% Triton X-100. The Triton X-100 can be subsequently removed by one or two chloroform extractions and a 5–10 mL aliquot of the upper aqueous phase added to the PCR reaction. Heating the sample to 95°C for 2–4  min prior to extracting with chloroform has been carried out in some labs (e.g., http://microbiology.ucdavis.edu/meekslab/xpro6.htm). 17. In addition to the methods for genomic DNA isolation given in Subheading 3.5, a rapid method for the isolation of DNA from cells on plates utilizing glass beads to break the cells is given in ref. 5. 18. For a maximum yield of genomic DNA, do not let the culture grow beyond an OD730  nm of 3.0 before commencing this procedure. In this first step, centrifuge bottles compatible

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with either the Sorvall GSA or Beckman JA14 rotors are recommended. 19. Alternatively at step 6 in Subheading 3.5.3, dry lysozyme can be added at 10 mg/mL followed by incubation at 37°C until lysis is visible. This is checked at 30 min intervals by taking a 100 mL aliquot and adding SDS to a final concentration of 1%. The tube is then vortexed intermittently for 5 s. Lysed cells stick to the bottom of the tube when it is inverted. Following lysis, proteinase K, and SDS are added to the suspension at a final concentration of 200  mg/mL and 0.5%, respectively. The sample is then incubated at 60°C for 30 min, brought to an SDS concentration of 1% and incubated at 37°C for a further 20 min before proceeding to step 8. This procedure can substantially increase the yield of genomic DNA (S. Ermakova-Gerdes, personal communication). 20. Phenol-resistant polypropylene screw-capped tubes compatible with either the Sorval GSA or Beckman JA17 rotors are recommended for steps 8–14 in Subheading 3.5.3. 21. An RNAase A treatment at 30 mg/mL at 37°C for 30 min is optional and will depend on the intended application. 22. In the case of the glucose tolerant strain of Synechocystis PCC 6803, steps 1–7 of the procedure outlined in Subheading 3.6.1 can be adapted for mixotrophic growth curves in the presence of 5  mM glucose or heterotrophic growth curves where, in addition to 5 mM glucose, 20 mM atrazine or diuron are added. 23. Before taking a reading, it is necessary to top-up the culture to 150 mL with sterile ddH2O to correct for evaporation. In addition, when the culture exceeds an OD730 nm of 0.4, it is necessary to dilute the sample to obtain an accurate reading. 24. The stock solutions for 2,5-dimethyl-p-benzoquinone and 2,3,5,6-tetramethyl-p-phenylenediamine are made up in ethanol. 25. A simple extraction method for the isolation of thylakoid membranes is described in ref. 21. 26. Particularly in the case of membrane-spanning proteins, the use of sequence containing the promoter for homologous recombination should be avoided. Synechocystis PCC 6803 promoters may be recognized in E. coli and the expression of genes encoding hydrophobic foreign proteins may impose a selective pressure to inactivate expression. Thus, upon reintroduction of the mutated gene into Synechocystis PCC 6803, the gene may be found to have been silenced. This problem can be circumvented by designing separate deletion-mutant strains for the 5¢ and 3¢ halves of the target gene.

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Acknowledgment The author wishes to thank Professor Wim Vermaas as many of the procedures described in this chapter are based on methods that the author originally acquired in the Vermaas laboratory at Arizona State University. The methods for the isolation of genomic DNA are modified from an original procedure designed by Dr. J. G. K. Williams. References 1. Rippka, R., Deruelles, J., Waterbury, J. B., Herdman, M., and Stanier, R. Y. (1979) Generic assignments, strain histories and properties of pure cultures of cyanobacteria. J. Gen. Microbiol. 111, 1–61. 2. Grigorieva, G. and Shestakov, S. (1982) Transformation in the cyanobacterium Synechocystis sp. 6803. FEMS Microbiol. Lett. 13, 367–370. 3. Yoshihara, S., Geng, X. X., Okamoto, S., et al. (2001) Mutational analysis of genes involved in pilus structure, motility, and transformation competency in the unicellular motile cyanobacterium Synechocystis sp. PCC 6803. Plant Cell Physiol. 42, 63–73. 4. Williams, J. G. K. (1988) Construction of specific mutations in photosystem II photosynthetic reaction center by genetic engineering methods in Synechocystis 6803. Meth. Enzymol. 167, 766–778. 5. Vermaas, W. F. J. (1998) Gene modifications and mutation mapping to study the function of photosystem II. Meth. Enzymol. 297, 293–311. 6. Kaneko, T., Sato, S., Kotani, H., et al. (1996) Sequence analysis of the genome of the unicellular cyanobacterium Synechocystis sp. strain PCC6803. II. Sequence determination of the entire genome and assignment of potential protein-coding regions. DNA Res. 3, 109–136. 7. Ikeuchi, M. and Tabata, S. (2001) Synechocystis sp. PCC 6803 – a useful tool in the study of the genetics of cyanobacteria. Photosynth. Res. 70, 73–83. 8. MacKinney, G. (1941) Absorption of light by chlorophyll solutions. J. Biol. Chem. 140, 315-322. 9. Allen, J. F. and Holmes, N. G. (1986) Electron transport and redox titration, in: Photosynthesis, Energy Transduction: A Practical Approach

10.

11.

12.

13.

14.

15.

(Hipkins, M. F. and Baker, N. R., eds.), IRL Press, Oxford, pp. 103–141. Jansson, C., Debus, R. J., Osiewacz, H. D., Gurevitz, M., and McIntosh, L. (1987) Construction of an obligate photoheterotrophic mutant of the cyanobacterium Synechocystis 6803. Plant Physiol. 85, 1021–1025. Chu, H.-A., Nguyen, A. P., and Debus, R. J. (1994) Site-directed photosystem II mutants with perturbed oxygen-evolving properties. 2. Increased binding or photooxidation of manganese in the absence of the extrinsic 33-kDa polypeptide in vivo. Biochemistry 33, 6150–6157. Morgan, T. R., Shand, J. A., Clarke, S. M., and Eaton-Rye, J. J. (1998) Specific requirements for cytochrome c-550 and the ­manganese-stabilizing protein in photoautotrophic strains of Synechocystis sp. PCC 6803 with mutations in the domain Gly-351 to Thr-436 of the chlorophyll-binding protein CP47. Biochemistry 41, 14,437–14,449. Gay, P., Le Coq, D., Steinmetz, M., Ferrari, E., and Hoch, J. A. (1983) Cloning structural gene sacB, which codes for exoenzyme levansucrase of Bacillus subtilis: expression of the gene in Escherichia coli. J. Bacteriol. 153, 1424–1431. Cai, Y. P. and Wolk, C. P. (1990) Use of a conditionally lethal gene in Anabaena sp. Strain PCC-7120 to select for double recombinants and to entrap insertion sequences. J. Bacteriol. 172, 3138–3145. Ermakova-Gerdes, S. and Vermaas, W. (1999) Development of a psbA-less/psbD-less strain of Synechocystis sp. PCC 6803 for simultaneous mutagenesis of the D1 and D2 proteins of photosystem II, in The Phototrophic Prokaryotes (Peschek, G. A. and Loffelhardt, W., eds.), Kluwer, Dordrecht, pp. 51–60.

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16. Kunkel, T. A. (1985) Rapid and efficient sitespecific mutagenesis without phenotypic selection. Proc. Natl Acad. Sci. USA 82, 488–492. 17. Kunkel, T. A., Roberts, J. D., and Zakour, R. A. (1987) Rapid and efficient site-specific mutagenesis without phenotypic selection. Meth. Enzymol. 154, 367–382. 18. Vandeyar, M. A., Weiner, M. P., Hutton, C. J., and Batt, C. A. (1988) A simple and rapid method for the selection of oligodeoxynucleotide-directed mutants. Gene 65, 129–133. 19. Vermaas, W., Charite, J., and Eggers, B. (1990) System for site-directed mutagenesis in the psbD1/C operon of Synechocystis sp.

PCC 6803, in Current Research in Photosynthesis. Vol. I (Baltscheffsky, M., ed.), Kluwer, Dordrecht, pp. 231–238. 20. Eaton-Rye, J. J. and Vermaas, W. F. J. (1991) Oligonucleotide-directed mutagenesis of psbB, the gene encoding CP47, employing a deletion mutant strain of the cyanobacterium Synechocystis sp. PCC 6803. Plant Mol. Biol. 17, 1165–1177. 21. Bentley, F. K., Luo, H., Dilbeck, P., Burnap, R. L., and Eaton-Rye, J. J. (2008) Effects of inactivating psbM and psbT on photodamage and assembly of photosystem II in Synechocystis sp. PCC 6803. Biochemistry 47, 11,637–11,646.

Chapter 23 A Simple Method for Chloroplast Transformation in Chlamydomonas reinhardtii Vellupillai M. Ramesh, Scott E. Bingham, and Andrew N. Webber Abstract Photosystem I (PSI) is a multisubunit pigment-protein complex that uses light energy to transfer electrons from plastocyanin to ferredoxin. Application of genetic engineering to photosynthetic reaction center proteins has led to a significant advancement in our understanding of primary electron transfer events and the role of the protein environment in modulating these processes. Chlamydomonas reinhardtii provides a system particularly amenable to analyze the structure–function relationship of Photosystem I. C. reinhardtii is also a particularly favorable organism for chloroplast transformation because it contains only a single chloroplast and grows heterotrophically when supplemented with acetate. Chlamydomonas has, therefore, served as a model organism for the development of chloroplast transformation procedures and the study of photosynthetic mutants generated using this method. Exogenous cloned cpDNA can be introduced into the chloroplast by using this biolistic gene gun method. DNA-coated tungsten or gold particles are bombarded onto cells. Upon its entry into chloroplasts, the transforming DNA is released from the particles and integrated into the chloroplast genome through homologous recombination. The most versatile chloroplast selectable marker is aminoglycoside adenyl transferase (aadA), which can be expressed in the chloroplast to confer resistance to spectinomycin or streptomycin. This article describes the procedures for chloroplast transformation. Key words: Photosystem I, Chloroplast transformation, Chlamydomonas reinhardtii, Site-directed mutagenesis, Electron transfer

1. Introduction Photosystem I is a multisubunit pigment–protein complex that uses light energy to initiate electron transfer from plastocyanin to ferredoxin. It consists of 13 polypeptide subunits. The two largest subunits, PsaA and PsaB, form a heterodimeric core which binds approximately 100 chlorophyll a molecules and the electron transfer cofactors P700, A, A0, A1, and Fx (1–3). Application of

Robert Carpentier (ed.), Photosynthesis Research Protocols, Methods in Molecular Biology, vol. 684, DOI 10.1007/978-1-60761-925-3_23, © Springer Science+Business Media, LLC 2011

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genetic engineering to photosynthetic reaction center proteins has led to a significant advancement in our understanding of primary electron transfer events and the role of the protein environment in modulating these processes (4). Much of this work has been restricted to purple bacteria and cyanobacteria in which manipulations of the genes encoding reaction centers is fairly straightforward. In eukaryotes, biolistic transformation of the chloroplast genome has several inherent complications as plants and many green algae are unable to grow in the absence of ­photosynthesis and most of them contain several to hundreds of ­chloroplasts with each organelle containing 80–100  copies of chloroplast DNA. Several of these limitations are overcome by using the green alga Chlamydomonas reinhardtii, which contains only a single chloroplast and grows heterotrophically when supplemented with acetate. Chlamydomonas has, therefore, served as a model organism for the development of chloroplast transformation procedures and the study of photosynthetic mutants generated using this method (2). Exogenous-cloned cpDNA can be introduced into the chloroplast by using this biolistic gene gun method (4, 5). DNA-coated tungsten or gold particles are bombarded onto cells. Upon its entry into chloroplasts, the transforming DNA is released from the particles and integrated into the chloroplast genome through homologous recombination (5–7). Because of the efficient homologous chloroplast recombination system, specific deletion or site-directed mutation could be created relatively easily (8). This article describes the procedures for highly efficient chloroplast transformation.

2. Materials 2.1. Growth

1. Strains: C. reinhardtii strains CC125 and CC 2696. 2. Medium: Cox Chlamydomonas medium (CC) (1×): Tris base (2.5  g), glacial acetic acid (1  mL), NH4NO3 (0.5  g), MgSO4⋅7H2O (0.1 g), CaCl2⋅2H2O (0.02 g), KH2PO4 (0.05 g), KCl (0.1 g), Hutner’s trace elements solution (1 mL). 3. Yeast extract should be added to the liquid CC medium (1×) at a final concentration of 0.1% before autoclaving. 4. Spectinomycin. 5. Hutner’s trace element solution: For 1  L of trace elements mix, dissolve each compound in the volume of water indicated. The EDTA should be dissolved in boiling water, and the FeSO4 should be prepared last to avoid oxidation. EDTA, disodium salt (50 g in 250 mL H2O, ZnSO4 ⋅ H2O – 22 g in 100 mL H2O, H3BO3 – 11.4 g in 200 mL⋅H2O),

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MnCl2 ⋅ 4H2O (5.06 g in 50 mL H2O), CoCl2 ⋅ 6H2O (1.61 g in 50  mL H2O), CuSO4 ⋅ 5H2O (1.57  g in 50  mL H2O), (NH4)6Mo7O24 ⋅ 4H2O (1.1 g in 50 mL H2O), FeSO4 ⋅ 7H2O (4.99 g in 50 mL water). Mix all solutions except EDTA. Bring this mixture to boil and add EDTA. The mixture should turn green. When everything is dissolved, cool to 70°C. Keeping temperature at 70°C, adjust the pH to 6.7 with 80–90 mL hot KOH (20%). Bring the final solution to 1 L total volume. It should be clear green initially. Stopper the flask with a cotton plug, and let it stand for 1–2 weeks, shaking it once a day. Filter the solution using two layers of Whatman # 1 filter paper. When the solution turns purple, store it in the refrigerator. 2.2. Particle Bombardment

1. 2.5 M CaCl2. 2. 0. 1 M spermidine. 3. 100% isopropyl alcohol (high-performance HPLC grade). 4. M10 Tungsten (Bio-Rad, Richmond, CA). 5. Mutated plasmid DNA. 6. Macrocarrier (Bio-Rad, Richmond, CA). 7. Biolistic Gun PDS-1000/He (Bio-Rad, Richmond, CA).

2.3. Minipreparation of DNA and PCR

1. TEN buffer: 10  mM Tris–HCl pH 8.0, 10  mM EDTA, 50 mM NaCl. 2. 20 mg/mL Pronase. 3. 10% Sodium dodecyl sulfate (SDS). 4. PCR primers – designed to flank the site of the introduced mutation and to amplify the region of interest. 5. Agarose and agarose gel electrophoresis unit.

3. Methods The following methods outline the detailed procedure for the chloroplast transformation: (1) Recipient strains and culture conditions; (2) Particle bombardment; (3) Selection and screening for homoplasmic transformants. 3.1. Recipient Strains and Culture Conditions (see Note 1)

C. reinhardtii strains Wt CC 125  mt+ and Wt CC 2696 were obtained from the Chlamydomonas culture collection at Duke University and used as recipient strains for site-directed mutation. Strain Wt CC125 contains both PSI and PSII, whereas strain CC 2696 lacks both chlorophyll a/b containing light-harvesting

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complexes and photosystem II in the thylakoid membrane as it carries the DS521 cab deficiency mutation and a psbA deletion mutation. Cells were maintained on 1.5% agar plates containing CC medium. For chloroplast transformation, a starter culture was initiated by growing the recipient cells heterotrophically in 250 mL flasks containing autoclaved 100 mL CC liquid medium at 25°C under the light intensity of 50  mm photons/m2 s with constant shaking for 2–3  days according to Harris (9). Then, 1–2 mL of starter culture was transferred to 1 L flasks containing sterilized 500  mL of CC medium and grown to a density of 1–1.5 × 106  cells/mL under the same condition as described above. Then, the cells were concentrated to 5 × 107 cells/mL and 2 × 107  cells were spread on plates containing spectinomycin (100 mg/mL) (see Note 2). 3.2. Particle Bombardment

1. Weigh 90 mg tungsten particles, then, transfer to a disposable glass tube.

3.2.1. Preparation of M10 Tungsten Particle

2. Keep the tube in a beaker and bake in oven at 180°C overnight. 3. Transfer tungsten particles to a 50 ml plastic conical tube and add 15 ml 70% isopropanol and vortex for 3 min. 4. Leave at room temperature for 15 min and collect the tungsten particles by centrifugation at 15,000 rpm in a Sorvall SS – 34 rotor for 5 min. 5. Wash the pellet three times with 5–10 mL in sterile deionized water followed by centrifugation. 6. Suspend the particles in 1.5 mL sterile 40% glycerol and aliquot 60 ml in sterile microfuge tubes and store at −20°C.

3.2.2. Precipitation of DNA onto Tungsten Particles (see Note 3)

1. Vortex the microfuge tubes containing 60  ml of tungsten particles and add 5  mg of DNA and continue to vortex for 30 s. 2. Add 60 ml of 2.5 M CaCl2 while vortexing. 3. Add 12 ml of 0.1 M spermidine while vortexing and continue to vortex for 1 min. 4. Leave at room temperature for 15 min 5. Centrifuge for 10 s and discard the supernatant 6. Carefully suspend the particles in 250 ml of 100% isopropyl alcohol by repeated pipetting until there is no visible aggregate and vortex for 2 min (see Note 4). 7. Centrifuge for 10 s and discard the supernatant. 8. Resuspend the particles in 60 ml of 100% isopropyl alcohol by repeated pipetting.

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1. Load the 60  ml of DNA-precipitated tungsten particles on the thin macrocarrier and allow the particles to air dry. 2. Use a 1,350 psi rupture disk. 3. Place the macrocarrier 5–8  in. from the rupture disk (see Note 5). 4. Place the Petri plates containing the recipient cells on the bottom self of the chamber. 5. Evacuate the chamber at least to 28.5  in. Hg and start the bombardment. 6. Incubate the plates under dim light for 2  weeks until the single colonies appear. Then, single colonies are randomly picked and streaked on to new plates and incubated under dim light. Using the above conditions, transformation efficiencies between 1 × 10−5 and 1 × 10−4 can be obtained.

3.3. Selection and Screening for Homoplasmic Transformants 3.3.1. PCR Analysis

Selection was performed using marker recycling method as described by Redding et  al. (10). It is important to select for transformant cells in which every copy of cpDNA contains the introduced mutation. PCR can be used to screen for homoplasmy of each colony if the donor plasmid introduces a unique restriction site in the region amplified. Primers are designed to flank the site of the introduced mutation and then used to amplify the region of interest. The PCR product is then digested with the restriction enzyme and size fractionated by 2% agarose gel electrophoresis (Fig.  1). PCR analysis was carried out using the

Fig. 1. Analysis of PCR product from wild type and mutant cells. A 400 bp fragment of PsaB was amplified from total cellular DNA using primers that flank the site of the silent mutation in the donor plasmid that generates an Eae I site. The product was either loaded directly onto the 2% agarose gel (lanes 1, 3) or digested with Eae I (lanes 2, 4–7 ) before electrophoresis. Complete digestion with Eae I (lanes 4 and 7) indicates that all copies of the wild type PsaB gene have been replaced with mutant copy. Wild type, lanes 1 and 2; Mutants containing the Eae I site, lanes 3–7. Note that Eae I did not cut the PCR product from wild type cells (lane 2).

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mini-prepared DNA from a patch of plated cells as described below. Single colony selection is repeated for two to three rounds to obtain complete homoplasmic transformants. 3.3.2. Minipreparation of DNA from Chlamydomonas Cells for PCR Analysis

1. Transfer a patch of cells (0.5 cm−2) from plated to a microfuge tube containing 400  ml TEN buffer, and then suspend the cells by pipetting or vortexing. 2. Add 50 ml of pronase and 50 ml of 10% SDS, and then mix this solution by inverting two to three times and incubate for 1 h at 50°C. 3. Extract the above solution once with phenol/chloroform/ isoamylalcohol (25:24:1) and centrifuge for 5 min. 4. Remove the top layer containing dissolved DNA and precipitate with two volumes of 100% ethanol. 5. Pellet the DNA by centrifugation, wash once with 70% ethanol and then air dry. 6. Dissolve the DNA in 30 ml of sterile deionized water and use 1–2 ml of this DNA for the PCR reaction. Following the amplification, approximately 15–20 ml of the PCR reaction mixture can be digested with restriction enzyme and size fractionated by 2% agarose gel electrophoresis. Figure 1 shows an example of PCR-amplified DNA from wild type cells (lane 1, wild type; lane 2, wild type digested (no introduced enzyme site) and two mutants (lane 3, undigested mutant; lanes 4–6, digested) that have a unique Eae I site introduced in the transforming plasmid. As shown in Fig.  1, the DNA from wild type cells does not cut with Eae I (Fig. 1, lane 2), whereas DNA from the mutants cuts completely (lanes 4 and 6). If the mutants are not homoplasmic, then the DNA would not cut completely with Eae I. This assay is a very sensitive screening method for homoplasmy. Erickson (11) reported that PCR can be used to detect one wild type copy at a dilution of 1 in 20,000.

4. Notes 1. Several procedures for biolistic transformation of Chlamydomonas chloroplasts have been published (4, 11, 12). Efficiency of transformation is sensitive to the conditions used. The following three factors (1) Growth stage of recipient cells, (2) Preparation of tungsten particles, and (3) Conditions of bombardment considerably affect the efficiency of chloroplast transformation. Hence, we have modified slightly the earlier method (4), especially in the preparation of tungsten particles as described in the methods section. Preparation of tungsten

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­ articles by our modified method and dissolving the p ­DNA-precipitated tungsten particles in isopropyl alcohol was found to increase the chloroplast efficiency as isopropyl alcohol makes the tungsten particles very well separated and prevents from reaggregation. 2. Growth stage of recipient cells is very important because when cells are grown to a density over 1.5 × 106  cells/mL, there is a significant reduction in the transformation efficiency. 3. Quality of DNA is a parameter affecting transformation efficiency. Therefore, it is recommended that the CsCl-purified DNA (13) should be used. 4. For resuspending the DNA-coated tungsten particles, 100% isopropyl alcohol should be used as it allows the tungsten particles to spread on the macrocarrier uniformly without any aggregation. 5. During particle bombardment, the distance between the rupture disk and macrocarrier and the vacuum pressure in the bombardment chamber should be maintained as described in the method.

Acknowledgments This work was supported by USDA # (2001-35318-11137) to A.N. Webber. References 1. Brettel, K. (1997) Electron transfer and arrangement of the redox cofactors in photosystem. Biochimica Biophysica Acta 1318: 322–373. 2. Webber, A.N., Su, H., Bingham, S.E., Kass, H., Krabben, L., Kuhn, M., Jordan, R., Schlodder, E., and Lubitz, W. (1996) Sitedirected mutations affecting the spectroscopic characteristics and midpoint potential of the primary donor in photosystem I. Biochemistry 35: 12857–12863. 3. Webber, A.N. and Ramesh, V. (2006) Mutagenesis of Ligands to Photosystem I Cofactors, in Photosystem I: The Light-Driven Plastocyanin: Ferredoxin Oxidoreductase, Vol. 24 (Golbeck, J.H., ed.), Springer, Dordrecht, pp. 193–204. 4. Lee, H., Bingham, S.E., and Webber, A.N. (1998) Specific mutagenesis of reaction center proteins by chloroplast transformation of Chlamydomonas reinhardtii. Methods in Enzymology 297: 310–320.

5. Boynton, J.E., Gillham, N.W., Harris, E.H., Hosler, J.P., Johnson, A.M., Jones, A.R., Randolph-Anderson, B.L., Robertson, D., Klein, T.M., Shark, K.B., and Sanford, J.C. (1988) Chloroplast transformation in Chlamydomonas with high velocity microprojectiles. Science 240: 1534–1538. 6. Blowers, A.D., Bogorad, L., Shark, K.B., and Sanford, J.C. (1989) Transcriptionalanalysis of endogenous and foreign genes in chloroplast transformants of Chlamydomonas. Plant Cell 2: 1059–1070. 7. Newman, S.M., Boynton, J.E., Gillham, N.W., Randolphanderson, B.L., Johnson, A.M., and Harris, E.H. (1990) Transformation of chloroplast ribosomal-RNA genes in Chlamydomonas – Molecular and genetic-characterization of integration events. Genetics 126: 875–888. 8. Rochaix, J.D., Fischer, N., and Hippler, M. (2000) Chloroplast site-directed mutagenesis of photosystem I in Chlamydomonas: Electron

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transfer reactions and light sensitivity. Biochimie 82: 635–645. 9. Harris, E.H. (1989) Chlamydomonas Source Book, Academic Press, San Deigo, CA, USA. 10. Redding, K., MacMillan, F., Leibl, W., Brettel, K., Hanley, J., Rutherford, A.W., Breton, J., and Rochaix, J.-D. (1998) A Systemic survey of conserved histidines in the core subunits of photosystem I by site-directed mutagenesis reveals the likely axial ligands of P700. EMBO Journal 17: 50–60.

11. Erickson, J.M. (1995) In “Oxygenic Photosynthesis: The Light Reactions” (Ort, D.R. and Yocum, C.F., eds.) Kluwer Academic Publishers, The Netherlands, p. 589. 12. Boynton, J.E. and Gillham, N.W. (1993) Chloroplast transformation in Chlamydomonas. Methods in Enzymology 217: 510–536. 13. Sambrook, J., Fritsch, E.F., and Maniatis, T. (1989) Molecular Cloning. Cold Spring, Harbor, NY.

Chapter 24 Rapid Isolation of Intact Chloroplasts from Spinach Leaves David Joly and Robert Carpentier Abstract In this chapter, a rapid method to isolate intact chloroplasts from spinach leaves is described. Intact chloroplasts are isolated using two short centrifugation steps and avoiding the use of percoll gradient. Intactness of chloroplast is evaluated by the inability of potassium ferricyanide to enter inside the chloroplasts and to act as an electron acceptor for photosystem II. Key words: Chloroplasts, Leaves, Spinach, Potassium ferricyanide, Oxygen evolution

1. Introduction A wide variety of organisms and preparation types can be used to study photosynthetic electron transport mechanisms. Reaction centres complexes and photosystem submembrane fractions can be isolated to analyse precise or specific electron transport steps and reactions. On the opposite, experiments can be performed with whole leaves. The latter approach, when possible, is also appropriate since it allows the study of photosynthesis in an intact system and can therefore give information that is more physiologically relevant. However, using intact leaves has also some limitations. There may be significant variation between measurements because the photosynthetic responses can vary with growth conditions, leaf age, probed portion of the leaf, position of the leaf on the plant, dark adaptation, etc. Thus, a large number of repetitions must be made to obtain reliable results. Moreover, the possibility to use inhibitors of specific electron transport reactions is limited because infiltration of chemicals, when it is successful, results in an unknown effective concentration inside chloroplasts.

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On the other hand, thylakoid membranes can easily be prepared, facilitate the use of inhibitors, and yield to highly reproducible results between each measurement due to a homogenous preparation. However, molecules in the stroma (NADP pool, ferredoxin, Calvin-Benson cycle enzymes, etc.) are lost during the preparation. This results in an inefficient photosystem I operation in both linear and cyclic electron transport and practical loss of physiological relevance. Consequently, the preparation of intact chloroplasts represents a compromise between intactness of leaves and homogeneity of isolated thylakoids. The protocol presented here is mainly based on Robinson et  al. (1) with some modification (2). The procedure is quick with two short centrifugation steps. The intact chloroplasts obtained are suitable for photosynthetic electron transport studies.

2. Materials 1. Spinach leaves. 2. Kitchen blender. 3. Miracloth. 4. Buffer A (grinding buffer): 50 mM Hepes NaOH (pH 6.9), sorbitol 0.33  M, EDTA 2  mM, MgCl2 1  mM, MnCl2 1 mM. 5. Buffer B (resuspension buffer): 50  mM Hepes NaOH (pH 7.6), sorbitol 0.33  M, EDTA 2  mM, MgCl2 1  mM, MnCl2 1 mM, 10 mM KCl, 1 mM NaCl. 6. Buffer C (2× resuspension buffer): 100  mM Hepes NaOH (pH 7.6), sorbitol 0.66  M, EDTA 4  mM, MgCl2 2  mM, MnCl2 2 mM, 20 mM KCl, 2 mM NaCl. 7. 200 mM potassium ferricyanide solution. 8. 1 M d,l-Glyceraldehyde solution. 9. 250 mM NH4Cl solution. 10. Spectrophotometer. 11. Clark-type oxygen electrode.

3. Methods 3.1. Preparation of Intact Chloroplasts

1. Select 40 g of dark green leaves. Rinse leaves in cold water (see Note 1). 2. Grind leaves in about 150–175 mL of Buffer A.

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3. Filter the homogenate twice on two layers of Miracloth tissue. 4. Centrifuge the filtrate 60 s at 750 × g. 5. Discard the supernatant and resuspend the pellet in about 30 mL of Buffer B. 6. Centrifuge the filtrate 60 s at 750 × g. 7. Discard the supernatant and resuspend the pellet in a small volume of Buffer B (1–1.5 mL). 3.2. Chlorophyll Assay

Total chlorophyll concentration of the chloroplast preparation is measured according to Porra et al. (3): 1. Add 10 mL of chloroplast preparation in 5 mL of 80% acetone and vortex the mixture. 2. Centrifuge the mixture 2 min at 2,500 × g. 3. Carefully transfer the supernatant in the spectrophotometer cuvette. 4. Measure the absorbance of the supernatant at 647 and 664 nm using the spectrophotometer. 5. Calculate the chlorophyll concentration using the following equation (see Note 2): Chlorophyll concentration (mg/mL) = [(17.76 × A 647) + (7.34 × A664)]/2 Final chlorophyll concentration in this preparation is generally around 1 mg/mL (see Note 3).

3.3. Evaluation of Chloroplasts Intactness

Methods to evaluate the intactness of chloroplast preparations are based on the nonpermeability of the chloroplast membrane to ferricyanide molecules. Ferricyanide is a photosystem II electron acceptor. Therefore, it can support photosystem II activity under continuous saturating illumination only in ruptured (nonintact) chloroplasts. The intactness of chloroplast preparations is then evaluated by comparing photosystem II activity under continuous illumination in the presence of ferricyanide before and after an osmotic shock. The osmotic shock allows a comparison with all chloroplasts ruptured. The commonly used method is the measurement of ferricyanidesupported oxygen evolution (4) (see Note 4).

3.3.1. Preparation of Samples for Oxygen Evolution Measurement Without Osmotic Shock

The reaction medium (total volume: 2 mL) contains: 1. 1,920 mL of Buffer B. 2. 25 ml of chloroplast preparation to a final chlorophyll concentration of 12.5 mg/mL (for a 1 mg/mL preparation). 3. 20 mL of 1 M d,l-glyceraldehyde (final concentration: 10 mM) (see Note 5).

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4. 20 mL of 250 mM NH4Cl (final concentration: 2.5 mM) (see Note 6). 5. 15  mL of 200  mM potassium ferricyanide (final concentration: 1.5 mM) freshly prepared. 3.3.2. Preparation of Samples for Oxygen Evolution Measurement with Osmotic Shock

Incubate the following reaction mixture for 1 min. (total volume: 1 mL) for the osmotic shock: 1. 25 mL of chloroplast preparation to a final chlorophyll concentration of 12.5 mg/mL (for a 1 mg/mL preparation). 2. 920 mL of distilled water. Add the following reagents to the mixture: 1. 1,000 mL of Buffer C. 2. 20  mL of 1  M d,l-glyceraldehyde (final concentration: 10 mM). 3. 20 mL of 250 mM NH4Cl (final concentration: 2.5 mM). 4. 15  mL of 200  mM potassium ferricyanide (final concentration: 1.5 mM).

3.3.3. Chloroplast Intactness Assay and Calculation

1. Measure the oxygen evolution rate (see Note 7) of the chloroplast preparation under saturating white light for 1  min without and with osmotic shock. 2. The percentage of intact chloroplast can be calculated with the following equation: % of intact chloroplasts = 100 (O2 FeCN+osm − O2 FeCN)/O2 FeCN+osm where O2 FeCN+osm and O2 FeCN are the ferricyanide-supported oxygen evolution rate measured with or without osmotic shock, respectively. Typical chloroplasts preparations should yield to a percentage of intact chloroplasts higher than 80% (see Note 8).

4. Notes 1. All materials used for chloroplast isolation should be maintained at 4°C. 2. The chlorophyll concentration in the 80% acetone sample tube in mg/L (given by (17.76 × A647) + (7.34 × A664)) is divided by 2 to obtained the chlorophyll concentration of the chloroplast preparation in mg/mL. The division by 2 comes from the dilution factor of 500 in this chlorophyll assay (10 mL in 5 mL) and the conversion of units from L to mL.

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3. Experiments using intact chloroplasts must be carried out with freshly prepared samples for an optimal photosynthetic activity and chloroplast intactness. Therefore, freezing chloroplast preparation before usage should be avoided. 4. Alternatively, the photoreduction of ferricyanide can be followed by absorbance measurement at 410 nm with and without osmotic shock (4). 5. d,l-Glyceraldehyde prevents CO2-supported oxygen evolution by inhibiting Calvin–Benson cycle and does not directly affect photosystem II activity (5, 6). 6. NH4Cl is an electron transport uncoupler and thus allows the measurement of a maximal oxygen evolution rate. 7. Method for the measurement of oxygen evolution rate is described in Chapter 18 (see Subheading 3.2.2). 8. Quantum yield of photosystem II as measured by the Fv/Fm ratio of fluorescence induction is generally between 0.77 and 0.81. The presence of a clear I-step in the fluorescence induction traces is also an indicator of the intactness of the chloroplast preparation (2). For more information about fluorescence induction measurement, please refer to Chapter 18 (see Subheadings 3.2.4.2 and 3.2.4.3). References 1. Robinson, J. M., Smith, M. G., and Gibbs, M. (1980) Influence of hydrogen peroxide upon carbon dioxide photoassimilation in the spinach chloroplast: I. Hydrogen peroxyde generated by broken chloroplasts in an “intact” chloroplast preparation is a causal agent of the Warburg effect. Plant Physiol. 65, 755–759. 2. Joly, D. and Carpentier, R. (2009) Sigmoidal reduction kinetics of the photosystem II acceptor side in intact photosynthetic materials during fluorescence induction. Photochem. Photobiol. Sci. 8, 167–173. 3. Porra, R. J., Thompson, W. A., and Kriedemann, P. E. (1989) Determination of accurate extinction coefficients and simultaneous-equations for assaying chlorophyll-a and chlorophyll-b extracted with 4 different solvents – Verification of the concentration of

chlorophyll standards by atomic-absorption spectroscopy. Biochim. Biophys. Acta 975, 384–394. 4. Lilley, R. M., Fitzgerald, M. P., Rienits, K. G., and Walker, D. A. (1975) Criteria of intactness and the photosynthetic activity of spinach chloroplast preparations. New Phytol. 75, 1–10. 5. Hollinderbäumer, R., Ebbert, V., and Godde, D. (1997) Inhibition of CO2-fixation and its effect on the activity of Photosystem II, on D1-protein synthesis and phosphorylation. Photosynth. Res. 52, 105–116. 6. Stokes, D. M. and Walker, D. A. (1972) Photosynthesis by isolated chloroplasts. Inhibition by dl-glyceraldehyde of carbon dioxide assimilation. Biochem. J. 128, 1147–1157.

Chapter 25 Mechanical Isolation of Bundle Sheath Cell Strands and Thylakoids from Leaves of C4 Grasses Elz˙ bieta Romanowska and Eugeniusz Parys Abstract Bundle sheath (BS) strand cells and BS thylakoids from C4 plants represent a unique system for various studies using a combination of physiological, biochemical, and molecular approaches. We have developed procedures for mechanical disruption of leaf tissues in order to isolate metabolically active bundle sheath strand cells and thylakoids practically free from cross-contamination coming from mesophyll cells. The procedures are described in detail together with useful practical suggestions. Using mechanical disruption we have shown the supramolecular organization of the dimeric LHCII-PSII in BS thylakoids of maize. Key words: C4 plants, Mechanical leaf disruption, Bundle sheath cells, Bundle sheath thylakoids, Agranal chloroplasts, PSII dimer

1. Introduction A characteristic feature of C4 plants is the differentiation of the photosynthetic tissues into two distinct cell types: mesophyll (M) and bundle sheath (BS) cells (1). For understanding their cooperation, particularly the structural–functional relationships between chloroplasts, two techniques have been developed to separate mesophyll and bundle sheath cells: mechanical disruption (2–4) and enzymatic digestion of leaf tissue (5–8). When these two methods were compared, it appeared that enzymatic digestion resulted in significant depletion of major polypeptides of PSII (9). Therefore, the mechanical method is more suitable for isolation of bundle sheath strand cells and their chloroplast thylakoids. Using this method we have found that PSII is present in agranal BS chloroplasts of maize, exists as a dimer, and forms LHCII-PSII supercomplexes. Moreover, almost the same set of

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photosynthetic membrane complexes was present in both the granal and agranal chloroplasts (10). The success in isolation of metabolically active bundle sheath cells or their chloroplast thylakoids is principally dependent on two critical steps: leaf tissue preparation and grinding procedure. Although the bundle sheath cell wall is thick and rigid, a sudden application of mechanical force may lead to injury of cell compartments, especially chloroplasts. Another problem resulting from the BS cell wall properties is its extreme resistance to breaking required to release the organelles. Two different mechanical grinding procedures that we have developed are presented in detail together with practical hints.

2. Materials 2.1. Seeds and Accessories

1. Zea mays L. seeds. 2. Panicum miliaceum seeds. 3. Panicum maximum seeds. 4. Perlite. 5. 10-L plastic flower-box. 6. Transparent plastic foil. 7. Knop’s solution or commercial nutrient.

2.2. Leaf Grinding Medium (see Note 4)

1. 0.35 M mannitol or sorbitol. 2. 20 mM HEPES–KOH, pH 7.6. 3. 2 mM MgCl2. 4. 2 mM KH2PO4. 5. 2 mM sodium isoascorbate.

2.3. Strand Resuspension Medium (see Note 7)

1. 0.35 M Mannitol or sorbitol. 2. 20 mM HEPES–KOH, pH 7.6. 3. 10 mM KCl. 4. 2 mM MgCl2. 5. 2 mM KH2PO4.

2.4. Chloroplast Isolation Medium (see Notes 11 and 12)

1. 0.4 M Mannitol. 2. 50 mM HEPES–NaOH, pH 7.5. 3. 5 mM MgCl2. 4. 10 mM NaCl. 5. 1 mM Ethylenediamine tetraacetic acid (EDTA).

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1. 50 mM HEPES–NaOH, pH 7.5. 2. 5 mM MgCl2. 3. 10 mM NaCl. 4. 1 mM Ethylenediamine tetraacetic acid (EDTA).

2.6. Thylakoid Wash Medium (see Notes 11 and 12)

1. 50 mM HEPES–NaOH, pH 7.5.

2.7. Thylakoid Suspension Medium

1. 50 mM HEPES–NaOH, pH 7.5.

2. 10 mM NaCl. 3. 5 mM MgCl2. 2. 0.33 M Mannitol (see Note 18). 3. 10 mM NaCl. 4. 4 mM MgCl2.

2.8. Tools and Equipment

1. Fresh razor blades. 2. Beaker (50 mL, 100 mL), cylinder (50 mL, 100 mL), graduated tube with stopper (15–20  mL), crystallizer (500 and 900 mL). 3. Kitchen strainer (7–8 and 15–18 cm diameter). 4. Muslin, nylon net (1  mm, 600  mm, 80  mm, and 20  mm aperture). 5. Homogenizer or domestic blender, 1.5-L volume (see Note 3). 6. Homogenizer (Waring Laboratory, variable speed, 0.2-L volume) or domestic blender 0.4-L volume (see Note 3). 7. Potter homogenizer, 3 mL. 8. Autotransformer (0–250 V). 9. Microscope. 10. Laboratory mortar and pestle, 80% acetone, CaCO3, sand.

3. Methods 3.1. Seed Germination and Plant Growth Conditions

1. Soak maize (Zea mays L.) seeds in water overnight before planting (21–24 seeds) in perlite (soaked and drained) in10-L plastic flower-box. Cover the seed layer with about 1 cm of wet perlite. 2. Seeds of millet (Panicum miliaceum) and guinea grass (Panicum maximum) may be sown directly on wet perlite and covered thinly with wet perlite. Cover the boxes with transparent plastic foil and place under low light (about 100 mE/m2 s) in a growth cabinet.

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3. When shoots emerge from the perlite remove the cover. For both Panicum species reduce the amount of seedlings to about 40 per box. Cultivate plants under photosynthetic photon flux density on the upper leaves level of about 350 mE/ m2 s and 14-h photoperiod and night/day temperatures of about 20/25°C. 4. Fertilize every other day with Knop’s solution or commercial nutrient. Because the species differ in growth rates, fully expanded leaves suitable for isolation of bundle sheath cells are obtained from about 3–4-, 4–5-, and 7–8-week old plants of Z. mays, P. miliaceum, and P. maximum, respectively. 3.2. Isolation of Bundle Sheath Strand Cells

1. Harvest leaves from plants after the dark period (see Note 1) and excise middle parts (about 5–6  cm from the tip and 10–15-cm long, about 13–15 g of fresh weight) of the third and fourth leaves. Carry out all the following operations in a cold room (4–5°C). 2. Rinse leaf parts with tap water and remove the midribs (see Note 2) with a sharp razor blade. Put about 10 g of this tissue in cool water for 15–20  min under week light (30– 40 mE/m2 s). 3. Slice tissue transversally into 1–2 mm strips (see Note 3) with a razor blade on a plate in a layer of the grinding medium (see Note 4). Alternatively, very sharp scissors may be used for slicing tissue (but cut without bruising tissue) immediately into grinding medium (4–5°C, 70–80 mL) in a blender. 4. Connect blender (1.5  L) to an autotransformer (0–250  V) and homogenize once for 10 s at 60% of line voltage and then at 40% of line voltage for 10 s (for maize usually 5–6 times while for both Panicum species 2–3 times) until microscopic examination of the pulp shows single strands without attached mesophyll cells (see Notes 5 and 6). 5. Filter the homogenate through a large kitchen strainer (with 1 mm nylon net placed inside) into the larger crystallizer and then filter the suspension successively through nets (placed in the smaller strainer) with 600 mm and 80 mm openings. 6. Transfer strands collected on the 80-mm net to a 100-mL beaker with 70–80 mL of resuspension medium (see Note 7), and after stirring, collect strands on the 80-mm net. Remove excess medium by putting the net on filter paper. Repeat the whole procedure twice. 7. Suspend strands in 50 mL of resuspension medium, transfer to a 50-mL cylinder, and after stirring for 2–3 min, allow to settle for about 8–10 min. 8. Remove by suction the supernatant fluid together with a green layer above the settled strands, add resuspension

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medium, and repeat the procedure until chloroplasts and epidermal materials in settled strands are scarce. 9. Add 20–25  mL of resuspension medium to settled strands and transfer on the 80-mm net. Wash the strands with resuspension medium until no chloroplasts occur in the filtrate. Transfer the net with settled strands into a 100-mL beaker and remove strands by inversion of net and washing with resuspension medium; allow strands to settle. Obtained strands are free from mesophyll chloroplasts while epidermal material is present only occasionally. 10. Remove excess liquid and transfer concentrated suspension of pure strands into a 15–20 mL graduated tube. 11. After stirring, transfer 100–200 ml (depending on strand density) of concentrated suspension of strands with a widemouthed pipette the aperture into a precooled mortar. Add 2 mL of 80% acetone and a small amount of CaCO3 and sand and grind vigorously to release chlorophyll from cells (see Note 8). Transfer the homogenate to a centrifuge tube, rinse the mortar with about 2 mL of 80% acetone, and add to the centrifuge tube. After centrifugation for 5  min at 3,000 × g take the clear green supernatant and adjust to a known volume. Measure the absorbance (A) at 645 and 663  nm with 80% acetone as reference. Calculate the concentration of total chlorophyll (Total Chl) in the cuvette according to formula: Total Chl (mg/mL) = 20.2 A645 + 8.02 A663 (11) and then, taking into account all the dilutions, calculate total chlorophyll content in original suspension of strands. Adjust volume of the suspension of bundle sheath cell strands in the tube to chlorophyll concentration of about 50–60 mg/mL. 12. Remove CO2 by gassing the suspension with nitrogen for 2–3 min (see Note 9), stopper the tube, and store at 0–4°C in darkness until used for assays (see Note 10). Typical results are shown in Table 1. 3.3. Isolation of Bundle Sheath Thylakoids

1. Harvest about 30 g of leaf tissue as described in Subheading 3.2, step 1 and then proceed as in Subheading 3.2, steps 2 and 3. 2. Connect blender (1.5  L) to an autotransformer, add sliced tissue to 150  mL of chilled (4–5°C) chloroplast isolation medium (see Notes 11 and 12), and homogenize 3 times for 5 s at 75% of line voltage (see Note 13). 3. Filter homogenate through four layers of muslin, softly squeeze the pulp and discard filtrate (contains mesophyll chloroplasts) (see Note 14). Homogenize the residue 3 times for 30  s at 75% of line voltage in 1.5-L blender with 300–400 mL of chilled (4–5°C) water (do not use deionized or distilled water) or thylakoid wash medium (see Note 15).

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Table 1 Respiration rates of bundle sheath cell strands isolated from leaves of C4 plants O2 uptake (mmol/min/mg Chl) Treatment

Z. mays

P. miliaceum

Before illumination

0.15 ± 0.01

0.16 ± 0.08

+NADH

0.24 ± 0.04

0.23 ± 0.02

+Malate

0.16 ± 0.02

0.22 ± 0.01

+Glycine

0.16 ± 0.03

0.20 ± 0.01

+Succinate

0.17 ± 0.04

0.19 ± 0.01

After illumination

0.16 ± 0.01

0.30 ± 0.02

+NADH

0.23 ± 0.02

0.32 ± 0.02

+Malate

0.23 ± 0.01

0.28 ± 0.01

+Glycine

0.19 ± 0.01

0.29 ± 0.01

+Succinate

0.21 ± 0.01

0.22 ± 0.01

The data represent the mean values ± SE of seven independent experiments. Oxygen uptake rate was measured before and after illumination (10 min, 800 mmol photons/ m2 s) in the presence of 5 mM bicarbonate and, in separate experiments, after addition of metabolites. Malate, glycine, and succinate were at 10 mM and NADH at 1 mM. Assay solution (resuspension medium) of 2 mL contained bundle sheath strand cells equivalent to 30 mg/mL chlorophyll. Oxygen uptake was measured at 25°C using a Clark-type electrode (TriOximatic EO200, WTW GmbH Weilheim, Germany), and it was monitored in the range 250–150 mM O2. Rate of O2 evolution during illumination of P. miliaceum bundle sheath cells in the presence of 5  mM bicarbonate was 1.20 ± 0.04 mmol O2/min/mg Chl, whereas bundle sheath cells of Z. mays did not evolve O2 during illumination. Light was provided by a slide projector (150 W).

Add 300–400 mL of water or wash medium and again homogenize 3 times for 30 s. 4. Filter the homogenate through four layers of 20-mm net and wash collected sediment briefly with cold water or thylakoid wash medium. Check the purity of strands under microscope. If mesophyll cells are still visible repeat homogenization and washing (steps 3 and 4) (see Note 16). 5. Transfer strands from 20-mm nylon net into chilled 0.2-L blender, add about 40  mL of chilled chloroplast isolation medium, and homogenize once for 30 s at maximal voltage. 6. Filter homogenate trough six layers of 20-mm nylon net and collect the filtrate into a crystallizer. 7. Repeat steps 5 and 6 three times or more. Check purity of BS chloroplast filtrate using anti-PEPC test and/or PEPC activity. Typical result is shown in Fig. 1.

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Fig. 1. Purity of bundle sheath chloroplast preparations from Panicum miliaceum leaves. The same amount of protein (2 mg) was loaded in each lane and proteins were separated on SDS-PAGE. Soluble fractions of mesophyll (M) and bundle sheath (BS) chloroplast extracts were analyzed by immunoblotting with anti-PEPC. Two independent isolations from BS calls are shown. Contamination from mesophyll cells in bundle sheath preparation is less than 1%. Similar purity of bundle sheath chloroplast preparation was obtained for Zea mays and Panicum maximum (9, 13).

8. Pour the filtrate to 50-mL centrifuge tubes and centrifuge at 7,000 × g for 5 min (4°C). 9. Remove supernatant and add small volume (e.g. 0.3–0.5 mL) of chilled osmotic shock medium into chloroplast pellet in each tube and resuspend using a small soft painting-brush or a pipette. Fill up the centrifuge tubes with osmotic shock medium (about 40–50 mL to each), and after stirring, centrifuge at 8,000 × g for 10 min (4°C). 10. Discard the supernatant, add about 1  mL of thylakoid suspension medium to one of the tubes with BS thylakoid pellet, resuspend using a small brush, combined transfer into next tube with thylakoid pellet, and so on. Then transfer the BS thylakoids into small (1.5 mL) tube and centrifuge at 6,000 × g for 10 min (4°C). Discard the supernatant, add 0.5–1.0 mL of the above medium to the pellet, roughly resuspend and transfer into 2–3 mL Potter homogenizer. After homogenization, transfer uniform suspension of BS thylakoids to preweighed 1.5-mL tube with stopper and determine the suspension volume by weighing. 11. After stirring, take 5  mL of thylakoid suspension and add 1,495 mL of 80% acetone into 2-mL centrifuge tube, stir, and centrifuge for 2 min at 1,000 × g. Transfer clear supernatant to spectrophotometer cuvette and measure absorbance (A) at 645 nm and 663 nm with 80% acetone as reference. Calculate the concentration of chlorophyll a (Chl a) and chlorophyll b (Chl b) in the cuvette using the equations: Chl a (mg/ mL) = 12.7 A663 − 2.69 A645, Chl b (mg/mL) = 22.9 A645 − 4.68 A663 (11). Taking into account the dilutions, calculate the total chlorophyll content in original suspension of BS thylakoids. Adjust the volume of thylakoid suspension with thylakoid suspension medium (if necessary) to a chlorophyll concentration of 0.5–0.6  mg/mL (see Note 17). Determine the chlorophyll a/b ratio.

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Table 2 PSII and PSI activity in bundle sheath thylakoids isolated from leaves of C4 plant PSII activity (mmol DCPIP/mg Chl/h) Zea mays Panicum miliaceum

PSI activity (mmol O2/ mg Chl /h)

36 ± 3

1,278 ± 72

116 ± 9

818 ± 55

Thylakoid samples corresponding to 20 mg/mL of chlorophyll were used. DCPIP was 0.1 mM. The data represent the mean values ± SE of four to five independent experiments. For details see in ref. 9

Fig. 2. Immunodetection of maize bundle sheath thylakoid phosphoproteins. Thylakoid sample corresponding to 1.5 mg of chlorophyll was loaded in the well. Proteins were detected with anti-PThr antibody (from Cell Signaling Technology). D non-illuminated leaves, L illuminated leaves.

12. Freeze thylakoids in liquid nitrogen and store at −80°C until use. BS thylakoids represent an excellent system to study physiological, biochemical, and structural aspects of photosynthetic apparatus activity (see Note 18). Examples of some results obtained from such examinations are presented in Table 2 and Figs. 2 and 3.

4. Notes 1. High content of starch granules in chloroplasts after the light period may lead to there injury. 2. It is important to remove the thick midrib from leaves, especially from maize, because it contains large amounts of C4 acids. If leaves of millet or guinea grass have a thin midrib, its removal may be omitted.

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Fig. 3. Protein pattern of mesophyll (M) and bundle sheath (BS) thylakoids isolated from Zea mays leaves. (a) SDS-PAGE in 12–20% polyacrylamide gradient. Samples 15 mg of chlorophyll were loaded in the well and gel was stained with Coomassie. (b) Immunodetection analysis of selected PSII proteins: D2 and Lhcb1, 2, 3 and 6. Two micrograms of chlorophyll was loaded in the well, 15% SDS-PAGE was used. Proteins were transferred onto membrane and detected with specific antibodies. Additional information on structural organization and protein distribution in BS thylakoids is in refs. 10, 14.

3. The cutting of leaf tissue to 1–2-mm strips is necessary for efficient grinding and elimination of mesophyll cells. 4. Sodium isoascorbate is unstable in solution; prepare fresh and add to grinding medium (80 mL) just before grinding. 5. The blades of the blender must be very sharp to produce strand cells with high metabolic activity. Even in a new blender the blades require sharpening usually. 6. Exact values of line voltage, duration, and number of grinding cycles depend on blender type, particularly on the size and shape of the cup, arrangement of blades, etc. and must be determined experimentally. The number of homogenization cycles, especially those at 40% of line voltage, greatly depends on plant species, light intensity during growth, and age of leaves. Always increase voltage using an autotransformer progressively during grinding. 7. Prepare 250–300 mL of resuspension medium (0–4°C). 8. Grind strands at dim light and protect the extracted chlorophyll from strong light in order to prevent its photo-oxidative loss.

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9. Removal of CO2 from strand suspension is necessary to minimize their photosynthetic metabolism (e.g. O2 evolution) recorded without added substrates. 10. The isolated bundle sheath strand cells, which are stored on ice in darkness, maintain metabolic function for many hours. During experimentation, they are usually very active for at least 2–3 h. These cells, unlike protoplasts, are extremely permeable to a variety of metabolites (12), and thus represent a unique system to study various aspects of cell metabolism. 11. For the detection of thylakoid phosphoproteins, add NaF (phosphatase inhibitor) to all the buffers throughout the isolation procedure at a final concentration of 10 mM. Prepare stock solution and add fresh just before use. 12. For protection of proteins from proteolysis add protease inhibitors: phenylmethylsulfonylfluoride (PMSF), amino-ncaproic acid (EACA), and benzamidine to media at final concentrations of 0.2  mM, 5  mM, and 1  mM, respectively. Prepare stock solutions and add just before use. 13. Notice that this method of isolation of bundle sheath strand cells differs from that described in Subheading 3.2. The former method describes the procedure to obtain bundle sheath cells with retained metabolic activities. They are collected immediately from filtrate after homogenization of leaf slices and then purified to remove contaminations, especially epidermal materials that may interfere with oxygen electrode performance during assays. Because the yield of chloroplast release, and therefore thylakoids, from bundle sheath cells is very low since their cell walls are resistant to breaking, it is necessary to obtain a large amount of strands. For this reason the bundle sheath strands (Subheading 3.2) are prepared by successive homogenization and washing of sediments until the residue (strands) are completely free of mesophyll cells. 14. This filtrate may be used for preparation of mesophyll chloroplasts. Remember, however, that these chloroplasts are injured; therefore, they are not able to fix CO2 photosynthetically but may be used for isolation of, e.g., thylakoids. 15. Only in the case of maize, water may be used instead of the thylakoid wash medium for further homogenization and washing strands. For both Panicum species, use thylakoid wash medium instead of water, since after water washing, there is a significant loss of PSI and PSII activity. 16. Homogenization could be longer if necessary, until microscopic examination shows single strands without attached mesophyll cells. 17. Chlorophyll a/b determined in BS maize thylakoids should be higher than 4.5.

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18. For native polyacrylamide gel electrophoresis of chlorophyll– protein complexes in resuspension medium for thylakoids, omit mannitol.

Acknowledgments We thank our PhD students Anna Drożak, Berenika Pokorska, and Hubert Jastrzębski involved for years in the studies on C4 plant photosynthesis. Theirs results are presented in the protocol. Studies were financed by grants from the Ministry of Science and Higher Education of Poland. References 1. Hatch, M.D. (1987) C4 photosynthesis: a unique blend of modified biochemistry, anatomy and ultrastructure. Biochim. Biophys. Acta 895, 81–106. 2. Ghirardi, M.L. and Melis, A. (1983) Localization of photosynthetic electron transport components in mesophyll and bundle bundle sheath chloroplasts of Zea mays. Arch. Biochem. Biophys. 224, 19–28. 3. Agostino, A., Furbank, R.T., and Hatch, M.D. (1989) Maximizing photosynthetic activity and cell integrity in isolated bundle cell strands from C4 species. Aust. J. Plant Physiol. 16, 279–290. 4. Ivanov, B.N., Sacksteder, C.A., Kramer, D.M., and Edwards, G.E. (2001) Light-induced ascorbate-dependent electron transport and membrane energization in chloroplasts of bundle sheath cells of the C4 plant maize. Arch. Biochem. Biophys. 385, 145–153. 5. Kanai, R. and Edwards, G.E. (1973) Separation of mesophyll protoplasts and bundle sheath cells from maize leaves for photosynthetic studies. Plant Physiol. 51, 1133–1137. 6. Schuster, G., Ohad, I., Martineau, B., and Taylor, W.C. (1985) Differentiation and development of bundle sheath and mesophyll thylakoids in maize. Thylakoid polypeptide composition, phosphorylation, and organization of photosystem II. J. Biol. Chem. 260, 11866–11873. 7. Bassi, R., Marquardt, J., and Lavergne, J. (1995) Biochemical and functional properties of photosystem II in agranal membranes from maize mesophyll and bundle sheath chloroplasts. Eur. J. Biochem. 233, 709–719.

8. Lu, Y.-K. and Stemler, A.J. (2002) Extrinsic photosystem II carbonic anhydrase in maize mesophyll chloroplasts. Plant Physiol. 128, 643–649. 9. Romanowska, E., Drożak, A., Pokorska, B., Shiell, B.J., and Michalski, W.P. (2006) Organization and activity of photosystems. J. Plant Physiol. 163, 607–618. 10. Romanowska, E., Kargul, J., Powikrowska, M., Finazzi, G., Nield, J., Drożak, A., and Pokorska, B. (2008) Structural organization of photosynthetic apparatus in agranal chloroplasts of maize. J. Biol. Chem. 283, 26037–26046. 11. Hipkins, M.F. and Baker, N.R. (1986) Spectroskopy. In: Hipkins, M.F. and Baker, N.R., eds. Photosynthesis energy transduction, a practical approach. IRL Press, Oxford, Washington DC, p. 51–101. 12. Weiner, H., Burnell, J.N., Woodrow, I.E., Heldt, H.W., and Hatch, M.D. (1988) Metabolite diffusion into bundle sheath cells from C4 plants. Relation to C4 photosynthesis and plasmodesmatal function. Plant Physiol. 88, 815–822. 13. Romanowska, E. and Drożak, A. (2006) Comparative analysis of biochemical properties of mesophyll and bundle sheath chloroplasts from various subtypes of C4 plants grown at moderate irradiance. Acta Biochim. Pol. 55, 709–719. 14. Pokorska, B., Zienkiewicz, M., Powikrowska, M., Drożak, A., and Romanowska, E. (2009) Differential turnover of the photosystem II reaction centre D1 protein in mesophyll and bundle sheath chloroplasts of maize. Biochim. Biophys. Acta. 1787, 1161–1169.

Chapter 26 Isolation of Ribulose-1,5-Bisphosphate Carboxylase/Oxygenase from Leaves A. Elizabete Carmo-Silva, Csengele Barta, and Michael E. Salvucci Abstract Ribulose-1,5-bisphosphate carboxylase/oxygenase (Rubisco) is a multifunctional enzyme that catalyzes the fixation of CO2 and O2 in photosynthesis and photorespiration, respectively. As the rate-limiting step in photosynthesis, improving the catalytic properties of Rubisco has long been viewed as a viable strategy for increasing plant productivity. Advances in biotechnology have made this goal more attainable by making it possible to modify Rubisco in planta. To properly evaluate the properties of Rubisco, it is necessary to isolate the enzyme in pure form. This chapter describes procedures for rapid and efficient purification of Rubisco from leaves of several species. Key words: Calvin cycle, Carbamylation, Carbon metabolism, CO2 fixation, Enzyme activation, Rate zonal centrifugation, Rubisco activase

1. Introduction As the most abundant protein on the planet (1) and the ratelimiting enzyme in photosynthesis, ribulose-1,5-bisphosphate carboxylase/oxygenase (Rubisco) needs little introduction. The large investment of leaf nitrogen in this one enzyme attests to both its importance for photosynthetic CO2 assimilation and its sluggish nature as a catalyst. Because of its peculiar reaction mechanism, Rubisco has a propensity for catalyzing unproductive side-reactions that further restrict its ability to catalyze CO2 assimilation efficiently (2, 3). Under certain circumstances, the products of these side-reactions, and even the substrate, RuBP, can act as tight-binding inhibitors of Rubisco that block catalysis until released by the action of Rubisco activase (4, 5). Since photosynthetic as well as water- and nitrogen-use efficiency can all be traced to the catalytic properties of Rubisco, improving Robert Carpentier (ed.), Photosynthesis Research Protocols, Methods in Molecular Biology, vol. 684, DOI 10.1007/978-1-60761-925-3_26, © Springer Science+Business Media, LLC 2011

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Rubisco has long been viewed as a viable strategy for increasing plant productivity (5, 6). A major impediment to this goal has been the inability to synthesize and assemble the higher plant enzyme in a recombinant system. Recently, advances in chloroplast transformation, both in higher plants (7) and in the green alga Chlamydomonas (8), have made it possible to modify Rubisco in planta. Analysis of these modified Rubisco proteins requires purification of the enzyme from leaves or algal cells. The abundance of Rubisco in leaves, i.e., up to 40 or 50% of the soluble protein (1), facilitates purification of the enzyme from most higher plant species. For our studies of the interaction with Rubisco activase, we have developed a 1-day procedure that uses rate zonal centrifugation and anion-exchange chromatography to separate Rubisco from contaminating proteins by mass and charge. The final preparation is highly active even after storage for several years as a frozen ammonium sulfate suspension.

2. Materials 2.1. Extraction of Rubisco from Leaves

1. Plant material: 30 g (fresh weight, FW) of leaf tissue of the species under study. We routinely store leaves frozen at −80°C in preweighed batches (see Note 1). 2. Basic extraction buffer: 50 mM Tris–HCl, pH 7.6, 20 mM MgCl2, 20 mM NaHCO3, and 0.2 mM EDTA. The buffer is filtered through a 0.2-mm filter and can then be stored for several weeks at 4°C (see Notes 2 and 3). 3. Rubisco extraction buffer: Basic extraction buffer plus 1 mM phenylmethanesulphonylfluoride (PMSF), 10 mM leupeptin, 5 mM dithiothreitol (DTT), and 2% (w/v) polyvinylpolypyrrolidone (PVPP, insoluble). Add these components immediately prior to use. 4. Blender (1 L) and varistat. 5. Cheesecloth and miracloth. 6. Refrigerated high-speed centrifuge with fixed-angle rotor for 50-mL tubes and 250-mL bottles (Thermo Fisher Scientific Inc., Waltham, MA or equivalent).

2.2. Precipitation with Ammonium Sulfate

1. Ammonium sulfate, solid and saturated (~4.1  M) solution, adjusted to pH 7.0 with NH4OH. 2. Separatory funnel.

2.3. Rate Zonal Centrifugation using Sucrose Gradients

1. Sucrose gradients: 0.2–0.8  M sucrose in half-strength (½) basic extraction buffer containing 5 mM DTT. Fill to within 2 mL of the top of the ultracentrifuge tube (see Note 4).

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2. Disposable polyallomer tubes (Optiseal, 32.6  mL capacity, Beckman–Coulter, Palo Alto, CA). 3. Peristaltic pump. 4. Gradient maker (at least 30 mL capacity). 5. Preparative ultracentrifuge with large capacity rotor (70 Ti fixed-angle rotor, Beckman–Coulter). 6. UV detector and fraction collector. 2.4. Anion-Exchange Chromatography on HiTrap Q Sepharose

1. Anion-exchange buffer A: ½ Basic extraction buffer containing 2 mM DTT. 2. Anion-exchange buffer B: ½ Basic extraction buffer containing 2 mM DTT and 1 M KCl. 3. Anion-exchange columns, Q-Sepharose HiTrap, 3 × 5  mL (GE-Healthcare/Amersham Biosciences, Piscataway, NJ, or equivalent). 4. Low, medium, or high pressure chromatographic system with UV-detector and fraction collector (ÄKTA, GE-Healthcare/ Amersham Biosciences, or equivalent).

2.5. Concentration and Storage

1. Ammonium sulfate, saturated (~4.1 M) solution, adjusted to pH 7.0 with NH4OH. 2. Liquid N2 (LN2). 3. Glass vials (20 mL).

2.6. Determination of Rubisco Concentration

1. UV–Vis spectrophotometer.

2.7. Centrifugal Desalting for Assay of Activity

1. Rubisco activation buffer: 100 mM Tricine–NaOH, pH 8.0, 10 mM MgCl2, 10 mM NaHCO3, and 50 mM DTT.

2. Quartz cuvette.

2. Columns, 0.8 × 5 cm, packed with 2-mL Sephadex G-50-fine. 3. Refrigerated high-speed centrifuge with swinging-bucket rotor (Thermo Fisher Scientific Inc., or equivalent). 4. Screw-cap microfuge tubes (0.5 mL).

3. Methods All procedures for enzyme extraction and purification are conducted at 4°C and as rapidly as possible in order to minimize proteolytic activity and maximize protein recovery. 3.1. Extraction of Rubisco from Leaves

1. Harvest leaves in the light (at least 2 h after the start of the photoperiod) and remove the mid-ribs, if practical. Divide the tissue into batches of known fresh weight, quickly freeze

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in LN2 and store at −80°C. In our experience, about 30  g (FW) of Arabidopsis leaf tissue (rosettes) yields about 75 mg of Rubisco protein. 2. Prepare Rubisco extraction buffer by adding PMSF, leupeptin, and DTT to 200 mL of basic extraction buffer. Add this solution to a prechilled 1-L blender containing 4 g of PVPP and mix by blending for 15 s at high speed and then adjust the blender to low speed with a varistat (see Note 1). 3. Quickly powder 30 g (FW) of frozen leaf tissue and gradually add it to the blender until all the powder has been mixed with Rubisco extraction buffer. 4. Increase the speed of the blender to high speed and thoroughly grind the leaf tissue with several 15-s bursts. 5. Filter the homogenate through four layers of cheesecloth and one layer of miracloth suspended over a plastic funnel to remove cell walls and other unbroken material. To recover as much extract as possible, gently squeeze the suspension by hand (use gloves). The cheese- and miracloth are washed extensively with Milli-Q water and stored hydrated at 4°C before use. 6. Transfer the filtrate to ice-cold 250-mL centrifuge bottles. 7. Centrifuge for 12 min at 23,500 × g in a fixed-angle rotor to remove cell walls and membranes plus PVPP. Discard the green/white pellet and save the supernatant. (Save a small aliquot of the crude supernatant for later analysis.) 3.2. Precipitation with Ammonium Sulfate

1. Measure the volume of the supernatant and add saturated ammonium sulfate to 45%. The volume of saturated ammonium sulfate required for precipitation is calculated as follows: Volume of saturated ammonium sulfate for 45% saturation =  (volume of the supernatant/0.55) − volume of the supernatant. Add the saturated ammonium sulfate drop-wise to the supernatant with continuous stirring using a separatory funnel in a ring stand. Stir for 30 min. 2. Transfer the suspension to 250-mL bottles and centrifuge for 13 min at 27,500 × g in a fixed-angle rotor to remove precipitated material (mostly membranes). Discard the pellet and keep the supernatant. 3. Slowly add solid ammonium sulfate to the straw-colored supernatant to achieve 57% saturation. About 78.6 g/L are required to increase the ammonium sulfate concentration from 45 to 57%. Stir for a total of 30 min. 4. Centrifuge the suspension for 13 min at 23,500 × g in a fixedangle rotor to collect the precipitated protein. 5. Suspend the protein pellet in about 8–10  mL of Rubisco extraction buffer minus PVPP (see Note 5).

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3.3. Isolation by Rate Zonal Centrifugation Using Sucrose Gradients (see Notes 6 and 7)

343

1. Divide the Rubisco solution into 1.5  mL aliquots. (Save a small aliquot of the pregradient material for later analysis.) 2. Carefully load 1.5 mL of the solution to each sucrose gradient. 3. Overlay the samples with Rubisco extraction buffer minus PVPP, plug the filled tubes with the stoppers, and load into a fixed-angle rotor for ultracentrifugation. 4. Centrifuge for 2.5 h at 435,000 × g (rmax) to separate Rubisco from lower molecular mass proteins. 5. Fractionate each gradient using a siphoning tube lowered to within 2 cm of the bottom of the tube and connected to a peristaltic pump. Collect fractions while monitoring the absorbance at 280 nm. The peak corresponding to Rubisco is recovered in the early fractions corresponding to the bottom 1/3 of the gradient (Fig. 1).

Fig. 1. UV profile of proteins in the 45–57% ammonium sulfate fraction separated by rate zonal centrifugation in sucrose gradients. The trace shows a typical peak of Rubisco near the bottom of the gradient, well-resolved from other proteins.

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6. Pool the fractions containing Rubisco. (Save an aliquot of the pooled sucrose gradient fraction separately for later analysis by SDS-PAGE). 3.4. Purification by Anion-Exchange Chromatography on HiTrap Q Sepharose

1. Load the Rubisco-containing fraction on a 10–15-mL HiTrap Q Sepharose column previously equilibrated with five column volumes of Anion-exchange buffer A. 2. Elute the column at 3  mL/min with a 200–300-mL linear gradient from 0 to 0.5 M KCl. The gradient is produced by mixing Anion-exchange buffer A (no KCl) with Buffer B (1 M KCl). Rubisco elutes from the anion-exchange column with ca. 0.35 M KCl. 3. Pool the peak fractions. (Save an aliquot of the pooled anionexchange sample separately for later analysis.)

3.5. Concentration and Storage

1. Measure the volume of the Rubisco solution recovered from the anion-exchange column. 2. Slowly add saturated ammonium sulfate to a final saturation of 60%. 3. Centrifuge for 10  min at 16,500 × g in a swinging-bucket rotor to collect the precipitated Rubisco protein. 4. Suspend the pellet in a minimum volume of the supernatant. 5. Flash-freeze the protein suspension as droplets by pipetting drop-wise into a 20-mL glass vial containing LN2. Store the final preparation as frozen beads at −80°C.

3.6. Determination of Rubisco Amount and Purity

The amount of purified Rubisco can be calculated based on its absorbance at 280  nm. The purity of the preparation can be assessed by SDS-PAGE analysis of the aliquots collected during the purification process (Fig. 2). 1. Determine the Rubisco concentration by measuring the absorbance at 280 nm (9). Use 10 mL of the pooled anionexchange fractions in 0.5  mL of ½ basic extraction buffer with 2 mM DTT and 0.35 M KCl (see Note 8). 2. Analyze a subsample of each aliquot taken at the various steps of purification (i.e., crude supernatant, pregradient, sucrose gradient, and anion-exchange fractions) by SDS-PAGE (10).

3.7. Desalting Rubisco in Preparation for Assay

The activity of Rubisco is determined from the RuBP-dependent incorporation of 14CO2 into acid-stable products (see accompanying Chapter 29). Prior to determining activity, the Rubisco preparation is desalted by centrifugal gel filtration (adapted from (11)). 1. Transfer the frozen droplets of purified Rubisco to a microfuge tube and thaw on ice. Collect the precipitated protein by centrifugation for 10 min at 10,000 × g. Discard the supernatant

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Fig. 2. Coomassie blue stained gel showing standard molecular weight markers (M) and the polypeptides in the various fractions from the Rubisco purification: CS, crude supernatant after extraction; AS, 45–57% ammonium sulfate fraction prior to loading on sucrose gradients; SG, Rubisco peak from sucrose-gradient; AX, Rubisco peak from anion-exchange column, after separation by SDS-PAGE.

and suspend the pellet in 150 mL of Rubisco activation buffer. Incubate for 1 h at 4°C to fully carbamylate the enzyme (see Note 9). 2. Equilibrate 2 mL of Sephadex G-50-fine with three column volumes of Rubisco activation buffer containing 5 instead of 50 mM DTT. 3. Place the column on top of a 0.5-mL screw-cap microfuge tube and place the entire assembly inside a 15-mL Corex glass tube. 4. Centrifuge the column at 400 × g for 2  min in a swingingbucket rotor to remove excess buffer. Discard the eluant from the microfuge tube. Centrifuge the column for an additional 1 min and again discard the eluant. 5. Carefully add 100–250 mL of fully carbamylated Rubisco to the top of the dry column bed. 6. Centrifuge the column and collection tube assembly for 2 min at 400 × g. 7. Recover the desalted sample from the microfuge tube. 8. Determine Rubisco concentration from the absorbance at 280 nm (see Note 8). 9. Assay Rubisco activity as described in the accompanying Chapter 29.

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4. Notes 1. This procedure has been used for isolation of Rubisco from tobacco, spinach, Arabidopsis, and Xanthium. For Chlamy­ domonas, prepare extracts by sonication instead of extracting in a blender. 2. The buffer used for extraction should promote recovery and stability of Rubisco. A pH of 7.6 is used with high concentrations of Mg2+ and CO2 to keep the enzyme in the active (carbamylated) state. Protease inhibitors (PMSF and leupeptin), and DTT are included, as well as PVPP to adsorb phenolic compounds. Prepare the basic extraction buffer with all components except protease inhibitors, DTT, and PVPP and add these components immediately prior to use from concentrated stocks (PMSF and leupeptin) or as solids (DTT and PVPP). Add PVPP directly to the blender and then add the extraction buffer with the other components. 3. For leaf tissue with high phenolic content, use 0.1 M borate buffer at pH 7.8 in place of Tris–HCl. The use of borate buffer for plant species like cotton and creosote improves the recovery of protein by preventing cross-linking (12, 13). 4. For the sucrose gradients, prepare 0.2 and 0.8  M sucrose solutions in half-strength basic Rubisco extraction buffer containing 5 mM DTT. Prepare each gradient by mixing the two sucrose solutions using a gradient maker and peristaltic pump. The gradients are prepared the day before the purification and stored at 4°C overnight. 5. Depending on the density, it may be necessary to increase the volume of extraction buffer used to suspend the protein pellet to ensure that the solution is not too dense for the sucrose gradients. This problem often occurs when concentrations of ammonium sulfate greater than 60% are used for protein precipitation. 6. Because of its large mass (i.e., 530,000D) and compact structure, Rubisco migrates as an 18S particle on sucrose gradients (14). 7. Preparative gel-filtration chromatography through a semirigid matrix like Sephacryl S-300 can be used in place of rate zonal centrifugation. However, in our experience, the separation of Rubisco from other proteins is much cleaner on sucrose gradients than on gel filtration columns. A chromatography column with a bed volume of about 400  mL is required to accommodate 12  mL, the volume of sample applied to eight sucrose gradients. 8. Use a 1-mL quartz cuvette containing 0.5 mL of half-strength basic extraction buffer with 2 mM DTT and 0.35 M KCl to

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zero the spectrophotometer. For determining the concentration of spinach or Arabidopsis Rubisco, use the formula, Rubisco concentration (mg/mL) = A280 × 0.61 (9). For tobacco, use A280 × 0.71 (15). 9. Sulfhydryl groups of Rubisco can oxidize to form disulfides and these modifications can compromise the catalytic activity of the enzyme (recently reviewed in (16)). Therefore, we routinely incubate Rubisco with a high concentration of reducing agent by including DTT in our activation buffer (15).

Acknowledgments This research was funded by the US Department of Agriculture, Agricultural Research Service and the US Department of Energy, grant no. DE-FG02-08ER20268. Mention of a trademark, proprietary product, or vendor does not constitute a guarantee or warranty of the product by the United States Department of Agriculture and does not imply its approval to the exclusion of other products or vendors that may also be suitable. References 1. Ellis, R. J. (1979) The most abundant protein in the world. Trends Biochem Sci 4, 241–4. 2. Andrews, T. J. (1996) The bait in the Rubisco mousetrap. Nat Struct Biol 3, 3–7. 3. Pearce, F. G. and Andrews, T. J. (2003) The relationship between side reactions and slow inhibition of ribulose-bisphosphate carboxylase revealed by a loop 6 mutant of the tobacco enzyme. J Biol Chem 278, 32526–36. 4. Portis, A. R. Jr. (2003) Rubisco activase – Rubisco’s catalytic chaperone. Photosynth Res 75, 11–27. 5. Spreitzer, R. J. and Salvucci, M. E. (2002) Rubisco: structure, regulatory interactions, and possibilities for a better enzyme. Annu Rev Plant Biol 53, 449–75. 6. Parry, M. A. J., Madgwick, P. J., Carvalho, J. F. C., and Andralojc, P. J. (2007) Prospects for increasing photosynthesis by overcoming the limitations of Rubisco. J Agric Sci 145, 31–43. 7. Whitney, S. M. and Sharwood, R. E. (2008) Construction of a tobacco master line to improve Rubisco engineering in chloroplasts. J Exp Bot 59, 1909–21. 8. Satagopan, S. and Spreitzer, R. J. (2008) Plantlike substitutions in the large-subunit carboxy terminus of Chlamydomonas Rubisco increase CO2/O2 Specificity. BMC Plant Biol 8, 85.

9. Wishnick, M. and Lane, M. D. (1971) Ribulose diphosphate carboxylase from spinash leaves. Meth Enzymol 23, 570–7. 10. Chua, N.-H. (1980) Electrophoretic analysis of chloroplast proteins. Meth Enzymol 69, 434–46. 11. Penefsky, H. S. (1972) Reversible binding of Pi by beef heart mitochondrial adenosine triphosphate. J Biol Chem 252, 2891–9. 12. King, E. E. (1971) Extraction of cotton leaf enzymes with borate. Phytochemistry 10, 2337–41. 13. Crafts-Brandner, S. J. and Salvucci, M. E. (2000) Rubisco activase constrains the photosynthetic potential of leaves at high temperature and CO2. Proc Natl Acad Sci USA 97, 13430–5. 14. Kawashima, N. and Wildman, S. G. (1970) Fraction I protein. Annu Rev Plant Physiol 21, 325–58. 15. McCurry, S. D., Gee, R., and Tolbert, N. E. (1982) Ribulose-1,5-bisphosphate carboxylase/oxygenase from spinach, tomato, or tobacco leaves. Meth Enzymol 90, 515–21. 16. Moreno, J., García-Murria, M. J., and MarínNavarro, J. (2008) Redox modulation of Rubisco conformation and activity through its cysteine residues. J Exp Bot 59, 1605–14.

Chapter 27 Quantifying the Amount and Activity of Rubisco in Leaves David S. Kubien, Christopher M. Brown, and Heather J. Kane Abstract The CO2-fixing enzyme Rubisco plays a crucial biological role as a primary determinant of both plant yield and the response of the biosphere to global change. Here, we describe techniques for measuring the amount and activity of Rubisco in higher plants. To accommodate a range of experimental capabilities, we describe basic radioisotopic methods as well as non-radioactive techniques. The required calculations are included. We discuss problems that commonly arise during the extraction and assay of Rubisco. Key words: Photosynthesis, 14CO2, Spectrophotometry, 14C-CABP, Quantitative immunoblotting

1. Introduction Ribulose-1, 5-bisphosphate carboxylase/oxygenase (Rubisco, E.C. 4.1.1.39) is the most abundant protein on Earth (1), comprising as much as half of the protein in leaves. Rubisco catalyses the carboxylation of ribulose-1, 5-bisphosphate (RuBP), producing two molecules of 3-phosphoglycerate (3-PGA). This irreversi­ ble first step of photosynthesis is the entry point for carbon into the biosphere. The amount and kinetics of Rubisco are principal determinants of the rate and efficiency of CO2 fixation in the leaves of C3 plants (2, 3). Even with a carbon concentrating metabolism, such as that found in the leaves of C4 species, Rubisco activity can be the rate limiting step of photosynthesis under certain conditions (4). Predicting the effects of global climate change on photosynthesis depends on our ability to model the process, and all useful mathematical models of photosynthesis directly incorporate the kinetics and amount of Rubisco (5). This chapter describes the quantification of Rubisco from higher plant leaves. We describe basic procedures for sample collection, extraction, and activation of Rubisco, and then describe Robert Carpentier (ed.), Photosynthesis Research Protocols, Methods in Molecular Biology, vol. 684, DOI 10.1007/978-1-60761-925-3_27, © Springer Science+Business Media, LLC 2011

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Fig. 1. Spectrophotometric coupled-enzyme assay for Rubisco activity. Rubisco produces 3-phosphoglycerate (3-PGA), which is phosphorylated by 3-PGA kinase (PGK). Glyceraldehyde-3-P dehydrogenase (GAPDH) reduces the bis-PGA to an aldehyde (Ga-3-P). Triose-phosphate isomerase (TIM) produces di-hydroxyacetone phosphate (DHAP) from Ga-3-P; glycerol-P dehydrogenase (GlyPDH) reduces DHAP. Rubisco produces two molecules of 3-PGA for each RuBP carboxylated; four NADH are oxidised per RuBP.

two methods for measuring both the amount and maximum carboxylation activity of the enzyme. Rubisco content is determined by the stoichiometric binding of radio-labelled 2¢-carboxyarabinitol-1, 5-bisphosphate (14C-CABP) to the catalytic sites of the enzyme (6, 7), or by polyacrylamide gel electrophoresis (PAGE) followed by quantitative immuno-detection methods (8, 9). The maximum carboxylation activity of Rubisco in vitro (e.g. Vcmax) may be determined by 14CO2 uptake, followed by liquid scintillation counting of the acid-stable products (10–13) Alternatively, activity can be measured continuously, using an NADH-linked coupled enzyme system and a spectrophotometer (14, 15) (Fig. 1). In either case, Rubisco activity is determined in the presence of near-saturating CO2 levels, to determine carboxylation capacity.

2. Materials All chemicals used are available from commercial sources, and the highest grade chemicals and water are recommended. Prepare sufficient volumes of all stock solutions for the whole experiment, and store at 4°C. Radiochemicals are available from MP Biomedicals (http://www.mpbio.com) and Perkin has taken over the GE radioisotope products. Observe local rules for radioisotope safety and waste disposal. 2.1. Radiochemicals

1. 2 mM 14C-CABP (see Note 1). 2. 315 mM NaH14CO3.

2.2. Extraction Buffer

1. HEPES buffer stock: 100  mM HEPES–KOH, pH 7.5, 20 mM MgCl2, 1 mM Na2–EDTA. 2. ACA-BAM (protease inhibitor solution): 200 mM 6-amino caproic acid, 40 mM benzamidine. 3. Extraction buffer (EB): 5  mM DTT, 10  mg/mL PVPP, 2 mg/mL BSA, 2 mg/mL PEG, 2% (v/v) Tween-80, 12 mM 6-amino caproic acid, 2.4 mM benzamidine, in HEPES stock (see Note 2).

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2.3. Radioisotopic Activity Assay Buffer

351

1. Assay buffer (BME): 100 mM Bicine-NaOH, pH 8.2, 20 mM MgCl2, 1 mM Na2-EDTA. 2. Rubisco activating solution (BMEC): 100 mM NaH12CO3 in BME. 3. Rubisco activity assay buffer: 500  mM RuBP, 10  mM NaH14CO3 in BME (see Note 3).

2.4. Spectro­ photometric CoupledEnzyme Assay Buffer

1. Assay buffer (EME): 100 mM EPPS–NaOH, pH 8.2, 20 mM MgCl2, 1 mM Na2–EDTA. 2. ATP/phosphocreatine: 10 mM/50 mM. 3. Coupling enzymes (CE): (see Table 1). 4. Coupling enzyme buffer (EED): 50 mM EPPS–NaOH, pH 7.8, 1 mM EDTA, 1 mM DTT. 5. 10 mM NADH in EME. 6. 250 mM NaH12CO3 in EME.

2.5. Quantitative Immunoblotting

See Note 4 for discussion of this technique. 1. Quantitative immunoblotting requires a precise standard. A quantified Rubisco standard (0.15  pmol RbcL/mL in 1× loading buffer) is available from (Agrisera). 2. 4× blue sample loading buffer (Invitrogen). 3. Pre-stained molecular weight marker (e.g. Precision Plus, Bio-Rad). In addition, a molecular weight marker that binds

Table 1 Coupling enzymes for spectrophotometric Rubisco activity assay. All enzymes were purchased from Sigma, in December 2007. The final volume was 8.0 mL, including 20% (v/v) glycerol. The amount per assay assumes 40 mL of coupling enzymes per 2 mL assay Enzyme

Cat. #

Amount

Units/mg

Units

Units/2 mL assay

Creatine P-kinase

C3755

54 mg

185

10,000

50

Carbonic anhydrase

C3934

40 mg

>2,500

>100,000

>500

Glyceraldehyde-3-P dehydrogenase

G9263

90 mg

112

10,000

50

3-Phosphoglycerate kinase

P7634

5.4 mg (0.934 mL)

1,845

10,000

50

Triose-P Isomerase/ Glycerol-3-P dehydrogenase

G1881

50.4 mg (8.32 mL)

1,590/159

80,000/8,000

400/40

a

a

Indicates that the enzyme(s) is shipped as a suspension in (NH4)2SO4

a

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the secondary antibody and generates a chemiluminescent signal (e.g. Invitrogen Magic Marks LC5600) may run separately or blended with the pre-stained marker at a ratio of 1:30 mL. 4. Transfer buffer: Concentrated commercial stock (e.g. Invitrogen NP0006) diluted to 1× with water, 10% (v/v) methanol, 500 mM DTT (1 mL of 0.5 M DTT/L of buffer). 5. Anti-RbcL primary antibodies from rabbit or hen are commercially available (Agrisera AS03 037 or AS01 017). The slightly lower affinity of the IgY from egg yolks makes it very suitable for quantifying highly abundant Rubisco in plants. 6. Secondary antibody: horseradish peroxidase (HRP) conjugated anti-rabbit or anti-chicken, depending on the primary antibody used. 7. Tris-buffered saline (TBS), 5× stock: 100 mM Tris base, pH 7.6, 4% (w/v) NaCl. Dilute with water before use to give 1× TBS. 8. Immuno-blot washing solution (TBS-T): 0.1% Tween-20 in 1× TBS. 9. Blocking solution: commercial system as per manufacturer’s instructions, in TBS-T. Other common blocking systems [e.g. 5% (w/v) non-fat milk powder in TBS-T] can work well. 10. Chemiluminescent detection system. 11. PVDF or nitrocellulose membranes cut to the exact size of the gel. 12. Whatman filter paper, cut to the exact size of the gel. 13. Protein standard assay (e.g. Lowry, BCA). 14. PAGE and blotting systems: (e.g. Invitrogen NuPAGE Bis– Tris pre-cast gels with MOPS or MES running buffers). 2.6. Chromatography buffer and columns

See Note 4 for discussion of this technique. 1. Column elution buffer: 20 mM EPPS-NaOH (pH 8.0), 75 mM NaCl, degassed. 2. Low-pressure chromatography columns (0.7 x 30 cm, with stopcocks). 3. Sephadex G-50 (fine).

3. Methods Tissue grinders, EB, and assay buffers should be prepared and kept on ice before sample grinding. Equilibrate a water bath or heating plate (for 14CO2 assays) or spectrophotometer (if so equipped) to the desired assay temperature (normally 25°C). If measuring activity by 14CO2 uptake a heating plate (80°C) is

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353

needed for sample drying (use one with holes that fit the vials). Drying the samples at higher temperature saves time, but caramelization of the samples can lead to colour quenching during scintillation counting (20). 3.1. Radioisotopes 3.1.1. 14C-CABP

CABP is an analogue of the 6-carbon intermediate of the Rubisco carboxylation reaction and binds to carbamylated Rubisco catalytic sites. A working solution of 14C-CABP is prepared from Na14CN or K14CN (50–62  mCi/mmol) (6). Dissolve 1  mCi (37 MBq) of 14CN in 10 mM Na2CO3 (pH 10.0), to give a final concentration of 100 mM 14CN (approximately, 190 mL of Na2CO3 required, depending on radioactivity of the 14CN). In a glass scintillation vial, add 3  mL 10  mM Na2CO3, 150  mL of 100  mM RuBP, and the 14CN solution, and incubate at room temperature overnight (~16 h). Stop the reaction with 500 mL of 2 N HCl, and evaporate to dryness on a heating block at 40°C under a gentle stream of N2. Suspend in BME buffer (see Subheading 2.3 to give a final concentration of 2  mM 14C-CABP (about 7  mL). Store 200 mL aliquots at −20°C. Thaw and maintain at 4°C before use.

3.1.2. NaH14CO3

Prepare a working solution of NaH14CO3 by diluting a concentrated (50–62  mCi/mmol) commercial stock. Make 40  mM Bicine, acidify with HCl, and degas with N2 on ice for 30 min. Bring to pH 9.0 with 50% NaOH, and degas for an additional 30  min on ice. Make 375  mM NaH12CO3 using the degassed Bicine-NaOH. Combine 4 mL of 375 mM NaH12CO3 and 1 mCi (37  MBq, approximately 1  mL of solution, depending on the activity) of commercial NaH14CO3 solution. The final concentration needs to be determined precisely, but it will be about 315 mM HCO3- (total 12C and 14C) depending on concentration of the commercial stock NaH14CO3. Store 1 mL aliquots of this working solution at 4°C.

3.2. Sampling

Collect leaf disc samples from illuminated leaves, using a cork borer of known cross-sectional area; seal in a microcentrifuge tube (with the cap punctured) and immediately freeze in LN2. Samples can be stored at −80°C for a few weeks, if necessary, but refreeze them in LN2 prior to extraction.

3.2.1. Sample Collection

3.2.2. Extraction

Tenbroek glass-in-glass homogenizers are convenient for sample grinding. If using commercial protease inhibitor cocktail (see Note 2), add the appropriate volume to each grinder, and set on ice. Add ice-cold EB (~2 mL/cm2) and the frozen leaf disc to the homogenizer, and grind thoroughly for about 30 s. Briefly centrifuge the extract at 4°C (<30 s, it is usually sufficient to bring a standard bench-top centrifuge up to maximum speed, ~14,000 × g, and then stop it), and discard the pellet. If determining Rubisco content by immunoblotting, collect an aliquot, freeze in LN2, and proceed to 3.5 or store at −80°C until use.

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3.2.3. Activation

Rubisco requires Mg2+ and CO2 for catalysis. For activity measurements or 14C-CABP quantification, add extract to BMEC in a 9:1 ratio (e.g. 900 mL extract to 100 mL BMEC, giving 10 mM NaHCO3 final concentration) to activate the Rubisco. Incubate the enzyme at 25°C for 10–20 min.

3.3. Activity Assays

In this assay, Rubisco is supplied with 14CO2 (as bicarbonate) and RuBP for a specified interval (usually 30–60 s). The acid-stable products are counted by liquid scintillation counting. The 14CO2 activity assay produces small amounts of radioactive volatiles, and must be done in a fumehood.

3.3.1. 14CO2 Uptake

1. Determine the specific activity (SA) of the working NaH14CO3 solution, while preparing the assay buffer. Add 990  mL of 500 mM NaOH to one scintillation vial and 300 mL to each of three vials. Add 10 mL of the working NaH14CO3 to the first vial (final volume 1 mL) and vortex thoroughly. Take 10 mL of this solution to each of the other three vials, add scintillation cocktail, vortex thoroughly, and set aside for liquid scintillation counting. 2. Prepare three blanks by adding 250 mL of activity assay buffer and 250 mL of 2 N HCOOH to glass vials, and place on the drying block at 80°C. When dry, add 200 mL of H2O to dissolve the residue, add scintillation cocktail, vortex thoroughly, and set aside for scintillation counting. 3. Assay each sample in duplicate or triplicate. Add 250 mL of assay buffer to three glass sample vials, and equilibrate in a water bath or on heating plate (25°C) for 60–90 s. Initiate the reaction by adding 50  mL of activated leaf extract (see Subheading 3.2.3) to the vials, and swirl gently to mix. Stop the reactions after 30 s with 250 mL 2 N HCOOH. It is convenient to assay replicates simultaneously by adding extract to sample vials at 0, 10, and 20 s, and terminating the reactions at 30, 40, and 50 s. Dry the samples at 80°C. Add 200 mL of water; once the residue dissolves add scintillation cocktail, vortex thoroughly, and set aside for scintillation counting. Keep the remaining activated extract on ice ready for incubating with 14C-CABP (see Subheading 3.4.1). 3.3.2. Spectrophotometric Coupled-Enzyme Assay

In this assay, Rubisco is supplied with 12CO2 (as bicarbonate) and RuBP, and produces 3-PGA. By including specific coupling enzymes and an ATP-generating system in the assay, 3-PGA is reduced to triose-phosphates and glycerol-phosphate (Fig.  1). The oxidation of NADH during the assay is continuously monitored by changes to A340nm; Rubisco activity is calculated from the rate of NADH oxidation.

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1. Prepare a stock solution of coupling enzymes in advance (see Table 1). Mix the enzymes that are suspended in (NH4)2SO4, centrifuge (10,000 × g, 4°C, 20 min), and dissolve the pellet in 6 mL EED buffer. Add the remaining enzymes and dialyse overnight against EED buffer at 4°C. Centrifuge as above, measure the supernatant volume and add glycerol to 20% final concentration (~1.6 mL). Freeze small aliquots (200 mL) in LN2, and store at −80°C. 2. The final assay volume is 2 mL. Add 1,550 mL of the EME buffer to a cuvette, and blank the spectrophotometer at 340 nm. Add 40 mL 10 mM of NADH (final concentration 200 mM), 80 mL of 250 mM NaH12CO3 (10 mM), 200 mL of ATP/phosphocreatine (1 and 5 mM, respectively), 40 mL of coupling enzymes, 40 mL of 25 mM RuBP (500 mM), and mix gently. The final concentration of each coupling enzyme is shown in Table 1. 3. Initiate the reaction by adding 50  mL of the activated leaf extract (see Subheading  3.2.3) and monitor A340 continuously. Assays may be repeated, with higher or lower amounts of Rubisco as appropriate (with corresponding adjustment to the volume of EME), depending on the rates observed. 3.4. Quantifying Rubisco Content with 14C-CABP 3.4.1 Sample Preparation 3.4.2. Chromatographic Separation of RubiscoBound and Unbound 14 C-CABP

Thoroughly vortex a thawed aliquot of 14C-CABP. Add 2 mL of 14 C-CABP to a microcentrifuge tube, and add 50 mL of activated plant extract (see Subheading  3.2.3). Incubate at 25°C for 20–30 min, and then keep on ice until ready to load columns. Size exclusion chromatography is used to separate 14C-CABPRubisco complexes from unbound 14C-CABP (7). Pack low pressure chromatography columns (0.7 × 30 cm, with stopcocks) with Sephadex G-50, pre-swollen and de-gassed in buffer (20  mM EPPS-NaOH, 75 mM NaCl, pH 8). Fractions are gravity-eluted, at approximately 0.5 mL/min (see Table 2). 1. Open the column outlet valve, and drain the buffer to the top of the Sephadex gel. Gently load the sample, open the valve, and drain the sample into the gel. Wash the sample into the gel with 200 mL of column buffer, followed by three 750 mL aliquots of column buffer. 2. Collect the next seven 750 mL washes in plastic scintillation vials, as shown in Table 2. Add scintillation cocktail to each sample, vortex thoroughly, and count by liquid scintillation. The first collected fraction is considered a background count. The next two fractions contain the high molecular weight Rubisco fraction with bound 14C-CABP, while the remaining fractions contain low molecular weight components, including unbound 14C-CABP.

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Table 2 Fraction collection for chromatographic separation of Rubisco-bound and free 14C-CABP. The counts from the two bound 14C-CABP fractions (samples 6 and 7) should be less than about 10% of the total counts from fractions 6–11 (see Note 1) Step

#

mL

Collect

1

Load

100

No

 1

200

No

 2

750

No

 3

750

No

 4

750

No

 5

750

Yes

Background

 6

750

Yes

Bound 14C-CABP

 7

750

Yes

Bound 14C-CABP

 8

750

Yes

 9

750

Yes

10

750

Yes

11

750

Yes

12

20 mL

Yes

2

3

Sample

Free 14C-CABP

Waste

3. After collecting seven fractions continue to wash the columns with 20 mL of storage buffer (column buffer containing 0.2% (w/v) NaN3). Collect this in a plastic 20 mL scintillation vial for disposal. 3.5. Quantitative Immuno-Detection

A variety of PAGE and blotting systems are available. The procedures described here are optimal for the reagents and antisera outlined. It is advisable to conduct a pilot experiment, to establish the sensitivity of RbcL detection in your system, and to calibrate your sample and standard loads.

3.5.1. SDS-PAGE and Western Blotting

1. Thaw the sample(if required, see Subheading  3.2.2) and determine protein content using a standard assay. Prepare samples for SDS-PAGE with 0.2  mg (total protein)/mL, 50 mM DTT, 1× loading buffer, and ultrapure H2O to a final volume of 50 mL. Prepare Rubisco standards to the appropriate concentration, in 1× loading buffer with 50 mM DTT. 2. Heat samples and standards to 65°C for 5 min prior to loading. Allow to cool for 1–2 min. Centrifuge briefly to bring all material to bottom of tube.

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3. Load samples (1 mg total protein) and standards on NuPAGE Bis–Tris gels. With the power supply set on constant voltage, run the gel at 200 V for 40 min. 4. Immerse PVDF membrane in methanol for 30  s. Rinse and soak in transfer buffer until ready to set up the transfer. For nitrocellulose membranes, soak in transfer buffer prior to use. 5. Set up transfer apparatus according to manufacturer instructions. Transfer protein to membrane at 30 V for 1 h if doing a single blot, 75  min if transferring two blots in the same transfer cell. 3.5.2. Antibody Incubations and Signal Detection

Incubate and wash the blot at room temperature on a shaking platform. 1. Incubate blot in blocking buffer for 1 h. 2. Incubate blot with primary antibody (diluted 1:20,000 in blocking buffer) for 1 h. 3. Wash the blot with two short rinses in TBS-T, followed by a 15-min wash and three 5-min washes. 4. Incubate blot with secondary antibody (1:50,000) for 1  h, and wash as described in step 3. 5. Develop the blot as per manufacturer’s instructions. 6. Capture image of the blot with an imager or by exposing to X-ray film to measure chemiluminescence. X-ray film is sensitive but has a narrower dynamic range than most imagers.

3.6. Calculations

3.6.1. Amount of Rubisco

Radioisotopic calculations are shown with units DPM (decays per minute, 1 DPM = 60  Bq), and square brackets indicate solute concentrations. 1. 14C-CABP specific activity: 14

CABP =

14 14 æ X mCi* ö æ mmol K CN ö æ 8 mmol CABP** ö è mmol K 14CN ø çè mmol 14CABP ÷ø çè 8 mmol sites ÷ø 9 8 mmol sites æ ö æ 2.22x10 DPM ö æ 68.75mg ö çè ÷ç ÷ø è nmol sites ø 6 550 x 10 mg rubisco ø è mCi

14

(

CABP = 2216.5 ´ DPM/nmol sites

)

The specific activity of 14C-CABP depends on the activity of the 14 CN*; for 53.3  mCi/mmol K14CN the specific activity of 14 C-CABP is 118319 DPM/nmol sites. Butz and Sharkey (21) found that 14C-CABP occupies only 6.5 of the 8 active sites. This is disputed (S.M. Whitney, personal communication), but can be incorporated to this calculation**.

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2. Rubisco concentration: æ Bound 14C-CABP (DPM) ö mmol/m 2 = ç 14 CABP (DPM/nmol) ÷ø è æ ö EV (mL) çè (Sample (mL)) (LA (cm 2 )) ÷ø

(10000 cm

2

/m 2

)(0.001mmol/nmol )

Where EV is the extraction volume, Sample is the volume of extract, and LA is leaf area. Units are mmol of Rubisco catalytic sites per leaf area. Typically, the “Bound CABP” is the sum of radioactivity in collected fractions 2 and 3, after correcting for the background by subtracting the radioactivity given by fraction 1 (see Table 2). 3.6.2. Activity of Rubisco: 14 CO2 Fixation

1. NaH14CO3 specific activity: SA (DPM/nmol) =



(Ave DPM ) éë NaH CO3 ùû (mM)* 0.1 14

Divide the average DPM from the three specific activity vials (see Subheading 3.3.1) by the concentration of 14CO2 added to each vial. 2. Rubisco activity (Vcmax): Vcmax (mmol/m 2 s1 ) =

(sample (DPM) - blank (DPM))(EV )(10000 (cm 2 /m 2 ))(D) (DF) (SA (DPM/nmol))(Time (s ))(LA (cm 2 ))(Sample-vol (mL)) Time is the length of the assay (typically 30 s). The correction for Rubisco discrimination (D) against 14C is 1.05. The dilution factor (DF) accounts for the change to the bicarbonate concentration of the assay buffer caused by the sample. 3. Dilution factor (DF): DF =

[HT](RB) + [ACT](RS) [HT](RB)

HT is the concentration of 14CO2 in the assay buffer, RB is the volume of buffer in the assay, RS is the sample volume, and ACT is the concentration of bicarbonate in the activated extract.

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359

4. Catalytic turnover, kcat: The maximum activity of Rubisco in leaves depends on the concentration of catalytic sites, and the turnover number (kcat, the first order rate constant for carboxylation, (5). kcat = Vcmax (mmol/m2 s)/Rubisco (mmol/m2). 3.6.3. Activity of Rubisco: Spectrophotometric Assay

Rubisco activity (Vcmax): Determine the rate of RuBP consumption (mmol/s) from a linear region of the spectrophotometer trace, assuming an extinction coefficient of 6.18  mmol/mL/cm for NADH oxidation at 340 nm. Rubisco carboxylation of RuBP produces two molecules of PGA; the coupled-enzyme assay consumes four NADH per RuBP consumed (Fig. 1). The rate of RuBP consumption is: RuBP - use (mmol/s) =



(At0 - At1 ) Vol 6.18 ´ 4 ´ (t 0 - t1 )

Where A is absorbance (340 nm) at time 0 or time 1, Vol is assay volume (mL), and t is time (s). The time interval should be within the first 30–120  s after the addition of activated extract. The activity of Rubisco is then: Vcmax (mmol/m 2s) =

(RuBP - use (mmol/s))(EV (mL))(10000 (LA (cm 2))(Sample (mL))

cm 2 /m 2

)

4. Notes These methods are useful with a range of agricultural and wild plant species and can be easily adapted to prokaryotes, algae, and non-vascular plants. For prokaryotic organisms, the presence of alternative CO2 fixing reactions necessitates the use of activity blanks containing Rubisco but not RuBP. The choice of methods depends on the purpose of the experiment. Detailed studies of Rubisco biochemistry inevitably utilise 14CO2, and the amount of Rubisco protein is most precisely determined with 14 C-CABP. It is probably insufficient to determine kcat based on immuno-detection of Rubisco, although we have found good agreement between immunoblotting and 14C-CABP-binding assays in moss and algal extracts (Brown and Kubien, unpublished data). The spectrophotometric assay is suitable for all activity measurements (15), although 14CO2 uptake gave higher activities than spectrophotometric approaches in field-grown plants (16).

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An advantage to the spectrophotometric and immuno-detection approaches is that they produce no radioactive waste. 1. CABP is an analogue of the 6-carbon intermediate of the Rubisco carboxylation reaction and binds almost irreversibly to carbamylated Rubisco catalytic sites. 14C-CABP is a 40–50% mixture of 2-carboxy-d-arabinitol-1, 5-bisphosphate (CABP), and 2-carboxy-d-ribitol-1, 5-bisphosphate (CRBP); the correct name of this mixture is carboxypentitol bisphosphate (CPBP). We use the term CABP loosely, to mean either the mixture (CPBP) or chromatographically purified CABP. For most purposes, CPBP is suitable for use in these assays. For greatest accuracy, the total amount of bound counts should constitute only 10–15% of the total counts recovered (see Table  2). Reduce the amount of leaf extract and repeat the assay if the amount of counts in the high molecular weight fractions exceeds this 10–15% rule.   Note that the chromatographic separation is now widely used in place of immuno-precipitating and filtration-collecting the 14C-CABP-Rubisco complexes. The chromatographic approach is faster, produces much smaller volumes of radioactive waste, and eliminates animal care issues associated with antibody production. 2. This buffer is a starting point and requires optimization for each species. For some species, a 100  mM Bicine–NaOH (pH 8.0) buffer may be preferable to the HEPES stock. Low Rubisco activity or yield can result from protease activity, the inhibitory effects of secondary metabolites, or inadequate membrane solubilization. Increasing the amount of antiprotease compounds may be required to increase yield. The ACA-BAM may be replaced with approximately 4% (v/v) of protease inhibitor cocktail (e.g. Sigma P9599), or used at lower concentration in combination with 2% (v/v) protease cocktail; in either case, add the protease cocktail to the tissue grinder(s) directly. It may also be advisable to replace the insoluble PVPP with soluble PVP, to ensure that Rubisco does not bind with PVPP and “precipitate” out of solution. For some species, Triton may disrupt membranes more effectively than Tween. The extraction volume/leaf area ratio also requires optimization, but 2 mL/cm2 is a good starting point. If determining site concentration by quantitative immunodetection, remove BSA from the extraction buffer. Make the EB on the day of use, and keep on ice. 3. It is crucial to use only high-purity RuBP for activity assays, as standard grade reagents typically contain inhibitory epimers, such as xylulose bisphosphate (XuBP), that bind to the active site of Rubisco. For the 14CO2 assays, a classic way to

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ensure high purity substrate is to synthesise RuBP in the reaction vessel by including Ru-5-P kinase, ribose-5-phosphate, phospho-ribose isomerase, and ATP in the assay buffer (4, 13). However, the procedure for purifying large quantities of Ru-5-P kinase from leaves is labour intensive, and Rubisco must be excluded from the preparation. If conducting large numbers of assays, it might be economical to produce RuBP enzymatically and purify it by liquid chromatography (17). A 25 mM RuBP stock solution (at acidic pH) is convenient for regular use. Assay buffers must be freshly prepared in small aliquots (e.g. only enough for the measurements and blanks). Keep assay buffer sealed and on ice as much as possible. 4. Immunoblotting (Western blotting) can detect the presence, absence, or the abundance of target proteins among samples. The relationship between the target protein abundance and the intensity of the immunoblot signal that it generates is often non-linear and does not have a zero intercept. However, using a quantified protein standard to produce a standard curve allows absolute quantification of the target. Campbell et al. (8) described a bioinformatic strategy to design antibodies directed toward a small region of the target protein that is highly conserved in a wide range of taxa. These global antibodies are suitable for quantitative immunoblotting even on protein samples from distantly related organisms. Rubisco has been immuno-quantified in phytoplankton (18, 19) and a variety of higher plants, including Spartina (9).   We typically load 1 mg total protein for the measurements of RbcL. As Rubisco is abundant in higher plants and the rabbit anti-RbcL antiserum from Agrisera has a high affinity, low protein loads provide the best results. For quantitative immunoblotting, high loads (>5 mg) can cause samples to behave differently from standards. At least three different standard amounts, ranging from, for example, 0.05 pmol to 1.0 pmol RbcL, should be loaded to generate a standard curve.   For quantitative immunoblotting, dilute antibodies give more consistent results than higher concentrations. We use primary and secondary antibodies at a dilution of 1:20,000– 1:50,000 in 2% ECL Advance blocking buffer (GE Healthcare) made up in TBS-T. The efficacy of antibody binding and washing steps is sensitive to temperature.

Acknowledgments We thank Amanda Cavanagh and Jonathan Neilson (UNB) for helpful comments on the manuscript.

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References 1. Ellis, R.J. (1979) The most abundant protein in the world. Trends Biochem. Sci. 4, 241–244. 2. Björkman, O. (1968) Carboxydismutase activity in shade and sun adapted species of higher plants. Physiol. Plant. 21, 1–10. 3. Laing, W.A., Ogren, W., and Hageman, R. (1974) Regulation of soybean net photosynthetic CO2 fixation by the interaction of CO2, O2, and ribulose-1, 5 disphosphate carboxylase. Plant Physiol. 54, 678–685. 4. Kubien, D.S., von Caemmerer, S., Furbank, R.T., and Sage, R.F. (2003) C4 photosynthesis at low temperature: a study using transgenic plants with reduced amounts of Rubisco. Plant Physiol. 132, 1577–1585. 5. von Caemmerer, S. (2000) Biochemical Models of Leaf Photosynthesis. CSIRO Publishing. 6. Collatz, G.J., Badger, M., Smith, C., and Berry, J.A. (1979) A radioimmune assay for RuP2 carboxylase protein. Carneige Inst. Wash. Year B. 78, 171–175. 7. Ruuska S., Andrews T.J., Badger M.R., Hudson G.S., Laisk A., Price G.D., and von Caemmerer S. (1998) The interplay between limiting processes in C3 photosynthesis studied by rapid-response gas exchange using transgenic tobacco impaired in photosynthesis. Aust. J. Plant Physiol. 25, 859–870. 8. Campbell, D.A., Cockshutt, A.M., and Porankiewicz-Asplund, J. (2003) Analysing photosynthetic complexes in uncharacterized species or mixed microalgal communities using global antibodies. Physiol. Plant. 119, 322–327. 9. Morash, A.J., Campbell, D.A., and Ireland, R.J. (2007) Macromolecular dynamics of the photosynthetic system over a seasonal developmental progression in Spartina alterniflora. Can. J. Bot. 85, 476–483. 10. Rabin, N.G. and Trown, P.W. (1964) Mechanism of action of carboxydismutase. Nature 202, 1290–1293. 11. Paulsen, J.M. and Lane, M.D. (1966) Spinach ribulose diphosphate carboxylase I. Purification and properties of the enzyme. Biochem. 5, 2350–2357.

12. Badger, M.R. and Andrews, T.J. (1974) Effects of CO2, O2 and temperature on a high-affinity form of ribulose disphosphate carboxylase-oxygenase from spinach. Biochem. Biophys. Res. Comm. 60, 204–210. 13. Seemann, J.R., Badger, M.R., and Berry, J.A. (1984) Variations in the specific activity of ribulose-1, 5-bisphosphate carboxylase between species utilising different photosynthetic pathways. Plant Physiol. 74, 791–794. 14. Lilley, R. McC. and Walker, D.A. (1974) An improved spectrophotometric assay for ribulosebisphosphate carboxylase. Biochim. Biophys. Acta. 358, 226–229. 15. Sharkey, T.D., Savitch, L.V., and Butz, N.D. (1991) Photometric method for routine determination of kcat and carbamylation of Rubisco. Photosyn. Res. 28, 41–48. 16. Keys, A.J. and Parry, M.A.J. (1990) Ribulose bisphosphate carboxylase/oxygenase and carbonic anhydrase. In: Methods in Plant Biochemistry. Vol. 3 (Dey, P.M. and Harborne, J.B., eds.), pp. 1–14. 17. Butz, N.D. and Sharkey, T.D. (1989) Activity ratios of ribulose-1, 5-bisphosphate carboxylase accurately reflect carbamylation ratios. Plant Physiol. 89, 735–739. 18. Reid, C.D., Tissue D.T., Fiscus, E.L., and Strain, B.R. (1997) Comparison of spectrophotometric and radioisotopic methods for the assay of Rubisco in ozone-treated plants. Phys. Plant. 101, 398–404. 19. Kane, H.K., Wilkin, J.M., Portis, A.R., and Andrews, T.J. (1998) Potent inhibition of ribulose-bisphosphate carboxylase by an oxidized impurity in ribulose-1, 5-bisphosphate. Plant Physiol. 117, 1059–1069. 20. Brown, C.M., MacKinnon, J.D., Cockshutt, A.M., Villareal, T., and Campbell, D.A. (2008) Flux capacities and acclimation costs in Trichodesmium from the Gulf of Mexico. Mar. Biol. 154, 413–422. 21. Brown, C.M., Campbell, D.A., and Lawrence, J.E. (2007) Resource dynamics during infection of Micromonas pusilla by virus MpV-Sp1. Environ. Microbiol. 9, 2720–2727.

Chapter 28 Purification of Rubisco Activase from Leaves or after Expression in Escherichia coli Csengele Barta, A. Elizabete Carmo-Silva, and Michael E. Salvucci Abstract Rubisco activase is a molecular chaperone that modulates the activation state of Rubisco by catalyzing the ATP-dependent removal of tightly-bound inhibitory sugar-phosphates from Rubisco’s catalytic sites. This chapter reports methods developed for the purification of native and recombinant Rubisco activase from leaves and bacterial cells, respectively. Key words: AAA+ protein, ATP, Arabidopsis thaliana, CO2 fixation, Calvin Cycle, Molecular chaperone, Rubisco activase

1. Introduction Ribulose-1,5-bisphosphate carboxylase/oxygenase (Rubisco) activase is a soluble AAA+ chloroplast protein that interacts with Rubisco to restore the activity to catalytic sites (reviewed in (1–3)). Like other AAA+ proteins, Rubisco activase is thought to be a mechano-chemical motor protein that uses the energy from ATP hydrolysis to contort the structure of its target protein, Rubisco (4). The alteration in Rubisco structure caused by Rubisco activase promotes the dissociation of tightly-bound sugar-phosphates that form dead-end complexes in the active site (5). This action is required for normal photosynthesis and growth of higher plants (6, 7), as well as green (8) and possibly other groups of algae. Rubisco activase activity is dependent on both the ATP/ADP ratio and the redox status of the chloroplast and is extremely sensitive to high temperature. Consequently, this protein serves two very important regulatory roles in photosynthesis: 1) adjusting

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the rate of CO2 fixation to the rate of electron transport activity (9) and 2) limiting CO2 assimilation during heat stress (10). Rubisco activase was first isolated by fractionating chloroplast stromal extracts by rate zonal centrifugation on sucrose gradients followed by anion-exchange and gel filtration chromatography (11). Once the ATP requirement and the ability of ATP to protect against proteolysis and instability were recognized (12, 13), the need for using isolated chloroplasts as a source of clean stromal and thylakoid fractions was eliminated, and the advantage of faster isolation and higher yield from the extraction of leaves was quickly realized (14). The findings that Rubisco activase catalyzes ATP hydrolysis and precipitates from solution with only 35–40% ammonium sulfate facilitated development of methods for isolation of Rubisco activase (15, 16). This chapter provides details of the procedures that have been developed and optimized for the purification of native Rubisco activase from leaves and the recombinant protein from Escherichia coli cells. The purification protocol involves the precipitation of Rubisco activase with ammonium sulfate and purification by gelfiltration and ion-exchange chromatography. The procedure is designed to be completed in a single day, producing active Rubisco activase from leaves or cell pellets with a minimum of proteolytic degradation.

2. Materials 2.1. Isolation of Rubisco Activase from Arabidopsis thaliana Leaves (see Note 1)

1. Frozen leaf tissue, 100 g FW, stored at −80°C. 2. Leaf Extraction Buffer: 50 mM HEPES-KOH, pH 7.0, 5 mM MgCl2, 1 mM EDTA, 1 mM ATP, 50 mM 2-mercaptoethanol, 20 mM ascorbate, 1 mM phenylmethylsulfonate (PMSF), 10  mM leupeptin, and 2% (w/v) polyvinylpolypyrrolidone (PVPP) (see Note 2). 3. Blender and varistat. 4. Cheesecloth and miracloth. 5. Refrigerated high-speed centrifuge with fixed angle and swinging bucket rotors (Thermo Fisher Scientific Inc., Waltham, MA, or equivalent). 6. Plastic centrifuge tubes, 50 and 15 mL capacity. 7. Ammonium sulfate, saturated (~4.1 M) solution, adjusted to pH 7.0 with NH4OH. 8. Separatory funnel. 9. Leaf Resuspension Buffer: 50  mM HEPES-KOH, pH 7.0, 2  mM MgCl2, 100  mM KCl, 5  mM dithiothreitol (DTT),

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1 mM ATP, 1 mM PMSF, and 10 mM leupeptin. Add DTT, ATP, PMSF, and leupeptin fresh. 10. Leaf Gel Filtration Buffer: 20  mM HEPES-KOH, pH 7.2, 2 mM MgCl2, 50 mM KCl, 2 mM DTT, and 0.2 mM ATP. Add DTT and ATP fresh. 11. Anion-Exchange Buffer A: 20  mM HEPES-KOH, pH 7.2, 5 mM MgCl2, and 2 mM DTT. Add DTT fresh. 12. Anion-Exchange Buffer B: Anion-Exchange Buffer A containing 1 M KCl. 13. Preparative gel filtration column, Toyopearl HW-55S, 2.6 × 66 cm. 14. Anion-exchange columns, Q-Sepharose HiTrap, 3 × 5  mL (GE-Healthcare/Amersham Biosciences, Piscataway, NJ, or equivalent). 15. Low, medium, or high pressure chromatographic system with UV-detector and fraction collector (ÄKTA, GE-Healthcare/ Amersham Biosciences or equivalent). 16. Chart recorder or computer with A/D converter. 17. Glass tubes, 20 mL capacity. 18. UV-Vis spectrophotometer. 19. Ultrafiltration stirred cell, 60  mL capacity, with 43  mm YM-30 membrane (Millipore Corp., Billerica, MA). 20. Compressed N2 gas tank. 21. ATP, 100  mM, adjusted to pH 7.0 with NaOH. Store at −80°C. 22. Screw cap tubes, 1.5 mL capacity. 23. Bradford protein assay reagent (Bio-Rad Laboratories, Hercules, CA). 24. Bovine serum albumin (BSA), 10 mg/mL stock. 2.2. Large-Scale Expression and Purification of Recombinant Rubisco Activase 2.2.1. Cell Growth and Collection

1. BL21(DE3)pLysS E. coli strain transformed with Rubisco activase cDNA cloned in pET-23a or d(+) vector (EMD Chemicals/Novagen, Gibbstown, NJ). 2. LB-Miller Broth liquid medium (EMD Chemicals/Novagen) containing 10 g peptone from casein, 5 g yeast extract and 10  g NaCl for 1  L media. Autoclave at 121°C for 20  min. Store at 4°C. 3. M9ZB medium (for 1  L): 1  g NH4Cl, 3  g KH2PO4, 6  g Na2HPO4, 10 g Bacto-tryptone, and 5 g NaCl. 4. Carbenicillin, 50 mg/mL, filtered through a 0.25 mm sterile syringe filter. Store at −20°C. 5. Chloramphenicol, 50 mg/mL, in absolute ethanol. Store at −20°C.

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6. Glucose, 40% (w/v), filtered through a 0.25 mm sterile syringe filter. Store at 4°C. 7. MgSO4, 1 M, filtered through a 0.25 mm sterile syringe filter or autoclaved. Store at 4°C. 8. Isopropyl-b-d-thio-galactopyranoside (IPTG), 1  M, filtered through a 0.25 mm sterile syringe filter. Store at −20°C. 9. Orbital-plane shaker platform (Innova 2100, New Brunswick Scientific, Edison, NJ, or equivalent). 10. Falcon tubes, 50 mL capacity. 11. Fermentor/bioreactor, 15  L capacity (Bioflo 110, New Brunswick Scientific or equivalent) (see Note 3). 12. Air pump (small aquarium pump or equivalent). 13. UV-Vis spectrophotometer. 14. Peristaltic pump (MasterFlexII, Cole-Parmer Instrument Company, Vernon Hills, IL, or equivalent). 15. Centrifuge bottles (1 L) and tubes (50 mL). 16. Refrigerated high-speed centrifuge with fixed angle rotor for 1 L bottles and swinging bucket rotor for 50 mL Falcon tubes. 17. Protein Wash Buffer: 25  mM HEPES-KOH, pH 7.2, and 8 mM MgCl2. 2.2.2. Purification of Recombinant Rubisco Activase from E. coli

1. E. coli cell pellets from a 3-L culture. 2. Cell Extraction Buffer: 50 mM HEPES-KOH, pH 7.0, 5 mM MgCl2, 1 mM EDTA, 0.1% (w/v) Triton X-100, 2 mM ATP, 5  mM DTT, 20  mM ascorbate, 1  mM PMSF, and 10  mM leupeptin (see Note 2). 3. Sonifier with standard tip (Branson S450, Branson Ultrasonics Corp., Danbury, CT, or equivalent). 4. Water-jacketed glass beaker and refrigerated circulating water bath. 5. Refrigerated high-speed centrifuge with fixed angle and swinging bucket rotors. 6. Plastic centrifuge tubes, 50 and 15 mL capacity. 7. Ammonium sulfate, saturated (~4.1 M) solution, adjusted to pH 7.0 with NH4OH. 8. Separatory funnel. 9. Refrigerated ultracentrifuge with a small volume swinging bucket rotor (SW 55 Ti rotor, Beckman-Coulter, Palo Alto, CA, or equivalent). 10. Gel Filtration (GF) Buffer: 50 mM HEPES-KOH, pH 7.2, 10  mM MgCl2, plus 10  mM 2-mercaptoethanol. Add 2mercaptoethanol immediately prior to use. 2-Mercaptoethanol is toxic, handle with gloves, under a fume hood.

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11. Anion-Exchange Buffer A: half strength GF buffer containing 2 mM DTT. Add DTT fresh. 12. Anion-Exchange Buffer B: Anion-Exchange Buffer A containing 1 M KCl. 13. See Subheading 2.1, items 13–24.

3. Methods This section describes procedures for the isolation of native Rubisco activase from leaves and for the expression and purification of recombinant Rubisco activase from E. coli (see Fig. 1 for a flow diagram). Unless stated otherwise, all steps are performed at 4°C and as quickly as possible to minimize proteolysis and subsequent loss of activity. The entire purification procedure can be completed in 1 day starting from leaves or bacterial cell pellets. leaf tissue pellet

E. coli cell pellet

extraction & filtration centrifugation

pellet

soluble extract supernatant

ammonium sulfate (40%) centrifugation

soluble extract

precipitate

supernatant

supernatant

suspend in 35% ammonium sulfate centrifugation

ammonium sulfate (37.5%) centrifugation

precipitate dissolve in buffer pellet

precipitate pellet

sonication centrifugation

dissolve in buffer centrifugation

ultracentrifugation

supernatant

supernatant gel-filtration chromatography

anion-exchange chromatography add ATP flash freeze

purified Rubisco activase Fig. 1. Flow-chart outlining the procedure for the purification of Rubisco activase from leaves and Escherichia coli cells.

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3.1. Rubisco Activase Isolation from A. thaliana Leaves (see Note 1)

1. Extract 100  g FW of frozen leaf tissue in 400  mL of Leaf Extraction Buffer in a blender for 2–3 min using 30 s blender bursts with 30 s intervals between bursts. 2. Filter the homogenate through four layers of cheesecloth and one layer of miracloth suspended over a plastic funnel to remove cell walls and other unbroken material. To recover as much extract as possible, gently squeeze the suspension by hand (use gloves). The cheese- and miracloth are washed extensively with Milli-Q water and stored hydrated at 4°C before use. 3. Centrifuge the filtrate for 20  min at 30,000 × g to remove starch, membranes, and insoluble cell debris. Discard the pellet. Measure the volume of the supernatant. 4. Add saturated ammonium sulfate to the supernatant to reach a saturation level of 40%. Use the following equation: Volume of saturated ammonium sulfate for 40% saturation = (volume of the supernatant/0.6) – volume of the supernatant. Add saturated ammonium sulfate drop-wise to the supernatant with continuous stirring using a separatory funnel suspended in a ring stand. Stir for 30 min. 5. Collect the precipitate by centrifugation at 23,000 × g for 13 min. 6. Suspend the protein pellet in 20  mL of Leaf Resuspension Buffer containing 35% ammonium sulfate. Stir for 30 min. 7. Collect precipitated protein by centrifugation at 23,000 × g for 15 min. 8. Suspend the protein pellet in 10  mL of Leaf Resuspension Buffer (see Note 4) and centrifuge at 30,000 × g for 15 min. 9. Load the supernatant on a Toyopearl HW-55S gel filtration column previously equilibrated with at least three column volumes of Leaf Gel Filtration Buffer (see Note 5) and fractionate at 1 mL/min. Collect 10 mL fractions while monitoring the absorbance at 280 nm. 10. Identify fractions with the highest Rubisco activase activity by determining ATPase activity (for details see Chap. 29 “Rubisco Activase Activity Assays” by Barta et al. in the current volume) (see Note 6). Pool fractions with the highest activity. 11. Load pooled fractions on a 15  mL (3 × 5  mL) Q-Sepharose HiTrap anion-exchange column, equilibrated with four column volumes of Anion-Exchange Buffer A (see Note 7). Separate proteins by elution with a 0–0.5 M linear KCl gradient over 300 mL at a flow rate of 3 mL/min. The gradient is formed by mixing Anion-Exchange Buffers A and B. Collect 10 mL fractions while monitoring the absorbance at 280 nm.

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12. Measure ATPase activity and pool fractions with the highest activity. 13. If necessary, concentrate pooled fractions under N2 in a 60  mL ultrafiltration stirred cell fitted with a YM-30 membrane. 14. Supplement the concentrated protein with 0.2 mM ATP, and divide into aliquots in screw cap microfuge tubes. Flash freeze in liquid N2 and store at −80°C (see Note 8). 15. For use, slowly thaw an aliquot of the frozen protein on ice. 16. If necessary, desalt the protein solution by centrifugal gel filtration (for details see accompanying Chap. 29 on “Rubisco Activase Activity Assays” by Barta et al.). 17. Determine protein concentration from the absorbance at 595 nm by a dye-binding assay (17) using BSA as standard. 18. Determine the purity of the protein by SDS-PAGE analysis (18) and assay activity (for details see accompanying Chap. 29 on “Rubisco Activase Activity Assays” by Barta et al.). 3.2. Large-Scale Expression and Purification of Recombinant Rubisco Activase

Using T7-based expression plasmids like pET-23, soluble recombinant Rubisco activase can be produced in E. coli from a cDNA encoding the mature form of Rubisco activase (see Note 9). The purity and stability of recombinant Rubisco activase isolated from bacterial cultures is usually greater than the protein from leaves due to the lower proteolytic activity associated with bacterial compared with leaf extracts.

3.2.1. Cell Growth and Collection

For the production and purification of recombinant Rubisco activase, the BL21(DE3)pLysS competent cell strain is transformed with the pET-23a or d(+) plasmid harboring the Rubisco activase cDNA. The construct is engineered to produce the mature form of Rubisco activase; consequently, the nucleotides that encode for the N-terminal chloroplast transit peptide are replaced by the ATG start codon (19). The BL21(DE3)pLysS strain is deficient in both lon and ompT proteases and was optimized for stabilizing the pET recombinants. However, similar expression vectors and equivalent host cell lines may be used. Cell growth and harvest requires 3 days. Day 1 1. Add 6 × 5  mL of sterile LB-Miller Broth medium supplemented with 50 mg/mL carbenicillin and 50 mg/mL chloramphenicol to six 50  mL sterile Falcon tubes. Inoculate the cultures with a small volume of cells using sterile toothpicks (see Note 10). Grow cells for 16 h at 37°C under continuous shaking at 300 rpm in a temperature-controlled orbital-plane shaker.

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2. Autoclave 6 L of M9ZB culture medium in a 15 L glass fermentor for 30 min at 121°C. Let the solution cool overnight. Day 2 3. Supplement the M9ZB medium with 50 mg/mL carbenicillin, 50  mg/mL chloramphenicol, 0.4% (w/v) glucose, and 1 mM MgSO4. 4. Centrifuge starter cultures in 50 mL Falcon tubes at 3,300 × g for 15 min at 4°C. Suspend cell pellets in 5 mL of the supplemented M9ZB medium. 5. Inoculate a 6-L culture in the fermentor with the starter cultures. Grow cells at 37°C under continuous stirring at 225 rpm while bubbling with air through a sterile 0.25 mm filter. 6. When the cells reach an optical density of 0.6–0.8 at 600 nm, immediately cool the culture to 25°C. Remove a 10 mL aliquot for the analysis of “uninduced” expression. Induce Rubisco activase expression by adding 0.5 mM IPTG to the cell culture. 7. Maintain the culture at 25°C for 12 h under continuous air bubbling and stirring. Day 3 8. Transfer the culture to four 1 L centrifuge bottles on ice using a peristaltic pump. Remove a 10 mL aliquot for the analysis of “induced” expression. 9. Collect the cells by centrifuging at 5,000 × g for 10  min at 4°C. Repeat using the same 1 L bottles until the entire culture content is collected in the four bottles. 10. Suspend the accumulated cell pellets in about 100  mL of Protein Wash Buffer. Pool the cell suspension and distribute into two 50 mL Falcon tubes. 11. Centrifuge at 5,000 × g for 10 min at 4°C. The wet cell pellets are stored at −80°C until extraction. 12. Analyze soluble polypeptides in the uninduced and induced samples by SDS-PAGE (18) (see Note 11). 3.2.2. Purification of Recombinant Activase from E. coli

Recombinant activase is purified from washed cell pellets equivalent to a 3-L culture, i.e., half of the cells collected from the 6-L culture (see Subheading  3.2.1). Unless indicated otherwise, all steps are performed at 4°C. 1. Thaw and lyse the frozen cells by suspending the pellet in 130 mL of Cell Extraction Buffer for 5 min on ice (see Note 12). Transfer the lysate to a temperature-controlled water-jacketed glass beaker at 4°C for sonication.

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2. Rupture the cells and DNA by sonicating the suspension. Use five cycles of sonication, 1  min each, at 50% of maximum power to avoid excess foaming. A 30 s pause between each cycle prevents warming of the suspended cells. 3. Centrifuge the solution for 20 min at 23,000 × g to remove unbroken cells and insoluble cell debris. 4. Measure the volume of the supernatant and add saturated ammonium sulfate to 37.5%. Use the following equation: Volume of saturated ammonium sulfate for 37.5% saturation = (volume of the supernatant/0.625) – volume of the supernatant. Add the saturated ammonium sulfate drop-wise to the supernatant while stirring continuously using a separatory funnel in a ring stand. Stir for 30 min. 5. Collect precipitated material by centrifugation at 20,000 × g for 13 min. 6. Suspend pellets in a total volume of 10 mL of Cell Extraction Buffer. 7. Centrifuge the solution at 237,000 × g (rav) for 30 min in a swinging bucket ultracentrifuge rotor to remove membrane vesicles (see Notes 6 and 13). Pool the supernatants. 8. Load the supernatant on the Toyopearl HW-55S gel filtration column and follow steps 9–18 described in Subheading 3.1 to isolate and analyze Rubisco activase, but using the Gel Filtration and Anion-Exchange Buffers described in Subheading 2.2.2.

4. Notes 1. The Rubisco activase holoenzyme from Arabidopsis and many other plant species contains two isoforms, a ~47 kDa a- and a ~42 kDa b-isoform. In contrast, Rubisco activase from some plant and algal species, including tobacco, maize, and Chlamydomonas, is composed of only the b − isoform (11, 16). We use an identical purification procedure to purify Rubisco activase from spinach, Arabidopsis and tobacco. 2. All solutions used for chromatographic separation are filtered through a 0.22 mm Millipore GSWP filter upon preparation. To avoid acidic hydrolysis of ATP, ascorbate is added to the extraction and resuspension buffers prior to ATP. Add the ascorbate to the basic buffer solution (containing HEPES, MgCl2 and EDTA), then readjust the pH to 7.0 with 6 N KOH

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before adding the remaining components, with the exception of PVPP. After adding ATP, adjust the pH to 7.0 with 1 N KOH. Add solid PVPP immediately prior to extraction. Storage of the complete Leaf Extraction Buffer is not recommended. The addition of 1% (w/v) casein will reduce proteolysis in leaf extracts; however, a more rigorous purification protocol may be required to remove casein from the preparation. 3. As an alternative to using a fermentor, 1  L cultures can be prepared in 3 L fernbach flasks. The cultures are induced and harvested identically to the large-scale cultures. 4. The inclusion of ATP in the extraction and resuspension buffers causes Rubisco activase to associate and migrate as a higher-ordered oligomeric protein on the gel filtration column (20). 5. The volume of sample is limited by the size of the preparative gel filtration column. We try to load a sample volume equivalent to less than 4% of the bed volume. 6. Membrane vesicles also precipitate between 0 and 40% ammonium sulfate and membrane-bound enzymes in these vesicles catalyze considerable NADH oxidation and/or show ATPase activity. This activity generally elutes slightly ahead of the Rubisco activase in the turbid fractions associated with the void volume of the gel filtration column. To minimize contamination from this source, we recommend centrifuging the suspended material prior to chromatography and verifying the presence of the Rubisco activase polypeptides by SDSPAGE in the gel filtration fractions. 7. The size of the column used for anion-exchange chromatography depends on the amount of protein in the sample. For example, when purifying the recombinant Rubisco activase from a 3 L culture of the highest expressing clones (yielding about 150 mg protein), a column volume of 15 mL is used. Rubisco activase elutes as a single peak between 0.19 and 0.26 M KCl (11). 8. Inclusion of ATP and ADP during storage improves Rubisco activase stability (13). We generally add 0.2 mM ATP to the pooled anion-exchange fractions, then flash freeze the protein in aliquots and store at −80°C. Rubisco activase stored in this way maintains activity for at least 5 years. 9. We use Rubisco activase constructs that are identical to the native, mature protein, except for the presence of a N-terminal Met. Since the protein does not contain a His-tag or other similar modification, the purification requires classical rather than affinity-based techniques.

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10. Transformed E. coli cells stored as frozen glycerol stocks are used as inoculums for starter cultures. 11. To analyze the expression of Rubisco activase: (a) Collect cells from 10 mL aliquots of induced and uninduced cultures by centrifuging for 10  min at 5,000 × g and 4°C. Rinse pellets with Protein Wash Buffer (see Subheading 2.2.1). (b) Suspend cells in 2 mL of 50 mM HEPES-KOH, pH 7.2, 5  mM MgCl2, 5  mM DTT, 0.2  mM ATP, and 1  mM PMSF. Vortex thoroughly. (c) Lyse cells by adding 200  mL of Pop-Culture Reagent (Novagen/EMD Chemicals, Gibbstown, NJ, USA) and 2 mL of Benzonase Nuclease (Novagen/EMD Chemicals). Vortex and incubate on ice for 10 min. (d) Save a 20  mL aliquot from the suspension as “total protein.” (e) Centrifuge the extract for 5 min at 12,000 × g and 4°C. (f) Save a 20  mL aliquot from the supernatant as “soluble protein.” (g) Analyze protein expression in the total and soluble fractions from the uninduced and induced samples by SDSPAGE (19). 12. The BL21(DE3)pLysS cell strain harbors the pLysS plasmid that expresses the T7 lysozyme, known to facilitate cell lysis by freeze-thawing. 13. As an alternative to ultracentrifugation, suspended material may be centrifuged at 20,000 × g for 13 min in a fixed angle rotor to remove cell debris and microsomes generated during sonication.

Acknowledgments The authors acknowledge Nancy Parks and Gwen Coyle (USDAARS, Maricopa, AZ) for their technical assistance. This research was funded by the US Department of Agriculture, Agricultural Research Service and the US Department of Energy, grant no. DE-FG02-08ER20268. Mention of a trademark, proprietary product, or vendor does not constitute a guarantee or warranty of the product by the United States Department of Agriculture and does not imply its approval to the exclusion of other products or vendors that may also be suitable.

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References 1. Salvucci, M. E., Portis, A. R. Jr., and Ogren, W. L. (1985) A soluble chloroplast protein catalyses ribulose-bisphosphate carboxylase/ oxygenase activation in vivo. Photosynth Res 7, 193–201. 2. Portis, A. R. Jr. (2003) Rubisco activase – Rubisco’s catalytic chaperone. Photosynth Res 75, 11–27. 3. Andrews, T. J. (1996) The bait in the Rubisco mousetrap. Nat Struct Biol 3, 3–7. 4. Portis, A. R. Jr., Li, C. S., Wang, D. F., and Salvucci, M. E. (2008) Regulation of Rubisco activase and its interaction with Rubisco. J Exp Bot 59, 1597–604. 5. Wang, Z. Y. and Portis, A. R. Jr. (1992) Dissociation of ribulose-1,5-bisphosphate bound to ribulose-1,5-bisphosphate carboxylase oxygenase and its enhancement by ribulose1,5-bisphosphate carboxylase oxygenase activase mediated hydrolysis of ATP. Plant Physiol 99, 1348–53. 6. Salvucci, M. E., Portis, A. R. Jr., and Ogren, W. L. (1986) Light and CO2 response of ribulose-1,5-bisphosphate carboxylase/oxygenase activation in Arabidopsis leaves. Plant Physiol 80, 655–59. 7. Hammond, E. T., Andrews, T. J., and Woodrow, I. E. (1998) Regulation of ribulose-1,5-bisphosphate carboxylase/oxygenase by carbamylation and 2-carboxyarabinitol 1-phosphate in tobacco: Insights from studies of antisense plants containing reduced amounts of Rubisco activase. Plant Physiol 118, 1463–71. 8. Pollock, S. V., Colombo, S. L., Prout, D. L., Godfrey, A. C., and Moroney, J. V. (2003) Rubisco activase is required for optimal photosynthesis in the green alga Chlamydomonas reinhardtii in a low CO2 atmosphere. Plant Physiol 133, 1854–61. 9. Salvucci, M. E., Portis, A. R. Jr., Heber, U., and Ogren, W. L. (1987) Stimulation of thylakoid energization and ribulose-bisphosphate carboxylase/oxygenase activation in Arabidopsis leaves by methyl viologen. FEBS Lett 221, 215–20. 10. Salvucci, M. E. and Crafts-Brandner, S. J. (2004) Relationship between the heat tolerance of photosynthesis and the thermal stability of Rubisco activase in plants from contrasting

11.

12.

13.

14.

15.

16.

17.

18. 19.

20.

thermal environments. Plant Physiol 134, 1460–70. Salvucci, M. E., Werneke, J. M., Ogren, W. L., and Portis, A. R. Jr. (1987) Purification and species distribution of Rubisco activase. Plant Physiol 84, 930–6. Streusand, V. J. and Portis, A. R. Jr. (1987) Rubisco activase mediates ATP-dependent activation of ribulose bisphosphate carboxylase. Plant Physiol 85, 152–4. Robinson, S. P. and Portis, A. R. Jr. (1989) Adenosine-triphosphate hydrolysis by purified Rubisco activase. Arch Biochem Biophys 268, 93–9. Robinson, S. P., Streusand, V. J., Chatfield, J. M., and Portis, A. R. Jr. (1988) Purification and assay of Rubisco activase from leaves. Plant Physiol 88, 1008–14. Shen, J. B., Orozco, E. M., and Ogren, W. L. (1991) Expression of the two isoforms of spinach ribulose 1,5-bisphosphate carboxylase activase and essentiality of the conserved lysine in the consensus nucleotide-binding domain. J Biol Chem 266, 8963–8. Salvucci, M. E., Rajagopalan, K., Sievert, G., Haley, B. E., and Watt, D. S. (1993) Photoaffinity labeling of ribulose-1.5-bisphosphate carboxylase/oxygenase activase with ATP g-benzophenone. Identification of the ATP g-phosphate binding domain. J Biol Chem 268, 14239–44. Bradford, M. M. (1976) A rapid and sensitive method for the quantitation of microgram quantities of proteins utilizing the principle of protein-dye binding. Anal Biochem 72, 248–59. Chua, N.-H. (1980) Electrophoretic analysis of chloroplast proteins. Methods Enzymol 69, 434–46. van de Loo, F. J. and Salvucci, M. E. (1996) Activation of ribulose-1,5-bisphosphate carboxylase/oxygenase (Rubisco) involves Rubisco activase Trp16. Biochemistry 35, 8143–48. Wang, Z. Y., Ramage, R. T., and Portis, A. R. Jr. (1993) Mg2+ and ATP or adenosine 5¢-[g-thio]triphosphate (ATP-g-S) enhances intrinsic fluorescence and induces aggregation which increases the activity of spinach Rubisco activase. Biochim Biophys Acta 1202, 47–55.

Chapter 29 Rubisco Activase Activity Assays Csengele Barta, A. Elizabete Carmo-Silva, and Michael E. Salvucci Abstract Ribulose-1,5-bisphosphate (RuBP) carboxylase/oxygenase (Rubisco) activase functions as a mechano-chemical motor protein using the energy from ATP hydrolysis to contort the structure of its target protein, Rubisco. This action modulates the activation state of Rubisco by removing tightly-bound inhibitory sugar-phosphates from Rubisco’s catalytic sites, thereby restoring the sites to catalytic competence. This chapter reports methods developed for assaying the two activities of Rubisco activase: ATP hydrolysis and Rubisco activation. Key words: AAA+ protein, ATP hydrolysis, Calvin Cycle, CO2 fixation, Rubisco activase, Rubisco, RuBP

1. Introduction Rubisco is prone to dead-end inhibition caused by the unproductive binding of the substrate, ribulose-1,5-bisphosphate (RuBP), substrate analogs like carboxyarabinitol-1-phosphate or catalytic misfire products to active sites (1, 2). The restoration of activity (i.e., Rubisco activation) requires a molecular chaperone, Rubisco activase, to promote conformational changes that facilitate the dissociation of inhibitory sugar-phosphates from Rubisco’s sites (1–4). Rubisco activase also catalyzes ATP hydrolysis, and this activity is essential for Rubisco activation (4–6). However, ATP hydrolysis is not strictly coupled to Rubisco activation, since the rate of ATP hydrolysis is unaffected by the presence of inhibited Rubisco (5). The first studies of Rubisco activase used illuminated chloroplast lysates supplemented with RuBP and an electron acceptor to measure its activity by the ability to restore catalytic competence to inhibited Rubisco (7, 8). After recognizing the requirement Robert Carpentier (ed.), Photosynthesis Research Protocols, Methods in Molecular Biology, vol. 684, DOI 10.1007/978-1-60761-925-3_29, © Springer Science+Business Media, LLC 2011

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for ATP hydrolysis (5), assays were developed for measuring Rubisco activase activity either by the rate of ATP hydrolysis, measured spectrophotometrically from ADP production or colorimetrically from Pi release, or by the rate and extent of Rubisco activation in defined systems that included ATP and inactive Rubisco complexed with RuBP (ER) (9). This chapter describes methods for assaying both activities of Rubisco activase: ATP hydrolysis and Rubisco activation.

2. Materials 2.1. ATP Hydrolysis

1. Purified Rubisco activase (see Note 1) and Rubisco activase (see Note 1). 2. ATPase Assay: 100  mM Tricine–KOH, pH 8.0, 10  mM MgCl2, 20  mM KCl, 5  mM dithiothreitol (DTT), 2  mM phosphoenolpyruvate (PEP), 5% (w/v) polyethylene glycol (PEG) 3350, 2 mM ATP, 0.3 mM NADH, 5 U/mL pyruvate kinase (PK), and 5.75  U/mL lactate dehydrogenase (LDH) (see Note 2). 3. UV-Vis spectrophotometer with thermally regulated cuvette holder and 1 mL optical glass or disposable polymethyl methacrylate cuvettes.

2.2. In Vitro Rubisco Activation

1. Purified Rubisco (see Note 3) and Rubisco activase (see Note 1). 2. Activation Mix: 100  mM Tricine–NaOH, pH 8.0, 10  mM MgCl2, 10 mM NaHCO3, and 50 mM DTT. 3. Rubisco ER Desalting Buffer: 50  mM Tricine–NaOH, pH 8.0, and 0.1 mM EDTA, prepared CO2-free by purging with N2 prior to adjusting the pH with 60% (w/v) NaOH. 4. RuBP, sodium salt, ~50 mM, and pH 7.0. Store at −80°C in small aliquots. 5. Plastic columns, 0.8 × 5  cm, containing 2  mL of Sephadex G-50-fine. 6. Corex glass tubes, 15 mL capacity. 7. Refrigerated high-speed centrifuge with swinging bucket rotor (Thermo Fisher Scientific Inc., Waltham, MA, or equivalent). 8. UV-Vis spectrophotometer and quartz cuvettes. 9. Rubisco Activase first Stage Assay: 50 mM Tricine–NaOH, pH 8.0, 10 mM MgCl2, 10 mM NaHCO3, 40 U/mL creatinephosphokinase, 4  mM creatine-phosphate, 2  mM DTT, 5  mM ATP, 5% (w/v) PEG-3350, and 4  mM RuBP (see Note 4).

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10. Rubisco 14C Assay: 100 mM Tricine–NaOH, pH 8.0, 10 mM MgCl2, 9.8 mM NaHCO3, 0.2 mM NaH14CO3 (50 Ci/mol), and 0.4 mM RuBP. 11. Reaction Quench Mix: 1 N HCl/4 N HCOOH. 12. Speed-Vac centrifugal vacuum concentrator (Thermo Fisher Scientific Inc., or equivalent) equipped with an acid vapor trap. 13. 0.3 N KOH. 14. 0.1 N HCl. 15. Scintillation cocktail (BioSafe II, Research International Corporation, Mount Prospect, IL).

Products

16. Liquid scintillation counter (TriCARB 2200CA, PerkinElmer Life and Analytical Science Inc., Waltham, MA, or equivalent).

3. Methods 3.1. Assay for ATP Hydrolysis

Rubisco activase catalyzes the hydrolysis of ATP and also promotes the ATP-dependent removal of inhibitory sugar-phosphates from Rubisco active sites. As shown in Fig. 1, the assay for ATPase activity is based on coupling ADP production to NADH oxidation using PK and LDH (see Note 5). 1. Verify the performance of the coupling reactions by adding 1 mmol of ADP to the ATPase assay in the absence of Rubisco activase and measuring the rate of NADH oxidation. 2. Initiate ATPase assays at the desired temperature (usually 30°C) by adding 10–20 mg of Rubisco activase (in 20 mL) to 480 mL of ATPase Assay Mix in a 1 mL optical glass or disposable cuvette. Monitor the ATP-dependent decrease in absorbance at 340 nm continuously for 10 min. 3. Calculate the rate of ATP hydrolysis from the DA340/min using a molar extinction coefficient for NADH of 6.22 mM−1 cm−1. One unit (U) of activity corresponds to 1  mmol ATP hydrolyzed/min (see Note 6).

ATP PEP

RUBISCO ACTIVASE

PK

ADP + Pi pyruvate

NAD +

NADH

LDH

lactic acid

Fig.  1. Coupled enzyme assay for measuring the rate of ATP hydrolysis by Rubisco activase.

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+ activase

Rubisco Activity

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Rubisco-RuBP (inactive)

minus activase or ATP Time

various times

Activase ADP ATP RuBP CO2

Rubisco (active) 14CO + 2

Mg2+

Mg2+ 14

1st stage

RuBP

C-PGA

2nd stage

Fig. 2. Two-stage assay for measuring Rubisco activation by Rubisco activase (modified from ref. 15). In the first stage, the inactive Rubisco–RuBP complex is “activated” by the interaction with Rubisco activase. The rate or extent of activation is determined by measuring Rubisco activity in the second stage (6).

3.2. Assay for Rubisco Activation

3.2.1. Formation of the Inactive Rubisco–RuBP Complex

The ability of Rubisco activase to promote Rubisco activation is assayed in a two-stage assay (Fig. 2) that measures Rubisco activity after incubating inhibited Rubisco (i.e., ER) with ATP and Rubisco activase (see Note 7). Rubisco activity is measured radiometrically by the RuBP-dependent rate of 14CO2 incorporation into acid-stable products (see Note 8). The amount of catalytically competent Rubisco is determined after incubation with Rubisco activase for various lengths of time and calculated from the specific activity of fully carbamylated Rubisco. 1. Slowly thaw Rubisco on ice (see procedure for Rubisco purification in Chap. 26). 2. Collect the precipitated protein by centrifugation at 10,000 × g for 10 min at 4°C. 3. Suspend the protein pellet in 150  mL of Activation Mix to fully carbamylate the enzyme. Incubate at 4°C for 1 h. 4. Equilibrate a 2  mL column of Sephadex G-50-fine with at least three column volumes of Rubisco ER Desalting Buffer. Place the column on top of a 500  mL screw-cap microfuge tube and place the entire assembly inside a 15 mL Corex glass tube.

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5. Centrifuge the column at 400 × g and 4°C for 2  min in a swinging bucket rotor to remove excess buffer. Discard eluant from the microfuge tubes. Centrifuge the column for an additional 1 min and again discard the eluant. 6. Carefully pipet 100–250 mL of fully carbamylated Rubisco on top of the dry column bed. 7. Centrifuge the column and collection tube assembly for 2 min at 400 × g and 4°C. 8. Recover the desalted sample from the microfuge tube. 9. Determine Rubisco concentration from the absorbance at 280  nm using the formula: Rubisco concentration (mg/ mL) = A280 × 0.61 (10). 10. Incubate Rubisco with 0.5  mM RuBP for 20  min at room temperature or at least 4  h at 4°C to form ER, the decarbamylated Rubisco–RuBP complex. The ER is stable for several days when stored at 4°C. 3.2.2. Two-Stage Rubisco Activation Assay (Fig. 2)

1. Prepare the mix for the Rubisco Activase first Stage Assay on ice without Rubisco activase and Rubisco–RuBP complex. Prepare separate reactions without RuBP, for the fully carbamylated controls, and with RuBP but without ATP and/ or Rubisco activase, for the spontaneous activation controls (see Note 9). 2. Immediately prior to assay, prepare the mix for the Rubisco 14 C Assay and dispense into 7-mL minivials (450 mL/assay). Store capped vials at 4°C until use. 3. Incubate first stage reactions at the desired temperature for 1  min and second stage Rubisco assays at 30°C for at least 3 min. 4. Add Rubisco activase to the first stage reaction to a final concentration of 0.1 mg/mL. 5. After 30 s, initiate the first stage reaction by adding 0.5 mg/ mL ER to the reaction mix (see Note 10). 6. At various times after the addition of ER (e.g., 0.5, 1.5, and 6 min; see Note 11), transfer a 50 mL aliquot of the first stage reaction to the Rubisco assay to initiate the reaction. 7. After 30 s, add 100 mL of the Reaction Quench Mix to the Rubisco assay to stop the reaction by acidification. Zero-time controls are prepared by adding the acid quench mix to the reaction prior to the addition of the first stage reaction aliquot. 8. Take the acidified Rubisco assays to dryness in a Speed-Vac Concentrator using high heat (65°C) for about 1  h (see Note 12).

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9. Add 500 mL of 0.1 N HCl to the dried contents of the vials and hydrate for 5 min. Suspend the samples by vigorous mixing using a vortex mixer. 10. Add 5 mL of scintillation cocktail, mix well, and determine 14 C dpm using a liquid scintillation counter. 11. Determine the specific activity of the 14C by adding 20 mL of the Rubisco Assay to 480 mL of 0.3 N KOH and then adding 5 mL of scintillation cocktail. 12. Calculate Rubisco activity as mmol of CO2 incorporated from the 14C dpm and the specific activity (i.e., 20 mL of the Rubisco assay contains 0.2 mmol total inorganic carbon). The percent of catalytically active sites is determined from the activity of the fully carbamylated control.

4. Notes 1. Methods for the purification of Rubisco activase from leaves and after expression in E. coli are described in the Chap. 28 (“Purification of Rubisco Activase from Leaves or after Expression in Escherichia coli” by Barta et al.). 2. Dissolve Tricine, MgCl2, KCl, and PEG in Milli-Q water. To avoid acidic hydrolysis of ATP, adjust the pH to 8.0, and then add the indicated amounts of ATP and PEP. This “incomplete ATPase reaction mix” can be stored indefinitely at −20°C. Prior to use, complete the assay mix by adding the indicated amounts of DTT, NADH, and PK/LDH enzymes. The complete assay mix may be stored frozen for several months at −20°C and freeze-thawed repeatedly. 3. Purified Rubisco is generally stored at −80°C in droplets as a frozen ammonium sulfate precipitate (for details see Chap. 26 “Isolation of Ribulose-1,5-Bisphosphate Carboxylase/ Oxygenase from Leaves” by Carmo-Silva et al.). 4. The volume of first stage reaction mix is adjusted for the number of time points assayed. Inclusion of RuBP prevents spontaneous activation of Rubisco. 5. ATP hydrolysis by Rubisco activase can also be measured by determining the rate of Pi release from ATP. We use the method of Chifflett (11) to determine Pi colorimetrically using a microplate reader. 6. Typical rates for purified Rubisco activase vary from 0.5 to 2.0 U/mg protein. 7. In addition to activating Rubisco, Rubisco activase will prevent deactivation of Rubisco in the presence of RuBP. Thus,

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it is also possible to assay Rubisco activase by its ability to prevent the decline in activity that accompanies decarbamylation and ER formation under catalytic conditions (12). 8. Rubisco activation can be measured spectrophotometrically in a single-stage assay (13). However, the spectrophotometric assay requires several linking enzymes, some with specific cofactor requirements. For our studies of Rubisco–Rubisco activase interaction, we prefer the two-stage assay since it allows the measurement of Rubisco activation under defined conditions. Also, attempts to use the highly sensitive enzymecycling method of Sulpice et al. (14) to measure Rubisco activation in leaves were thwarted by difficulty in obtaining one of the linking enzymes, i.e., PGA kinase, from the only supplier. 9. In the absence of Rubisco activase or ATP, the high level of RuBP present in the assay prevents spontaneous carbamylation of Rubisco. The increase in Rubisco activity in the absence of ATP and/or Rubisco activase (i.e., spontaneous activation controls) is typically less than 5–8% of the activity of the standard assay described with ATP and Rubisco activase (Fig. 2). 10. When performing first and second stage activation assays, it is important to adjust the timing of the reactions to allow timeshifted initiation and quenching of the second stage reactions. 11. With the concentrations of Rubisco activase and Rubisco concentrations suggested in the procedure (i.e., 0.1 and 0.5  mg/mL, respectively), 5  min is generally sufficient for complete activation of Rubisco in the first stage (12). With highly active or more concentrated enzymes, first stage assays of longer duration can consume all of the RuBP and/or ATP before completion. Purging assays with air-levels of CO2 in the presence of carbonic anhydrase can be used to reduce RuBP consumption during assays of longer duration (12). 12. Other types of drying systems can be used depending on the local regulations regarding the release of 14CO2 into the atmosphere. Potassium hydroxide in the acid trap of the Speed-Vac Concentrator adsorbs the unreacted 14CO2 during dry down of the assay vials. After several years, the trap is discarded as hazardous waste.

Acknowledgments The research was funded by the US Department of Agriculture, Agricultural research Service and the US Department of Energy, grant no. DE-FG02-08ER20268. Mention of a trademark,

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proprietary product, or vendor does not constitute a guarantee or warranty of the product by the United States Department of Agriculture and does not imply its approval to the exclusion of other products or vendors that may also be suitable. References 1. Spreitzer, R. J. and Salvucci, M. E. (2002) Rubisco: structure, regulatory interactions, and possibilities for a better enzyme. Annu Rev Plant Biol 53, 449–75. 2. Andrews, T. J. (1996) The bait in the Rubisco mousetrap. Nat Struct Biol 3, 3–7. 3. Portis, A. R. Jr. (2003) Rubisco activase – Rubisco’s catalytic chaperone. Photosynth Res 75, 11–27. 4. Wang, Z. Y. and Portis, A. R. Jr. (1992) Dissociation of ribulose-1,5-bisphosphate bound to ribulose-1,5-bisphosphate carboxylase/oxygenase and its enhancement by ribulose-1,5-bisphosphate carboxylase/oxygenase activase-mediated hydrolysis of ATP. Plant Physiol 99, 1348–53. 5. Robinson, S. P. and Portis, A. R. Jr. (1989) Adenosine triphosphate hydrolysis by purified Rubisco activase. Arch Biochem Biophys 268, 93–9. 6. Salvucci, M. E., Rajagopalan, K., Sievert, G., Haley, B. E., and Watt, D. S. (1993) Photoaffinity labeling of ribulose-1,5-bisphosphate carboxylase/oxygenase activase with ATP-g-benzophenone. J Biol Chem 268, 14239–44. 7. Salvucci, M. E., Portis, A. R. Jr., and Ogren, W. L. (1985) A soluble chloroplast protein catalyzes ribulose-bisphosphate carboxylase/ oxygenase activation in vivo. Photosynth Res 7, 193–201. 8. Portis, A. R. Jr., Salvucci, M. E., and Ogren, W. L. (1986) Activation of ribulosebisphosphate carboxylase/oxygenase at physiological CO2 and ribulosebisphosphate concentrations by Rubisco activase. Plant Physiol 82, 967–71.

9. Salvucci, M. E. (1992) Subunit interactions of Rubisco activase: polyethylene glycol promotes self-association, stimulates ATPase and activation activities, and enhances interactions with Rubisco. Arch Biochem Biophys 298, 688–96. 10. Wishnick, M. and Lane, M. D. (1971) Ribulose diphosphate carboxylase from leaves. Meth Enzymol 23, 570–77. 11. Chifflet, S., Torriglia, A., Chiesa, A., and Tolosa, S. (1988) A method for the determination of inorganic phosphate in the presence of labile organic phosphate and high concentrations of protein: application to lens ATPases. Anal Biochem 168, 1–4. 12. Crafts-Brandner, S. J. and Salvucci, M. E. (2000) Rubisco activase constrains the photosynthetic potential of leaves at high temperature and CO2. Proc Natl Acad Sci U S A 97, 13430–35. 13. Esau, B. D., Snyder, G. W., and Portis, A. R. Jr. (1996) Differential effects of N- and C-terminal deletions on the two activities of Rubisco activase. Arch Biochem Biophys 326, 100–5. 14. Sulpice, R., Tschoep, H., Von Korff, M., Bussis, D., Usadel, B., Hohne, M., WituckaWall, H., Altmann, T., Stitt, M., and Gibon, Y. (2007) Description and applications of a rapid and sensitive non-radioactive microplatebased assay for maximum and initial activity of D-ribulose-1,5-bisphosphate carboxylase/ oxygenase. Plant Cell Environ 30, 1163–75. 15. Salvucci, M. E. and Ogren, W. L. (1996) The mechanism of Rubisco activase: insights from studies of the properties and structure of the enzyme. Photosynth Res 47, 1–11.

Chapter 30 Quantification of Rubisco Activase Content in Leaf Extracts Wataru Yamori and Susanne von Caemmerer Abstract Rubisco activase functions to promote and maintain the catalytic activity of Rubisco. Studies with the activase-lacking Arabidopsis rca mutant (Salvucci et al. Photosynth Res 7:193–201, 1985; Salvucci et al. Plant Physiol 80:655–659, 1986), antisense activase tobacco, Arabidopsis and Flaveria bidentis plants (Mate et al. Plant Physiol 102:1119–1128, 1993; Eckardt et al. Plant Physiol 113:575–586, 1997; von Caemmerer et  al. Plant Physiol 137:747–755, 2005) have shown that photosynthesis at atmospheric levels of CO2 is severely impaired when plants lack activase because Rubisco becomes sequestered in an inactive form. Activase protein has been detected in all plant species, including C3 and C4 plants and green algae (Salvucci et al. Plant Physiol 84:930–936, 1987). Rubisco activase is essential in all these photosynthetic organisms for photosynthesis and plant growth. The physiological importance of Rubisco activase is reinforced by recent studies indicating that it plays a role in the response of photosynthesis to temperature. In this chapter, we describe how to extract and quantify Rubisco activase content in leaf. Key words: Photosynthesis, Rubisco, Rubisco activase, SDS-PAGE, Western blotting

1. Introduction The catalytic sites of Rubisco must be activated for CO2 fixation to take place. This requires the carbamylation of a lysine residue of the catalytic sites to allow the binding of Mg2+ and RuBP (7). Rubisco can also deactivate when RuBP binds to decarbamylated sites of Rubisco, sugar phosphates (e.g., 2-carboxyarabinitol 1-phosphate) bind to Rubisco catalytic sites, or through the formation and binding of the catalytic misfire products (e.g., xylulose 1,5-bisphosphate, 3-ketoarabinitol bisphosphate and pentadiulose 1,5-bisphosphate) by misprotonation of RuBP at the catalytic site (8–10). These interfere with both the process of carbamylation and full activity of the carbamylated enzyme. The activation and maintenance of Rubisco activity are facilitated by Rubisco activase (11, 12).

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Activase has been detected in all plant species examined, including both C3 and C4 plants (6). In many plants, activase is composed of two isoforms, larger form (a) and shorter form (ß), which are the products of alternative splicing of a single pre-mRNA in many plants (e.g., Arabidopsis, rice, and spinach) (12). In Arabidopsis, the isoforms are of 46 kDa and 43 kDa. Barley also expresses two isoforms of activase that are derived from alternative pre-mRNA splicing, but in addition contains a second activase gene for ß form of activase that is not alternatively spliced (13). In cotton, the two isoforms are encoded by separate genes (14). The two isoforms of activase differ in length by the presence of an extra about 30 amino acids at the C-terminus (6, 15). Although both two forms are active, only the a form is subjected to redox regulation via thioredoxin (16). On the other hand, some plants (e.g., tobacco, tomato, and maize) express only the non-redox-regulated ß form of activase (6). Especially, tobacco has at least three separate activase genes, but all of them encode the ß form (17). Thus, the functional significance of producing two forms of activase is unknown. Therefore, it might be better to quantify two activase isoforms separately, when activase contents are quantified in plants. This protocol describes (1) extraction of soluble proteins, including Rubisco activase, (2) separation of the extracted proteins by SDS-PAGE, (3) western blotting of the separated protein to a nitrocellulose membrane, and (4) subsequent immunodetection of Rubisco activase and quantification of Rubisco activase contents.

2. Materials 2.1. Extraction of Soluble Protein

1. Extraction buffer: 50  mM Hepes-KOH, pH 7.8, 10  mM MgCl2, 1 mM EDTA, 5 mM DTT, 0.1% triton X100 (v/v). Store at 4°C. 2. Complete Protease Inhibitor Cocktail: (Sigma, St Louis, Missouri, MO, USA) Store in aliquots at −20°C. 3. Rubisco activase: for the purification of Rubisco activase, see Chapter 28. 4. 4× sample-buffer: 250 mM Tris–HCl, pH 6.8, 40% glycerol, 8% SDS, 0.2% Bromophenol-blue, 200 mM DTT. With respect to DTT, add DTT right before use. Store at room temperature.

2.2. SDSPolyacrylamide Gel Electrophoresis

1. Invitrogen NuPAGE Gel System: (Invitrogen, Carlsbad, CA, USA). 2. Gel for quantifications of several photosynthetic components: (Invitrogen, Carlsbad, CA, USA). We often use a NuPAGE

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4–12% Bis-Tris Gel for quantifications of several photosynthetic components. 3. Running buffer: 50  mM MES, 50  mM Tris Base, 3.5  mM SDS, 0.8 mM EDTA, pH 7.3. We selected this buffer for a NuPAGE 4–12% Bis-Tris Gel according to the protocol of Invitrogen. We usually stock this for 10× running buffer at room temperature and dilute this with MilliQ water to onetenths before use. 4. Prestained molecular weight marker: Blue Plus2, (Invitrogen, Carlsbad, CA, USA). 2.3. Western Blotting for Rubisco Activase

1. XCell II blot module for western blotting: (Invitrogen, Carlsbad, CA, USA). 2. Nitrocellulose membrane: Hybond-C, (Amersham, UK). 3. 3.Transfer Buffer: 25 mM Bis-Tris, 25 mM Bicine, 0.8 mM EDTA, pH 7.2. Stock as 10× transfer buffer at room temperature and dilute with MilliQ water to one-tenths before use. 4. Tris-Buffered Saline (TBS): 20  mM Tris–HCl, pH 7.5, 150  mM NaCl. Stock for 10× TBS buffer at room temperature and dilute with MilliQ water to one-tenths before use. 5. Tris-Buffered Saline-Tween (TBST): 20 mM Tris–HCl, pH 7.5, 150  mM NaCl, 0.05% (v/v) tween 20. Stock for 10× TTBS buffer at room temperature and dilute with MilliQ water to one-tenths before use. 6. Blocking Solution: TBST with 10% skim milk powder. 7. Activase antibody (Primary antibody): original activase antibody raised against spinach activase with an N-terminal glutathione S-transferase fusion in rabbits in our laboratory. Or the antibody can now be purchased from companies (e.g., Santa Cruz Biotechnology Inc.) or order to produce an activase antibody suitable for your plants in lots of companies (see Note 1). 8. Secondary antibody: IgG-HRP goat anti-rabbit conjugate (Bio-Rad, Hercules, CA, USA) because our activase antibody is produced in rabbits (see Note 2).

2.4. Detection and Quantification of Activase

1. Attophos AP fluorescent substrate system: (Promega, Madison, WI, USA). 2. PhosphorImager: (Molecular Dynamics, Sunnyvale, CA, USA). 3. ImageQuant computer software: (Molecular Dynamics, Sunnyvale, CA, USA).

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3. Methods 3.1. Extraction of Soluble Protein

1. Extract each leaf disc (1.0 cm2) on ice in a mortar and pestle or a glass homogenizer in 800 ml of extraction buffer containing 25 ml (1.5%) of Complete Protease Inhibitor Cocktail. 2. Centrifuge the samples in a microcentrifuge tube at 16,000 × g for 1 min in a cooled centrifuge. Take 230 ml of the supernatant and add 70 ml SDS (10%) in a microcentrifuge tube, and then heat at 65°C for 10 min. For quantifications of activase, a dilution series is required. Purified activase is needed to quantify the absolute contents of activase as a standard (e.g., in the range from 0 to 30 ng). On the other hand, if you want to quantify the relative activase contents to the control plant, you can use the activase content in your control plant as a standard (e.g., in the range from 20 to 100%). In the same way, take 230 ml of the standard and add 70 ml SDS (10%) in an eppendorf tube, and then heat at 65°C for 10 min. 3. Add 45 ml of the extract (or the standard for a dilution series) to 15 ml of 4× sample buffer in a microcentrifuge tube. 4. Pierce cap in the eppendorf tube, heat at 100°C for 5 min, and then centrifuge at 16,000 × g for 1 min. After cooling to room temperature, the samples are ready for separation by SDS-polyacrylamide gel electrophoresis (SDS-PAGE).

3.2. SDS-PAGE

These instructions assume the use of Invitrogen NuPAGE Gel System. In this system, a wide range of gels for protein separations, including different gel formulations, percentages and formats are available, and it is easy to choose the appropriate gels suitable for your experiments. It is, of course, possible to make gels by yourself and run the gel, based on your system for SDS-PAGE. 1. Carefully slide off the comb from a gel. Remove the white tape from the bottom of the gel and gently wash packing buffer off with MilliQ water. 2. Insert the gel(s) so that the wells face inward. If only running one gel, there is a dummy gel that can be used on the other side. Push forward the white clamp provided with the gel rig to seal the compartment. Fill the inner compartment with running buffer so that the level is above the inner plate. Check that there are no leaks to the outer compartment. Wash wells out with running buffer using a syringe. Fill the outer compartment with the running buffer. 3. Load 12 mL sample per well with a fine pipette tip, layering gently from the bottom. Load 7  mL prestained molecular weight marker in a separate lane.

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4. Place a lid on the chamber, and run in 4°C refrigerator at 200 V constant for about 40 min. Turn off the power supply, when the Bromophenol-blue dye of the sample-buffer reaches the bottom of the gel. 3.3. Western Blotting for Rubisco Activase

The samples that have been separated by SDS-PAGE are transferred to supported membranes electrophoretically. These directions assume the use of XCell II blot module for western blotting (wet transfer) and nitrocellulose membranes. There are several membranes for blotting, including nitrocellulose membranes, nylon membranes and polyvinilidene difluoride (PVDF). It is better to use the nitrocellulose membranes in this protocol because activase protein is a soluble protein, and it is not necessary to infiltrate the membrane with methanol first. To use western blotting (wet transfer) instead of western blotting (semi-dry transfer) would be easy because it has less mistakes, although it takes more time. 1. Cut the nitrocellulose membrane and filter paper to about 8.0 × 7.5 cm for the gel size. Write the sample name on the membrane which faces the gel with a pencil. 2. Take the gel out of the gel plates, transfer the gel to transfer buffer in a clean square Petri dish, and allow it to soak for 10 min. Get ready to pre-wet filter papers and membranes in transfer buffer in second clean square Petri dish. 3. Place two pads on the negative electrode and then add a filter paper. Wet the surface of the filter paper with transfer buffer. Pick up the gel and position it on the filter paper ensuring that there are no bubbles between the paper and the gel. The bottom of the gel should sit at the base of the apparatus. 4. Place a wet membrane on the gel by starting at one edge and laying it down so that no bubbles are trapped between it and the gel. Bubbles can be removed by strocking the membrane with a gloved finger or a small test tube. Mark membrane so that orientation is obvious. 5. Place a wet filter paper on top of the membrane. 6. If only one gel is to be transferred, add enough pads to the sandwich so that there is no room for movement inside the apparatus. If transferring two gels, add one pad then assemble the next paper/gel/membrane/paper sandwich and proceed as before. 7. Add pads until the negative side of the electrode is almost full, and then place the positive electrode on top and wedge into the gel rig. Tighten with the white clamp provided with the gel rig. 8. Fill the inner compartment with transfer buffer, ensuring that there are no leaks. Fill the outer chamber with tap water to

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prevent the gel rig from overheating. Assemble the lid and run the transfer in the fridge at 20 V for overnight (or at 25 V constant for about 3 h). It is vitally important to ensure that the proteins are completely transferred from the gel to the membrane. If used, the colored molecular weight markers should be clearly visible on the membrane. 9. Remove the nitrocellulose membrane from the blotting apparatus, and place the membrane at the right side up in a clean square Petri dish. Wash the membrane for 5 min in TBS. 10. Incubate the membrane in blocking solution for 1  h on a shaker at room temperature. 11. Wash 1 × 10 min (at one time for 10 min) and 2 × 3 min in 40–50 ml TBST on shaker. 12. Incubate the membrane for 45  min with activase antibody (primary antibody) in 20 ml TBST with 1% skim milk powder on shaker. Primary antibody was used at 1:3,000 dilutions (6.7 mL antibody + 20 mL TBST). 13. Wash 1 × 10 min and 2 × 3 min in 40–50 ml TBST on shaker. 14. Incubate for 45 min in secondary antibody with 20 mL TBST on shaker. Secondary antibody was used at 1:5,000 dilutions (4.0 mL antibody + 20 mL TBST). 15. Wash 3 × 7 min in 40–50 ml TBST on shaker. 16. Finally, wash the membrane in TBS to remove the Tween-20 from the membrane surface prior to the next step. 3.4. Detection and Quantification of Activase

Activase bands on the membrane are visualized by enhanced chemiluminescence detection of immunoblots with the fluorescence detector. These instructions assume the use of Attophos AP fluorescent substrate system for the development of the activase bands (see Note 3) and the use of PhosphorImager for the detection of the bands (see Note 4). However, these instructions can be arranged by yourself, since there are lots of kits (e.g., HRP) and systems (e.g., X-ray film as a detector) for this stage. 1. Mix 100 mL 10× Attophos and 900 mL Attophos buffer with a pipette tip in a clean square Petri dish. 2. Remove extra buffer from the membrane with paper towels. Lower the membrane into the Attophos Buffer Mix so that the protein side is down and soaked. Air bubbles are undesirable. Let it sit out of direct light for 5–15 min. 3. Pick up the membrane and remove extra Attophos Buffer Mix from the membrane with paper towels. 4. Place the membrane in the system of PhosphorImager and detect the fluorescence at excitation wavelength 430 nm and emission wavelength 560 nm. A care is taken to ensure that the signal has not saturated.

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Fig.  1. Detection and quantification of activase (a) Immunoblot analysis of known amounts of purified tobacco activase (lane 1: 30 ng, lane 2: 15 ng, lane 3: 10 ng, lane 4: 5 ng) and samples (lane 5: wild type of tobacco, lanes 6 and 7: transformants of antiactivase tobacco). (b) A standard curve to quantify the activase content of leaf samples. Closed circles show a data for known amounts of purified tobacco activase and the solid line represents the correlation line. Open circle: wild type of tobacco, open triangle and open square: transformants of anti-activase tobacco.

5. The density of the bands was quantified by an ImageQuant computer software. It is also possible to use a free software for quantifications of the band intensities (e.g., Scion Image). Rubisco activase concentration in the leaf samples is determined by interpolating from the linear relationship between band intensity and Rubisco activase standard concentration (Fig. 1), (see Note 5).

4. Notes 1. The primary antibody can be saved for subsequent experiments by the addition of 0.05% final concentration of sodium azide (conveniently done by dilution from a 10% stock solution; exercise caution since azide is highly toxic.) and storage at 4°C. These primary antibodies can be used for about three times over several months, with the only adjustment required being the increasing length of exposure to scan. 2. Take care to match the correct animal for primary and secondary antibodies. Use new secondary antibody in every experiments. 3. We selected Attophos as the substrate for the development of the activase bands for two reasons: it has low background fluorescence due to the relatively large difference between its excitation (430–440 nm) and emission (550–560 nm) wavelengths, and there is little or no photo-bleaching upon

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repeated readings so that fluorescence can be followed for long periods of time. 4. The activase bands are detected by PhosphorImager (Molecular Dynamics, Sunnyvale, CA) in our laboratory. Any fluorometer that can be set to an excitation wavelength of 430–440  nm and an emission wavelength greater than 525  nm (the range of 550–560  nm optimal) may be used. Suitable instrument sources include BioTek, LabSystems, Molecular Devices and PerSeptive Biosystems. An X-ray film detection system could also be used. 5. Although crystal structure of activase is not proved, it is thought that the activase appears to be an oligomer with as many as 16 subunits (12). It might be better to show the activase content as a unit of milligram (mg). When you want to analyze activase content on the basis of molecular weight, you need to assume the structure of activase: for examples, the previous studies assumed the activase structure as a monomer or tetramer for the calculation of the molecular weight. References 1. Salvucci, M.E., Portis, A.R. Jr., and Ogren, W.L. (1985) A soluble chloroplast protein catalyzes ribulosebisphosphate carboxylase/ oxygenase activation in vivo. Photosynth Res 7, 193–201. 2. Salvucci, M.E., Portis, A.R. Jr., and Orgen, W.L. (1986) Light and CO2 response of ribulose1,5-bisphosphate carboxylase/oxygenase activation in Arabidopsis leaves. Plant Physiol 80, 655–659. 3. Mate, C.J., Hudson, G.S., von Caemmerer, S., Evans, J.R., and Andrews, T.J. (1993) Reduction of ribulose-bisphosphate carboxylase activase levels in tobacco (Nicotiana tabacum) by antisense RNA reduces ribulose bisphosphate carboxylase carbamylation and impairs photosynthesis. Plant Physiol 102, 1119–1128. 4. Eckardt, N.A., Snyder, G.W., Portis, A.R. Jr., and Ogren, W.L. (1997) Growth and photosynthesis under high and low irradiance of Arabidopsis thaliana antisense mutants with reduced ribulose-1,5-bisphosphate carboxylase/oxygenase activase content. Plant Physiol 113, 575–586. 5. von Caemmerer, S., Hendrickson, L., Quinn, V., Vella, N., Millgate, A.G., and Furbank, R.T. (2005) Reductions of Rubisco activase by antisense RNA in the C4 plant Flaveria bidentis reduces Rubisco carbamylation and leaf photosynthesis. Plant Physiol 137, 747–755.

6. Salvucci, M.E., Werneke, J.M., Portis, A.R. Jr., and Ogren, W.L. (1987) Purification and species distribution of Rubisco activase. Plant Physiol 84, 930–936. 7. Andrews, T.J. and Lorimer, G.H. (1987) Rubisco: Structure, mechanisms, and prospects for improvement, in The Biochemistry of Plants: A Comprehensive Treatise, Vol. 10, Photosynthesis (Hatch, M.D. and Boardman, N.K., eds.), Academic Press, New York, pp. 131–218. 8. Portis, A.R. Jr. (2003) Rubisco activase: Rubisco’s catalytic chaperone. Photosynth Res 75, 11–27. 9. Salvucci, M.E. and Crafts-Brandner, S.J. (2004) Inhibition of photosynthesis by heat stress: the activation state of Rubisco as a limiting factor in photosynthesis. Physiol Plant 120, 179–186. 10. Parry, M.A.J., Keys, A.J., Madgwick, P.J., Carmo-Silva, A.E., and Andralojc, P.J. (2008) Rubisco regulation: a role for inhibitors. J Exp Bot 59, 1569–1580 11. Andrews, T.J. (1996) The bait in the Rubisco mousetrap. Nat Struct Biol 3, 3–7. 12. Spreitzer, R.J. and Salvucci, M.E. (2002) Rubisco: interactions, associations and the possibilities of a better enzyme. Annu Rev Plant Biol 53, 449–475. 13. Rundle, S.J. and Zielinski, R.E. (1991) Organization and expression of two tandomly

Quantification of Rubisco Activase Content in Leaf Extracts oriented genes encoding ribulosebisphosphate carboxylase/oxygenase activase in barley. J Biol Chem 266, 4677–4685. 14. Salvucci, M.E., van de Loo, F.J., and Stecher, D.S. (2003) Two isoforms of Rubisco activase in cotton, the products of separate genes not alternative splicing. Planta 216, 736–744. 15. Werneke, J.M., Zielinski, R.E., and Ogren, W.L. (1988) Structure and expression of spinach leaf cDNA-encoding ribulosebisphosphate

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carboxylase oxygenase activase. Proc Natl Acad Sci U S A 85, 787–791. 16. Zhang, N. and Portis, A.R. Jr. (1999) Mechanism of light regulation of Rubisco: a specific role for the larger Rubisco activase isoform involving reductive activation by thioredoxin-f. Proc Natl Acad Sci U S A 96, 9438–9443. 17. Qian, J. and Rodermel, S.R. (1993) Ribulose1,5-bisphosphate carboxylase/oxygenase activase cDNAs from Nicotiana tabacum. Plant Physiol 102, 683–684.

Index A Antenna protein............................................. 171–172, 228 Arabidopsis thaliana (A. thaliana) allelism..................................................................... 262 create mapping population...................................... 263 establish isogenic line.............................................. 262 markers............................................................ 263, 265 mutagenesis in......................................................... 260 mutant screens................................................. 260–262 positional cloning............................................ 263–269 ATP hydrolysis............................... 363, 364, 375–378, 380

B Bundle sheath cells................................................ 327–337

C Cell culture............................. 259, 296–300, 304–305, 370 Chlamydomonas reinhardtii (C. reinhardtii) chloroplasts transformation DNA and PCR minipreparation, selection and screening............................................... 318 particle bombardment......................... 313, 316–319 PCR, selection and screening.................... 317–318 recipient strains and culture conditions............................................ 315–316 growing............................................. 314–315, 318–319 Chlorophyll (Chl) determination, Chl concentration................ 4–7, 13, 18, 39, 42, 86, 162, 176, 209, 254, 333 Chloroplast biogenesis................................................. 2–3, 257–271 cell culture............................................................... 259 intact chloroplasts.................................... 142, 146–147, 156, 174, 178, 210, 321–325 isolation.................................... 142, 324, 328, 331, 332 lipids determination, fatty acid compositions and lipids content................................................. 100–102 extraction.................................. 95, 97–98, 102, 132 isolation and identification.......................... 95–103 protein targeting, membranes.......................... 139–156 transformation, Chlamydomonas reinhardtii

DNA and PCR minipreparation, selection and screening...................................................... 318 particle bombardment......................... 315–317, 319 PCR, selection and screening of homoplasmic transformants....................................... 317–318 recipient strains and culture conditions...... 315–316 Chromatography............................................. 2, 4, 5, 8–10, 18, 19, 23, 25, 30, 32–35, 37, 42–43, 45, 46, 69–70, 72, 82–85, 88–91, 95–97, 100–103, 106–108, 110, 117, 132, 144, 152–154, 179, 180, 184, 219, 240, 255, 278, 285–286, 290, 291, 340, 341, 344, 346, 352, 355–356, 360–361, 364, 365, 367, 371–372 Circular dichroism (CD), LHCII......................... 132–135 Core particles isolation.............................................. 29–39 CP43 and CP47, isolation and purification............ 105–111 C4 plants................................................ 327, 331–332, 334, 337, 384 Cyanobacterium Cyanobase......... 281–282, 295–296, 298, 301–302, 308 Mastigocladus laminosus............................. 66, 68–69, 72 Nostoc sp. PCC 7120................................. 67, 69, 71–72 Synechocystis sp........................................................... 80 Synechocystis sp. PCC 6803 cell culture.................................................. 299–300 core particles.................................................. 29–39 gene interruptions and deletions, construction of..................................... 295–311 Synechocystis sp. PCC 7002, gene expression creating a site-specific, targeting vector for.............................................. 278–279 preparations....................................................... 279 transformation........................................... 279–280 Thermosynechococcus vulcanus........................... 42–44, 49 Cytochrome b6f complex grana and stroma membranes (B3 and T3)......... 53–63 isolation............................................................... 53–63 purification and crystallisation....................... 54, 65–75 Cytochrome c6, purification....................................... 79–92

E Electron paramagnetic resonance (EPR) free radicals...................................................... 190–198 reactive oxygen species (ROS)......................... 190–198

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Electron transfer............................................ 29, 66, 71–72, 79, 171, 187–188, 218, 238, 313–314 Electron transport.................................................... 11–12, 53, 66, 139, 155, 189, 202, 228, 306, 307, 309, 321, 322, 325, 363–364 Escherichia coli (E. coli)....................................... 2–3, 80–81, 84–90, 277–279, 288, 307–308, 310, 363–373, 380

F Ferredoxin........................................................ 72, 313, 322 Fluorescence induction.......................................... 221, 226, 234–238, 242, 325 Fourier transform infrared (FTIR), proteoliposomes.................. 218, 221, 226, 229–233

G Gene expression creating a site-specific, targeting vector for...... 278–279 Synechococcus sp. PCC 7002 preparations....................................................... 279 transformation........................................... 279–280 Gene interruptions and deletions construction cyanobase, use.................................................. 298, 301 oxygen evolution, basic phenotypic characterization......................................... 306–307 polymerase chain reaction (PCR)............ 301–304, 309 Synechocystis sp. PCC 6803 in cyanobacterium...................................... 295–311 transformation.................................... 298, 302–303 verification, mutant genotype................... 299, 303–305 Green algae, Monoraphidium braunii......................... 80, 81, 84, 85

H Higher plants............................... 1–10, 41, 53–54, 98, 172, 201–215, 217–218, 340, 349–350, 361, 363–364 High-performance liquid chromatography (HPLC)......................................... 32, 37, 106, 110, 115, 119, 123, 174, 180, 219, 240, 289–290, 315 Homologous recombinaison............................ 30, 274–276, 278, 279, 281, 282, 295–302, 310, 314

I Immobilization, photosynthetic materials albumin-glutaraldehyde (BSA-Glu)............... 249–251, 253–255 poly(vinylalcohol) (PVA-SbQ)........................ 248–250 Immunoblotting.................................... 172, 174–175, 214, 278, 290–291, 333, 351–353, 359, 361, 388–389 Intact chloroplasts isolation........................... 142, 146–147, 174, 178, 321–325 Ion exchange chromatography anionic........................................... 19, 88–90, 107–108, 340, 341, 344, 367, 372 cationic......................................................... 90–91, 108

L Light-harvesting chlorophyll-a/b complex (LHCIIb) LHCII circular dichroism (CD) measurements..... 132–135 isolation..................................................... 127–137 lamellar aggregates..................................... 127–137 lipid macro-assemblies............................... 127–137 native and recombinant preparation................ 118–124 Lipids chloroplast determination of fatty acid compositions and contents of lipids.................... 100, 102–103 extraction............................................... 95–98, 102 isolation and identification.......................... 95–103 proteoliposomes.............................................. 217–219, 221–222, 228–232, 234, 238–241

M Mapping........................................................ 172, 259–260, 263, 267–270, 298 Membrane protein........................................... 1, 17, 41–42, 95, 113, 160, 165, 184, 193, 249 Mutagenesis......................................................... 259–262, 267–269, 271, 274, 295–296, 307–308

O Oxygen evolution........................ 30, 36–38, 190, 210, 218, 220–221, 226, 228–229, 242, 306–307, 323–325 Oxygen-evolving complex (OEC).............................. 1–10, 29, 41–42, 106, 236

P Photosystem II (PSII) core complexes............................ 2–4, 7, 17, 36, 41, 110 CP43 and CP47 isolation and purification.......................................... 28, 105–111 extinsic proteins......................................................... 37 HPLC, core particles isolation............................ 32, 37 isolation......................................... 1–10, 17–26, 29–39, 42–44, 105–111, 203, 225, 236, 237, 307, 327 oxygen evolution............. 30, 36–38, 218, 221, 226, 228 oxygen-evolving complex (OEC) core complex................. 2–4, 7, 36, 41–50, 105–107 core particles isolation............................ 1–2, 29, 42 OEC subunit proteins isolation....................... 1–10 oxygen-evolving PSII membranes isolation................................................... 3, 5–7 purification, oxygen-evolving PSII complexes..................................................... 7–8 photoinhibition heat treatment.................................................... 210 photoinhibitory illumination............................. 210 protease....................................... 203, 207, 213–215 protein oxidation analysis................... 207, 212–213

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SDS/Urea-polyacrylamide gel electrophoresis (SDS/Urea-PAGE)..................................... 206 western blot analysis and fluorography...... 211–212 proteoliposomes, reconstitution into characterization......................................... 217–242 fluorescence induction............................... 221, 226, 234–238, 242 Fourier transform infrared (FT-IR) spectroscopy.................. 118, 221, 226, 229–231 lipid bilayer vesicles................................... 218, 219, 221, 222, 232, 239 oxygen evolution................................ 218, 220–221, 226, 228–229, 242 proteoliposomes......................................... 217–242 PS II particles............................................ 221, 223, 225, 226, 228–229, 232, 238, 241–242 sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE)..................... 221, 226, 227 thylakoid membranes................................ 217–218, 223–225, 235, 237–239, 241–242 reaction center (RC) complexes from plants 5-Chl and 6-Chl........................... 19–21, 23–24, 26 Cyt b559.................................................... 17, 24–25 D1-D2..................................................... 17, 24–25 isolation......................................................... 17–26 purity of preparation................................ 19, 24–25 Photosystem I (PSI) submembrane fractions isolation.......................................... 11–14, 250, 253 Pigment-protein complex................................ 29, 105–106, 108, 110, 113, 128, 134, 313–314 Plasmid...................................... 81, 92, 141, 145, 273–292, 297–298, 300–302, 308, 315, 317, 318, 369, 373 Plastocyanin purification........................................... 79–93 Plastoquinone................................. 11–12, 29, 37, 201, 235 Polymerase chain reaction (PCR).......................... 260, 263, 266–270, 276–279, 281–283, 288–289, 291, 301–304, 309, 315, 317–318 Protein targeting, chloroplast membranes............. 139–156 Protein transport................................ 2, 142–144, 153–154

R Reaction center (RC)....................................... 2, 17–26, 29, 30, 105, 109, 128, 171–172, 201–202, 228, 235, 247, 313–314 Reactive oxygen species (ROS) applications, spin trapping EPR spectroscopy............................................... 193–194 Ribulose–1,5-bisphosphate carboxylase/oxygenase (Rubisco) isolation of....................................... 339–347, 364–365, 367–369, 380 purification of........................... 340, 363–373, 380, 384 quantifying the amount and activity of............ 349–361

rubisco activase activity of.................... 339, 363, 368, 369, 375–383 quantification of......................... 349–350, 383–390

S Sodiumdodecylsulfate–polyacrylamide gel electrophoresis (SDS-PAGE).......................................... 24–25, 48, 58–60, 68, 70, 106, 110, 115, 120, 143–144, 148–153, 163, 165, 172–173, 176, 177, 179, 213, 221, 226–228, 285–286, 291, 333, 335, 344–345, 356–357, 369, 370, 372, 373, 384, 386–387 Spinach.................................................. 3–8, 11, 12, 17–18, 20, 25, 53–63, 67–72, 75, 80–82, 84–88, 106, 107, 117, 130, 136, 175, 179, 190, 205, 209–214, 251–255, 321–325, 346–347, 371, 384, 385 Spin trapping......................................................... 190–198

T Thin-layer chromatography (TLC), chloroplasts lipids............................................. 95–100, 102, 103 Thylakoid membranes electron paramagnetic resonance (EPR), free radicals....................................................... 190–198 extrinsic proteins............................................... 2, 48, 56 grana............................................................ 1–2, 53–63 immobilization of photosynthetic materials albumin-glutaraldehyde (BSA-Glu).................. 249 poly(vinylalcohol) (PVA-SbQ).......................... 249 intact chloroplasts.................................... 142, 146–147, 156, 174, 178, 210, 322 isolation..................................... 33, 42–44, 68, 69, 113, 174–176, 178, 190–194, 196, 224, 225, 307, 310 lipids......................................... 129–130, 132, 218, 241 phosphoproteins determination identification, phosphorylation sites.......... 171–185 immunodetection with protein specific antibodies..................................................... 177 labeling, thylakoid phosphoproteins with 32P-ATP....................................... 178–179 mass spectrometric analyses of.................. 173–174, 179–182 P-Thr antibodies method........... 174–176, 178, 183 preparation....................... 45, 68–69, 161–162, 179, 252 proteins............................................................ 165, 184 proteins characterization......... 2, 79, 160, 172, 248–249 protein targeting.............................................. 139–156 proteomic analysis........................................... 159–169 spin trapping EPR spectroscopy applications, ROS........................................................... 193–194

W Western blot.......................................... 173, 203, 206–207, 211–212, 356–357, 361, 384, 385, 387–388