Photosynthesis: Molecular Approaches to Solar Energy Conversion [47, 1 ed.] 3030674061, 9783030674069

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Advances in Photosynthesis and Respiration 47 Including Bioenergy and Related Processes

Jian-Ren Shen Kimiyuki Satoh Suleyman I. Allakhverdiev   Editors

Photosynthesis: Molecular Approaches to Solar Energy Conversion

Photosynthesis: Molecular Approaches to Solar Energy Conversion

Advances in Photosynthesis and Respiration Including Bioenergy and Related Processes VOLUME 47 Series Editors THOMAS D. SHARKEY Biochemistry and Molecular Biology, Michigan State University,  East Lansing, MI, USA JULIAN J. EATON-RYE Department of Biochemistry, University of Otago,  Dunedin, New Zealand Founding Editor GOVINDJEE University of Illinois at Urbana-Champaign, Urbana, IL, USA The book series Advances in Photosynthesis and Respiration – Including Bioenergy and Related Processes provides a comprehensive and state-of-the-art account of research in photosynthesis, respiration, bioenergy production and related processes. Virtually all life on our planet Earth ultimately depends on photosynthetic energy capture and conversion to energy-rich organic molecules. These are used for food, fuel, and fiber. Photosynthesis is the source of almost all Bioenergy on Earth. The fuel and energy uses of photosynthesized products and processes have become an important area of study and competition between food and fuel has led to resurgence in photosynthesis research. This series of books spans topics from physics to agronomy and medicine; from femtosecond processes through season-long production to evolutionary changes over the course of the history of the Earth; from the photophysics of light absorption, excitation energy transfer in the antenna to the reaction centers, where the highly-efficient primary conversion of light energy to charge separation occurs, through intermediate electron transfer reactions, to the physiology of whole organisms and ecosystems; and from X-ray crystallography of proteins to the morphology of organelles and intact organisms. In addition to photosynthesis in natural systems, genetic engineering of photosynthesis and artificial photosynthesis is included in this series. The goal of the series is to offer beginning researchers, advanced undergraduate students, graduate students, and even research specialists, a comprehensive, up-to-date picture of the remarkable advances across the full scope of research on photosynthesis and related energy processes. This series is designed to improve understanding of photosynthesis and bioenergy processes at many levels both to improve basic understanding of these important processes and to enhance our ability to use photosynthesis for the improvement of the human condition. For more information, please contact the Series Editors Thomas D.  Sharkey, Michigan State University, East Lansing, MI, U.S.A.  E-mail: [email protected]; phone 1-517-353-3257 or Julian J.  Eaton-Rye, Department of Biochemistry, University of Otago, New Zealand, E-mail: [email protected]. A complete list of references listed per volume can be found following this link: http:// www.life.uiuc.edu/govindjee/Reference-Index.htm Founding Editor Govindjee, Professor Emeritus of Biochemistry, Biophysics and Plant Biology Advisory Editors Elizabeth Ainsworth (USA); Basanti Biswal (India); Robert E. Blankenship (USA); Ralph Bock (Germany); Wayne Frasch (USA); Johannes Messinger (Sweden); Masahiro Sugiura (Japan); Davide Zannoni (Italy); and Lixin Zhang (China)

More information about this series at http://www.springer.com/series/5599

Photosynthesis: Molecular Approaches to Solar Energy Conversion Edited by

Jian-Ren Shen

Research Institute for Interdisciplinary Science, Okayama University, Okayama, Japan

Kimiyuki Satoh

Faculty of Science, Okayama University, Okayama, Japan and

Suleyman I. Allakhverdiev

Institute of Plant Physiology, Russian Academy of Sciences, Moscow, Russia

Editors Jian-Ren Shen Research Institute for Interdisciplinary Science Okayama University Okayama, Japan

Kimiyuki Satoh Faculty of Science Okayama University Okayama, Japan

Suleyman I. Allakhverdiev Institute of Plant Physiology Russian Academy of Sciences Moscow, Russia

ISSN 1572-0233     ISSN 2215-0102 (electronic) Advances in Photosynthesis and Respiration ISBN 978-3-030-67406-9    ISBN 978-3-030-67407-6 (eBook) https://doi.org/10.1007/978-3-030-67407-6 © Springer Nature Switzerland AG 2021 This work is subject to copyright. All rights are reserved by the Publisher, whether the whole or part of the material is concerned, specifically the rights of translation, reprinting, reuse of illustrations, recitation, broadcasting, reproduction on microfilms or in any other physical way, and transmission or information storage and retrieval, electronic adaptation, computer software, or by similar or dissimilar methodology now known or hereafter developed. The use of general descriptive names, registered names, trademarks, service marks, etc. in this publication does not imply, even in the absence of a specific statement, that such names are exempt from the relevant protective laws and regulations and therefore free for general use. The publisher, the authors, and the editors are safe to assume that the advice and information in this book are believed to be true and accurate at the date of publication. Neither the publisher nor the authors or the editors give a warranty, expressed or implied, with respect to the material contained herein or for any errors or omissions that may have been made. The publisher remains neutral with regard to jurisdictional claims in published maps and institutional affiliations. This Springer imprint is published by the registered company Springer Nature Switzerland AG The registered company address is: Gewerbestrasse 11, 6330 Cham, Switzerland

From the Series Editors Advances in Photosynthesis and Respiration Including Bioenergy and Related Processes Volume 47: Photosynthesis: Molecular Approaches to Solar Energy Conversion

Photosynthesis represents the central process in nature that has established the diversity of life on Earth. Each year, photosynthesis is responsible for using solar energy to convert about 100 gigatons of carbon dioxide into the energy-rich carbohydrates that ultimately support almost all ecosystems. The key step for the conversion of solar energy into chemical energy takes place in molecular machines known as photosystems. These are found in biological membranes in the chloroplasts of plants and algae and in similar membranes in photosynthetic bacteria. One of these photosystems, known as Photosystem II, uses light-energy to obtain electrons from water that are utilized in the reactions leading to fixing carbon into sugars while at the same time being the source of the oxygen we breathe and the oxygen of the protective ozone layer of our planet. In volume 47 of the Advances in Photosynthesis and Respiration (AIPH) series, Jian-Ren Shen, Kimiyuki Satoh, and Suleyman Allakverdiev have brought together an impressive team of experts providing a detailed appraisal of current knowledge on the molecular systems that operate in biological solar energy conversion. Inspired by the outstanding career of the late Professor Vyacheslav Vasilevich Klimov, and as a tribute to his many contributions to our understanding of Photosystem II, the

chapters focus on photosystems in plants, algae, and cyanobacteria and their light-harvesting systems. The importance of this field for the development of artificial photosynthesis and photosynthetic hydrogen production for clean energy technologies is also emphasized in this book. Authors of Volume 47 Reflecting the international impact of Professor Klimov’s career and the international focus of the AIPH book series, this volume has authors from 14 countries: Australia (6), China (2), Germany (2), Hungary (3), India (2), Iran (1), Israel (2), Japan (17), Russia (10), Slovak Republic (2), Spain (3), Ukraine (1), UK (1), and the USA (2). There are 54 authors (including the 3 editors) who are all recognized experts in their fields. Alphabetically (by last names), they are Parveen Akhtar, Seiji Akimoto, Fusamichi Akita, Miguel Alfonso, Suleyman I.  Allakhverdiev, James Barber, Vinzenz Bayro-Kaiser, Marian Brestic, Min Chen, Thomas Friedrich, Vera Grechanik, Győző Garab, Miguel A. Hernández-Prieto, Harvey J.M.  Hou, Kentaro Ifuku, Hiroshi Isobe, Takahashi Kawakami, Andrey A.  Khorobrykh, Petar H. Lambrev, María A.  Luján, Sai Kiran Madireddi, Mahir v

From the Series Editors

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D.  Mamedov, David Mauzerall, Koichi Miyagawa, Ruyo Nagao, Yoshiki Nakajima, Nathan Nelson, Takumi Noguchi, Ron J. Pace, Simon Petrie, Rafael Picorel, XiaoChun Qin. Galina Riznichenko, Andrew Rubin, Kimiyuki Satoh, Franz-Josef Schmitt, Alexey Yu. Semenov, Jian-Ren Shen, Rob Stranger, Rajagopal Subramanyam, Michihiro Suga, Oksana Sytar, Richard Terrett, Alexander N.  Tikhonov, Tatsuya Tomo, Peyman Mohammadzadeh Toutounchi, Anatoly Tsygankov, Yoshifumi Ueno, Wen-Da Wang, Kizashi Yamaguchi, Shunsuke Yamanaka, Denis V.  Yanykin, Makio Yokono, and Mareck Zivcak. We are grateful for their efforts in making this important volume. Our Books We list below information on the volumes that have been published thus far (see http://www.springer.com/series/5599 for the series website). Electronic access to individual chapters depends on subscription (ask your librarian), but Springer provides free downloadable front matter as well as indexes for all volumes. The available websites of the books in the series are listed below. • Volume 45 (2020) Photosynthesis in Algae: Biochemical and Physiological Mechanisms, edited by Anthony W.D.  Larkum from Australia, Arthur R.  Grossmann from the USA and John A.  Raven from the UK.  Seventeen chapters, 514 pp, Hardcover ISBN 978-3-030-33396-6, eBook ISBN 9783-030-33397-3 [http://www.springer.com/gp/ book/9783030333966] • Volume 44 (2018) The Leaf: A Platform for Performing Photosynthesis, edited by William W.  Adams III from the USA and Ichiro Terashima from Japan. Fourteen chapters, 575 pp, Hardcover ISBN 978-3-31993592-8, eBook ISBN 978-3-319-93594-2

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[ h t t p : / / w w w. s p r i n g e r. c o m / g p / book/9783319935928] Volume 43 (2018) Plant Respiration: Metabolic Fluxes and Carbon Balance, edited by Guillaume Tcherkez from Australia and Jaleh Ghashghaie from France. Eighteen chapters, 302 pp, Hardcover ISBN 978-3-31968701-8, eBook ISBN 978-3-319-68703-2 [ h t t p : / / w w w. s p r i n g e r. c o m / u s / book/9783319687018] Volume 42 (2016) Canopy Photosynthesis: From Basics to Applications, edited by Kouki Hikosaka from Japan, Ülo Niinemets from Estonia, and Neils P.R. Anten from the Netherlands. Fifteen chapters, 423 pp, Hardcover ISBN 978-94-017-7290-7, eBook ISBN 978-94-017-7291-4 [http://www. springer.com/book/9789401772907] Volume 41 (2016) Cytochrome Complexes: Evolution, Structures, Energy Transduction, and Signaling, edited by William A. Cramer and Tovio Kallas from the USA. Thirty-five chapters, 734 pp, Hardcover ISBN 978-94-017-7479-6, eBook ISBN 97894-017-7481-9 [http://www.springer.com/ book/9789401774796] Volume 40 (2014) Non-Photochemical Quenching and Energy Dissipation in Plants, Algae and Cyanobacteria, edited by Barbara Demmig-Adams, Győző Garab, William W.  Adams III, and Govindjee from USA and Hungary. Twenty-eight chapters, 649 pp, Hardcover ISBN 978-94-017-9031-4, eBook ISBN 978-94-017-9032-1 [http:// w w w. s p r i n g e r. c o m / l i f e + s c i e n c e s / plant+sciences/book/978-94-017-9031-4] Volume 39 (2014) The Structural Basis of Biological Energy Generation, edited by Martin F.  Hohmann-Marriott from Norway. Twenty-four chapters, 483 pp, Hardcover ISBN 978-94-017-8741-3, eBook ISBN 97894-017-8742-0 [http://www.springer.com/ life+sciences/book/978-94-017-8741-3] Volume 38 (2014) Microbial BioEnergy: Hydrogen Production, edited by Davide Zannoni and Roberto De Phillipis, from Italy.

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Eighteen chapters, 366 pp, Hardcover ISBN 978-94-017-8553-2, eBook ISBN 978- 94017-8554-9 [http://www.springer.com/ life+sciences/plant+sciences/ book/978-94-017-8553-2] Volume 37 (2014) Photosynthesis in Bryophytes and Early Land Plants, edited by David T. Hanson and Steven K. Rice, from USA.  Eighteen chapters, approx. 342 pp, Hardcover ISBN 978-94-007-6987-8, eBook ISBN 978-94-007-6988-5 [http://www. springer.com/life+sciences/plant+sciences/ book/978-94-007-6987-8] Volume 36 (2013) Plastid Development in Leaves during Growth and Senescence, edited by Basanti Biswal, Karin Krupinska and Udaya Biswal, from India and Germany. Twenty-eight chapters, 837 pp, Hardcover ISBN 978-94-007-5723-33, eBook ISBN 978-94-007-5724-0 [http://www.springer. com/life+sciences/plant+sciences/ book/978-94-007-5723-3] Volume 35 (2012) Genomics of Chloroplasts and Mitochondria, edited by Ralph Bock and Volker Knoop, from Germany. Nineteen chapters, 475 pp, Hardcover ISBN 978-94-0072919-3 eBook ISBN 978-94-007-2920-9 [http://www.springer.com/life+sciences/ plant+sciences/book/978-94-007-2919-3] Volume 34 (2012) Photosynthesis  - Plastid Biology, Energy Conversion and Carbon Assimilation, edited by Julian J.  Eaton-Rye, Baishnab C. Tripathy, and Thomas D. Sharkey, from New Zealand, India, and USA.  Thirtythree chapters, 854 pp, Hardcover, ISBN 97894-007-1578-3, eBook ISBN 978-94-007-1579-0 [http://www.springer. com/life+sciences/plant+sciences/ book/978-94-007-1578-3] Volume 33 (2012): Functional Genomics and Evolution of Photosynthetic Systems, edited by Robert L.  Burnap and Willem F.J.  Vermaas, from USA.  Fifteen chapters, 428 pp, Hardcover ISBN 978-94-007-1532-5, Softcover ISBN 978-94-007-3832-4, eBook ISBN 978-94-007-1533-2 [http://www.

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springer.com/life+sciences/ book/978-94-007-1532-5] Volume 32 (2011): C4 Photosynthesis and Related CO2 Concentrating Mechanisms, edited by Agepati S. Raghavendra and Rowan Sage, from India and Canada. Nineteen chapters, 425 pp, Hardcover ISBN 978-90-4819406-3, Softcover ISBN 978-94-007-3381-7, eBook ISBN 978-90-481-9407-0 [http:// w w w. s p r i n g e r. c o m / l i f e + s c i e n c e s / plant+sciences/book/978-90-481-9406-3] Volume 31 (2010): The Chloroplast: Basics and Applications, edited by Constantin Rebeiz, Christoph Benning, Hans J. Bohnert, Henry Daniell, J.  Kenneth Hoober, Hartmut K.  Lichtenthaler, Archie R.  Portis, and Baishnab C.  Tripathy, from USA, Germany, and India. Twenty-five chapters, 451 pp, Hardcover ISBN 978-90-481-8530-6, Softcover ISBN 978-94-007-3287-2, eBook ISBN 978-90-481-8531-3 [http://www. springer.com/life+sciences/plant+sciences/ book/978-90-481-8530-6] Volume 30 (2009): Lipids in Photosynthesis: Essential and Regulatory Functions, edited by Hajime Wada and Norio Murata, both from Japan. Twenty chapters, 506 pp, Hardcover ISBN 978-90-481-2862-4, Softcover ISBN 978-94-007-3073-1 eBook ISBN 978-90-4812863-1 [http://www.springer.com/ life+sciences/plant+sciences/ book/978-90-481-2862-4] Volume 29 (2009): Photosynthesis in Silico: Understanding Complexity from Molecules, edited by Agu Laisk, Ladislav Nedbal, and Govindjee, from Estonia, The Czech Republic, and USA. Twenty chapters, 525 pp, Hardcover ISBN 978-1-4020-9236-7, Softcover ISBN 978-94-007-1533-2, eBook ISBN 978-14020-9237-4 [http://www.springer.com/ life+sciences/plant+sciences/ book/978-1-4020-9236-7] Volume 28 (2009): The Purple Phototrophic Bacteria, edited by C.  Neil Hunter, Fevzi Daldal, Marion C.  Thurnauer and J.  Thomas Beatty, from UK, USA and Canada.

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Forty-eight chapters, 1053 pp, Hardcover ISBN 978-1-4020-8814-8, eBook ISBN 9781-4020-8815-5 [http://www.springer.com/ life+sciences/plant+sciences/ book/978-1-4020-8814-8] Volume 27 (2008): Sulfur Metabolism in Phototrophic Organisms, edited by Christiane Dahl, Rüdiger Hell, David Knaff and Thomas Leustek, from Germany and USA.  Twenty-four chapters, 551 pp, Hardcover ISBN 978-4020-6862-1, Softcover ISBN 978-90-481-7742-4, eBook ISBN 9781-4020-6863-8 [http://www.springer.com/ life+sciences/plant+sciences/ book/978-1-4020-6862-1] Volume 26 (2008): Biophysical Techniques Photosynthesis, Volume II, edited by Thijs Aartsma and Jörg Matysik, both from The Netherlands. Twenty-four chapters, 548 pp, Hardcover, ISBN 978-1-4020-8249-8, Softcover ISBN 978-90-481-7820-9, eBook ISBN 978-1-4020-8250-4 [http://www. springer.com/life+sciences/plant+sciences/ book/978-1-4020-8249-8] Volume 25 (2006): Chlorophylls and Bacteriochlorophylls: Biochemistry, Biophysics, Functions and Applications, edited by Bernhard Grimm, Robert J.  Porra, Wolfhart Rüdiger, and Hugo Scheer, from Germany and Australia. Thirty-seven chapters, 603 pp, Hardcover, ISBN 978-140204515-8, Softcover ISBN 978-90-481-7140-8, eBook ISBN 978-14020-4516-5 [http://www.springer.com/ life+sciences/plant+sciences/ book/978-1-4020-4515-8] Volume 24 (2006): Photosystem I: The Light-Driven Plastocyanin:Ferredoxin Oxidoreductase, edited by John H. Golbeck, from USA. Forty chapters, 716 pp, Hardcover ISBN 978-1-40204255-3, Softcover ISBN 978-90-481-7088-3, eBook ISBN 978-14020-4256-0 [http://www.springer.com/ life+sciences/plant+sciences/ book/978-1-4020-4255-3] Volume 23 (2006): The Structure and Function of Plastids, edited by Robert R.  Wise and J.  Kenneth Hoober, from

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USA.  Twenty-seven chapters, 575 pp, Softcover, ISBN: 978-1-4020-6570–6; Hardcover ISBN 978-1-4020-4060-3, Softcover ISBN 978-1-4020-6570-5, eBook ISBN 978-1-4020-4061-0 [http://www. springer.com/life+sciences/plant+sciences/ book/978-1-4020-4060-3] Volume 22 (2005): Photosystem II: The Light-Driven Water:Plastoquinone Oxidoreductase, edited by Thomas J.  Wydrzynski and Kimiyuki Satoh, from Australia and Japan. Thirty-four chapters, 786 pp, Hardcover ISBN 978-1-4020-4249-2, eBook ISBN 978-1-4020-4254-6 [http:// w w w. s p r i n g e r. c o m / l i f e + s c i e n c e s / plant+sciences/book/978-1-4020-4249-2] Volume 21 (2006): Photoprotection, Photoinhibition, Gene Regulation, and Environment, edited by Barbara DemmigAdams, William W.  Adams III and Autar K. Mattoo, from USA. Twenty-one chapters, 380 pp, Hardcover ISBN 978-14020-3564-7, Softcover ISBN 978-1-4020-9281-7, eBook ISBN 978-1-4020-3579-1 [http://www. springer.com/life+sciences/plant+sciences/ book/978-1-4020-3564-7] Volume 20 (2006): Discoveries in Photosynthesis, edited by Govindjee, J.  Thomas Beatty, Howard Gest and John F.  Allen, from USA, Canada and UK.  One hundred and eleven chapters, 1304 pp, Hardcover ISBN 978-1-4020-3323-0, eBook ISBN 978-1-4020-3324-7 [http://www. springer.com/life+sciences/plant+sciences/ book/978-1-4020-3323-0] Volume 19 (2004): Chlorophyll a Fluorescence: A Signature of Photosynthesis, edited by George C. Papageorgiou and Govindjee, from Greece and USA.  Thirty-one chapters, 820 pp, Hardcover, ISBN 978-1-4020-3217-2, Softcover ISBN 978-90-481-3882-1, eBook ISBN 978-1-4020-3218-9 [http://www. springer.com/life+sciences/ biochemistry+%26+biophysics/ book/978-1-4020-3217-2] Volume 18 (2005): Plant Respiration: From Cell to Ecosystem, edited by Hans Lambers

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and Miquel Ribas-Carbo, from Australia and Spain. Thirteen chapters, 250 pp, Hardcover ISBN978-14020-3588-3, Softcover ISBN 978-90-481-6903-0, eBook ISBN 978-14020-3589-0 [http://www.springer.com/ life+sciences/plant+sciences/ book/978-1-4020-3588-3] Volume 17 (2004): Plant Mitochondria: From Genome to Function, edited by David Day, A.  Harvey Millar and James Whelan, from Australia. Fourteen chapters, 325 pp, Hardcover, ISBN: 978-1-4020-2399-6, Softcover ISBN 978-90-481-6651-0, eBook ISBN 978-1-4020-2400-9 [http://www. springer.com/life+sciences/cell+biology/ book/978-1-4020-2399-6] Volume 16 (2004): Respiration in Archaea and Bacteria: Diversity of Prokaryotic Respiratory Systems, edited by Davide Zannoni, from Italy. Thirteen chapters, 310 pp, Hardcover ISBN 978-14020-2002-5, Softcover ISBN 978-90-481-6571-1, eBook ISBN 978-1-4020-3163-2 [http://www. springer.com/life+sciences/plant+sciences/ book/978-1-4020-2002-5] Volume 15 (2004): Respiration in Archaea and Bacteria: Diversity of Prokaryotic Electron Transport Carriers, edited by Davide Zannoni, from Italy. Thirteen chapters, 350 pp, Hardcover ISBN 978-1-40202001-8, Softcover ISBN 978-90-481-6570-4, eBook ISBN 978-1-4020-3163-2 [http:// w w w. s p r i n g e r. c o m / l i f e + s c i e n c e s / biochemistry+%26+biophysics/ book/978-1-4020-2001-8] Volume 14 (2004): Photosynthesis in Algae, edited by Anthony W.D.  Larkum, Susan Douglas and John A.  Raven, from Australia, Canada and UK.  Nineteen chapters, 500 pp, Hardcover ISBN 978-0-7923-6333-0, Softcover ISBN 978-94-010-3772-3, eBook ISBN 978-94-007-1038-2 [http://www. springer.com/life+sciences/plant+sciences/ book/978-0-7923-6333-0] Volume 13 (2003): Light-Harvesting Antennas in Photosynthesis, edited by Beverley R.  Green and William W.  Parson, from Canada and USA.  Seventeen chapters,

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544 pp, Hardcover ISBN 978-07923-6335-4, Softcover ISBN 978-90-481-5468-5, eBook ISBN 978-94-017-2087-8 [http://www. springer.com/life+sciences/plant+sciences/ book/978-0-7923-6335-4] Volume 12 (2003): Photosynthetic Nitrogen Assimilation and Associated Carbon and Respiratory Metabolism, edited by Christine H.  Foyer and Graham Noctor, from UK and France. Sixteen chapters, 304 pp, Hardcover ISBN 978-07923-6336-1, Softcover ISBN 978-90-481-5469-2, eBook ISBN 978-0-30648138-3 [http://www.springer.com/ life+sciences/plant+sciences/ book/978-0-7923-6336-1] Volume 11 (2001): Regulation of Photosynthesis, edited by Eva-Mari Aro and Bertil Andersson, from Finland and Sweden. Thirty-two chapters, 640 pp, Hardcover ISBN 978-0-7923-6332-3, Softcover ISBN 978-94017-4146-0, eBook ISBN 978-0-306-48148-2 [http://www.springer.com/life+sciences/ plant+sciences/book/978-0-7923-6332-3] Volume 10 (2001): Photosynthesis: Photobiochemistry and Photobiophysics, edited by Bacon Ke, from USA.  Thirty-six chapters, 792 pp, Hardcover ISBN 978-07923-6334-7, Softcover ISBN 978-0-79236791-8, eBook ISBN 978-0-306-48136-9 [http://www.springer.com/life+sciences/ plant+sciences/book/978-0-7923-6334-7] Volume 9 (2000): Photosynthesis: Physiology and Metabolism, edited by Richard C. Leegood, Thomas D. Sharkey and Susanne von Caemmerer, from UK, USA and Australia. Twenty-four chapters, 644 pp, Hardcover ISBN 978-07923-6143-5, Softcover ISBN 978-90-481-5386-2, eBook ISBN 978-0-306-48137-6 [http://www. springer.com/life+sciences/plant+sciences/ book/978-0-7923-6143-5] Volume 8 (1999): The Photochemistry of Carotenoids, edited by Harry A.  Frank, Andrew J. Young, George Britton and Richard J. Cogdell, from USA and UK. Twenty chapters, 420 pp, Hardcover ISBN 978-0-79235942-5, Softcover ISBN 978-90-481-5310-7, eBook ISBN 978-0-306-48209-0 [http://

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w w w. s p r i n g e r. c o m / l i f e + s c i e n c e s / plant+sciences/book/978-0-7923-5942-5] Volume 7 (1998): The Molecular Biology of Chloroplasts and Mitochondria in Chlamydomonas, edited by Jean David Rochaix, Michel Goldschmidt-Clermont and Sabeeha Merchant, from Switzerland and USA. Thirty-six chapters, 760 pp, Hardcover ISBN 978-0-7923-5174-0, Softcover ISBN 978-94-017-4187-3, eBook ISBN 978-0-30648204-5 [http://www.springer.com/ life+sciences/plant+sciences/ book/978-0-7923-5174-0] Volume 6 (1998): Lipids in Photosynthesis: Structure, Function and Genetics, edited by Paul-André Siegenthaler and Norio Murata, from Switzerland and Japan. Fifteen chapters, 332 pp. Hardcover ISBN 978-0-7923-5173-3, Softcover ISBN 978-90-481-5068-7, eBook ISBN 978-0-306-48087-4 [http://www. springer.com/life+sciences/plant+sciences/ book/978-0-7923-5173-3] Volume 5 (1997): Photosynthesis and the Environment, edited by Neil R. Baker, from UK.  Twenty chapters, 508 pp, Hardcover ISBN 978-07923-4316-5, Softcover ISBN 978-90-481-4768-7, eBook ISBN 978-0-30648135-2 [http://www.springer.com/ life+sciences/plant+sciences/ book/978-0-7923-4316-5] Volume 4 (1996): Oxygenic Photosynthesis: The Light Reactions, edited by Donald R.  Ort and Charles F.  Yocum, from USA. Thirty-four chapters, 696 pp, Hardcover ISBN 978-0-7923-3683-9, Softcover ISBN 978-0-7923- 3684–6, eBook ISBN 978-0306-48127-7 [http://www.springer.com/ life+sciences/plant+sciences/ book/978-0-7923-3683-9] Volume 3 (1996): Biophysical Techniques in Photosynthesis, edited by Jan Amesz and Arnold J.  Hoff, from The Netherlands. Twenty-four chapters, 426 pp, Hardcover ISBN 978-0-7923-3642-6, Softcover ISBN 978-90-481-4596-6, eBook ISBN 978-0-30647960-1 [http://www.springer.com/ life+sciences/plant+sciences/ book/978-0-7923-3642-6]

• Volume 2 (1995): Anoxygenic Photosynthetic Bacteria, edited by Robert E.  Blankenship, Michael T. Madigan and Carl E. Bauer, from USA. Sixty-two chapters, 1331 pp, Hardcover ISBN 978-0-7923-3682-8, Softcover ISBN 978-0-7923-3682-2, eBook ISBN 978-0-30647954-0 [http://www.springer.com/ life+sciences/plant+sciences/ book/978-0-7923-3681-5] • Volume 1 (1994): The Molecular Biology of Cyanobacteria, edited by Donald R. Bryant, from USA.  Twenty-eight chapters, 916 pp, Hardcover, ISBN 978-0-7923-3222-0, Softcover ISBN 978-0-7923-3273-2, eBook ISBN 978-94-011-0227-8 [http://www. springer.com/life+sciences/plant+sciences/ book/978-0-7923-3222-0]

Further information on these books and ordering instructions are available at http:// www. springer.com/series/5599. Special 25% discounts are available to members of the International Society of Photosynthesis Research, ISPR http://www. photosynthesisresearch.org/. See http:// www.springer.com/ispr. Future Advances in Photosynthesis and Respiration and Other Related Books The readers of the current series are encouraged to watch for the publication of the forthcoming books (not necessarily arranged in the order of future appearance): • Photosynthesis and Climate Change (working title) (Editors: Katie M.  Becklin, Joy K. Ward, and Danielle A. Way) • Cyanobacteria (Editor: Donald Bryant) • Modeling Photosynthesis and Growth (Editors: Xin-Guang Zhu and Thomas D. Sharkey)

In addition to the above books, the following topics are under consideration: Algae, Cyanobacteria: Biofuel and Bioenergy Artificial Photosynthesis

From the Series Editors ATP Synthase: Structure and Function Bacterial Respiration II Evolution of Photosynthesis Green Bacteria and Heliobacteria Interactions Between Photosynthesis and Other Metabolic Processes Limits of Photosynthesis: Where Do We Go from Here? Photosynthesis, Biomass, and Bioenergy Photosynthesis Under Abiotic and Biotic Stress

If you have any interest in editing/co-editing any of the above listed books, or being an author, please send an e-mail to Tom Sharkey ([email protected]) and/or to Julian EatonRye ([email protected]). Suggestions for additional topics are also welcome. Instructions for writing chapters in books in our series are available by sending e-mail requests to one or both of us. We take this opportunity to thank and congratulate Jian-Ren Shen, Kimiyuki Satoh, and Suleyman Allakhverdiev for their outstanding editorial work; they have collectively done an excellent job, not only in editing, but also in organizing this book for all of us, and for their highly professional dealing with the reviewing process. We thank all

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54 authors of this book (see the list given earlier and on the following pages); without their authoritative chapters, there would be no such volume. We give special thanks to Mr. Prasad Gurunadham at Straive, India, for directing the typesetting of this book; his expertise has been crucial in guiding the final steps that have bought this book to completion. We also thank Zuzana Bernhart, Andre Tournois, and Mariska van der Stigchel (Springer) for their friendly working relation with us that led to the production of this book and for their ongoing organization and assistance with the AIPH series. July 1, 2021 Julian J. Eaton-Rye Department of Biochemistr University of Otago Dunedin, New Zealand email: [email protected] Thomas D. Sharkey Department of Biochemistry and Molecular Biology Michigan State University East Lansing, MI, USA email: [email protected]

Series Editors

A 2017 informal photograph of Govindjee (right) and his wife Rajni (left) in Champaign-Urbana, Illinois; Photograph by Dilip Chhajed.

Govindjee  who uses one name only, was born on October 24, 1932, in Allahabad, India. Since 1999, he has been professor emeritus of biochemistry, biophysics, and plant biology at the University of Illinois at Urbana-Champaign (UIUC), Urbana, IL, USA. He obtained his B.Sc. (chemistry, botany, and zoology) and M.Sc. (botany and plant physiology) in 1952 and 1954, respectively, from the University of Allahabad. He learned plant physiology from Shri Ranjan, who was a student of Felix Frost Blackmann (of Cambridge, UK). Then, Govindjee studied photosynthesis at the UIUC, under two giants in the field, Robert Emerson (a student of Otto Warburg) and Eugene Rabinowitch (who had worked with James Franck), obtaining his Ph.D. in biophysics in 1960. Govindjee is best known for his research on excitation energy transfer, light emission (prompt and delayed fluorescence, and thermoluminescence), primary photochemistry, and electron transfer in Photosystem II (PS II, water-plastoquinone oxidoreductase). His research, with many others, includes the discovery of a short-wavelength form of chlorophyll (Chl) a functioning in PS II; of the

two-light effect in Chl a fluorescence; and, with his wife Rajni Govindjee, of the twolight effect (Emerson Enhancement) in NADP+ reduction in chloroplasts. His major achievements, together with several others, include an understanding of the basic relationship between Chl a fluorescence and photosynthetic reactions; a unique role of bicarbonate/carbonate on the electron acceptor side of PS II, particularly in the protonation events involving the QB binding region; the theory of thermoluminescence in plants; the first picosecond measurements on the primary photochemistry of PS II; and the use of fluorescence lifetime imaging microscopy (FLIM) of Chl a fluorescence in understanding photoprotection by plants against excess light. His current focus is on the history of photosynthesis research and in photosynthesis education. He has served on the faculty of the UIUC for approximately forty years. Govindjee’s honors include: Fellow of the American Association of Advancement of Science (AAAS); distinguished lecturer of the School of Life Sciences, UIUC; fellow and lifetime member of the National xiii

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Academy of Sciences (India); president of the American Society for Photobiology (1980–1981); Fulbright scholar (1956), Fulbright senior lecturer (1997), and Fulbright specialist (2012); honorary president of the 2004 International Photosynthesis Congress (Montréal, Canada); the first recipient of the Lifetime Achievement Award of the Rebeiz Foundation for Basic Biology, 2006; recipient of the Communication Award of the International Society of Photosynthesis Research, 2007; and of the Liberal Arts and Sciences Lifetime Achievement Award of the UIUC, (2008). Further, Govindjee has been honored many times: (1) in 2007, through 2 special volumes of Photosynthesis Research, celebrating his 75th birthday and for his 50-year dedicated research in photosynthesis (guest editor: Julian J. Eaton-Rye); (2) in 2008, through a special International Symposium on “Photosynthesis in a Global Perspective”, held in November 2008, at the University of Indore, India; this was followed by a book Photosynthesis: Basics and Applications (edited by S. Itoh, P. Mohanty and K.N.  Guruprad); (3) in 2012, through Photosynthesis  – Plastid Biology, Energy Conversion and Carbon Assimilation, edited by Julian J. Eaton-Rye, Baishnab C. Tripathy, and Thomas D. Sharkey; (4) in 2013, through special issues of Photosynthesis Research (volumes 117 and 118), edited by Suleyman Allakhverdiev, Gerald Edwards, and JianRen Shen celebrating his 80th (or rather 81st) birthday; (5) in 2014, through celebration of his 81st birthday in Třeboň, the Czech Republic (O.  Prasil [2014] Photosynth Res 122: 113–119); and (6) in 2016, through the

Series Editors prestigious Prof. B.M. Johri Memorial Award of the Society of Plant Research, India. In 2018, Photosynthetica published a special issue to celebrate his 85th birthday (Editor: Julian J. Eaton-Rye). Govindjee’s unique teaching of the Z-scheme of photosynthesis, where students act as different intermediates, has been published in two papers (1) P.K. Mohapatra and N.R. Singh [2015] Photosynth Res 123:105– 114); (2) S.  Jaiswal, M.  Bansal, S.  Roy, A, Bharati, and B, Padhi [2017] Photosynth Res 131: 351—359. Govindjee is a coauthor of a classic and highly popular book Photosynthesis (with E.I.  Rabinowitch, 1969) and of a historical book Maximum Quantum Yield of Photosynthesis: Otto Warburg and the Midwest Gang” (with K. Nickelsen, 2011). He is editor (or coeditor) of many books including: Bioenergetics of Photosynthesis (1975); Photosynthesis, 2 volumes (1982); Light Emission by Plants and Bacteria (1986); Chlorophyll a Fluorescence: A Signature of Photosynthesis (2004); Discoveries in Photosynthesis (2005); and Non-Photochemical Quenching and Energy Dissipation in Plants, Algae and Cyanobacteria (2015). Since 2007, each year, a Govindjee and Rajni Govindjee Award is given to graduate students, by the Department of Plant Biology (odd years) and by the Department of Biochemistry (even years), at the UIUC, to recognize excellence in biological sciences. For further information on Govindjee, see his website at http://www.life.illinois.edu/ govindjee.

Series Editors

Thomas D. (Tom) Sharkey  obtained his bach-

elor’s degree in biology in 1974 from Lyman Briggs College, a residential science college at Michigan State University, East Lansing, Michigan, USA. After 2 years as a research technician, Tom entered a Ph.D. program in the Department of Energy Plant Research Laboratory at Michigan State University under the mentorship of Klaus Raschke and finished in 1979. Postdoctoral research was carried out with Graham Farquhar at the Australian National University, in Canberra, where he coauthored a landmark review on photosynthesis and stomatal conductance. For 5 years he worked at the Desert Research Institute, Reno, Nevada. After Reno, Tom spent 20 years as professor of botany at the University of Wisconsin in Madison. In 2008, Tom became professor and chair of the Department of Biochemistry and Molecular Biology at Michigan State University. In 2017, Tom stepped down as department chair and moved to the MSU-DOE Plant Research Laboratory, completing a 38-year sojourn back to his beginnings. Tom’s research interests center on the exchange of gases between plants and the atmosphere and carbon metabolism of photosynthesis. The biochemistry and biophysics underlying carbon dioxide uptake and isoprene emission from plants form the two major research topics in his laboratory.

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Among his contributions are measurement of the carbon dioxide concentration inside leaves, an exhaustive study of short-term feedback effects in carbon metabolism, and a significant contribution to elucidation of the pathway by which leaf starch breaks down at night. In the isoprene research field, his laboratory has cloned many of the genes that underlie isoprene synthesis and he has published many important papers on the biochemical regulation of isoprene synthesis. Tom’s work has been cited over 26,000 times according to Google Scholar in 2017. He has been named an Outstanding Faculty member by Michigan State University, and in 2015, he was named a University Distinguished Professor. He is a fellow of the American Society of Plant Biologists and of the American Association for the Advancement of Science. Tom has co-edited three books, the first on trace gas emissions from plants in 1991 (with Elizabeth Holland and Hal Mooney), volume 9 of this series (with Richard Leegood and Susanne von Caemmerer) on the physiology of carbon metabolism of photosynthesis in 2000, and volume 34 (with Julian J. Eaton-Rye and Baishnab C. Tripathy) entitled Photosynthesis: Plastid Biology, Energy Conversion and Carbon Assimilation. Tom has been co-editor of this series since volume 31.

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Julian  J.  Eaton-Rye  is a professor in the Department of Biochemistry at the University of Otago, New Zealand. He received his undergraduate degree in botany from the University of Manchester in the UK in 1981 and his Ph.D. from the University of Illinois in 1987, where he worked with Govindjee on the role of bicarbonate in the regulation of electron transfer through Photosystem II.  Before joining the Biochemistry Department at Otago University in 1994, he was a postdoctoral researcher focusing on various aspects of Photosystem II protein biochemistry with Professor Norio Murata at the National Institute of Basic Biology in Okazaki, Japan, with Professor Wim Vermaas at Arizona State University, and with Dr. Geoffrey Hind at Brookhaven National Laboratory. His current research interests include structure-function relationships of Photosystem II proteins both in biogenesis and electron transport as well as the role of additional protein factors in the assembly of Photosystem II.  Julian has

Series Editors

been a consulting editor for the Advances in Photosynthesis and Respiration series since 2005, and edited volume 34 (with Baishnab C. Tripathy and Thomas D. Sharkey) entitled Photosynthesis: Plastid Biology, Energy Conversion and Carbon Assimilation. He is also an associate editor for the New Zealand Journal of Botany and an associate editor for the Plant Cell Biology part of Frontiers in Plant Science. He edited a Frontiers Research Topic titled Assembly of the Photosystem II Membrane-Protein Complex of Oxygenic Photosynthesis (with Roman Sobotka) in 2016 and this is available as an eBook [ISBN 978-2-88945-233-0]. Julian has served as president of the New Zealand Society of Plant Biologists (2006–2008) and as president of the New Zealand Institute of Chemistry (2012). He has been a member of the International Scientific Committee of the Triennial International Symposium on Phototrophic Prokaryotes (2009–2018) and is currently the secretary of the International Society of Photosynthesis Research.

Foreword What a wonderful collection of chapters dedicated to the memory of Slava Klimov. The editors are to be congratulated, and particularly Suleyman Allakhverdiev for drawing together chapters covering a wide spectrum of aspects of the light reactions of photosynthesis. Suleyman studied under the direction of Slava for his PhD and is one of his most successful students. Slava Klimov contributed greatly to photosynthesis research but perhaps his most important and lasting contribution was the discovery that pheophytin is the primary electron acceptor in the Photosystem Two (PS II) reaction center (1). In fact it was the study of PS II which dominated his research focus as recorded in the memorial paper published in Photosynthesis Research in 2017 soon after he passed away (2). What could be more important than unravelling the molecular mechanisms of one of the most important enzymes on our planet which has been the “Engine of Life” for the past three billion years? In fact life on our planet occurred about four billion years ago following half a billion years of geological and chemical evolution. It took the form of single-cell prokaryotic chemotrophs obtaining their energy from chemicals which could act as electron donors. After 1.5 billion years there were a wide variety of anaerobic prokaryotes which fell into two groups, archaea and bacteria. Darwinian principles had allowed these prokaryotes to be widely distributed in many different types of environments and evolve complex systems to capture and utilize

energy for the reductive process of cellular biochemistry. Sooner or later, however, the energy sources (oxidizable substrates) available to these chemotrophic prokaryotes became limiting and some developed the capacity to supplement their metabolic energy requirements by absorbing solar radiation. This heralded the “Big Bang of Evolution” since this strategy gave rise to phototrophic organisms which could use sunlight to totally drive their bioenergetics. This was achieved by using solar energy to extract “hydrogen” (in the form of high energy electrons and protons) from water and release oxygen into the atmosphere. From then on life prospered and diversified on an enormous scale driven by Darwinian evolution. It was from this success that humans evolved as a very special species with abilities far beyond any other animals that have ever lived on planet Earth. Yet we are on a very dangerous course which some believe could result in the elimination of our species in a matter of a few hundred years if we carry on the way we do today, that is, continue to use fossil fuel as a major energy source. Biology solved its energy problem by using sunlight to split water and extract the stored energy by recombining “hydrogen” with oxygen via respiration. We must do the same by using energy from the sun captured directly via photovoltaic cells and indirectly by hydro-, wind-, and wave-power to give electricity and hydrogen as the two main energy carriers. The latter complements batteries for renewable energy storage and must come from the electrolysis

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of water. There is an urgent need to progress by expanding and developing the appropriate technologies and putting in place the infrastructure, particularly for the generation of hydrogen and its storage, transport, and use. The studies of Slava Klimov and the many other contributions represented in this book serve as a blue print for tackling the most urgent problem facing organized humankind, namely, anthropogenic climate change. References 1. Klimov VV, Klevanik AV, Shuvalov VA, Krasnovsky AA (1977) Reduction of pheophytin in the primary light reaction of photosystem II. FEBS Lett 82:183–186

Foreword 2. Allakhverdiev SI, Zharmukhamedov SK, Rodionova MV, Shuvalov VA, Dismukes C, Shen JR, Barber J, Samuelsson G, Govindjee (2017) A scientist par excellence, a great human being, a friend, and a Renaissance man. Photosynth Res 136:1–16

James Barber Imperial College London, London, UK

Professor James Barber passed away on January 5, 2020 at the age of 79. He kindly wrote this forward for us while he was alive. The editors would like to acknowlege his great contributions in the field of photosynthesis research and pray for the repose of his soul.

Preface Photosynthesis has been the subject of extensive research for several decades involving a number of scientists in different fields, in order to elucidate and understand the mechanisms underlying the solar energy conversion reactions and to utilize them artificially to obtain renewable energy for the realization of a sustainable society. In recent years, photosynthesis research has increasingly developed into a deeper understanding of the molecular and atomic level of the components involved. The present book, Photosynthesis: Molecular Approaches to Solar Energy Conversion, focuses on such aspects as solar energy harvesting, utilization, and novel pigments absorbing and utilizing far-red light, from a molecular to an atomic level. Owing to the rapid development of X-ray crystallography and cryoelectron microscopy, the structures of photosystem I (PS I), photosystem II (PS II), and their complexes with various light-harvesting complexes (LHCs), have been solved, and they are the subject of this book. Chapter 1 describes the structure of the PS II core, its conservation and differences, from various organisms, and the mechanism of water oxidization achieved by recent pump-probe X-ray free electron laser experiments. This experimental result is followed by two theoretical chapters (Chaps. 2 and 3) describing the theoretical calculations of the Mn4CaO5 cluster and the mechanism of water oxidation. Chapter 2 favors the high oxidation scenario, whereas Chap. 3 favors the low oxidation scenario, of the Mn ions within the Mn4CaO5 cluster. Chapter 4 deals with the artificial utilization of solar energy by modeling natural water oxidization. A molecular mechanism for asymmetric electron transfer on the donor side of PS II is described in Chap. 12.

Due to the low energy density that solar energy has on the surface of the earth, light energy has to be collected and transferred to the reaction center of PS II and PS I to initiate the charge separation and a series of electron transfer reactions. The pigments involved in light-­ energy harvesting and transfer are reviewed in Chap. 5, with the emphasis on two recently characterized, far-red light absorbing pigments, chlorophyll d and f. Subsequently, the light-harvesting complexes II and I from various oxygenic photosynthetic organisms are described in Chaps. 6 and 7. These Chapters describe the structure and function of LHCII and LHCI from various organisms from red algae to diatoms and higher plants, and give a consensus overview of the LHC proteins and their associated pigments. In particular, the functions of chlorophyll d and f, absorbing light energy in the far-red region that was thought of little use before, have been “hot spots” of recent extensive studies, and their functions and energy transfer properties are considered further in Chaps. 8 and 9. The light energy absorbed will need to be distributed over the two PSs differently based on the light intensity, and there are a number of mechanisms to regulate the energy balance between the two PSs. Two major mechanisms for energy balancing, spill-over and state transitions, are described in Chaps. 10 and 11. Photosynthetic light-dependent reactions occur in the thylakoid membranes of cyanobacteria, various algae and higher plants, and the properties of membrane structure and organization will have great effects on the photosynthetic performance. There are a number of chapters dealing with the membrane dynamics and regulation, which include temperature-dependent regulation xix

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of photosynthesis studied by EPR (Chap. 13), plasticity of PS II (Chap. 14), role of lipids and fatty acids in the assembly and maintenance of photosynthetic complexes during PS II turnover (Chap. 15). Evolution and function of extrinsic proteins in water oxidization is described in Chap. 16, and effects of trehalose on the functional properties of PS II is reviewed in Chap. 17. Dynamic processes of the thylakoid membranes are discussed in Chap. 18, and its specific thermodynamic information detected by photoacoustics is described in Chap. 19. Accumulation of metabolites in plants under different light quality is reviewed in Chap. 20. Finally, artificial utilization of photosynthesis research is reviewed in Chaps. 21 and

Preface 22. Chapter 21 considers hydrogen production by natural PSs in an economic background, whereas Chap. 22 compares the ways to produce hydrogen by different approaches. By bringing a number of scientists working on the molecular to atomic levels of photosynthetic systems from various fields, the book provides an important overview on the current status of photosynthesis research. Jian-Ren Shen Okayama, Japan Kimiyuki Satoh Okayama, Japan Suleyman I. Allakhverdiev Moscow, Russia

A Tribute to Vyacheslav (Slava) Vasilyevich Klimov (1945–2017)

Vyacheslav Vasilyevich (V.V.) Klimov (or Slava, as most of us called him) was born on January 12, 1945, and passed away on May 9, 2017. He began his scientific career at the Bach Institute of Biochemistry of the USSR Academy of Sciences, Moscow, Russia, and then he became associated with the Institute of Photosynthesis, Pushchino, Moscow Region, for about 50 years. He worked in the field of biochemistry and biophysics of photosynthesis. He is known for his studies on the molecular organization of photosystem II (PS II). He was an eminent scientist in the

field of photobiology, a well-respected professor, and, above all, an outstanding researcher. Furthermore, he was one of the founding members of the Institute of Photosynthesis in Pushchino, Russia. To most, Slava Klimov was a great human being. He was one of the pioneers of research on the understanding of the mechanism of light energy conversion and of water oxidation in photosynthesis. Slava had many collaborations all over the world, and he is (and will be) very much missed by the scientific community and friends in Russia as well as xxi

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A Tribute to Vyacheslav (Slava) Vasilyevich Klimov (1945–2017)

around the world. We present here a brief biography and some comments on his research in photosynthesis. We remember him as a friendly and enthusiastic person who had an unflagging curiosity and energy to conduct outstanding research in many aspects of photosynthesis, especially that related to PS II.* *For more details see: Allakhverdiev SI, Zharmukhamedov SK, Rodionova MV, Shuvalov VA, Dismukes GC, Shen JR, Barber J, Samuelsson S, Govindjee (2018) Vyacheslav (Slava) Klimov (1945-­2017): A scientist par excellence, a great human being, a friend, and a Renaissance man. Photosynth Res, v. 136, p.  1-16. The text below is an edited excerpt from this article. Vyacheslav (Slava) Vasilyevich Klimov was born on January 12, 1945, in the village of Karavainka, Stalingrad (later Volgograd) in the Union of Soviet Socialist Republics (USSR). His parents were teachers: his father (Vasiliy Alexandrovich Klimov) was a history teacher and his mother (Elizaveta Ivanovna Klimova) was a primary school teacher. In 1963, after graduating from Gorno-Balykley secondary school, also in the Volgograd region, Slava Klimov entered the Department of Biology and Soil Science of M.V.  Lomonosov Moscow State University. In 1968, after graduating cum laude, he did postgraduate work on “Photoinduced Changes of Chlorophyll a Fluorescence Yield during Photosynthesis” in the same department. This research was done under the mentorship of Academician (Prof.) Alexander A.  Krasnovsky, a worldrecognized leader in photobiochemistry, and (Prof.) Navasard V. Karapetyan. Later, after Slava Klimov obtained the Candidate degree (equivalent to a PhD), he joined research group(s) led by A.A.  Krasnovsky and Vladimir (Vlad) A.  Shuvalov and worked first as a junior, and later as a senior, research investigator at the Institute of Photosynthesis of AN SSSR in Pushchino, Moscow Region. Then, in 1982, Slava Klimov was appointed as the head of the research laboratory for

“Photosynthetic Water Oxidation and Oxygen Evolution” at the Institute of Basic Biological Problems (formerly Institute of Photosynthesis), Russian Academy of Science, RAS, where he worked until the last days of his life. In 1986, Slava Klimov obtained his Doctor of Science (specialization: Biochemistry) from the A.N.  Bach Institute of Biochemistry, RAS, for his work on “Light Reactions of Electron Transfer in Photosystem II of Plants and Algae.” Slava Klimov was not only a brilliant scientist in the field of photosynthesis, but he was equally involved in educating young scientists. He became Professor of Biochemistry, teaching initially at the Pushchino State University and then at the Pushchino Branch of Moscow State University (MSU). In 1991, he became one of the laureates of the USSR State Prize for Science, awarded to the school of Academician Krasnovsky (A.A.  Krasnovsky, Yu.E.  Erokhin, V.B.  Evstigneev [posthumously], N.V. Karapetyan, A.V. Klevanik, V.V. Klimov and V.A.  Shuvalov) for studies on the Photobiochemistry of Chlorophylls. In addition, Slava Klimov was named to the prestigious Soros Professorship several times. As mentioned, Slava Klimov’s research centered on the molecular mechanism of light energy conversion and water oxidation during photosynthesis. He formulated and experimentally proved fundamentally new ideas about the mechanism of light energy conversion in oxygen-evolving PS II (Klimov and Krasnovsky 1981) that were widely accepted and included in contemporary reviews, monographs, and university courses on photosynthesis and advanced plant physiology. During his scientific career, Slava Klimov pioneered, along with his coworkers, the investigation of pheophytin participation in primary charge separation at the reaction center of PS II (see Klevanik et  al. 1977; Klimov et al. 1977, 1978, 1979a, b, 1980a, b, c, 1986). Furthermore, he and his collaborators revealed the quantitative composition and heterogeneity of the manganese cluster

A Tribute to Vyacheslav (Slava) Vasilyevich Klimov (1945–2017) of WOC, the water-oxidizing complex (Klimov et  al. 1982, 1985, 1990; Allakhverdiev et al. 1983, 1989a). Under Slava Klimov’s guidance, the concept of water photooxidation through a two-electron mechanism with the production of peroxide as an intermediary product was experimentally justified (Ananyev et al. 1992; Klimov et al. 1993b). In addition, the role of bicarbonate as a significant component for the formation and functioning of the WOC was revealed (Klimov et al. 1995a, b, 1997a, b; Wincencjusz et  al. 1996; Allakhverdiev et  al. 1997b; Hulsebosch et al. 1998; Yruela et al. 1998). Slava Klimov, together with others, put forward and experimentally proved the hypothesis regarding the crucial role of Mn-bicarbonate complexes in the evolutionary origin of oxygenic photosynthesis (Klimov et  al. 1995a, b; Dismukes et  al. 2001). For discoveries on the unique role of bicarbonate bound to non-heme-iron involved in electron transport (between the platoquinone electron acceptors QA and QB) and plastoquinone protonation, see Shevela et al. (2012). Furthermore, Slava was involved in the discovery of a new class of photosynthesis inhibitors (see Allakhverdiev et  al. 1989b, 1997a; Klimov et al. 1992, 1993a, 1995c, for details). In contrast to the known inhibitors, the action of this class of chemicals was shown to be based on the formation of a short electron cycle around the PS II reaction center. Being a powerful tool for the investigation of charge separation and recombination in PS II, such inhibitors can also be considered as potential eco-friendly herbicides since they inhibit reactions specific to plants (but not animals). In the laboratory headed by Slava Klimov, PS II associated carbonic anhydrase, important for both WOC functioning and stability, was also revealed (Villarejo et  al. 2002; Shutova et  al. 2008; Shitov et al. 2009, 2011; Karacan et al. 2012, 2014, 2016; also see Rodionova et al. 2017). Furthermore, upon the removal of a

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Mn-cluster from the WOC, an increase in photoinhibition was described by Klimov et al. (1990). On the chlorophyll (Chl) a fluorescence front, a new hypothesis for the origin of PS II variable Chl a fluorescence, as a recombination luminescence, was put forward (Klimov et al. 1978; Allakhverdiev et al. 1994a), and the redox potential values of the PS II primary electron donors and acceptors were first determined in his lab (Klimov et  al. 1979a, b; Allakhverdiev et  al. 2010, 2011). Slava Klimov had collaborations at many universities and research centers in the Netherlands, USA, United Kingdom, Canada, Japan, Sweden, and Spain. His colleagues and friends remember him as a kindhearted, cheerful person, a wise leader, and trusted comrade ready to help in any situation. We note that he had worked with a large number of scientists including Robert Carpentier (Canada), Norio Murata (Japan), James Barber (UK), Charles Dismukes (USA), Bacon Ke (USA), Göran Samuelsson (Sweden), Hans van Gorkom (the Netherlands), Arnold Hoff (the Netherlands), Gernot Renger (Germany), Rafael Picorel (Spain). Reminiscences Jian-Ren Shen It was really a sad news to hear that Prof. Slava Klimov passed away suddenly in May 2017. I met and talked with him many times at international conferences, and knew his great work along with his colleagues on the discovery of pheophytin in PS II, when I was a graduate student. I followed his numerous studies on the structure and function of PS II since then. He was a great person, and I have a wonderful memory of meeting him when I was in Pushchino to attend the “Photosynthesis Research for Sustainability” conference in 2014. His passing away is a great loss to the photosynthesis community, and I will miss him very much.

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A Tribute to Vyacheslav (Slava) Vasilyevich Klimov (1945–2017)

Suleyman I. Allakhverdiev It is obvious that the contribution of the laboratory headed by Slava Klimov had a great impact on the investigation of photosynthetic water oxidation and the mechanism of light energy conversion. Among Slava Klimov’s students, there were 18 Candidates of Science (PhD) and 2 Doctors of Science (Dr. Sci.), and I (SIA) was his first PhD student; Slava was not only my teacher, but a good friend. On October 2, 1977, I came from Baku (Azerbaijan) to Pushchino (Moscow Region, USSR) for my PhD thesis, and I joined the same group led by A.A. Krasnovsky and Vladimir A.  Shuvalov. I worked on the investigation of pheophytin in photosynthetic reaction centers (RCs) under Slava Klimov and academician A.A.  Krasnovsky as my supervisors. At that time two papers on pheophytin were published (Klevanik et  al. 1977; Klimov et  al. 1977). But this work was criticized by other researchers, who stated that it was an artifact. These were really very exciting times! Together, we tried to show that the participation of pheophytin in reaction centers of PS II is not an artifact. At that time, we published several papers in Russian (see Klimov et  al. 1978, 1979a, b, 1980a), and then Slava visited Bacon Ke’s lab in the USA (see Klimov et al. 1980b, c) where he performed additional experiments. The experimental evidence for pheophytin participation, and the energetics and kinetics of electron transport in PS II in the presence of pheophytin, was summarized in my PhD thesis. In 1984, I defended my thesis (in Physics and Mathematics [specialties-Biophysics]): “Photoreduction of Pheophytin in Reaction Centers of Photosystem II in Higher Plants and Algae” at the Institute of Biophysics, USSR Academy of Sciences, Pushchino, Moscow region, Russia. Then, in 2002, I obtained my Doctor of Science (highest/top degree in science) in Plant Physiology and Photobiochemistry from the Institute of Plant Physiology, RAS, Moscow, on “Functional Organization and Inactivation of Photosystem II.”

During 1977–1986, together with Slava Klimov, Sasha Klevanik, Vlad Shuvalov, and Professor Likhtenshtein’s group (at the branch of the Institute of Chemical Physics of the USSR Academy of Sciences in Chernogolovka, Moscow Region), we determined the number of manganese (Mn) atoms acting in the WOC of PS II.  It had been shown that the WOC on the PS II donor side contains four atoms of Mn. Reconstitution of the Mn-cluster after a complete removal of Mn from PS II preparations had been shown using Mn(II) as well as various artificial Mn-organic complexes (binuclear and/or tetranuclear). We studied the magnetic interaction of Mn with pheophytin and P680, and evaluated the distance between the main components of PS II. The immersion depths of the main components of PS II reaction center in thylakoid membranes were also analyzed (Klimov et  al. 1982, 1985, 1990; Allakhverdiev et  al. 1983, 1986, 1989a, b, 1994b; Kulikov et al. 1983). The effect of enhancement of PS II photoinhibition, upon the removal of Mn-cluster from the WOC, had been described by Klimov et al. (1990). In our joint work with Ivan Setlik’s group in Třeboň, we showed that under anaerobic and reducing conditions, photoinhibition occurs on the acceptor side of PS II at the level of QA and QB , whereas under aerobic conditions, it occurs on the acceptor and/or donor side of PS II; at the same time, separation and stabilization of charges in PS II RC remain unchanged (Allakhverdiev et  al. 1987, 1993; Klimov et al. 1990; Setlik et al. 1990). From 1988 to 1995, together with Slava Klimov, we spent more time on the bicarbonate effect on the electron donor side of PS II. Previously bicarbonate has been considered only as an important component for electron transfer between the plastoquinone electron acceptors, QA and QB; however, we found that while bound to the nonheme Fe (Shevela et al. 2012), removal of bicarbonate affects the PS II donor side reactions (see Klimov et  al. 1995a, b, 1997a, b).

A Tribute to Vyacheslav (Slava) Vasilyevich Klimov (1945–2017) Bicarbonate availability for the PS II donor side is especially significant for reactivation of the Mn-containing WOC after its removal by different treatments. It was suggested that bicarbonate may serve as a ligand to Mn, convert the aqua-ions of Mn (nonoxidized by PS II) into an easily oxidizable form Mn(HCO3), or act as a structural component important for the formation of a functionally active Mn cluster, or function in proton transfer (from water to the lumen). We didn’t risk publishing our results until 1995, and then investigations in the Netherlands (in the research groups of Arnold Hoff and Hans van Gorkom), in Spain (in the research group of Rafael Picorel), and in Sweden (in the research group of Göran Samuelsson) contributed to the shaping of our studies (see Klimov et al. 1995a, b, 1997a, b; 2003; Wincencjusz et  al. 1996; Allakhverdiev et  al. 1997a; Hulsebosch et al. 1998; Yruela et al. 1998; Shutova et al. 2008). Once again, I would like to emphasize that, for me, Slava was a teacher, a senior colleague, but also a good friend, serving as an adviser in both science and life in general. It is true to say that he was my very close friend and a dear teacher. I very much miss Slava Klimov for his stimulating attitude. Slava Klimov was a brilliant scientist. We are particularly impressed with his, and his colleague’s discovery of “pheophytin” and fundamental works about the functioning of PS II, including functioning of four atoms of Mn and the role of bicarbonate on the electron donor side of PS II. He was a wonderful human being and his whole life was devoted to science. References Allakhverdiev SI, Klevanik AV, Klimov VV, Shuvalov VA, Krasnovsky AA (1983) Estimation of the number of manganese atoms functioning in the donor side of photosystem II. Biofizika 28:5–8. (in Russian)

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Allakhverdiev SI, Shafiev MA, Klimov VV (1986) Effect of reversible extraction of manganese on photooxidation of chlorophyll P 680  in photosystem II preparations. Photobiochem Photobiophys 12:61–65 Allakhverdiev SI, Setlikova E, Klimov VV, Setlik I (1987) In photoinhibited photosystem II particles pheophytin photoreduction remains unimpaired. FEBS Lett 226:186–190 Allakhverdiev SI, Kulikov AV, Klimov VV, Bogatyrenko VR, Likhtenstein GI (1989a) Determination of the immersion depth of chlorophyll P680, pheophytin and secondary electron donor Z in pea subchloroplast preparations of the photosystem II.  Biofizika 34:434–438. (in Russian) Allakhverdiev SI, Zharmukhamedov SK, Klimov VV, Vasilev SA, Korvatovsky BN, Pashchenko VZ (1989b) Effect of dinoseb and other phenolic compounds on fluorescence decay kinetics of photosystem II chlorophyll in higher plants. Biol Membr 6:1147–1153. (in Russian) Allakhverdiev SI, Komenda J, Feyziev YM, Nedbal L, Klimov VV (1993) Photoinactivation of isolated D1/ D2/cytochrome b 559 complex under aerobic and anaerobic conditions. Photosynthetica 28:281–288 Allakhverdiev SI, Klimov VV, Carpentier R (1994a) Variable thermal emission and chlorophyll fluorescence in photosystem II particles. Proc Natl Acad Sci U S A 91:281–285 Allakhverdiev SI, Karacan MS, Somer G, Karacan N, Khan EM, Rane SY, Padhye S, Klimov VV, Renger G (1994b) Reconstitution of the w ­ ater-­ oxidizing complex in manganese-depleted photosystem II complexes by using synthetic binuclear manganese complexes. Biochemistry 33:12210–12214 Allakhverdiev SI, Klimov VV, Carpentier R (1997a) Evidence for the involvement of cyclic electron transport in the protection of photosystem II against photoinhibition: influence of a new phenolic compound. Biochemistry 36:4149–4154 Allakhverdiev SI, Yruela I, Picorel R, Klimov V (1997b) Bicarbonate is an essential constituent of the water-oxidizing complex of photosystem II. Proc Natl Acad Sci U S A 94:5050–5054 Allakhverdiev SI, Tomo T, Shimada Y, Kindo H, Nagao R, Klimov VV, Mimuro M (2010) Redox potential of pheophytin a in photosystem II of two cyanobacteria having the different special pair chlorophylls. Proc Natl Acad Sci U S A 107:3924–3929 Allakhverdiev SI, Tsuchiya T, Watabe K, Kojima A, Los DA, Tomo T, Klimov VV, Mimuro M (2011) Redox potentials of primary electron acceptor quinone molecule (QA) and conserved energetics of photosystem II in cyanobacteria with chlorophyll

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A Tribute to Vyacheslav (Slava) Vasilyevich Klimov (1945–2017)

a and chlorophyll d. Proc Natl Acad Sci U S A 108:8054–8058 Ananyev GM, Wydrzynski T, Renger G, Klimov VV (1992) Transient peroxide formation by the manganese-containing, redox-active donor side of PS II upon inhibition of O2 evolution with lauroylcholine chloride. Biochim Biophys Acta 1100:303–311 Hulsebosch RJ, Allakhverdiev SI, Klimov VV, Picorel R, Hoff A (1998) Effect of bicarbonate on the S2 multiline EPR signal of the oxygen-­evolving complex in photosystem II membrane fragments. FEBS Lett 424:146–148 Karacan MS, Yakan C, Yakan M, Karacan N, Zharmukhamedov SK, Shitov A, Los DA, Klimov VV, Allakhverdiev SI (2012) Quantitative structure-­ activity relationship analysis of perfluoroisopro pyldinitrobenzene derivatives known as photosystem II electron transfer inhibitors. Biochim Biophys Acta 1817:1229–1236 Karacan MS, Zharmukhamedov SK, Mama ş S, Kupriyanova EV, Shitov AV, Klimov VV, Özbek N, Özmen Ü, Gündüzalp A, Schmitt FJ, Karacan N, Friedrich T, Los DA, Carpentier R, Allakhverdiev SI (2014) Screening of novel chemical compounds as possible inhibitors of carbonic anhydrase and photosynthetic activity of photosystem II. J Photochem Photobiol B 137:156–167 Karacan MS, Rodionova MV, Tunç T, Venedik KB, Mamaş S, Shitov AV, Zharmukhamedov SK, Klimov VV, Karacan N, Allakhverdiev SI (2016) Characterization of nineteen antimony (III) complexes as potent inhibitors of photosystem II, carbonic anhydrase, and glutathione reductase. Photosynth Res 130:167–182 Klevanik AV, Klimov VV, Shuvalov VA, Krasnovsky AA (1977) Reduction of pheophytin in the light reaction of photosystem II of higher plant. Dokl Akad Nauk SSSR 236:241–244. (in Russian) Klimov VV, Krasnovsky AA (1981) Pheophytin as the primary electron acceptor in photosystem 2 reaction centres. Photosynthetica 15:592–609 Klimov VV, Klevanik AV, Shuvalov VA, Krasnovsky AA (1977) Reduction of pheophytin in the primary light reaction of photosystem II.  FEBS Lett 82:183–186 Klimov VV, Allakhverdiev SI, Pashchenko VZ (1978) Measurement of the activation energy and lifetime of fluorescence of photo- system II chlorophyll. Dokl Akad Nauk SSSR 242:1204–1207. (in Russian) Klimov VV, Allakhverdiev SI, Demeter S, Krasnovsky AA (1979a) Photoreduction of pheophytin in Photosystem 2 of chloroplasts with respect to redox potential of the medium. Dokl Akad Nauk SSSR 249:227–230. (in Russian)

Klimov VV, Allakhverdiev SI, Krasnovsky AA (1979b) EPR signal at photoreduction of pheophytin in Photosystem 2 reaction centres of chloroplasts. Dokl Akad Nauk SSSR 249:485–488. (in Russian) Klimov VV, Allakhverdiev SI, Shutilova NI, Krasnovsky AA (1980a) Investigation of pheophytin photoreduction and chlorophyll P680 photooxidation with preparations of photosystem II from pea and Chlamydomonas reinhardii chloroplasts Fiziologiya Rastenii (Soviet). Plant Physiol 27:315– 326. (in Russian) Klimov VV, Dolan E, Ke B (1980b) EPR properties of an intermediary electron acceptor (Pheophytin) in PS II reaction centers at cryogenic temperatures. FEBS Lett 112:97–100 Klimov VV, Dolan E, Shaw E, Ke B (1980c) Interaction between the intermediary electron acceptor (Pheophytin) and a possible plastoquinone-iron complex in photosystem II reaction centers. Proc Natl Acad Sci U S A 77:7227–7231 Klimov VV, Allakhverdiev SI, Shuvalov VA, Krasnovsky AA (1982) Effect of extraction and readdition of manganese on light reactions of photosystem II preparations. FEBS Lett 148:307–312 Klimov VV, Allakhverdiev SI, Shafiev MA, Demeter S (1985) Effect of complete extraction and re-addition of manganese on thermoluminescence of pea photosystem II preparations. Biochim Biophys Acta 809:414–420 Klimov VV, Allakhverdiev SI, Ladygin VG (1986) Photoreduction of pheophytin in photosystem II of the whole cells of green algae and cyanobacteria. Photosynth Res 10:355–361 Klimov VV, Shafiev MA, Allakhverdiev SI (1990) Photoinactivation of the reactivation capacity of photosystem II in pea subchloroplast particles after a complete removal of manganese. Photosynth Res 23:59–65 Klimov VV, Zharmukhamedov SK, Allakhverdiev SI, Kolobanova LP, Baskakov YA (1992) New phenolic inhibitors of electron transfer in photosystem II. Biol Membr 9:565–575. (in Russian) Klimov VV, Zharmukhamedov SK, Allakhverdiev SI, Kolobanova LP, Baskakov YA (1993a) New group of inhibitors of electron transfer in photosystem II of plants: chemical structure and efficiency of inhibition. Biol Membr 10:565–570. (in Russian) Klimov VV, Ananyev GM, Zastryzhnaya OM, Widrzynski T, Renger G (1993b) Photoproduction of hydrogen peroxide in photosystem II membrane fragments: a comparison of four signals. Photosynth Res 38:409–416

A Tribute to Vyacheslav (Slava) Vasilyevich Klimov (1945–2017) Klimov VV, Allakhverdiev SI, Feyziev YM, Baranov SV (1995a) Bicarbonate requirement for the donor side of photosystem II. FEBS Lett 363:251–255 Klimov VV, Allakhverdiev SI, Baranov SB, Feyziev YM (1995b) Effects of bicarbonate and formate on the donor side of photosystem II.  Photosynth Res 46:219–225 Klimov VV, Zharmukhamedov SK, De Las Rivas J, Barber J (1995c) Effect of PSII inhibitor K-15 on photochemical reactions of the isolated D1/D2 cytochrome b559 complex. Photosynth Res 44(6):7–74 Klimov VV, Baranov SV, Allakhverdiev SI (1997a) Bicarbonate protects the donor side of photosystem II against photoinhibition and thermoinactivation. FEBS Lett 418:243–246 Klimov VV, Hulsebosch B, Allakhverdiev SI, Wincencjusz H, van Gorkom HJ, Hoff A (1997b) Bicarbonate may be required for ligation of manganese in the oxygen-evolving complex of photo- system II. Biochemistry 36:16277–16281 Klimov VV, Allakhverdiev SI, Nishiyama Y, Khorobrykh AA, Murata N (2003) Stabilization of the oxygen-evolving complex of photosystem II by bicarbonate and glycinebetaine in thylakoid and subthylakoid preparations. Funct Plant Biol 30:797–803 Odionova MV, Zharmukhamedov SK, Karacan MS, Venedik KB, Shitov AV, Tunc T, Mamas S, Kreslavski VD, Karacan N, Klimov VV, Allakhverdiev SI (2017) Evaluation of new Cu(II) complexes as a novel class of inhibitors against plant carbonic anhydrase, glutathione reductase and photosynthetic activity in photosystem II. Photosynth Res 133:139–153 Setlik I, Allakhverdiev SI, Nedbal L, Setlikova E, Klimov VV (1990) Three types of photosystem 2 photoinactivation. I.  Damaging processes on the acceptor side. Photosynth Res 23:39–48 Shevela D, Eaton-Rye JJ, Shen JR, Govindjee (2012) Photosystem II and unique role of bicarbonate:

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a historical perspective. Biochim Biophys Acta 1817:1134–1151 Shitov AV, Pobeguts OV, Smolova TN, Allakhverdiev SI, Klimov VV (2009) Manganese-dependent carboanhydrase activity of photosystem II proteins. Biochem Mosc 74:509–517 Shitov AV, Zharmukhamedov SK, Shutova TV, Allakhverdiev SI, Samuelsson G, Klimov VV (2011) A carbonic anhydrase inhibitor induces bicarbonate-­ reversible suppression of electron transfer in pea photosystem 2 membrane fragments. J Photochem Photobiol B 104:366–371 Shutova T, Kenneweg H, Buchta J, Nikitina J, Terentyev V, Chernyshov S, Andersson B, Allakhverdiev SI, Klimov VV, Dau H, Junge W, Samuelsson G (2008) The photosystem II-associated Cah3  in Chlamydomonas enhances the O2 evolution rate by proton removal. EMBO J 27:782–791 Ulikov AV, Bogatyrenko VR, Likhtenstein GI, Allakhverdiev SI, Klimov VV, Shuvalov VA, Krasnovsky AA (1983) Magnetic interaction of manganese with pheophytin anion-radical and chlorophyll cation-radical in reaction centers of photosystem II.  Biofizika 28:357–363. (in Russian) Villarejo A, Shutova T, Moskvin O, Forssén M, Klimov VV, Samuelsson G (2002) A photosystem II-associated carbonic anhydrase regulates the efficiency of photosynthetic oxygen evolution. EMBO J 21:1930–1938 Wincencjusz H, Allakhverdiev SI, Klimov VV, van Gorkom HJ (1996) Bicarbonate-­ reversible formate inhibition at the donor side of photosystem II. Biochim Biophys Acta 1273:1–3 Yruela I, Allakhverdiev SI, Ibara JV, Klimov VV (1998) Bicarbonate binding to the water-oxidizing complex in the photosystem II. A Fourier transform infrared spectroscopy study. FEBS Lett 425:396–400

Contents

Part I Natural and Artificial Water Oxidation 1

Structure, Electron Transfer Chain of Photosystem II and the Mechanism of Water Splitting  3 Jian-Ren Shen, Yoshiki Nakajima, Fusamichi Akita, and Michihiro Suga Summary   4 I. Introduction   4 II. Structure of PS II    5 III. Electron Transfer Chain of PS II   20 IV. Structure of the Mn4CaO5-­Cluster and the Mechanism of Water Oxidation  21 Concluding Remarks and Perspectives   33 Acknowledgements  33 References  33

2

Mechanism of Water Oxidation in Photosynthesis Elucidated by Interplay Between Experiment and Theory 39 Kizashi Yamaguchi, Shusuke Yamanaka, Hiroshi Isobe, Mitsuo Shoji, Takashi Kawakami, and Koichi Miyagawa Summary  40 I. Introduction  41 II. Structure and Bonding of MnxOy Clusters  42 III. High-Resolution XRD Structure of PS II   47 IV. Electronic and Spin Structures of the S2 State  51 V. System Structures of OEC of PS II   53 VI. Possible Intermediates in the S3 State  59 VII. Possible Mechanisms for Water Oxidation   66 VIII. Concluding Remarks  72 Acknowledgements  73 References  74

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Contents

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3

On the Nature of the Functional S-States in the Oxygen Evolving Centre of Photosystem II—What Computational Chemistry Reveals About the Water Splitting Mechanism 81 Rob Stranger, Simon Petrie, Richard Terrett, and Ron J. Pace Summary  81 I. Introduction  82 II. The Oxidation State Possibilities   84 III. X-Ray Structures, Extended X-Ray Absorption Fine Structure   85 IV. Substrate Exchange Kinetics   89 V. X-Ray Spectroscopy  93 VI. Low Paradigm Functional S-States   96 VII. Mechanism of Oxygen Evolution   96 VIII. Conclusions 100 Acknowledgements 101 References 101

4

Toward Molecular Mechanisms of Solar Water Splitting in Semiconductor/Manganese Materials and Photosystem II105 Harvey J. M. Hou Summary 105 I. Introduction 106 II. Photosystem II Water Splitting Chemistry  108 III. Manganese-based Photosystem II Functional Models  110 IV. Semiconductor/Manganese Systems Mimicking Photosytem II  119 V. Concluding Remarks  124 Acknowledgements 126 References 126

Part II Light-Harvesting Systems 5

Chlorophyll Species and Their Functions in the Photosynthetic Energy Conversion133 Tatsuya Tomo and Suleyman I. Allakhverdiev Summary 133 I. Introduction 134 II. The Diversity of Chlorophylls and Related Pigments  136 III. Red-Shifted Chlorophylls  142 Acknowledgments 156 References 157

6

Structure, Organization and Function of Light-Harvesting Complexes Associated with Photosystem II Wenda Wang and Jian-Ren Shen

163

Summary 164 I. Introduction 164 II. Compositions and Functions of Various Types of Light-Harvesting Complexes II  168 III. Structures of LHCII and FCPII  173 IV. Organization of LHC Antennae in the PS II-LHCII Supercomplexes 185

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V. Energy Transfer Pathways and Photoprotection  186 VI. Perspectives 189 Acknowledgements 190 References 190

7

Structure, Function, and Evolution of Photosystem I-Light Harvesting Antenna I Complexes195 Xiaochun Qin Summary 195 I. Introduction 196 II. Structure of Cyanobacterial PS I and Evolution of the PS I Core Complex  197 III. Structure of the PS I Supercomplex of Higher Plants  201 IV. Structure of the PS I-LHCR Supercomplex from Red Algae  205 V. Structure of the PS I-LHCI Supercomplex of Green Algae  209 VI. Chlorophyll Arrangement of PS I-LHCI and its Possible Effect on Excitation Energy Transfer Pathways  215 VII. Evolution of the PS I Complex  217 Acknowledgements 217 References 219

8

Light Harvesting Modulation in Photosynthetic Organisms223 Miguel A. Hernández-Prieto and Min Chen Summary 224 I. Introduction 224 II. Light-Harvesting Protein Complexes  226 III. Light Acclimation and Adaptation  234 IV. Characteristics and Extended Functions of Light-Harvesting Protein Complex Superfamily Members  236 Acknowledgements 240 References 240

9

Red-Shifted and Red Chlorophylls in Photosystems: Entropy as a Driving Force for Uphill Energy Transfer?247 Thomas Friedrich and Franz-Josef Schmitt Summary 248 I. Introduction 248 II. “Uphill” Energy Transfer and Anti-Stokes Luminescence  251 III. “Red” vs. “Red-Shifted” Chlorophylls  253 IV. How Entropy Gain Supports “Uphill” Energy Transfer  269 V. Conclusions 271 Acknowledgements 272 References 272

10 Modification of Energy Distribution Between Photosystems I and II by Spillover Revealed by Time-Resolved Fluorescence Spectroscopy277 Makio Yokono, Yoshifumi Ueno, and Seiji Akimoto Summary 277 I. Introduction 278 II. Analysis of Energy Transfer  280 III. Energy Transfer Involving Antenna  283 IV. Evolution of Spillover Mechanisms  287

Contents

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V. VI. VII.

Benefits of the Direct-Type and Bridged-Type Spillovers  289 Thylakoid Structure and Spillover  292 Position of Quenching Site: Reaction Center

or Peripheral Antenna  293 VIII. Concluding Remarks  294

Acknowledgements 295 References 295

11 Perception of State Transition in Photosynthetic Organisms303 Rajagopal Subramanyam and Sai Kiran Madireddi Summary 303 I. Introduction 304 II. Photosystem Architecture  306 III. State Transitions  308 IV. Redox Poise of the Plastoquinone Pool  309 V. Role of Kinases and Phosphatases  311 VI. Phosphorylation of Thylakoid Membrane Proteins  312 VII. Thylakoid Membrane Dynamics in State Transitions  312 VIII. State Transitions and Cyclic Electron Flow  313 IX. Abiotic Stress and State Transition  314 X. Concluding Remarks  315 Acknowledgments 315 References 316

Part III Photo-Induced Charge Separation and Primary Electron Transfer Processes 12 Molecular Mechanism of Asymmetric Electron Transfer on the Electron Donor Side of Photosystem II323 Takumi Noguchi Summary 323 I. Introduction 324 II. Asymmetric Charge Distribution on the Radical Cation of the Chlorophyll Dimer P680  326 III. Asymmetric Photoreactions of Redox-Active Tyrosines, YZ and YD 331 IV. Mechanism of Asymmetric Electron Transfer from Tyrosines to P680+ 334 V. Conclusions 336 Acknowledgements 336 References 337

Part IV Membrane Dynamics and Regulation of Excitation Energy/Electron Transfer Processes 13 Structure-Function Relationships in Chloroplasts: EPR Study of Temperature-­Dependent Regulation of Photosynthesis, an Overview343 Alexander N. Tikhonov Summary 343 I. Introduction 344 II. Electron and Proton Transport in Chloroplasts  346 III. Lipid-Soluble Nitroxide Radicals as Molecular Probes for Membrane Fluidity  354

Contents

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IV.

Temperature-Dependent Regulation of Electron and Proton Transport and ATP Synthesis in Chloroplasts  358 V. Discussion and Concluding Remarks  366 Acknowledgements 368 References 368

14 Plasticity of Photosystem II. Fine-Tuning of the Structure and Function of Light-­Harvesting Complex II and the Reaction Center375 Petar H. Lambrev, Parveen Akhtar, and Győző Garab Summary 376 I. Introduction 376 II. Plasticity of Light-Harvesting Complex II  377 III. Plasticity of Photosystem II  383 Acknowledgements 388 References 388

15 Role of Lipids and Fatty Acids in the Maintenance of Photosynthesis and the Assembly of Photosynthetic Complexes During Photosystem II Turnover395 Miguel Alfonso, María A. Luján, and Rafael Picorel Summary 395 I. Introduction 396 II. Biosynthesis of Glycerolipids and Fatty Acids Is a Genuine Plastid Process  397 III. Chloroplast Membrane Lipids Have Different Composition with Respect to the Rest of the Cell Membranes  399 IV. Thylakoid Lipids Are Enriched in Polyunsaturated Fatty Acids  402 V. Role of Lipids in the Maintenance of Photosynthetic Activity  404 VI. Role of Lipids and Fatty Acids in the Assembly and Turnover of Photosystem II  413 VII. Concluding Remarks  417 Acknowledgements 419 References 419

16 Evolution and Function of the Extrinsic Subunits of Photosystem II429 Kentaro Ifuku and Ryo Nagao Summary 429 I. Introduction 430 II. Localization of Extrinsic Subunits in Photosystem II Structures  431 III. Functions of Each Extrinsic Subunit  432 IV. Molecular Evolution of PsbP and PsbQ Family Proteins  439 V. Concluding Remarks  440 Acknowledgements 441 References 441

17 Effect of Trehalose on the Functional Properties of Photosystem II447 Denis V. Yanykin, Andrey A. Khorobrykh, Alexey Yu. Semenov, and Mahir D. Mamedov Summary 447 I. Introduction 448

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Contents II. Effects of Trehalose on the Oxygen-Evolving PS II Complexes  450 III. Effects of Trehalose on the Manganese-Depleted PS II Complexes  452 IV. Discussion 454 Acknowledgements 459 References 459

18 Dynamic Models for the Electron Transfer Processes in Thylakoid Membranes465 Galina Riznichenko and Andrew Rubin Summary 465 I. Introduction: Kinetic and Agent-Based Models  466 II. Modelling the Processes in Photosynthetic Membranes  468 III. Modeling the Fluorescence Kinetics after Illumination by a Saturating Laser Pulse  473 IV. Detailed Kinetic Model of the Processes in Photosynthetic Membranes 476 V. Simplified Models  477 VI. Direct Multiparticle Models of Brownian Dynamics for the Description of Electron Transfer Involving Mobile Carriers  478 VII. Productive and Futile (Non-­productive) Encounter Complexes  481 VIIl. Probabilistic Models of Monte Carlo Type  482 IX. Models of Electron Fluxes Switching in Microalgae that Release Molecular Hydrogen  486 X. Concluding Remarks and Perspectives  492 Acknowledgements 493 References 493

19 Photoacoustics Reveals Specific Thermodynamic Information in Photosynthesis499 Harvey J. M. Hou and David Mauzerall Summary 499 I. Introduction 500 II. Photoacoustic Measurements on PS I  504 III. Thermodynamics of Charge Separation and S-State Cycle in PS II  515 IV. Limitations and Potential Problems  525 V. Conclusions 526 Acknowledgements 528 References 528

20 Plasticity of the Photosynthetic Energy Conversion and Accumulation of Metabolites in Plants in Response to Light Quality533 Oksana Sytar, Marek Zivcak, Marian Brestic, Peyman Mohammadzadeh Toutounchi, and Suleyman I. Allakhverdiev Summary 534 I. Introduction 534 II. Spectral Effects on Photosynthesis  536 III. Accumulation of Photoprotective Compounds Under Different Light Spectra  546 IV. Concluding Remarks  554 Acknowledgements 554 References 555

Contents

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Part V Photosynthetic Hydrogen Production 21 Feasibility of Sustainable Photosynthetic Hydrogen Production567 Vinzenz Bayro-Kaiser and Nathan Nelson Summary 567 I. Global Energy Economy – A Matter of Magnitude  568 II. Global Thermodynamics – only Solar Energy  569 III. Green Microalgae – A Complex Energy Managing Machine  570 IV. H2 Economy – The Way to Go  573 V. Microalgal H2 – Engineering a Photosynthetic Biorefinery  574 VI. Conclusions 582 Acknowledgments 583 References 583

22 Recent Advances in Microalgal Hydrogen Production589 Vera Grechanik and Anatoly Tsygankov Summary 589 I. Introduction 590 II. Hydrogen Production by Microalgae  590 III. Hydrogenases 592 IV. Overcoming the Oxygen Toxicity for Hydrogen Production  594 V. Genetic Approaches Improving Hydrogen Production  598 VI. Conclusions 602 References 602

Author Index

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Subject Index

609

Editors

Jian-Ren Shen  is a Professor at the Department of Biology, Okayama University, Japan, and an adjunct Professor at the Key Laboratory of Photobiology, Institute of Botany, Chinese Academy of Sciences, China. He obtained his PhD from the University of Tokyo in 1990, and performed postdoctoral research in the RIKEN Institute, Japan with Dr. Yorinao Inoue (1990– 1993). He subsequently joined the group of Dr. Yorinao Inoue in RIKEN Institute to study the structure and function of photosystem II (PS II). In 2003, he moved to Okayama University as a professor. His research interest is mainly focused on the structure and function of PS II, and in 2011, he and his colleagues succeeded in analyzing the structure of cyanobacterial PS II at the atomic level. This work illustrated the atomic

structure of the Mn4CaO5 cluster, the oxygen-­ evolving complex, and provided the basis for elucidating the mechanism of water-splitting. Recently, he also led his group to solve the structures of a number of photosynthetic systems such as PS I-light harvesting complex I from a higher plant, PS II-fucoxanthin chlorophyll a/c proteins of diatoms, by either X-ray crystallography or cryo-­electron microscopy. These studies greatly advanced our understanding of the photosynthetic systems. Owing to these studies, Dr. Shen was awarded several prizes, including the Gregori Aminoff Prize from The Royal Swedish Academy of Sciences, The Asahi Prize, The Green Science Award from the Cabinet of Japanese Prime Minister, etc. He has published more than 200 papers and around 10 book chapters. xxxvii

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Kimiyuki  Satoh  is an Emeritus Professor at Okayama University, Japan, where he was a staff member in the Department of Biology for 34 years. He graduated from Okayama University and earned a PhD from the University of Tokyo in 1972. He worked as a visiting scholar in the laboratory of Warren L. Butler at the University of California at San Diego (1975–1977) and was a Visiting Professor in the group of Charles J. Arntzen at the Michigan State University (1981), in the USA. He was an Adjunct Professor at the National Institute for Basic Biology in Okazaki, Japan (1992–1997). His contributions to the field of photosynthesis research have been recognized over the years by receiving awards and being elected to aca-

Editors

demic board memberships. In 2009, he was awarded the Finsen Medal from the Association Internationale de Photobiologie (later the International Union of Photobiology) for his contribution to the identification of the PS II reaction center. He has served several academic societies, including the Secretary of International Society of Photosynthesis Research (2001–2004) and the President of Japanese Society of Plant Physiologists (2002– 2003). He was a Consulting Editor for the Advances in Photosynthesis and Respiration series during 1999–2005 and edited volume 22 (with Tom Wydrzynski), Photosystem II: The Light-Driven Water:Plastoquinone Oxidoreductase (2005).

Editors

Suleyman I. Allakhverdiev  is the Head of the Controlled Photobiosynthesis Laboratory at the K.A. Timiryazev Institute of Plant Physiology of the Russian Academy of Sciences (RAS), Moscow; Chief Research Scientist at the Institute of Basic Biological Problems RAS, Pushchino, Moscow Region; Professor at the M.V.  Lomonosov Moscow State University, Moscow; Professor at the Moscow Institute of Physics and Technology (State University), Moscow, Russia; Head of Bionanotechnology Laboratory at the Institute of Molecular Biology and Biotechnologies of the Azerbaijan National Academy of Sciences, Baku, Azerbaijan; Invited Professor at the College of Science, King Saud University, Riyadh, Saudi Arabia, Professor at the Al-Farabi Kazakh National University, Almaty, Kazakhstan; Professor at the Foshan

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University, China, and Invited-Adjunct Professor at the Department of New Biology, Daegu Gyeongbuk Institute of Science & Technology(DGIST), Daegu, Republic of Korea. He received both his BS and MS in Physics from the Department of Physics, Azerbaijan State University, Baku. He earned his Dr.Sci. Degree in Plant Physiology and Photobiochemistry from the Institute of Plant Physiology, RAS (2002, Moscow), and PhD in Physics and Mathematics (Biophysics) from the Institute of Biophysics, USSR (1984, Pushchino). His PhD advisors were Academician Alexander A.  Krasnovsky and Dr. Sci. Vyacheslav V. Klimov. He worked for many years (1990–2007) as visiting scientist at the National Institute for Basic Biology (with Prof. Norio Murata), Okazaki, Japan, and in the Department de Chimie-Biologie, Université du

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Québec at Trois Rivières (with Prof. Robert Carpentier), Québec, Canada (1988–1990). He has broad cooperation with world-class researchers, including N. Murata, R. Carpentier, A. Hoff, G. Renger, Govindjee, H. van Gorkom, R. Picorel, S.  Ramakrishna, H.  Nishihara, J.  Eaton-Rye, M. Brestic, T. Tomo, J.-R. Shen, and M. Najafpour. He is associate editor of International Journal of Hydrogen Energy (Elsevier), section editor of BBA Bioenergetics (Elsevier), associate editor of Photosynthesis Research (Springer), associate editor of Photosynthetica (Springer), associate editor of Functional Plant Biology (CSIRO) and associate editor of Heliyon (Cell), and member of the Editorial Board of more than ten international journals. He has been a guest editor of many (more than 30) special issues in international peer-reviewed journals (Photosynth. Res.; Biochim. Biophys. Acta; Photochem. Photobiol. Sci.; J.  Photochem. Photobiol., Int. J Hydrogen Energy; Frontiers in Plant Science, etc.). He also acts as a referee for major international journals and grant proposals. He has authored (or coauthored) more than 300 research papers, 37 book chapters, 7 patents, and 9 books. His work has been cited over 19,000 times, and his h-index = 70. In 2016 he has been recognized by Thomson Reuters-­WoS (Clarivate Analytics) as the most Highly Cited Russian researcher worldwide in Biology. In 2018, 2019 and 2020 he was recognized by Thomson Reuters-WoS (Clarivate Analytics) as one of the Highly Cited world-­class researchers selected for their exceptional research performance, and entered the list of the most highly cited scientists of the world (1% of the total number of researchers). He has organized 10 international conferences on “Photosynthesis and Hydrogen Energy Research for Sustainability.”

Editors Allakhverdiev S.I. has the priority of discovering, detecting, and explaining the participation of pheophytin in electron transfer in the PS II reaction center (his PhD thesis). He suggested and proved energy and kinetic schemes of electron transfer during photosynthesis. His work proved that in photosynthesis, water oxidation and oxygen evolution occur involving four manganese atoms. Those results became a valuable contribution to worldwide biological science and are included in all textbooks in the photosynthesis field. During the last 10 years, Allakhverdiev S.I. has also worked in a new scientific discipline – the development of artificial photosynthesis systems in order to obtain molecular hydrogen as an alternative energy source. Through this work, the possibility of PS II reconstruction was demonstrated. More than 50 organometallic complexes (based on manganese oxides and some other metals with carbon nanotubes, graphene, or graphene oxide) were developed, synthesized, and studied in order to find highly effective water oxidation catalysts. A number of these compounds are capable of performing photosynthetic water oxidation mimicking the natural process. They are the prototype for the incorporation of a natureinspired oxygen-evolving complex in developed artificial photosynthesis systems. Artificial systems based on modified PS I and PS II for generating electricity (solar cells) and molecular hydrogen using solar energy have been developed and constructed. His research interests include the structure and function of PS II, especially the water-oxidizing complex, artificial photosynthesis, hydrogen photoproduction, catalytic conversion of solar energy, plants under environmental stress, and photo-receptor signaling.

Contributors

Parveen  Akhtar  Biological Research Centre, Szeged, Hungary ELI-ALPS, ELI Nonprofit Ltd., Szeged, Hungary

Vera  Grechanik  Institute of Basic Biological Problems Russian Academy of Sciences, Pushchino, Moscow Region, Russia

Seiji  Akimoto  Graduate School of Science, Kobe University, Kobe, Japan

Miguel  A.  Hernández-Prieto  School of Life and Environmental Sciences, University of Sydney, Sydney, NSW, Australia

Fusamichi  Akita  Research Institute for Interdisciplinary Science, and Graduate School of Natural Science and Technology, Okayama University, Okayama, Japan Miguel Alfonso  Department of Plant Nutrition, Estación Experimental de Aula Dei (EEAD), Consejo Superior de Investigaciones Científicas (CSIC), Zaragoza, Spain Suleyman  I.  Allakhverdiev  Institute of Plant Physiology, Russian Academy of Sciences, Moscow, Russia James  Barber  Imperial London, UK

College

London,

Vinzenz  Bayro-Kaiser  Department of Biochemistry and Molecular Biology, The George S.  Wise Faculty of Life Sciences, Tel Aviv University, Tel Aviv, Israel Marian  Brestic  Department of Plant Physiology, Slovak University of Agriculture, Nitra, Slovak Republic Min  Chen  School of Life and Environmental Sciences, University of Sydney, Sydney, NSW, Australia Thomas  Friedrich  Department of Bioenergetics, Technische Universität Berlin, Berlin, Germany Győző  Garab  Biological Research Centre, Szeged, Hungary Faculty of Science, University of Ostrava, Ostrava, Czech Republic

Harvey  J.  M.  Hou  Laboratory of Forensic Analysis and Photosynthesis, Department of Physical/Forensic Sciences, Alabama State University, Montgomery, AL, USA Kentaro Ifuku  Graduate School of Agriculture, Kyoto University, Kyoto, Japan Hiroshi  Isobe  Research Institute for Interdisciplinary Science, Okayama University, Okayama, Japan Takashi  Kawakami  Graduate School of Science, Osaka University, Toyonaka, Osaka, Japan RIKEN Center for Computational Science, Kobe, Hyogo, Japan Andrey  A.  Khorobrykh  Institute of Basic Biological Problems, FRC PSCBR RAS, Pushchino, Moscow, Russia Petar H. Lambrev  Biological Research Centre, Szeged, Hungary María A. Luján  Department of Plant Nutrition, Estación Experimental de Aula Dei (EEAD), Consejo Superior de Investigaciones Científicas (CSIC), Zaragoza, Spain Sai  Kiran  Madireddi  Department of Plant Sciences, School of Life Sciences, University of Hyderabad, Hyderabad, India Mahir D. Mamedov  A.N. Belozersky Institute of Physical-Chemical Biology, Lomonosov Moscow State University, Moscow, Russia

xli

xlii

David  Mauzerall  Laboratory of Photobiology, The Rockefeller University, New York, NY, USA Koichi  Miyagawa  Institute for Scientific and Industrial Research, Osaka University, Toyonaka, Osaka, Japan Ryo  Nagao  Research Institute for Interdisciplinary Science, Okayama University, Okayama, Japan Yoshiki  Nakajima  Research Institute for Interdisciplinary Science, and Graduate School of Natural Science and Technology, Okayama University, Okayama, Japan Nathan  Nelson  Department of Biochemistry and Molecular Biology, The George S.  Wise Faculty of Life Sciences, Tel Aviv University, Tel Aviv, Israel Takumi Noguchi  Division of Material Science, Graduate School of Science, Nagoya University, Nagoya, Japan Ron  J.  Pace  Research School of Chemistry, College of Physical and Mathematical Sciences, Australian National University, Canberra, Australia Simon  Petrie  Research School of Chemistry, College of Physical and Mathematical Sciences, Australian National University, Canberra, Australia Rafael  Picorel  Department of Plant Nutrition, Estación Experimental de Aula Dei (EEAD), Consejo Superior de Investigaciones Científicas (CSIC), Zaragoza, Spain Xiaochun  Qin  School of Biological Science and Technology, University of Jinan, Jinan, China Galina Riznichenko  Lomonosov Moscow State University, Biological Faculty, Moscow, Russia Andrew  Rubin  Lomonosov Moscow State University, Biological Faculty, Moscow, Russia Kimiyuki  Satoh  Faculty of Science, Okayama University, Okayama, Japan Franz-Josef  Schmitt  Department of Physics, Martin-Luther-Universität Halle-Wittenberg, Halle, Germany

Contributors Alexey Yu. Semenov  A.N. Belozersky Institute of Physical-Chemical Biology, Lomonosov Moscow State University, Moscow, Russia Jian-Ren  Shen  Research Institute for Interdisciplinary Science, Okayama University, Okayama, Japan Mitsuo  Shoji  Center for Computational Sciences, Tsukuba University, Tsukuba, Ibaraki, Japan Rob  Stranger  Research School of Chemistry, College of Physical and Mathematical Sciences, Australian National University, Canberra, Australia Rajagopal Subramanyam  Department of Plant Sciences, School of Life Sciences, University of Hyderabad, Hyderabad, India Michihiro  Suga  Research Institute for Interdisciplinary Science, and Graduate School of Natural Science and Technology, Okayama University, Okayama, Japan Oksana Sytar  Department of Plant Physiology, Slovak University of Agriculture, Nitra, Slovak Republic Department of Plant Biology, Institute of Biology and Medicine, Taras Shevchenko National University of Kyiv, Kiev, Ukraine Richard Terrett  Research School of Chemistry, College of Physical and Mathematical Sciences, Australian National University, Canberra, Australia Alexander  N.  Tikhonov  Department of Biophysics, Faculty of Physics, M. V. Lomonosov Moscow State University, Moscow, Russia Tatsuya  Tomo  Tokyo University of Science, Tokyo, Japan Peyman Mohammadzadeh Toutounchi  Depar tment of Agronomy, Faculty of Agriculture, Urmia University, Urmia, Iran of Basic Anatoly  Tsygankov  Institute Biological Problems Russian Academy of Sciences, Pushchino, Moscow Region, Russia Yoshifumi  Ueno  Graduate School of Science, Kobe University, Kobe, Japan

Contributors Wenda Wang  Photosynthesis Research Center, Key Laboratory of Photobiology, Institute of Botany, Chinese Academy of Sciences, Beijing, China Kizashi  Yamaguchi  Institute of Scientific and Industrial Research, Osaka University, Toyonaka, Osaka, Japan RIKEN Center for Computational Science, Kobe, Hyogo, Japan Shusuke Yamanaka  Graduate School of Science, Osaka University, Toyonaka, Osaka, Japan

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Denis V. Yanykin  Institute of Basic Biological Problems, FRC PSCBR RAS, Pushchino, Moscow, Russia Makio  Yokono  Institute of Low Temperature Science, Hokkaido University, Sapporo, Japan Marek Zivcak  Department of Plant Physiology, Slovak University of Agriculture, Nitra, Slovak Republic

Part I Natural and Artificial Water Oxidation

Chapter 1 Structure, Electron Transfer Chain of Photosystem II and the Mechanism of Water Splitting Jian-Ren Shen*,

Research Institute for Interdisciplinary Science, Okayama University, Okayama, Japan

and Yoshiki Nakajima, Fusamichi Akita, and Michihiro Suga Research Institute for Interdisciplinary Science, and Graduate School of Natural Science and Technology, Okayama University, Okayama, Japan

Summary������������������������������������������������������������������������������������������������������������������������������������������� 4 I. Introduction������������������������������������������������������������������������������������������������������������������������������� 4 II. Structure of PS II���������������������������������������������������������������������������������������������������������������������� 5 A. Structure of Cyanobacterial PS II���������������������������������������������������������������������������������������� 5 1. Organization and Structure of Protein Subunits������������������������������������������������������������� 5 2. Arrangement of Pigments��������������������������������������������������������������������������������������������  13 3. Other Cofactors�����������������������������������������������������������������������������������������������������������  14 B. Structure of Red Algal PS II����������������������������������������������������������������������������������������������  15 C. Structure of Diatom PS II��������������������������������������������������������������������������������������������������  15 D. Structure of Green Algal PS II�������������������������������������������������������������������������������������������  17 E. Structure of Higher Plant PS II������������������������������������������������������������������������������������������  19 III. Electron Transfer Chain of PS II���������������������������������������������������������������������������������������������� 20 IV. Structure of the Mn4CaO5-­Cluster and the Mechanism of Water Oxidation���������������������������� 21 A. Structure of the Mn4CaO5-Cluster������������������������������������������������������������������������������������� 22 1. S1-State������������������������������������������������������������������������������������������������������������������������ 22 2. S2-State����������������������������������������������������������������������������������������������������������������������� 29 3. S3-State����������������������������������������������������������������������������������������������������������������������� 30 B. Mechanism of Water Splitting�������������������������������������������������������������������������������������������� 31 Concluding Remarks and Perspectives������������������������������������������������������������������������������������������� 33 Acknowledgements�������������������������������������������������������������������������������������������������������������������������� 33 References�������������������������������������������������������������������������������������������������������������������������������������� 33

*Author for correspondence, e-mail: [email protected] © Springer Nature Switzerland AG 2021 J.-R. Shen et al. (eds.), Photosynthesis: Molecular Approaches to Solar Energy Conversion, Advances in Photosynthesis and Respiration 47, https://doi.org/10.1007/978-3-030-67407-6_1

3

J.-R. Shen et al.

4

Summary Photosystem II (PS II) is a multisubunit enzyme embedded in the lipid environment of thylakoid membranes of plants, various algae, and cyanobacteria, and performs a series of electron transfer and water-splitting reactions using the energy from the sun. Recent structural studies have revealed the details of the protein subunits and cofactors of PS II from various organisms by both high resolution X-ray diffraction and cryo-electron microscopic analyses of PS II cores. These studies give rise to a picture of well-conserved PS II core with variations in some peripheral subunits among the different organisms. Pump-­probe time-resolved structural studies also revealed the mechanism of water-splitting taking place within PS II. We describe the structures of PS II from various organisms and their variations, and the detailed mechanism of water-splitting in cyanobacterial PS II.

I.

Introduction

Photosystem II (PS II) is one of the two photosystems existing in all oxygenic photosynthetic organisms, and performs a series of light-induced electron transfer reactions, leading finally to the splitting of water and evolution of molecular oxygen (Wydrzynski and Satoh 2005; Blankenship 2014). In fact, PS II is the first photosystem where electrons are extracted from water and transferred to photosystem I (PS I) through the intermediate cytochrome b6f complex and a mobile electron carrier plastocyanin or cytochrome c6, and the electrons are finally used to reduce nicotinamide adenine dinucleotide phosphate. The central part of PS II is designated PS II core, and is composed of around 17–19 membrane-spanning subunits located in the thylakoid membrane and around 3–5 membrane-extrinsic proteins located in the lumenal surface of the membrane. The Abbreviations: Bcr  – β-carotene; Chl  – Chlorophyll; cryo-EM  – cryo-­ Electron Microscopy; cyt  – cytochrome; DGDG  – Digalactosyldiacylglycerol; EPR  – Electron Paramagnetic Resonance; EXAFS  – X-ray spectroscopy including extended X-ray Absorption Fine Structure; LHCII  – Light-Harvesting Complex-II; LMW  – Low Molecular Weight; MGDG  – Monogalactosyldiacylglycerol; OEC – Oxygen-Evolving Complex; PG – Phosphatidylglycerol; Pheo – Pheophytin; PQ – Plastoquinone; PS II – Photosystem II; RC – Reaction Center; SQDG  – Sulfoquinovosyldiacylglycerol; SFX  – Serial Femtosecond X-ray crystallography; TMH – TransMembrane Helices; XFEL – X-ray Free Electron Lasers

membrane-spanning PS II core subunits are involved in binding of the electron transfer cofactors, intrinsic antenna pigments and/or stabilization of the whole complex, whereas the extrinsic proteins are required to stabilize the catalytic center for water-splitting, the oxygen-evolving complex (OEC). Within the thylakoid membrane, the PS II core is surrounded by a large number of light-harvesting complex-­II (LHCII) proteins, which serve to harvest light energy and transfer them to the PS II core. LHCII are membranespanning subunits in most eukaryotic algae and higher plants (Chap. 6), but in cyanobacteria and red algae, membrane-extrinsic, hydrophilic proteins called phycobilisomes are attached at the stromal surface of the membrane and serve to harvest the light energy. The components of the PS II core are rather conserved throughout the oxygenic photosynthetic organisms from prokaryotic cyanobacteria to higher plants, but the components and organization of their light-harvesting antennas vary significantly depending on the species of the organisms, which is important for the adaptation of various organisms to various different light environments (Chap. 6). However, recent structural analyses also revealed some variations in the components and structure of the PS II core, which will be described in this chapter. The PS II core is sometimes simplified as PS II. Structural analysis of the PS II core first started with samples purified from thermo-

1  Structure, Electron Transfer Chain of Photosystem II and the Mechanism… philic cyanobacteria Thermosynechococcus elongatus (Zouni et al. 2001; Ferreira et al. 2004) and Thermosynechococcus vulcanus (Kamiya and Shen 2003). These studies were performed by X-ray crystallography and provided the arrangement of most protein subunits and pigments, electron transfer cofactors within the PS II core. However, the maximum resolution achieved with these studies are at 2.9–3.0 Å resolution until 2009 (Loll et al. 2005; Guskov et al. 2009), which was not enough to solve the catalytic center for water-­splitting. This catalytic center is a Mn-cluster composed by 4 Mn ions and 1 Ca ion connected by several oxy-bridged oxygens. As will be seen in this chapter, since the shortest Mn-Mn distances are around 2.7  Å and the Mn-O distances are around 1.8–2.0 Å within this cluster, roughly saying, a resolution of 2.0  Å or higher will be required to distinguish the individual atoms within this cluster from the experimental electron density map. This was achieved by a structural analysis of the PS II core from T. vulcanus at a resolution of 1.9  Å (Umena et  al. 2011), which provided the complete structure of the whole PS II core as well as the detailed structure of the Mn4CaO5cluster, the catalytic center for water  splitting (Umena et  al. 2011; Suga et  al. 2015; Shen 2015). Since the PS II core is a huge membrane-­ protein complex, crystallization of it proved difficult except that from the thermophilic cyanobacteria. Only the PS II core from a moderately thermophilic red alga Cyanidium caldarium was later crystallized (Adachi et al. 2009) and its structure solved by X-ray crystallography (Ago et al. 2016). Recently, owing to the rapid development of single particle structural analysis by cryo-electron microscopy (cryo-EM), the structures of PS II core in complex with its light-harvesting antennas have been solved from various organisms including higher plants (Wei et  al. 2016; Su et  al. 2017), a green alga (Shen et al. 2019; Sheng et al. 2019) and a diatom (Nagao et  al. 2019; Pi et  al. 2019). These studies provided a wealth of informa-

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tion regarding structural organization of the PS II core and its variations in various organisms (Cao et al. 2020), enabling comparisons of the structure and function of PS II from different organisms possible. This chapter will describe the structure of the PS II core and its variations in different organisms, the electron transfer chain of PS II, and the mechanism of water splitting based on the recent structural and functional studies.

II. Structure of PS II As mentioned above, the overall structure of the PS II core is highly conserved from cyanobacteria to higher plants. However, recent studies also revealed some differences in the protein composition and number of pigments in PS II from different organisms. So far the structure of cyanobacterial PS II has been analyzed at the highest resolution, so we will describe structure of the cyanobacterial PS II in detail, and then the structures of PS II from other organisms in comparison with that of the cyanobacterial PS II. The differences in the protein components of PS II core from different organisms are listed in Table 1.1. A.

Structure of Cyanobacterial PS II

1. Organization and Structure of Protein Subunits

The overall structure of the cyanobacterial PS II analyzed at 1.9 Å resolution is shown in Fig.  1.1 (Umena et  al. 2011; Suga et  al. 2015). It exists in a dimeric form, and each monomer contains 20 subunits with a molecular mass of 350 kDa. Each PS II monomer contains 17 trans-membrane subunits and 3 membrane-extrinsic subunits associated at the lumenal side. The 17 trans-membrane subunits are D1, D2, CP47, CP43, PsbE, PsbF, PsbH, PsbI, PsbJ, PsbK, PsbL, PsbM, PsbTc, PsbX, PsbY, PsbZ, Psb30 (Ycf12). The genes for all these trans-­membrane sub-

J.-R. Shen et al.

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Table 1.1.  Psb protein subunits of PS II core complex from different groups of organisms

Gene psbA psbB psbC psbD psbE psbF ? psbH psbI psbJ psbK psbL psbM psbS psbTc psbW psbX psbY psbZ ycf 12 ? psbO psbP psbQ psbTn psbU psbV psb31

Subunit Mass (kDa) Membrane-spanning subunits D1 38.02 (S) CP47 56.28 (S) CP43 50.07 (S) D2 39.42 (S) 9.26 (S) α-cyt b559 β-cyt b559 4.41 (S) G protein ? H protein 7.70 (S) I protein 4.20 (S) J protein 4.10 (S) K protein 4.28 (S) L protein 4.37 (S) M protein 3.76 (S) S protein 29.20 (S) 3.85 (S) TC protein W protein 5.93 (S) X protein 4.23 (S) Y protein 3.67 (S) Z protein 6.54 (S) Psb30 4.14 (Sy) Psb34 ? Extrinsic proteins 33-kDa O 26.54 (S) protein 23-kDa P 20.21 (S) protein 16-kDa Q 16.52 (S) protein Tn protein 5.00 (A) U protein 10.49 (Sy) V protein 15.12 (Sy) Psb31 13.24 (Ch)

TMH 5 6 6 5 1 1 1 1 1 1 1 1 1 4 1 1 1 1 2 1 1

Cyanobacteria + + + + + + ─ + + + + + + ─ + ─ + + + + ─

Red algae + + + + + + + + + + + + + ─ + + + + + + +

Diatoms Green algae + + + + + + + + + + + + + ─ + + + + + + + + + + + + ? + + + + + + + + + + + + + + ─

Higher plants + + + + + + ─ + + + + + + + + + + + + ─ ─

0

+

+

+

+

+

0

±

±

─

+

+

0

±

±

±

+

+

0 0 0 0

─ + + ±

─ + + ±

─ + + +

± ─ ─ ─

+ ─ ─ ─

Proteins in the white background are present in all oxygenic photosynthetic organisms, and those in grey backgrounds are present only in some groups of the organisms. The molecular masses of the mature proteins are calculated from the amino acid sequences reported in the SwissProt database using the MacBioSpec program (Sciex Corp., Thornhill, ON, Canada) for spinach (S), Arabidopsis thaliana (A), Chaetoceros gracilis (Ch), and Synechocystis sp. PCC 6803 (Sy)

units are found in all oxygenic photosynthetic organisms from cyanobacteria to higher plants, and 16 subunits except Psb30 are found in the structures of PS II from all organisms (Table  1.1). Only Psb30 was not

found in the structure of higher plant PS II, and it is not known if this subunit binds loosely to higher plant PS II and was lost during isolation, or it is not a component of higher plant PS II.

Fig. 1.1.  Structure of cyanobacterial PS II. (a) Structure of the dimeric PS II with a view perpendicular to the membrane plane. (b) Structure of the dimeric PS II with a top view from the stromal side. (c) Distribution of water molecules over the PS II dimer and structure of the chlorophylls, β-carotenes, etc., in a PS II monomer. The structure was made with 3WU2 (Umena et al. 2011), and the protein subunits are indicated in the figure. In panel c, pigments and other cofactors are shown in one of the monomers. Chls and Bcrs are shown in green and yellow, respectively, and other cofactors were shown in different colors

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Trans-Membrane Subunits

Among the 17 trans-membrane subunits, D1, D2, CP47 and CP43 are large subunits encoded by psbA, psbD, psbB and psbC genes, respectively. The D1 and D2 subunits have molecular masses of 38  kDa and 39 kDa, respectively, and each of them consists of five trans-membrane helices (TMHs). They have a low sequence homology, but their overall structures are similar, and are arranged in a pseudo-C2 symmetry. These two subunits are located at the center of the PS II core (Fig. 1.1) and bind all of the electron transfer cofactors (see below); thus, they constitute the reaction center core of PS II.  The N-termini of both D1 and D2 are located at the stromal side, and their C-termini are located at the lumenal side. There are small hydrophilic regions exposed to the stromal surface, but larger regions are exposed to the lumenal surface which participate in binding and stabilization of the Mn4CaO5-cluster as well as formation of hydrogen-bond networks that may be related to the proton egress and substrate water inlet (see below). Each of the D1 and D2 subunits binds three chlorophylls (Chls), one pheophytin (Pheo), one plastoquinone (PQ). The three Chls are P680 (the reaction center Chl of PS II), Chlacc (the ‘accessory Chl’), and ChlZ, respectively, among which, P680 and Chlacc form a four-­Chls cluster of the PS II reaction center (RC) (see below). A unique feature of D1 is its extremely high turn-over rate, which could be as short as 30 min under high light illumination, and is considered a prerequisite for the photodamage and repair cycle (Aro et al. 1993; Li et al. 2018). Under high light illumination, D1 is preferentially photodamaged and cleaved by specific proteases, and the damaged D1 is removed from the PS II core complex. Then, newly synthesized D1 is inserted into the complex, resulting in the re-assembly and recovery of PS II from photodamage. This feature is unique for D1 and not found in D2 and any other PS II components.

J.-R. Shen et al. CP47 and CP43 have molecular masses of 56  kDa and 50  kDa, respectively, and each of them consists of six TMHs. They have almost no sequence homology, but the arrangement of their TMHs and some extramembrane loop regions are similar. Both Nand C-termini of the two subunits are located at the stromal side. Like D1 and D2, small regions of these subunits are exposed to the stromal surface, but larger regions are exposed to the lumenal surface. Especially, the E-loop exposed to the lumenal surface between helix V and IV of both CP47 and CP43 has 150–200 residues, forming a large hydrophilic area in the lumenal side. These E-loops provide part of the cover for the Mn4CaO5-cluster to prevent it from free access of solute molecules in the lumenal space, and function to stabilize binding of the Mn4CaO5-cluster and three extrinsic subunits, as well as the proton egress and water inlet (Umena et al. 2011; Shen 2015; Suga et al. 2015). Residues in these E-loops interacts extensively with the three extrinsic subunits and participate in hydrogen-bond networks that are related with the proton egress and water inlet, and one of the residues (CP43-E354) provides a ligand to the Mn4CaO5-cluster directly. In the transmembrane region, CP47 binds 16 Chls and CP43 binds 13 Chls, which function as the intrinsic light-­harvesting antenna system of PS II (see below). These Chls are largely ligated by His residues conserved among different organisms. CP47 and CP43 surround the two sides of D1/D2, with CP47 located at the D2 side and CP43 located at the D1 side. The binding of CP43 to the PS II core is weaker than that of CP47, hence CP43 is lost upon detergent solubilization of the thylakoid membranes in some cases, resulting in a CP47-RC sub-complex (Boehm et  al. 2011, 2012). This has been taken as evidence for the presence of an assembly intermediate, indicating that CP47 is first attached to the D1/D2 RC core during assembly, and CP43 attaches at a later stage.

1  Structure, Electron Transfer Chain of Photosystem II and the Mechanism… The remaining 13 trans-membrane subunits are collectively called low molecular weight (LMW) proteins with their molecular masses lower than 10 kDa, and most of them have molecular masses of around 5  kDa. Most of them have one TMH, except PsbZ which has two TMHs. These LMW subunits are distributed over the peripheral region of PS II, and play various roles such as binding and stabilization of some cofactors, stabilization of the dimeric form of PS II, stabilization of the binding of some other subunits within the PS II monomer, mediating interactions of the PS II core with LHCs, etc. However, the detailed functions of most LMW subunits are not clear. Among the 13 LMW subunits, PsbE and PsbF together binds cytochrome (cyt) b559, one of the two c-type cyts found in cyanobacterial PS II.  These two subunits are located at the periphery of D2 and found to be present in a PS II RC complex consisting of five subunits (D1, D2, PsbE, PsbF and PsbI) (Nanba and Satoh 1987), suggesting the tight association of these two subunits with the PS II RC core. The heme of cyt b559 is located close to the stromal surface of the membrane, and is coordinated by PsbE-­ His23 and PsbF-His24 (sequence numbering of T. vulcanus). PsbF has a molecular mass of 4  kDa, whereas PsbF has a molecular mass of 9  kDa; thus, PsbE is much longer than PsbF.  While PsbF has one TMH with almost no extra regions exposed to either side of the membrane, PsbE has a long C-terminal region exposed to the lumenal surface, which interacts with lumenal loop regions of D2. One of the roles of this cyt may be mediating cyclic (secondary) electron transfer around the PS II RC (see below) as a means to mitigate photodamage. The cyt is normally present in a high midpoint redox potential form with a redox potential of 370– 435  mV (Stewart and Brudvig 1998). However, upon some treatments of isolated PS II membranes or core complexes such as washing with high concentration salts, high or low pHs, the cyt changes to a low redox

9

potential  form with a midpoint potential of 0–80  mV (Stewart and Brudvig 1998; Mizusawa et al. 1999). The appearance of the low redox form seems to be correlated with the inactivation of oxygen evolution; however, their exact relationships remain to be established. In cyanobacterial cells, removal of the heme by changing its coordinating ligands does not prevent growth of cyanobacterial cells, but presence of the heme was shown to be required to maintain the maximum activity of PS II (Sugiura et al. 2015). However, deletion of either the psbE or psbF genes prevent assembly of the PS II complex (Shinopoulos and Brudvig 2012), resulting in a mutant unable to grow photoautotrophically. This indicates an important role of these subunits in maintaining the stable assembly of PS II. PsbI and PsbX are two subunits located at the periphery of D1/CP43 and D2/CP47, respectively, and they are related by the pseudo-C2 symmetry that relates D1/D2 (Fig. 1.1). PsbI is also one of the components of the isolated PS II RC consisting of five subunits (Ikeuchi and Inoue 1988), ­suggesting its tight association with D1 and function as an early assembly component of PS II.  It is located outside of ChlZD1 and therefore may stabilize the binding of the ChlZD1. Deletion of the psbI gene from either cyanobacteria (Ikeuchi et al. 1995; Kawakami et  al. 2011) or green algae (Künstner et  al. 1995) did not prevent photoautotrophic growth of the cells but destabilized the dimeric form of PS II, resulting in the increase of PS II monomer. On the other hand, PsbX does not bind to the PS II RC as tight as that of PsbI, and is lost in the isolated PS II RC. It is located close to PsbH and outside of ChlZD2, and the phytol tail of ChlZD2 is inserted into the middle space between the TMHs of PsbH and PsbX (Umena et  al. 2011). Therefore, both PsbX and PsbH may help stabilizing binding of the ChlZD2. Deletion of the psbX gene in cyanobacteria did not inhibit PS II assembly and photoautotrophic growth (Funk 2000; Katoh and

10

Ikeuchi 2001). However, functional studies suggested that PsbX may play a role in the electron transfer functions between QA and QB. This can be explained by the presence of a sulfoquinovosyl-­ diacylglycerol (SQDG) molecule in a position close to PsbX (Umena et  al. 2011). The opposite side of this lipid molecule is close to QB; thus, removal of PsbX may affect the binding of the SQDG molecule, thereby affecting the stability and property of QB, leading to alterations in the electron transfer between QA and QB (Nakajima et al. 2018). Furthermore, the end of the isoprenoid tail of QB is close to the TMH of PsbX.  Removal of PsbX may also affect the binding of QB directly, resulting in the effect on the electron transfer between QA and QB. PsbH has a molecular weight of 6 ~ 7 kDa and is located in a peripheral region between CP47 and D2 and close to PsbX (Fig. 1.1). The PsbH protein has a long N-terminal region consisting of 30 ~ 40 amino acids at the stromal surface, which is composed of a short helix at its N-terminus connected to the main TMH by a loop. This N-terminal region covers the stromal surface of D2, and forms extensive interactions with the stromal regions of D2. Thus, this N-terminal region may be required for the stable binding of PsbH and/or stabilization of the stromal regions of D2. In higher plants and green algae, there is a conserved Thr residue which undergoes reversible phosphorylation (Summer et al. 1997). The functional significance of the phosphorylation of PsbH is unclear. In cyanobacteria, the N-terminus is shorter and truncated, thus, it does not have the Thr residue and the cyanobacterial PsbH is not phosphorylated. Deletion of the psbH gene from cyanobacteria does not prevent the assembly of PS II and photoautotrophic growth, but destabilizes the PS II complex and prevents association of PsbX with PS II (Komenda et al. 2002; Iwai et al. 2006). The deletion mutant also became more sensitive to photoinhibition than the wild type, which seemed to be due to the partial inhibition of

J.-R. Shen et al. the D1 protein repair process rather than to an increase in the photochemical damage. In contrast, deletion of the psbH gene in the green alga Chlamydomonas reinhardtii prevented the assembly of PS II (O’Connor et al. 1998). PsbJ and PsbK are located at the peripheral region between D2 and CP43, with PsbJ close to D2 and Psb E/F, and PsbK close to CP43 and PsbZ (Fig.  1.1). PsbJ was suggested to be involved in a channel for the entry of quinones to the QB binding site within PS II (Guskov et al. 2009); however, this channel is not confirmed in the high resolution structure of PS II (Umena et al. 2011; Suga et  al. 2015). Deletion of psbJ in the cyanobacterium Synechocystis sp. PCC 6803 leads to diminished oxygen evolution rates of PS II (Lind et al. 1993; Regel et al. 2001) and a longer lifetime of reduced QA (Regel et  al. 2001; Ohad et  al. 2004), suggesting that PsbJ is involved in the regulation of forward electron transfer from QA to QB. The structural basis for this effect is not clear. PsbJ is neither close to the QA nor to QB, but there are a few hydrophobic molecules between PsbJ and QB, which include a monogalactosyldiacylglycerol (MGDG), a digalactosyldiacylglycerol (DGDG), and a β-carotene (Bcr) molecule (Umena et  al. 2011). Deletion of PsbJ may affect the binding of these hydrophobic molecules, thereby affecting the binding and stability of QB. In higher plants, deletion of psbJ results in loss of photoautotrophic growth, high light sensitivity and defects in the assembly of the extrinsic proteins PsbP and PsbQ (Regel et al. 2001; Hager et al. 2002; Swiatek et al. 2003; Ohad et al. 2004; Suorsa et al. 2004). In contrast to PsbJ, deletion of the psbK gene in cyanobacteria has very little effect on photoautotrophic growth and PS II activity (Ikeuchi et  al. 1991). However, deletion of the gene in a green alga destabilized the PS II complex, resulting in a deletion mutant unable to grow photoautotrophically (Takahashi et al. 1994).

1  Structure, Electron Transfer Chain of Photosystem II and the Mechanism… PsbY and Psb30 (Ycf12) are located outside of the PsbE and PsbK subunits, respectively (Fig.  1.1). Both of them are loosely associated with the PS II core, resulting in their easy loss during purification and/or crystallization. In fact, Psb30 is the last trans-membrane subunit that was identified in the isolated PS II core (Kashino et  al. 2007), and PsbY was absent in some of the crystal structures of PS II (Ferreria et  al. 2004; Umena et al. 2011; Suga et al. 2015). Psb30 has been found in the structures of PS II from cyanobacteria (Umena et  al. 2011), red algae (Ago et al. 2016), diatom (Nagao et al. 2019; Pi et al. 2019), and green algae (Shen et  al. 2019; Sheng et  al. 2019), but absent in the structure of higher plant PS II (Wei et al. 2016; Su et al. 2017), although a homologous gene is found in the genome of green plants. Deletion of the psbY gene in cyanobacteria does not inhibit the growth and water oxidation process significantly (Meetam et al. 1999; Kawakami et al. 2007), but this protein may be important for the redox control of cyt b559 in higher plants (von Sydow et al. 2016). On the other hand, deletion of the ycf12 gene in cyanobacteria resulted in little effect on the oxygen-evolving activity at normal light intensities (InoueKashino et al. 2008; Takasaka et al. 2010). PsbZ is the only LMW subunit that has two transmembrane helices. It is located at the peripheral region outside of PsbC (CP43), and in close contact with PsbK/Psb30 (Fig. 1.1). This protein probably functions to stabilize the binding of pigments, since there are Bcr molecules bound in a space outside of PsbK, and one Bcr and one Chl a molecule bound at a peripheral region of CP43; these pigments are covered by PsbZ. Deletion of the psbZ gene in cyanobacteria leads to a decrease in oxygen evolution and de-stabilization of the PS II core (Takasaka et  al. 2010), whereas the same deletion destabilizes the binding of LHCII to the PS II core in higher plants (Swiatek et  al. 2001). The latter suggests that PsbZ has been evolved to interact with LHCII in the green lineage

11

organisms, which is consistent with its role in stabilizing the binding of pigments associated with PsbK and CP43. Finally, there are three LMW subunits located in the monomer-monomer interface within the dimer; these are PsbL, PsbM and PsbTc (Fig. 1.1). This location suggests that these three subunits are involved in the stabilization of the PS II dimer. Functional studies, however, revealed different roles of the three subunits in stabilization of the PS II dimer and functioning of PS II. Inactivation of the psbL gene in both a cyanobacterium and tobacco severely retarded oxygen evolution and the mutant is unable to grow photoautotrophically (Anbudurai and Pakrasi 1993; Swiatek 2003; Suorsa et al. 2004). It has also been reported that PsbL is required for normal functioning of QA based on studies with isolated PS II complexes (Hoshida et  al. 1997). PsbL has a long N-terminal, hydrophilic loop (~15 residues) extending to the stromal surface of CP47 from the adjacent monomer. Mutagenesis studies of this N-terminal region suggested that  it influences forward electron transfer from QA through indirect interactions with the D–E loop of the D1 reaction center protein (Luo et al. 2014), although the N-terminal region of PsbL does not interact with D-E loop of D1 nor QA directly. In contrast to PsbL, deletion of the psbM gene in cyanobacteria results in slight decrease of the PS II dimer content and increase of monomer, but does not inhibit growth of the mutant (Bentley et  al. 2008; Kawakami et  al. 2011). PS II dimer is still formed in the PsbM-deletion mutant, and it is monomerized completely when psbTc, a chloroplast-encoded gene, is also deleted. Deletion of the psbTc gene results in monomerization of PS II to a larger extent than that caused by deletion of PsbM (Bentley et  al. 2008). A PsbTc-deletion mutant of Chlamydomonas grows photoautotrophically, whereas its growth is significantly impaired under strong light illumination, suggesting that it is required for preventing

12

PS II from photodamage under high light conditions (Onishi and Takahashi 2001). Extrinsic Subunits

Three extrinsic proteins PsbO, PsbU, PsbV are associated at the lumenal side of PS II and required for the maximum activity of oxygen evolution (Enami et  al. 2008). Among these three extrinsic proteins, PsbO is the largest one and has a molecular weight of ~26  kDa with a length of 241– 247 residues. In SDS polyacrylamide gel electrophoresis, this protein has an apparent molecular weight of 33  kDa, so it is also called 33-kDa protein. Since it functions to stabilize the Mn4CaO5-cluster, it is also known as the manganese-stabilizing protein. PsbO is present in PS II of all photosynthetic organisms (Enami et al. 2008), and plays the most important role among all of the extrinsic proteins. The structure of PsbO is mainly composed of eight antiparallel β-strands arranged in a cylindric form, with a small hollow in its middle with a diameter of ~4.0  Å (Umena et  al. 2011). Whether there is a role for this hollow is unknown. Large loops are found to join the β-strands between 1 and 2, 3 and 4, and between 5 and 6, and these loops provide residues interacting with D1, D2, CP43, and CP47 at the lumenal hydrophilic region. PsbO is located mainly beneath D1 at the lumenal side, but a small region of it also extends to the lumenal surface of D2. A large loop between β-strands 1 and 2 even extends to an area close to the adjacent monomer, and interacts with some lumenal residues of CP47 from the adjacent monomer (Fig.  1.1), suggesting that PsbO may also function to stabilize the PS II dimer. PsbO can be released from the thylakoid membrane by high concentrations of divalent cations or acidic or alkaline conditions, and plays an important role in stabilizing the Mn4CaO5-cluster and maintaining an optimal ion environment (Ca2+ and Cl−) for water oxidation to pro-

J.-R. Shen et al. ceed (Enami et  al. 2008; Bricker et  al. 2012). Some of the PsbO residues participate in hydrogen-bond networks connecting the Mn4CaO5-cluster with bulk solution of the lumen, which may function for the water inlet and/or proton egress (Umena et  al. 2011; Shen 2015). Deletion of the psbO gene in cyanobacteria reduces the oxygen-evolving activity to lower than half, and slows the growth, but does not inhibit water oxidation completely unless the psbV gene is also deleted (Burnap and Sherman, 1991; Philbrick et al. 1991; Shen et al. 1995a). However, the equivalent psbO deletion mutant in green alga and higher plants inhibits the assembly of a functional PS II core complex and abolishes the oxygen-evolving activity completely, resulting in the loss of photoautotrophically growth (Mayfield et al. 1987; Yi et al. 2005). The other two extrinsic proteins are PsbU and PsbV, with molecular weight of ~11 kDa and ~15 kDa, respectively. These two extrinsic proteins are also found in PS II of red algae and diatoms, but are absent in green algae and higher plants. PsbU is an all α-protein, and is located at the most outside surface at the lumenal side like a cap covering the regions surrounded by PsbO, PsbV, and the lumenal loops of CP47 and CP43. Its full binding to PS II requires PsbO and PsbV (Shen and Inoue, 1993; Enami et al., 2008). Deletion of the psbU gene in cyanobacteria results in a very slight reduction ( 4.1 was the ground state for Mn4O4(OH)2 with the 3343 valence structure (10bIL). Therefore, the energy levels (11a)− and the Ca(II)Mn4 O4(OH)2 with the obtained by the exact diagonalization are 3433 valence structure (11b)−. Their full compatible with the observed three-­ geometry optimizations by QM(DFT) /MM different EPR spectra. The EPR results for methods provided the R-opened structure for the IR irradiation (Boussac et  al. 1996; (11aR)− with the (3343) valence state and Boussac 2019) may be explained with the S  =  5/2 ground spin state, the L-opened sequential steps. structure for (11aL)− with the the (3343) valence state and S = 1/2 ground spin state

2  Mechanism of Water Oxidation in Photosynthesis Elucidated by Interplay… and the R-opened structure for (11bR)− with the (3433) valence state and the S  =  7/2 ground spin state. Therefore, assignments of the ground spin states for R- and L-isomers of (11a)− by the LOS model were just reverse as compared with the HOS model for (10a)+ (Yamaguchi et  al. 2016). The L-isomer (10bL)+ with S = 7/2 by the HOS model was corresponding to the R-isomer (11bR)− with S = 7/2 by the LOS model. The XFEL experiments for the S2 state for cyanobacteria (Kern et  al. 2018, Suga et  al. 2019) revealed the R-opened structure, (10aR)+ and/or (10bR). The EPR spectra for the S2 state (Cox et al. 2014) was consistent with the R-opened S = 1/2 and g = 2 structure for cyanobacteria. Thus, the computational results based on the HOS model are consistent with available experimental results for cyanobacteria obtained by EPR.  Table  2.1 summarizes the hyperfine constants obtained by EPR and the computational results by the HOS models. Our computational results (Yamaguchi et al. 2016) are also compatible with available EPR results at the low temperature. The (10aR)+ with the (4443) valence structure in the S2 state arises from the 9aR (3443) in the S1 state. The deprotonation of W2 was conceivable for (10aR)+, providing (10bR… Table 2.1.  The hyperfine constants (Aiso) (MHz) for 55 Mn ions in the S2 state of OEC of PS II Methods

Mn(IV)4(a) Mn(IV)3(b) Mn(IV)2(c) Mn(III)1(d)

EPRa EPRb EPRc QMd QM/ MMe QMf

−244 −255 −251 −264 −223

217 238 208 200 185

200 191 191 187 204

−297 −324 −312 −329 −374

−292 −303

201

195

204

193

−208 −219

QMf

From Hasegawa et al. (1999) From Charlot et al. (2005) c From Cox et al. (2011) d From Yamaguchi et al. (2016), adiabatic + ZPE e From Shoji et al. (2019a, b, c) f From Ames et al. (2011), 1d2′ f From Ames et al. (2011), 1d2″ a

b

53

H+B) at the room temperature, where B denotes the base as proton acceptor, for example the Asp61 anion (Yamaguchi et al. 2013), proton storage path (path III) forming the so-called Eigen complex (O5H11)+(Shoji et al. 2019a; Suga et al. 2019). The (10aL)+ with the (4443) valence structure in the S2 state arises from the 9bR (3443) in the S1 state in conjugation with proton shift from O(5)H to W2=OH− to afford W2=H2O.  The deprotonation of W2 was also conceivable for (10aL)+, providing (10bR…H+B). The examination of hydrogen bonding networks around the CaMn4O5 cluster is crucial for further discussions (Shoji et al. 2015a).

 . System Structures of OEC V of PS II A.

Proton Release and Water Inlet Pathways

The FTIR results by Noguchi (2008) and Noguchi et  al. (2012) have indicated the water insertion in the S2 to S3 transition. Before discussion of the S3 intermediate, we must investigate system structures of OEC of PS II revealed by HR XRD since one extra water molecule is inserted through the water inlet pathway (WIP). The HR XRD experiment (Umena et  al. 2011) elucidated that OEC of PS II has a dimer structure, where each monomer contains about 20 protein subunits that have 77 cofactors including more than 20 lipids, 35 chlorophyll a, 11 β-carotene, 2 plastoquinone, 2 pheophytin, CaMn4O5 complex, 2-heme Fe counters, 1 nonheme Fe center, 1 hydrogen carbonate, 2 chloride anions and over 1000 water molecules. It elucidated the 3D structure of the OEC of PS II catalyzing the water-­splitting reaction by sunlight. Indeed, HR XRD have revealed the biomolecular system structures consisted of hydrogen bonding networks, which are responsible for WIP and proton release (PRP) pathways for water oxidation in OEC of PS II.

54

Nowadays, quantum mechanics (QM)/ molecular mechanics (MM) methods (Karplus 2006; Warshell and Levitt 1976; Levitt 2001) are applicable for theoretical investigation of complex biological systems such as OEC of PS II (Shoji et al. 2013; Chen et al. 2018). In order to construct theoretical system models for OEC, we have performed theoretical calculations starting from the 3D heavy atom structures revealed by HR XRD. The hydrogen atoms are added to construct the hydrogen-bonding networks in the QM/MM geometry optimizations of the networks. Our large-scale QM/MM calculations (Shoji et al. 2013, 2015a, b) have indeed elucidated the network structures, hydrogen bonding O…O(N) and O-H distances and O(N)-H…O angles, together with Cl-O(N) and CL-H distances and O(N)-H---Cl angles. The obtained hydrogen bonding networks by QM/MM are fully consistent with the proposed WIP and PRP by HR XRD and available amino-acide mutation experiments (Debus 1992). Here, some of the QM/MM results are re-visited in relation to water oxidation in OEC of PS II. B. H-Bond Networks for Tyr161 and Ca Ion

Figure 2.5 illustrates the optimized hydrogen bonding networks around Tyr161 and Ca(II) ion. Tyr161 and His190 play significant roles for electron and proton transfers. His190 is connected with Asn298 and other amino acids. The O…N distance between the oxygen atom of Tyr161 and the nitrogen atom of His190 by HR XRD was found to be 2.46 Å, in consistent with the optimized value (2.59 Å) by QM/MM. The optimized N…H distance and O-H…N angle by QM/MM were, respectively, 1.53 Å and 170°, indicating the strong hydrogen bond (H-bond) between Tyr161 and His190. Water molecule W(7) exists in the nearest position to Tyr161, indicating the short O…O(7) distance (2.62 Å) between the oxygen anion of the OH group of Tyr161 and W(7), in consistent with the

K. Yamaguchi et al. optimized value (2.65 Å) by QM/MM. The optimized O…H distance and O(7)-H…O angle were, respectively, 1.66  Å and 171°, indicating the strong H-bond between Tyr161 and W(7). W(7) further forms strong H-bonds with water molecules W(6) and W(3), and the oxygen anion of carboxyl group of Glu189. The optimized O(7)…H distance and O(W3)H…O(7) angle between W(7) and W(3) by QM/ MM were, respectively, 2.06  Å and 173°. The tetrahedral structure around W(7) was formed by the hydrogen bonding, indicating the octet rule (Saito et al. 2012). The OH group of Tyr161 forms a hydrogen bond with water molecule W(4) that is coordinated to the Ca(II) ion. The optimized O…O(4) and O…H distances and O(4)-H…O angle by QM/MM were, respectively, 2.81 and 1.83 (Å) and 179°. This implies that the OH group of Tyr161 is geometrically fixed with strong H-bond between lone pair of OH of Tyr161 and HO of W(4). This H-bond structure plays an important role for suppression of radical reaction by harmful Tyr-­O• radical formed in the electron transfer step from Tyr-OH to P680+ cation radical. The water molecule W(6) forms hydrogen bond with the water molecule W(5) and the oxygen anion of carbonyl group of the backbone C=O group of Phe182. W(5) forms three other hydrogen bonds with W(3), water molecule W(2) and backbone C=O group of Asp170. W(2) is linked with the proton release pathway (PRP) (see below). The optimized geometrical parameters for these hydrogenbonding networks were consistent with those of HR XRD (Umena et al. 2011). Thus, QM/ MM were applied for elucidation of the hydrogen-bonding network; W(2)-W(5)-W(6)W(7)-Tyr161-His190 where W(7) and Tyr161 are, respectively, linked with W(3) and W(4) which is in turn linked with the water inlet pathway (WIP) (see later). C. Water Inlet Pathway (WIP)

Figure 2.6 illustrates the optimized water inlet pathway (WIP) by HR XRD and the

2  Mechanism of Water Oxidation in Photosynthesis Elucidated by Interplay…

55

Fig. 2.5.  The observed heavy atom distances for the hydrogen bonding networks around the Tyr161 and the Ca(II) ion by the HR XRD, where the corresponding optimized values by the QM/MM method are given in parentheses. W3 and W4 are coordinated to the Ca(II) ion as described by yellow vectors. The HR XRD (Umena et al. 2011), SFX XFEL (Suga et al. 2015) and QM/MM (Shoji et al. 2015a, b) studies elucidated the hydrogen bonding network, (WIP)-W4-Tyr-OH-­W7-W6-W5-W2-W9-W8-Asp61-(PRP) connecting the water inlet pathway (WIP) with the proton release pathway (PRP). The loops of W7-W3-W5-W6 and W3-W5-W2-O(5) play important roles for proton transfers relating to water oxidations catalyzed by the CaMn4O5 cluster

large-scale QM/MM method. The optimized O(1)-O(10) distance was 2. 77 Å in compatible with the short (2.67  Å) length by HR XRD.  The optimized O(1)…H distance and O(10)-H…O(1) angle were 1.95  Å and 156°, respectively. From Fig. 2.5, the water molecule W(10) forms three hydrogen bonds with water molecules W(20), W(21) and W(4) coordinated to the Ca(II) ion. The optimized O(10)…O(20) and O(10)…H distances and O(10)…H-O(20) angle between W(10) and W(20) were, respectively, 3.00, 2.05  Å and 164°. The optimized O(10)…O(21) and O(10)…H distances and O(10)…H-O(21) angle between W(10)

and W(21) were, respectively, 2.88 and 1.92 Å and 167°. The optimized O(10)…O(w4) and O(10)…H distances and O(10)…H-O(w4) angle between W(10) and W(w4) were, respectively, 2.88 and 2.07 Å and 147°. These hydrogen bonds act effectively to fix the water molecule W(10), where a tetrahedral structure is found to satisfy the octet rule. Therefore, oxygen dianion O(1) site of the CaMn4O5 core is protected by a strong hydrogen bond with W(10), that is also fixed with the hydrogen bond network. Water molecules W(20) and W(21) are connected with the water inlet channel revealed by HR XRD. This indicates their

56

K. Yamaguchi et al.

Fig. 2.6.  The observed heavy atom distances for the hydrogen bonding networks for water-inlet pathway (WIP) by the HR XRD, where the corresponding optimized values by the QM/MM method are given in parentheses. The HR XRD (Umena et al. 2011), SFX XFEL (Suga et al. 2015) and QM/MM (Shoji et al. 2015a, b) studies elucidated the WIP, which is channel A(C)-W20-W10-W21-W4-W3

important roles for supply of substrates waters for water oxidation. Thus, interplay between HR XRD and QM/MM elucidated the molecular system structure of the hydrogen-binding networks for water-inlet pathway (WIP). D. Proton Release Pathway (PRP)

Figure 2.7 illustrates the optimized proton release pathway (PRP). The water molecule W(1) coordinated to Mn4(a) forms a hydrogen bond (H-bond) with one of the oxygen atoms of the COO anion group of Asp61. The optimized H-bond para-meters eluci-

dated a very strong H-bond between W(1) and the COO anion group, fixing Asp61. The water molecule W(9) forms H-bond with the water molecule W(2) coordinated to the Mn4(a) ion, water molecule W(8) and carbonyl group of Asp181. The water molecule W(8) forms the H-bond with the other oxygen atom of the COO anion group of Asp61. The optimized H-bond parameters by QM/MM were consistent with these strong H-bond networks, also elucidating that Lys317 is strongly connected with W(8). Asp61 and Asp65 play significant roles for proton release. The water molecule W(12)

2  Mechanism of Water Oxidation in Photosynthesis Elucidated by Interplay…

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Fig. 2.7.  The observed heavy atom distances for the hydrogen bonding networks for proton-release pathway (PRP) by the HR XRD, where the corresponding optimized values by the QM/MM method are given in parentheses. The HR XRD (Umena et al. 2011), SFX XFEL (Suga et al. 2015) and QM/MM (Shoji et al. 2015a, b) studies elucidated the PRP, which is W2-W9-W8-Asp61-W12-W13 (W14)-Channel D

forms H-bonds with the other oxygen atom of the COO anion group of Asp61 and water molecule W(14). Similarly, water molecule W(13) forms hydrogen bonds with W(12) and W(14). Moreover, W(13) and W(14) form H-bond with Asp65. W(14) further forms H-bond with Glu312, Asp61 and Asp65 which are parts of proton release pathway (PRP) from W(2) to Glu312 connected with Channel D revealed by HR XRD. The optimized H-bonds parameters by QM/MM supported these H-bonds networks for PRP revealed by HR XRD (Umena et al. 2011). From Fig.  2.7, the chloride anion Cl(1) forms four H-bonds with water molecule W(15), backbone NH group of Glu333, NH3+

group of Lys317, and NH2 group of Asn181. The optimized H-bond parameters by QM/ MM indeed elucidated that Cl(1) plays significant roles for the formation of H-bond network involving Glu333, Asn181 and W(15), contributing to stabilization of the NH3+ group of Lys317. Thus, interplay between HR XRD and QM/MM elucidated the molecular system structure of the H-bonds network for the PRP. E. QM/MM Calculations of Other Networks

Full geometry optimizations by the large-­ scale QM/MM method have been per-

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formed for the H-bonds with dioxygen anions of the CaMn4O5 cluster in OEC of PS II: (a) H-bond network of O(4) with water molecules (W(11)-W(16)-W(17)…) for proton egress pathway (Path III) as illustrated in Fig.  2.8; (b) H-bond between O(3) and His337; (c) H-bond between O(2) and Arg357; (d) hydrogen bond network around

K. Yamaguchi et al. the chloride anion Cl(2); and (e) hydrophobic reaction field for O(5) with Val185 (see later). Recent SFX XFEL ­ experiments (Kern et al. 2018; Suga et al. 2019) revealed dynamical motion of W16 for Path III as illustrated in Fig.  2.8. The detailed results are given in other articles (Shoji et al. 2013, 2015a, 2019a, b, c).

Fig. 2.8.  The observed heavy atom distances for the hydrogen bonding networks for proton-storage pathway (PSP) by the HR XRD, where the corresponding optimized values by the QM/MM method are given in parentheses. The HR XRD (Umena et al. 2011), SFX XFEL (Suga et al. 2017, 2019) and QM/MM (Shoji et al. 2015a, b) studies elucidated the PRP: W2-W9-W8-Asp61-W11-W16-W17-W18-Eugen Complex. The SFX XFEL (Kern et al. 2018; Suga et al. 2019) elucidated dynamicl behavior of W16 in the S2 and S3 states

2  Mechanism of Water Oxidation in Photosynthesis Elucidated by Interplay… VI. Possible Intermediates in the S3 State

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inserted in this region for proton transfer from H2O(I) inserted at the Mn1(d) site to the base (W2  =  OH−) (Siegbahn 2013b). A. Water Insertion in the S3 State Therefore, Shoji et al. (2015b) examined a new insertion path of W(3) into Mn4(a) or The EXAFS experiments (Yano and Mn1(d) site by the QM/MM model, elucidatYachandra 2014) indicated large cluster ing two favorable reaction pathways for changes in the S2 to S3 transition. As men- water insertion in the S3 state. The O(5) site tioned above, Noguchi et  al. (2012) per- plays an important role for proton transfer formed the FTIR spectroscopy, also revealing (shuttle) for their L- and R-reaction paththe possibility of insertion of an extra water ways to generate the R-opened hydroxide molecule in the S2 to S3 transition of OEC of (12hoR) instead of the putative water molePS II. Isobe et al. (2012) examined the extra cule for proton transfer in front of the O(5) water insertion H2O(I) (I: inserted water) at site (Siegbahn 2013b). The HR XRD structhe S2 state by the QM model, indicating the ture provided definitive information for difficulty of water insertion because of the theoretical modeling of the catalytic site of repulsive interaction, and therefore they elu- OEC of PS II. One of the reaction pathways elucidated by cidated the necessity of one more oxidation of Mn(III) ion to Mn(IV) in the S3 state for Shoji et al. (2015b) is that W(3) is inserted at water insertion. Mn1(d) site along with the L-type six steps The proton release from W1(=H2O) of (ST(0)→ ST(5)) (L-reaction pathway). The 10bR(L) may occur before water insertion in other is that W(3) is also inserted at the Mn1(d) the S2 to S3 transition, providing the no water site along with the R-type five steps (ST(0)→ inserted cluster Ca(II) ST(−4)) (R-reaction pathway). The L path­ Mn(IV)4O5(OH)2(H2O)2 (W1=W2=OH−) way involves an extra step, namely the inter(12). The water insertion (H2O(I)) for 12 pro- nal conversion from the R-opened to vides the water inserted cluster Ca(II) L-opened structure before the W(3) insertion. Mn(IV)4 O5(OH)(O(I)H)(H2O)3 cluster (12ho) Therefore, ST(4(5)) and ST(−3(−4)) correwith the (4444) valence state (ho=hydroxide) spond to the same steps. The highest activain the S3 state. The left-­opened form of 12ho tion barrier was calculated to be 15.4  kcal/ with W2=OH− and HO(I) at the Mn4(a) site mole for the proton transfer step from (12hoL) was found to be more stable than the Mn4(a)-OH to O(5)-Mn1(d) for the L-reaction right-opened form of 12 with W2=OH− and pathway to 12hoR. On the other hand, the calHO(I) at the Mn1(d) site (12hoR) at the level of culated highest activation barrier was the small QM (QM-­S) model for OEC of PS 18.2 kcal/mole for the proton transfer process from O(5)H to W2(=OH−) for the R-reaction II by HR XRD (Isobe et al. 2012). Larger QM and QM/MM models than pathway to provide 12hoR. Thus, the QM the small QM model were necessary for (QM)/MM calculations indicated that the further theoretical investigations of the L-reaction pathway was more favorable than water insertion processes in the S3 state the R-reaction pathway because the activation since the hydrogen bonding networks and barrier from R-opened to L-opened structure specific amino acid residues may play (namely initial step) in the L-reaction pathimportant roles for regulation of the inser- way was 13.3 ( −2 -> −2 -> TS-2 to -3a -> −3a) by the QM/MM method (Shoji et al. 2015a, b). The −1 state is the S2Tyr-O• stage (trapped valence 4443) in the early S3 state. The TS−1 to −2 state is the transition state with the mixed-valence (MV) (4,3.5,4,3.5) configuration for the W3 insertion. The −2 state is the S3 intermediate (O(5)=OH− and Mn1(d)-OH) with the (4344) valence configuration. The −1 state is the S2Tyr-O• stage (trapped valence 4443) in the early S3 state. The TS−2 to −3a state is the transition state with the (4344) configuration for proton shift from O(5)H to OH− at the W2 site coupled with the one electron transfer to Tyr-O•. The −3 state is the S4 intermediate (O(5)=O2−, Mn1(d)-OH W2=H2O) with the (4444) valence configuration followed by the deprotonation of W2

2019) proposed that the equilibrium model for the S3 state on the basis of these computational results are in consistent with the Renger proposal (2012). Shoji et  al. (2017) performed the large-­ scale QM (380 atoms)/MM calculations of the above mentioned possible intermediates in the S3 state. Their QM/MM calculations elucidated that the L-opened structure (12L) was more stable than the R-opened structure (12R) without water insertion. On the other hand, 11hoR was calculated to be more stable

than 12hoL after water insertion, indicating the reverse tendency. The optimized Mn4(a)Mn3(b), Mn3(b)-Mn2(c), Mn2(c)-Mn1(d), Mn3(d)Mn1(d) and Mn4(a)-Mn1(d) distances by QM/ MM were 3.16(3.26), 2.76(2.79), 2.71(2.76), 2.87(2.91) and 5.03(5.23) (Å), respectively, for 12L, where the corresponding values for 12hoL are given in parentheses. On the other hand, they are 2.78, 2.80, 2.75, 3.53 and 5.32 (Å), respectively, for 12hoR. Therefore, Mn4(a)Mn3(b) and Mn3(b)-Mn1(d) distances were different between 12hoR and 12hoL. Shoji et  al.

Fig. 2.11.  The optimized ten S3 intermediates by the large-scale QM(380 atoms)/MM methods. (a) R-opened Mn-hydroxide (12ohR with the O(5)-O(I)H hydrogen bond). (b) R-opened Mn-hydroxide (12ohR with the Glu189CO2-O(I)H hydrogen bond). (c) L-opened Mn-hydroxide (12ohR with the O(5)-O(I)H hydrogen bond). (d) R-opened Mn-oxo, oxo (12oxoR). (e) R-opened Mn-oxyl, oxo (12oxylR). (f) L-opened Mn-oxo, oxo (12oxoR). (g) R-opened Mn-peroxide (12peroxideR). (h) L-opened Mn-peroxide (12peroxideL). (i) R-opened Mn-superoxide (12superoxideL). (j) L-opened Mn-superperoxide (12superoxideL)

2  Mechanism of Water Oxidation in Photosynthesis Elucidated by Interplay… (2019a) further performed the full geometry optimizations of the nine possible S3 intermediates by the large-scale QM/MM. B. XFEL Results for the S3 State

Recently, the serial femtosecond crystallography (SFX) (Kern et al. 2013; Kupitz et al. 2014) by the use of pulsed femtosecond X-ray Free-Electron Laser (XFEL) was developed to elucidate the short-lived S2 and S3 structures for OEC of PS II. Young et al. (2016) conducted the SFX experiment by XFEL for elucidation of the S3 intermediate in the Kok cycle of OEC of PS II. The SFX XFEL structure at 2.25  Å resolution indicated no evidence of the water insertion in the S2 to S3 transition. The observed Mn4(a)Mn3(b), Mn3(b)-Mn2(c), Mn2(c)-Mn1(d), and Mn2(c)-Mn4(d) distances by their SFX were 2.8, 2.7, 2.8 and 3.1 (Å), respectively. Judging from these distances, the SFX structure is considered to be the R-opened structure. However, the large-scale QM/MM calculations (Shoji et al. 2017) predicted the greater stability of 12L than 12R for no water insertion structure in contradiction to the SFX XFEL structure. Therefore, Yamaguchi et al. (2017) in turn suggested that the SFX XFEL structure might correspond to 12hoR, although the inserted OH− was not confirmed because of the low resolution. Suga et  al. (2017) performed the SFX XFEL experiment at room temperature for elucidation of structural changes in the S2 to S3 transition. An isomorphous difference Fourier map between the two-flash and darkadapted states by their SFX XFEL experiment revealed a positive peak around O(5), suggesting the insertion of a new oxygen atom (O(I)) close to O(5), and indeed the observed O(I)-O(5) distance of 1.5 Å is responsible for the O-O bond formation. However, this short O(I)-O(5) distance may involve the experimental uncertainty of SFX XFEL at the 2.35 Å resolution. Therefore, Suga et al. (2019) indeed continued the refined SFX XFEL experiments, elucidating the elonga-

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tion of the O(I)-O(5) distance from 1.5 Å to 1.9 Å because of the increase of the resolution of SFX XFEL.  They have shown that their SFX XFEL structure is compatible with the right-­opened oxyl-oxo structure (12oxylR) obtained by the large-scale QM method (Isobe et al. 2016). Thus, Suga et al. (2017) first discovered the water insertion in the S2 to S3 transition in OEC of PS II by the timeresolved SFX XFEL method. Kern et  al. (2018) performed the SFX XFEL experiment and simultaneous X-ray emission spectroscopy (XES) at room temperature, visualizing metastable intermediates in the four states Si (i = 0–4) of the Kok cycle. They denied the previous conclusion (no insertion of the extra water molecule) at the 2.35  Å resolution (Young et  al. 2016), indicating the insertion of water molecule in the S2 to S3 transition. The O(I)-O(5) distances elucidated by their refined SFX at 2.04 ~ 2.08 Å resolution were about 2.0 and 2.2 Å, respectively, for A and B-monomers of OEC of PS II. These O(I)-O(5) distances are compatible with those of the oxyl-oxo intermediate (12oxylR) with the (3444) valence state and oxo-oxo intermediate (12oxoR) with (4444) valence state revealed by the largescale QM/MM, respectively (see Fig. 2.11). Very recently, Ibrahim et  al. (2020) refined the O(I)-O(5) distances to be 2.05 and 2.10 (Å) for A- and B-monomers, respectively. Therefore, the average O(I)-O(5) distance by the Berkeley group is now about 2.1 Å, indicating no O-O bond formation in the S3 state. The observed Mn4(a)-Mn3(b), Mn3(b)-Mn2(c), Mn2(c)-Mn1(d), Mn3(b)-Mn1(d) and Mn4(a)-Mn1(d) distances for A(B)- monomers by the SFX (Ibrahim et  al. 2020) were 2.67 (2.80), 2.84(2.77), 2.78(2.75), 3.37(3.44) and 4.96(5.14) (Å), respectively. Judging from the Mn4(a)-Mn3(b) and Mn3(b)-Mn1(d) distances, the observed intermediates by SFX (2020) are regarded as R-opened types in compatible with the optimized geometries by QM (Isobe et  al. 2016). Ibrahim et  al. proposed the direct insertion of W(3) coordinated to the Ca(II) ion into the Mn1(d) site in accord with

K. Yamaguchi et al.

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the R-opened pathway revealed by Shoji et al. (2015b). This may mean the formation of the R-opened hydroxide (12hoR) with the long O(6)-O(5) distance (2.4  ~  2.5 Å) in the early stage. The observed Mn4(a)-O(5) distances were 2.14 and 2.29 (Å) for A- and B-monomers in the S3 state by SFX (Ibrahim et  al. 2020). These are longer than the standard lengths (2.1 Å (Table 3.2). There were no additional (exchangeable) water derived oxy species bound to the WOC cluster in S3, compared to the S1 state. Finally, Suga et  al. (2017) reported a further 2F XFEL structure of PS II at 2.35 Å. Unlike that of Young et al. (2016), an apparent difference (from S1) was seen. Although the O5 and WOC metal centre positions are very

similar in the two S3 structures (Fig.  3.1, Table 3.2), an additional oxy group was seen by Suga et al. (2017) (called O6), ~1.5 Å distant from the O5 position. The O–O separation could suggest formation of a peroxyl species, or the like. This is, however, unlikely (see below) and we proceed here with the assumption that the mean Mn oxidation levels in the 2.25 Å and 2.35 Å structures are the same. All the current atomic resolution PS II XRD structures defy precise quantitative interpretation within conventional high paradigm WOC modelling (see sections “VII. Mechanism of oxygen evolution” and “VIII.  Conclusions”). Some key bond

3  On the Nature of the Functional S-States in the Oxygen Evolving Centre… lengths are always ‘too long’ (see below) to be consistent with the levels of high valent (generally IV) Mn required to be present. As noted above, for the first S1 structures at 1.9 Å and 2.1 Å resolutions, these included the Mn1, 2 and Mn2, 3 distances, as well as the Mn3, 4, Mn–O5 and Mn4–W1, W2 distances. This discrepancy was initially attributed to Mn reduction in the WOC having occurred (to ~S−2 or lower) during the XRD data collection (Grundmeier and Dau 2012; Glöckner et al. 2013), due to the X-ray exposure levels used there. However, Vinyard et  al. (2013) have argued in detail against this. Pace, Stranger and co-workers have shown computationally, that the LOS paradigm readily explains the WOC structures in both the 1.9 Å and 1.95 Å S1 forms, as simple proton shifted tautomers (Gatt et al. 2012; Petrie et al. 2015a). Recently, they have shown that the same oxidation state assumption quantitatively explains the WOC geometries in the S3 state (Petrie et al. 2017a, 2018). The Young et al. 2.25 Å structure represents a single S3 population, while the Suga et  al. 2.35  Å structure contains roughly equal populations of two tautomeric forms, related by a single proton shift involving O5 and W2 (on Mn4), very similar to that seen in the S1 structures. In one 2.35 Å tautomeric form, the O5 group (as OH−) occupies the position seen in both the 2.25 Å and 2.35 Å structures, while in the other tautomer, the ‘O5’ group (now as water) occupies the O6 position (see Petrie et  al. 2018, for details). The relevant XRD and computational WOC structures are compared in Fig. 3.1. Crucially, these computational models reproduce the ‘long’ near Mn–O5 distances, consistently seen in all the XRD structures reported to date (Table 3.2). In HOS computational modelling, the O5 group is almost invariably a deprotonated, generally oxo species, bridging a Mn(III, IV) pair in S1 and a Mn(IV, IV) pair in S3 (see section “IV.  Substrate exchange kinetics”). Then the Mn–O bond lengths are typically

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~1.8–1.9  Å, as routinely seen in relevant model compounds (Pantazis et  al. 2012; Petrie et  al. 2015a). Further, because the Mn3, 4 pair in high paradigm always has one of these oxidation state combinations in S1⋯3, the Mn–Mn distance is generally ~2.7  Å. This is seen in the model compounds and indeed for the Mn1, 2 and Mn2, 3 distances in the XRD structures, which are di-μ oxo bridged (Fig.  3.1, Table  3.2) in S1 and S3. Additionally, in the high oxidation paradigm, the W1, 2 groups are invariably hydroxyl or oxo groups, ligating Mn(IV), particularly by S3. Then computationally, as well as from model compounds, the Mn–O bond lengths are significantly less than 2.0 Å. In low oxidation paradigm modelling of the 1.95 Å S1 (Petrie et al. 2015a, b; Terrett et al. 2016) and very recently the 2.25 Å and 2.35 Å S3 WOC structures (Petrie et al. 2017a, 2018), W1, 2 and O5 are generally hydroxyl or waters. Mn4 is a Mn(II) in S1 or Mn(III) in S3. Then the Mn4–W1, 2 bond lengths are always in the 2.1–2.4 Å range, while the Mn3, 4 bond length is ~2.9 Å in S1, as generally observed. From the above discussions, if the precise geometric details revealed around Mn4 and O5 are taken at face value, then the low Mn oxidation paradigm is, at the very least, strongly favoured, if not totally prescribed. So the question becomes, ‘How reliable are the XRD structural features, at the ~0.1 Å and below level?’ Mn EXAFS, extensively applied to the photosystem by several groups over many years, provides a precise, independent means to determine this. EXAFS (e.g., see Sauer et  al. 2008) is a radial, X-ray stimulated electron back scatter interference technique, giving precise distance measures, at the molecular level, but less well resolved angular information. For the WOC, the method most applied has been Mn K edge EXAFS.  This reveals Mn–B (B = back-scatterer) distances (to ≈0.01 Å), where B may be a first or higher shell ligand atom (typically O, N, C, etc.), or nearby heavy (metallic) scatterer (Mn, Ca, etc.). For systems of any molecular complexity (cer-

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tainly the WOC), modelling the EXAFS is required to extract structural information. This is generally reported as a series of ‘shells’, each shell corresponding to an average Mn–B distance, atom type (first row, C, N, O, etc., second row metal, Mn, Fe, Ca, etc.), with an average number of such scatterers for each shell. The distances are very precise for the closer shells, but atom identification and average scatterer numbers in each shell are less so. However, when coupled with XRD data of even moderate resolution, EXAFS becomes extremely informative. Standard resolution Mn EXAFS (~12 Å−1) is available for all S states, as well as extended range EXAFS (~16–17  Å−1) for S1 (see Table 3.2). An important question is the reliability of the Mn–O distances reported from the XRD. These are broadly of three types, bridging μ–oxo distances, Mn–ligand (generally carboxylate O−) distances and the Mn–O5 distances. For the first two there are 9–10 such interactions each, amongst the 4 Mn in the WOC (~2.5 scatterers per Mn). These will dominate the closer EXAFS scattering peaks. It is unlikely the fewer, longer (2–3 total) Mn–O5 interactions could be reliably extracted. From Table 3.2, both EXAFS groups resolved two shells within the Mn–O(N) scattering range (~1.8–2.0  Å), each with an effective scatterer number of ~2.5–3 per Mn. We assume that the shorter EXAFS distances correspond to the μO–Mn distances in the XRD structures, the longer to the OLig–Mn distances. Then Table  3.2 shows that the corresponding μO–MnAV and OLig–MnAV values from the EXAFS and XRD data match, in both S1 and S3, to within ~0.1 Å in all cases. Further, the second shell Mn–Mn distance estimates, from the XRD and EXAFS measures, are in similarly good agreement in both S states. This is particularly true for the Mn3–Mn4 distance in S1, compared to the higher resolution, extended range EXAFS data, as previously noted (Petrie et al. 2015a). The EXAFS gives only semi-­quantitative agreement with the third

R. Stranger et al. shell, generally Mn–Ca interactions, as these are of lower intensity, near the edge of reliable discrimination for the technique (Sauer et  al. 2008). Although some S state structures as crystallised might be simple tautomeric variants of forms found in functional turnover (see section “VI.  Low paradigm functional S-states” below), the strong conclusion from the concordance of the EXAFS and XRD data for the closer atomic interactions, is that we can trust the XRD results. In particular, for the Mn–O5, Mn–W1, 2 and Mn3–Mn4 distances. Finally, at the time of submission of this chapter, a new high resolution (~2.1 Å) study of crystallised PS II appeared from the Berkeley group (Kern et  al. 2018). This included structures of the S1 and S2 state forms, with time resolved results for the structural evolution during the period from double flashing to stabilised S3 formation. As previously, states beyond S1 were generated by single turnover laser flash advancement. The OEC structures from these studies are currently under active computational examination by us and will be reported in due course. However, some preliminary observations may be suggested here, at proof stage. The latest OEC structures from the Berkeley group are very similar, overall, to those seen earlier (from themselves and Shen et  al.). They show similar cluster geometries and O5 group locations (with ‘long’ Mn–O distances, as above), from S1 to S3. Interestingly however, in the stabilised S3 form (0.2 s after double turnover), a new density feature (designated Ox) is seen, with partial apparent occupancy (~ 0.4) in the OEC cavity, near O5. This is likely an oxy species and the apparent Ox–O5 separation is 120 ≈120 37 ± 2

Ea (kJ·mol−1)c

– – PC>APC>Chl a, from higher to lower. In response to light qualities, some cyanobacteria can modify the composition of PBS, which is called as complementary chromatic acclimation. A cyanobacterium Fremyella diplosiphon has PBSs composed of PE, PC, and APC under green light, and possesses those containing only PC and APC under red light (Grossman et al. 1993; StoweEvans and Kehoe 2004). After a laser pulse excitation of PE in a cyanobacterium Synechocystis sp. ATCC 27170 and a red alga Porphyridium cruentum, the Chl a fluorescence becomes dominant after 435 and 294  ps, respectively, indicating a sequential energy transfer of PE→PC→APC→Chl a (Yamazaki et  al. 1984, 1988). PE emits fluorescence even in the later time region, indicating some energy remains in PE.  Therefore, dissipation of energy not transferring to PC might be a cru-

cial process under stress conditions (Liu et al. 2008; Yokono et al. 2012). APC is capable of binding to both PS I and PS II; therefore, energy distribution between PS I and PS II can be controlled by switching PBS→PS I and PBS→PS II energy transfers (McConnell et al. 2002; Mullineaux and Emlyn-Jones 2005). For the PBS→PS I energy transfer, two pathways are possible: the PBS→PS I direct energy transfer (Liu et  al. 2013) and PBS→PS II→PS I energy transfer (Ueno et  al. 2016). Quenching of APC by the orange Car protein is another mechanism to regulate PBS→PS transfers (Kirilovsky 2007; Kirilovsky and Kerfeld 2012). In addition to Chl a, LHCs contain other types of Chl molecules, which transfer the energy to Chl a (Fig.  10.4) (Rüdiger and Schoch 1988; Scheer 2003). Green algae and land plants have Chl b, whereas brown algae, diatoms, and dinoflagellates possess Chl c. The divinyl(DV)-type Chls are found in a cyanobacterium Prochlorococcus marinus (Chisholm et al. 1988; Mimuro et al. 2011). Energy-transfer processes to Chl a in LHCs

10  Modification of Energy Distribution Between Photosystems I and II…

Fig. 10.4.  Molecular structures of chlorophylls (Chls): Chl a, Chl b, two types of Chl c (found in brown algae and diatoms), divinyl (DV)-Chl a, and DV-Chl b

have been elucidated by the fluorescence upconversion method. Time constants of the Chl b→Chl a energy transfers were analyzed to be ~700  fs in LHCII from a land plant Arabidopsis thaliana (Akimoto et al. 2005), 500  fs for Chl b→Chl a energy transfer in LHCII from the PS I–PS II double-deficiency strain C2 of a green alga Chlamydomonas reinhardtii (Eads et  al. 1989), ~400  fs in LHCII from a green alga Codium fragile (Akimoto et al. 2007), whereas those of Chl

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c→Chl a energy transfers were obtained to be 500–700  fs (intracomplex) and 4–6  ps (intercomplex) in Fx–Chl a/c proteins of a diatom Chaetoceros gracilis (Akimoto et al. 2014a), and 350–400  fs and 1.8–2.6  ps in intrinsic Peri–Chl a/c proteins from a dinoflagellate Symbiodinium sp. (Tanaka et  al. 2016). In prochlorophyte Chl-binding proteins, the energy transfer from DV-Chl b to DV-Chl a occurs in the same time regions as the above Chl b→Chl a energy transfers in Arabidopsis, Chlamydomonas, and Codium LHCIIs (Hamada et al. 2015). Furthermore, LHCs have characteristic Cars, which work as energy donors for Chl a (Fig.  10.5) (Goodwin and Britton 1988; Scheer 2003; Siefermann-Harms 1985): lutein (Lut) in land plants, siphonaxanthin (Siph) or Lut in green algae, fucoxanthin (Fx) in brown algae and diatoms, and peridinin (Peri) in dinoflagellates. Time constants of the Car→Chl a energy transfers were also examined by the fluorescence upconversion method, which resulted in ~90 and ~ 152 fs for Car→Chl a energy transfer in the wild type and the npq2 mutant of Arabidopsis LHCIIs, respectively (Holt et al. 2003), 220 fs for Car→Chl a energy transfer in the spinach LHCII (Peterman et al. 1997), ~50 fs for Lut (S2)→Chl a energy transfer in the wild type Arabidopsis LHCII (Akimoto et  al. 2005), 400  fs for Car (S1)→Chl a energy transfer in the Chlamydomonas mutant C2 LHCII (Walla et al. 2000), ~400 fs for Siph→Chl a energy transfer in the Codium LHCII (Akimoto et al. 2007), 300 fs for Fx→Chl a energy transfer in the Fx–Chl a/c proteins of C. gracilis (Akimoto et  al. 2014a), and 400 fs and 1.5 ps for Peri→Chl a energy transfers in the intrinsic Peri–Chl a/c proteins and the water-soluble Peri–Chl a proteins of Symbiodinium sp., respectively (Tanaka et al. 2016). Irrespective of the variety of the pigments found in LHCs that work as energy donors for Chl a, (DV-)Chl a receives the excitation energy within a few picoseconds in LHCs, irrespective of the types of the pigments.

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Fig. 10.5.  Molecular structures of carotenoids: lutein, siphonaxanthin, fucoxanthin, and peridinin

It should be noted that (DV-)Chl a does not monopolize the excitation energy in LHC by the energy transfer, but the energy is thermally equilibrated among (DV-)Chl a and other pigments through the transfer. After the equilibration, the time-resolved fluorescence spectra do not change in shape, independent of the excitation wavelengths (Nakayama et al. 1994). In this case, all the pigments sharing the energy should exhibit fluorescence kinetics having the common time constant(s), even if the kinetics are analyzed by Eq. 10.1 (not the global analysis by Eq. 10.7). LHCI functions as a peripheral antenna of PS I. The rate of LHCI→PS I energy transfer depends on not only the amount of LHCIs binding to PS I but also the energy level of low-energy Chls (Croce and van Amerongen 2013). Conversely, LHCII can bind to both PS I and PS II (Wientjes et  al. 2013; Wlodarczyk et  al. 2016); therefore, energy distribution between PS I and PS II can be controlled by switching LHCII→PS I and LHCII→PS II energy transfers (Lemeille and Rochaix 2010; Minagawa 2011). Various

LHCII-binding sites are discovered in PS I (Yadav et  al. 2017; Yokono et  al. 2019b). Larger amount of bound LHCII achieves larger excitation energy income to PS I (Bos et al. 2017). Recent studies suggest that only 10–15% of LHCII physically moves between PS I and PS II in land plants (GoldschmidtClermont and Bassi 2015). Therefore, land plants must use another method to deal with the excess excitation energy. Quenching functions by the LHC-like proteins, PsbS or LhcSR, are additional processes to regulate LHC→PS energy transfers. The quenching triggered by PsbS controls LHCII→PS II energy transfers in A. thaliana (Ware et  al. 2015; Correa-Galvis et al. 2016), whereas that by LhcSR regulates not only LHCII→PS II energy transfers but also LHCII→PS I energy transfers in C. reinhardtii (Allorent et al. 2013; Tokutsu and Minagawa 2013; Kim et  al. 2017; Kosuge et  al. 2018; Girolomoni et  al. 2019). In the diatom C. gracilis, the energy transfer to PS I is modified by the pH conditions (Nagao et al. 2019b).

10  Modification of Energy Distribution Between Photosystems I and II… IV. Evolution of Spillover Mechanisms Spillover is an excitation energy transfer from PS II to PS I. The energy transfer does not occur between distant molecules (Eq.  10.3) (Förster 1965); therefore, spillover mechanisms require closely coupled complexes composed of both PS I and PS II.  The direct coupling of PS I and PS II enables fast energy transfer from PS II to PS I (Yokono and Akimoto 2018; Yokono et al. 2015), whereas the antenna-bridged type connection could slow down the energy transfer in the case that too much antennas are sandwiched between PS I and PS II.  If the energy-transfer rate becomes too low, spillover cannot be a prominent process, and antenna will supply excitation energy to each PS I and PS II independently. If spillover occurs efficiently, a reaction center that has an excess energy can divert the energy to another reaction center that is situated under shortage of energy. In addition, PS I reaction center can convert excess excitation energy into heat (Tiwari et  al. 2016); therefore, the reaction centers are

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protected from the excess energy. In case the light intensity becomes low, the connection between PS I and PS II may be energetically decoupled rapidly (Yokono et  al. 2015), then the excitation energy transferred from the antenna is efficiently used to perform photosynthesis. This method does not require migration of energy over long distances in the antenna proteins, so it is useful to adapt the quickly changing light conditions. Cyanobacteria are the oldest oxygenic photosynthetic organisms that have both PS I and PS II. Their large light-harvesting antennas, PBSs, are located on their stromal surfaces of the thylakoid membranes. Liu et al. first isolated PBS–PS I–PS II megacomplex, where PS I and PS II separately binds to PBS antenna (Fig. 10.6a) (Liu et al. 2013). This is an antenna-bridged type connection, and excitation energy transfer occurs efficiently from the antenna to both PS I and PS II. PS I binds PBS via PsaA subunit, and there seems to be vacant space between PsaA and PS II in their megacomplex model. In the isolated megacomplex, spillover was not observed. On the other hand, the occurrence of spill-

b

c

Fig. 10.6.  Models of various connections between PS I and PS II with antenna complexes in (a) cyanobacteria (Liu et al. 2013), (b) red algae (Liu et al. 2013;

Pi et  al. 2018; Antoshvili et  al. 2019), and (c) green lineage (Tikkanen et al. 2010; Yokono et al. 2015)

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over in cells of cyanobacteria under PBSselective excitation was demonstrated from the measurements of delayed fluorescence (Ueno et al. 2016). These results suggest that cyanobacteria have megacomplex(es) containing PBS, PS I, and PS II that are capable of spillover. The PBS→PS II→PS I energy transfer could convey light energy to a higher-energy Chl than the PBS→PS I direct energy transfer (Ueno et  al. 2017). Further studies are required to reveal the structure of intact PBS–PS I–PS II megacomplex(es) and the more detailed pathway(s) of spillover in cyanobacteria. Energy balance between PS I and PS II is controlled with the modification of the spillover in response of the cyanobacterial cells to surrounding light conditions: long-term adaptation of A. platensis to light qualities and quantities (Akimoto et al. 2013) and short-term adaptation of Synechocystis sp. PCC 6803 to strong white- or single-colored light-­emitting diodes (Akimoto et  al. 2014b). During the long-term adaptation of A. platensis, the PBS→PS I and PBS→PS II energy transfers are also regulated (Akimoto et  al. 2013). In three strains of the PBSlacking cyanobacterium P. marinus, only the antenna→PS energy-transfer processes are modified under different light qualities and quantities (Hamada et al. 2017). Red algae acquired PBS from cyanobacteria, and show prominent spillover (Kowalczyk et al. 2013). It was found that the spillover is dominant when the amount of chromophore bound to PBS is increased (Yokono et  al. 2011). Like cyanobacteria, red algae could also utilize both PBS→PS I direct energy transfer and PBS→PS II→PS I energy transfer (Ueno et al. 2017). In the unicellular red alga Cyanidioschyzon merolae, spillover level changes depending on light qualities; the level is reduced under light that excites PS I (red light), whereas the opposite response is induced under yellow light, which excites PBS selectively (Ueno et  al. 2015). The PBS→PS I direct energy transfer is enhanced under yellow light (Ueno et al. 2015). In addition to PBS,

M. Yokono et al. red algal PS I acquired membrane spanning type light-harvesting antenna, Lhcr (Wolfe et  al. 1994). Recently, the structure of red algal PS I-Lhcr supercomplexes was determined (Pi et  al. 2018; Antoshvili et  al. 2019), where PsaA is covered by Lhcr, which may bridge the vacant space between PsaA and PS II in the megacomplex (Fig.  10.6b), and probably facilitate efficient spillover. Unlike cyanobacteria, green algae utilize various types of membrane-spanning lightharvesting antenna complexes (Iwai and Yokono 2017). Under different light qualities, unicellular green algae mainly regulate the LHC→PS energy transfers but not spillover (Ueno et al. 2019a). CO2 concentrations also do not affect the spillover level (Ueno et al. 2018). The antennas surround both PS I and PS II (Tokutsu et al. 2012; Drop et al. 2014; Ozawa et  al. 2018; Qin et  al. 2019; Shen et al. 2019; Sheng et al. 2019; Su et al. 2019; Suga et  al. 2019), and excitation energy transfer occurs efficiently from antenna to both PS I and PS II.  If a lot of antennas are accumulated around the PS I and PS II, the efficiency of spillover may be decreased because excitation energy at PS II should migrate among a lot of antennas before it reaches to PS I. In this case, state transition (couple/decouple of antennas from each of PS I and PS II) may be an effective regulatory mechanism than spillover (Hodges and Barber 1983; Ueno et al. 2018). On the other hand, if the amount of antenna accumulation is decreased due to environmental conditions including some stresses, the spillover could be an effective regulatory mechanism. Recently, a megacomplex, which is approximately 720 kDa and is composed of both PS I and PS II, was isolated from a green macroalga under salt stress (Gao et al. 2019). The supercomplex includes PsbS and LhcSR, as well as xanthophyll cycle pigments; therefore, spillover combined with the quenching function by these quenchers may play an important role in photoprotection under stress conditions.

10  Modification of Energy Distribution Between Photosystems I and II… A. thaliana utilizes the direct connection between PS I and PS II to dissipate excess excitation energy in PS I–PS II megacomplexes (Fig.  10.6c) (Yokono et  al. 2015; Yokono and Akimoto 2018). Two sizes of PS I–PS II megacomplexes were isolated from A. thaliana, a smaller one (~700 kDa) and a larger one (>2400 kDa). The smaller megacomplex contains one PS I monomer and one PS II monomer. The larger megacomplex may have two PS I monomers and one PS II dimer with various light harvesting antennas (Yokono and Akimoto 2018; Yokono et  al. 2015, 2019b). Both PS I–PS II megacomplexes show fast (~20  ps) energy transfer from PS II to PS I (Yokono et  al. 2019b), suggesting a direct connection between PS I and PS II. The amount of PS I–PS II megacomplexes increases under high light conditions, suggesting photoprotective roles that the complexes may play. V. Benefits of the Direct-Type and Bridged-Type Spillovers In the wild type A. thaliana, fast excitation energy transfer from PS II to PS I is observed (Fig.  10.7, black solid arrows) (Yokono et  al. 2015), which suggests the

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direct-type connection between PS I and PS II.  On the other hand, two Lhca mutants, Lhca1 and Lhca2, that lack LHCI show slow energy transfer to PS I (Fig. 10.7, gray solid arrows) (Yokono et  al. 2015). LHCI acts as the binding site of LHCII to PS I (Yadav et  al. 2017; Yokono and Akimoto 2018) in addition to other binding-site subunits such as PsaL (Kouřil et  al. 2005). Lhcb5 mutant also shows the slow energy transfer to PS I. Lhcb5 acts as the binding site of LHCII to PS II. The Tap38 mutant lacks thylakoid-associated phosphatase 38 required for dephosphorylation of LHCII, and also shows the slow energy transfer to PS I. Phosphorylation is an important regulation process of LHCII binding to PS I and PS II (Wollman 2001). In addition, lil8-1 mutant (Psb33) also shows the slow energy transfer to PS I (Kato et al. 2017). Lil8 regulates LHCII binding to PS I and PS II. All these mutants show high spillover level similar to wild type. Therefore, the slow energy transfer to PS I may indicate slow spillover via the LHCII-bridge located between PS I and PS II. The time constant is longer than 250 ps, which is consistent with a previous assumption that energy transfer between LHCII trimers may take ~150  ps (Caffarri et  al. 2011). Perturbations to the

Fig. 10.7.  Fluorescence decay-associated spectra of wild type and five mutants of Arabidopsis leaves measured at 77 K. Times at the right side are the time constants analyzed by Eq. 10.7. Black solid arrows are fast energy transfer to PS I. Gray solid arrows are slow energy transfer to PS I. Dotted arrows are PS I fluorescence in the delayed florescence (~20 ns), which indicates the existence of spillover

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regulation of LHCII-binding may cause irregular LHCII-binding to PS I and PS II. Under such conditions, the binding site for PS I and PS II may be covered by LHCII, thus, the bridged-type connection may be formed in A. thaliana. Note that the energy-­ transfer rate from LHCII to PS I may vary depending on the strength of LHCI-binding to PS I (Akhtar et al. 2016). Therefore, the slow energy transfer of LHCII to PS I observed in some mutants could also reflect loose LHCII-binding to PS I. Interestingly, both PS I–PS II megacomplexes isolated from A. thaliana by largepore clear-native polyacrylamide gel electrophoresis does not contain PsbS (Yokono et al. 2015), which is in contrast to the green macroalga (Gao et  al. 2019). Instead, the A. thaliana megacomplexes may dissipate excess excitation energy by nonphotochemical quenching at PS I (called PS I-NPQ) (Yokono et al. 2019b). Two mechanisms have been proposed for the PS I-NPQ: zeaxanthin-dependent charge transfer quenching around low-energy Chls (Ballottari et  al. 2014), and energy transfer from the low-energy Chls to P700+ (Shubin et al. 1995; Schlodder et al. 2011). Here, the low-energy Chls in PS I and LHCI have singlet electronically excited states with lower transition energies than P700. In the former mechanism, quenching of Chl-­excited states occurs through the formation of zeaxanthin radical cations within LHCI, and the effi-

ciency of this reaction is enhanced by the tight interaction between the two Chl molecules exhibiting the low-­energy form by the exciton coupling. In the latter mechanism, the low-energy Chl accepts excitation energy from other Chls, and transfers the energy to P700+ that may rapidly convert the energy into heat (Fig. 10.8). Both mechanisms require the presence of low-energy Chls within or around PS I, which were acquired by some green algae during evolution to resist to high light (Kunugi et al. 2016). The low-energy Chls in land plants possess lower transition energy than those in most green algae (Yokono et al. 2019b). The lower transition energy of Chl enhances efficiency of the latter PS I-NPQ mechanism, because the lower transition energy (fluorescence band in longer wavelength region) increases the Förster overlap integral with P700+ (absorption maxima at ~800 nm) (see Eq. 10.3). Therefore, A. thaliana may be able to dissipate excess energy in the direct-­type PS I–PS II megacomplexes, even without PsbS.  We note that previous reports suggest that PsbS is included in the PS I–PS II megacomplexes isolated from a moss Physcomitrella patens and A. thaliana by native polyacrylamide gel electrophoresis (Suorsa et  al. 2015; Furukawa et  al. 2019). Further study is required to reveal the diversity of quenching mechanisms in the PS I– PS II megacomplexes in moss and land plants. LHCII PSI-LHCI

RedChl P700

+

PSI-LHCI

PSII

P700+ RedChl

LHCII

Fig. 10.8.  Possible quenching pathways in PS I-PS II megacomplex. Large circles and small circles are trimer and monomer of LHCII, respectively. Gray levels of these circles reflect binding possibility of these LHCII at each binding site. The model is based on the result of Yokono et al. (2019b). Gray arrows are possible energytransfer pathways. Excitation energy in LHCII and PS II may be transferred to P700 via low-­energy Chl (Red Chl)

10  Modification of Energy Distribution Between Photosystems I and II… It was reported that the PS I photoinhibition is induced by fluctuating light in Δflv1(A) and/or Δflv3(A) mutants of cyanobacteria Synechocystis sp. PCC 6803 and Anabaena sp. PCC 7120 (Allahverdiyeva et al. 2013), and in land plants A. thaliana (Kono et  al. 2014) and Oryza sativa (Yamori et al. 2016). Besides the dissipation of excess excitation energy in PS II under the high-light conditions, the spillover has an important role in the protection of PS I against fluctuating light. Under the fluctuating light conditions, the light alternates between low and high intensities, it was found that the ratio of the PS I–PS II megacomplexes to the total PS II decreases in a diatom Phaeodactylum tricornutum, resulting in the reduction of excitation energy in PS I (Tanabe et al. 2020). The fluctuating light-induced suppression of spillover is not observed in C. glacilis, suggesting that the protective mechanism against the fluctuating light depends on species. In the C. glacilis cells, energy-transfer processes within and from the Fx–Chl a/c proteins are modified (Tanabe et al. 2020). By the direct-type connection (Fig. 10.9a), ~60% of excitation energy is trapped by PS I, even when the PS II is in open state (Yokono et  al. 2015). This may be because the PS II→PS I energy transfer (~20  ps) (Yokono et al. 2019b) is faster than the energy transfer from the PS II core antennas to the PS II reaction center (~50  ps) (Raszewski and Renger 2008). When PS II becomes closed, all the energy captured in the PS I–PS II megacomplex is trapped by PS I (Yokono et  al. 2015). Therefore, irrespective of the light conditions, the direct-type connection reduces the PS II excitation, and instead induces the PS I excitation. In combination with PS I-NPQ mechanisms, the direct-type connection can safely dissipate excitation energy. On the other hand, the PS II→PS I energy transfer via the bridged-type connection (Fig.  10.9b) may take time longer than 250 ps, which is slower than trapping at PS I and PS II. Therefore, the excitation energy

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Fig. 10.9.  Various connections in photosystems. Under stress conditions, PS II accepts more energy than that the following linear electron transfer pathways can process. Then PS II becomes a closed state (~500  ps). The direct-type connection enables fast energy transfer to PS I (~20  ps), then linear electron transfer is suppressed, and the energy is dissipated at PS I efficiently ( ms) (Buser et al. 1990; Faller et al. 2001), providing a much higher quantum yield in the former pathway. This difference in the quantum yield between the YZ and YD reactions is essential for the high quantum yield of water oxidation (> 0.9) at the Mn4CaO5 cluster (Suzuki et al. 2012), which donates an elec-

tron to YZ•. At the Mn4CaO5 cluster, two water molecules are oxidized to one oxygen molecule and four protons through a cycle of five intermediates called S states (S0–S4) (Joliot et al. 1969; Kok et al. 1970), in which each Sn state (n  =  0–3) is advanced to the next Sn + 1 state by electron transfer to YZ•. In contrast to quick rereduction of YZ• by the Mn4CaO5 cluster at the rate of 30μs – 2 ms (Haumann et al. 2005; Shimizu et al. 2018), YD• is highly stable even at room temperature (Styring and Rutherford 1987; Vass and Styring 1991), and hence YD is usually in its oxidized form under steady-state light condi-

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tions, which is another important factor to increase the quantum yields of YZ oxidation and water oxidation. A key to understand the mechanism of YZ and YD reactions is the fact that upon oxidation the phenolic proton of a tyrosine side chain is concomitantly released (Fig.  12.1) (Diner and Britt 2005; Styring et al. 2012). In such proton-­coupled electron transfer, the kinetics of the redox reaction is often regulated by proton transfer (Rhile et al. 2006; Hammes-­Schiffer 2009). In this chapter, the molecular mechanism of asymmetric electron transfer between P680 and the redox active tyrosines, YZ and YD, is discussed mainly based on our recent studies using light-induced Fourier transform infrared (FTIR) difference spectroscopy in combination with quantum chemical calculations and site-directed mutagenesis. Factors to determine the asymmetric reactions are examined from both sides of P680+ as an electron acceptor and YZ and YD as electron donors. For P680+, the symmetry of a charge distribution on P680+ is considered, while for YZ and YD, the difference in the mechanism of the proton transfer reaction is focused. II. Asymmetric Charge Distribution on the Radical Cation of the Chlorophyll Dimer P680

T. Noguchi ized on one Chl, the 131-keto C=O frequency upshifts by ~30  cm−1 upon oxidation (Nabedryk et al. 1990; Kitajima and Noguchi 2006), whereas when the charge is evenly distributed over the two Chl molecules, the C=O frequencies of both Chls upshift by the half of this value (Noguchi et  al. 1998; Okubo et  al. 2007). Thus, the extent of the shifts of the keto C=O frequencies upon cation formation reveals the distribution of a positive charge on the Chl dimer (Ivancich et al. 1998; Okubo et al. 2007). In the case of P680, because the 131-keto C=O groups of PD1 and PD2 are both free from hydrogen bonding (Fig.  12.2b), their C=O vibrations appear at ~1700 cm−1, the typical frequency of a free 131-keto C=O group of Chl (Krawczyk 1989), and overlap wtih each other. In P680+ of PS II core and membrane preparations, two 131-keto C=O bands appeared at ~1725 and  ~  1710  cm−1 as a result of larger and smaller upshifts upon oxidation, respectively (Fig.  12.2a) (Okubo et al. 2007; Nagao et al. 2017). It was thus suggested that 70–80% of a positive charge is distributed on one Chl that shows the higher frequency band at ~1725 cm−1 (Okubo et al. 2007). This FTIR detection of a charge distribution, however, does not tell which Chl, PD1 or PD2, mainly possesses a positive charge.

A. FTIR Detection of a Charge Distribution on P680+

B. Genetic Introduction of a Hydrogen Bond to the 131-keto C=O Group of PD1 and PD2

A charge distribution on the radical cation of P680, which consists of PD1 and PD2 attached to the D1 and D2 proteins, respectively, has been investigated using light-­induced FTIR difference spectroscopy (Okubo et al. 2007; Nagao et  al. 2017). By detecting the C=O stretching vibrations of the 131-keto C=O groups of PD1 and PD2 in a P680+/P680 FTIR difference spectrum, a charge distribution over the two Chl molecules can be estimated. When a positive charge is completely local-

To identify the Chl molecule (PD1 or PD2) that mainly possesses a positive charge in P680+, it is necessary to assign the two 131-­keto C=O bands of P680+ in the P680+/P680 FTIR difference spectrum (Fig. 12.2a) specifically to either PD1 or PD2. To this end, we introduced a hydrogen bond to the 131-­keto C=O of PD1 and PD2 by changing a nearby Val residue, D1-V157 and D2-V156, respectively, to His using a cyanobacterium Synechocystis sp. PCC 6803 (Nagao et al. 2017). Successful

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Fig. 12.2. (a) Light-induced P680+/P680 FTIR difference spectrum in the 131-keto C=O stretching region for the PS II core complexes from Synechocystis sp. PCC6803. The Chl molecule that shows a larger upshift of the C=O frequency upon P680+ formation mainly possesses a positive charge in P680+. (b) Structure of P680, which consists of PD1 and PD2, and amino acid residues near the 131-keto C=O groups (D1-V157 and D2-V156), along with YZ and YD. The 131-keto C=O groups (circled by red dotted lines) of PD1 and PD2 are free from hydrogen bonding in WT. A Val residue near the 131-keto C=O group is replaced with a His residue to introduce a hydrogen bond to PD1 and PD2 in the D1-V157H and D2-V156H mutants, respectively

introduction of a hydrogen bond in the D1-V157H and D2-V156H mutants was monitored by detecting YZ•/YZ and YD•/YD FTIR difference spectra for isolated PS II core complexes. Because YZ and YD are located close to the 131-keto C=O groups of PD1 and PD2, respectively (Fig. 12.2b), it has been suggested that a prominent differential signal centered at ~1700 cm−1 in the YZ•/YZ and YD•/YD FTIR difference spectra arises from the 131-keto C=O vibration of PD1 and PD2, respectively (Hienerwadel et  al. 1996; Berthomieu et  al. 1998). Indeed, the 1707/1699  cm−1 signal in the YZ•/YZ spectrum showed a downshift by 4/2 cm−1 upon D1-V157H mutation but without any change upon D2-V156H mutation (Fig.  12.3a), whereas the 1702/1695  cm−1 signal in the YD•/YD spectrum largely downshifted by 18/19  cm−1 upon D2-V156H mutation but without any change upon D1-V157H mutation (Fig. 12.3b) (Nagao et al. 2017). These observations confirmed (1) the assignments of the prominent differential signals at ~1700 cm−1 in the YZ•/YZ and YD•/YD spectra to the 131-keto C=O vibrations of PD1 and PD2, respectively, revealing that these signals are useful markers for selective examination

of the interactions of the keto C=O of PD1 and PD2 in P680 without interference of P680+ bands, and (2) the successful introduction of a hydrogen bond to the 131-keto C=O of PD1 and PD2 in the D1-H157H and D2-V156H mutants, respectively. The question remained in the above detection of a hydrogen bond interaction is why the shift of the 131-keto C=O frequency is so different between PD1 and PD2 (by 2–4 cm−1 and 18–19  cm−1, respectively). To answer this question, quantum mechanics/molecular mechanics (QM/MM) calculations were applied to the models of P680  in the wild type (WT) and the D1-V157H and D2-V156H mutants (Fig. 12.4) (Nagao et al. 2017). In the mutant models, the Val side chain was replaced with a His side chain. In the WT model, the calculated 131-keto C=O frequency of PD2 (1692  cm−1) was slightly lower than that of PD1 (1703 cm−1), suggesting a slightly asymmetric dimeric geometry even in neutral P680 (Fig.  12.4a). In the D1-V157H mutant, an introduced His side chain could not form a proper hydrogen bond with the PD1 keto C=O due to a relatively narrow space, providing a relatively long hydrogen bond distance (H⋯O) of 2.61 Å and an

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T. Noguchi

Fig. 12.3.  Light-induced FTIR difference spectra of (a) YZ and (b) YD upon photooxidation in the PS II core complexes from the D1-V157H (a, red lines) and D2-V156H (b, blue lines) mutants of Synechocystis sp. PCC6803 in comparison with the spectra of WT (black lines). YZ•/YZ FTIR difference spectra were measured with Mn-depleted PS II core complexes in the presence of ferricyanide at 250 K, while YD•/YD spectra were measured in the presence of ferricyanide/ferrocyanide (1:1) at 283 K

improper angle (N-H⋯O) of 96.4° (Fig. 12.4b). Because of this weak hydrogen bond, the 131-keto C=O frequency showed only a small downshift of 4  cm−1 upon D1-V157H mutation (Fig.  12.4b), which is in good agreement with the experimental downshift of 2–4 cm−1 (Fig. 12.3a). In contrast, in the D2-V156H mutant, a His side chain forms relatively strong hydrogen bond with a 1.94 Å distance and a 124.8° angle, showing a large downshift of the PD2 keto C=O frequency by 17  cm−1 (Fig.  12.4c), which is also in good agreement with the experimental downshift by 18–19  cm−1 (Fig.  12.3b). Thus, the immediate environment of P680 is originally asymmetric, and

this is the reason for the significant difference in the 131-keto C=O shifts between the D1-V157H and D2-V156H mutants. It is notable that in calculation of each mutant, the 131-keto C=O frequency of the other Chl without a hydrogen bond was unchanged, confirming the independent keto C=O vibrations between PD1 and PD2. C. Identification of the Charge Localized Chlorophyll from the Assignments of the 131-keto C=O Bands of P680+

The 131-keto C=O bands of P680+ in the P680+/P680 FTIR difference spectrum were assigned using the D1-V157H and

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Fig. 12.4. (a) Optimized geometry of P680 and nearby amino acid residues in WT obtained by QM/MM calculations. Only the QM region is shown. (b, c) Expanded views of the interactions of the genetically introduced His side chain in the optimized geometry of P680 in the (b) D1-D157H and (c) D2-D156H mutants. Magnesium, green; carbon, gray; oxygen, red; nitrogen, blue; hydrogen, cyan. The calculated frequencies and the shifts upon mutation (in parenthesis) of the 131-keto C=O stretching vibrations of PD1 and PD2 are shown together with hydrogen bond distances (H⋯O) and angles (N-H⋯O)

D2-V156H mutants (Fig. 12.5). The positive band at 1708  cm−1 shifted down to ~1689  cm−1 by −19  cm−1 upon D2-V156H mutation, whereas this band was unchanged upon D1-V157H mutation. In contrast, the band at 1726  cm−1 was hardly affected by D2-V156H mutation, whereas this band decreased its intensity and a new broad feature appeared at ~1720 cm−1 upon D1-V157H mutation (Fig.  12.5). These observations provided an assignment of the band at 1726 cm−1 to PD1, which forms a weak hydrogen bond in D1-V157H, and the band at

1708  cm−1 to PD2, which forms a strong hydrogen bond in D2-V156H (Nagao et  al. 2017). The remaining intensity at 1726 cm−1 in the D1-V157H mutant was attributed to the destruction of hydrogen bonding in some centers due to the unstable hydrogen bond interaction of PD1 in P680+. The assignment of the higher-frequency band at 1726 cm−1 in P680+ to the PD1 keto C=O indicates that a positive charge is mainly localized on PD1. The charge localization on PD1 was also supported by thermoluminescence (TL) measurement of D1-V157H

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Fig. 12.5.  Light-induced P680+/P680 FTIR difference spectra of the PS II core complexes from the (a) D1-V157H (red line) and (b) D2-V156H (blue line) mutants of Synechocystis sp. PCC6803 in comparison with the spectrum of WT (black lines). Spectra under dark (1 s) and during illumination (1 s) were repetitively measured with Mn-depleted PS II core complexes in the presence of ferricyanide and SiMo at 250 K, and light-minus-dark difference spectra were calculated using average spectra. The shifts of the 131-keto C=O bands of P680+ upon mutation are shown by green arrows. Assignments of the two 131-­keto C=O bands of P680+ to either PD1 or PD2 are also indicated

and D2-V156H cells (Nagao et al. 2017). A TL glow curve by S2QA− recombination in the D1-V157H cells showed a large upshift of the peak temperature by 17  °C with an intensity increase by a factor of 2.0 in comparison with WT cells, whereas a small upshift by 3  °C with a slight intensity increase by a factor of 1.2 was observed in the D2-V156H cells. It was estimated from these data that the redox potential (Em) of P680 was upshifted by ~18 and ~ 5 mV by introduction of a hydrogen bond to PD1 and PD2 in the D1-V157H and D1-V156H mutants, respectively. The larger Em change by the D1-side mutation in spite of the for-

mation of a much weaker hydrogen bonding than the D2-side mutation can be explained by the major localization of a positive charge on PD1. The same conclusion of charge localization on PD1 was previously obtained by mutations of the His ligands of PD1 and PD2 (D1-H198 and D2-H197, respectively) in combination with detection of visible absorption changes in the Qy and Soret regions (Diner et  al. 2001). However, the excited states of PD1 and PD2 are significantly coupled with each other (Raszewski et  al. 2005; Raszewski et al. 2008; Renger and Schlodder 2011), and hence the effects of mutation on

12  Molecular Mechanism of Asymmetric Electron Transfer on the Electron… the electronic bands is not necessarily straightforward. In contrast, the 131-keto C=O vibrations of PD1 and PD2 are independent of each other and the effect of mutation on PD1 and PD2 can be selectively examined using the 131-keto C=O frequency, which is highly sensitive to a hydrogen bonding interaction. Thus, it can be definitely concluded from the FTIR analysis of the D1-V157H and D1-V156H mutants that a positive charge is mainly localized on PD1 in P680+. The major charge localization on PD1 has also been proposed by theoretical calculations based on the high-resolution X-ray crystallographic structure of PS II (Saito et al. 2011; Narzi et al. 2016). Several D1/D2 residue pairs, which involve a charged amino acid residue on one side, were suggested to be responsible for the asymmetric charge distribution on P680+ (Saito et al. 2011). III. Asymmetric Photoreactions of Redox-Active Tyrosines, YZ and YD A. Proton-Coupled Electron Transfer Reactions of YZ and YD

When a tyrosine residue is oxidized, its phenolic proton is always released because of the extremely low pKa value (~ − 2) of oxidized tyrosine (Dixon and Murphy 1976). Two redox-active tyrosines in PS II, YZ and YD, also release their protons upon oxidation (Fig. 12.1), and hence proton-coupled electron transfer takes place at these tyrosines (Diner and Britt 2005; Styring et al. 2012). Thus, to clarify the mechanism of the significantly different electron transfer rates of YZ and YD, it is crucial to examine the hydrogen bonding structures of the reduced and oxidized forms of YZ and YD and the proton release processes upon oxidation. According to the high-resolution X-ray structure of PS II (Umena et  al. 2011), YZ and YD appear to have similar molecular interactions with surrounding amino acid

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residues and water molecules (Fig.  12.6), although the detailed hydrogen bonding structures are unknown in the absence of the information of protons. The phenolic oxygen of YZ and YD is in hydrogen-­bonding distance from the imidazole nitrogen of a neighboring His, D1-H190 and D2-H189, respectively, and from a water molecule, which is linked to a hydrogen bond network in the PS II proteins. The information of the hydrogen bonding structures of YZ and YD is contained in the YZ•/YZ and YD•/YD FTIR difference spectra. In addition to the typical bands of Tyr vibrations such as the CO stretching band of an oxidized form (1514 and 1504 cm−1 for YZ• and YD•, respectively) and the CO stretching/ COH bending band of a reduced form (1256 and 1251 cm−1 for YZ and YD, respectively) (Hienerwadel et al. 1996; Berthomieu et al. 1998; Takahashi and Noguchi 2007), a broad positive feature was observed around 2800 cm−1 in the YZ•/YZ difference spectrum, whereas no such feature was observed in the YD•/YD spectrum (Fig.  12.7a) (Nakamura et al. 2014). A large downshift by ~700 cm−1 upon H/D exchange and a slight downshift by ~10 cm−1 upon 15N labeling of the PS II proteins suggested the assignment of this broad band to the NH vibration of a His side chain coupled with YZ. For further assignment, density functional theory (DFT) calculations were applied for the models of a Tyr-His dyad and a free His side chain (Fig.  12.7b) (Nakamura et  al. 2014). Only the Nτ-H vibration of a protonated cation of His hydrogen-­ bonded with a Tyr• radical explained the experimental band at ~2800  cm−1 with a strong intensity. Other models of neutral His hydrogen-bonded with Tyr or Tyr•, and free His in neural and cationic forms showed much higher NH frequencies at 3600–3150  cm−1. Moreover, QM/MM calculation for the YZ• site involving surrounding water molecules and amino acid residues also showed a similar result to the Tyr•-HisH+ model in the DFT calculation. This broad band at ~2800  cm−1 was thus

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Fig. 12.6.  Hydrogen bond networks of the (a) YZ and (b) YD sites deduced from the X-ray crystallographic structure of PS II (PDB: 3ARC)

Fig. 12.7. (a) Light-induced (a) YZ•/YZ and (b) YD•/YD FTIR difference spectra of the PS II core complexes from Thermosynechococcus elongatus. A positive broad feature specific to YZ• is colored magenta and indicated by a red arrow. (b) Assignment of the broad band around 2800 cm−1 (blue line) by DFT calculations for model Tyr compounds and QM/MM calculation for the YZ site involving nearby amino acid residues and water molecules (red bars)

assigned to the Nτ-H stretching vibration of the protonated cation form of D1-H190, which is hydrogen bonded with YZ•. The broad bandwidth was attributed to a large proton polarizability of this hydrogen bond, which was proposed to be involved in the proton transfer process during water oxidation (Nakamura et al. 2014).

These FTIR results strongly suggest that the phenolic proton of reduced YZ is hydrogen bonded to the Nτ of D1-H190 and is shifted to D1-H190 upon YZ oxidation to form a protonated HisH+ (Fig. 12.8a). Here, the C=O group of D1-N298 functions as a hydrogen bond acceptor of the Nπ-H of D1-H190. In contrast, on the D2 side, the

12  Molecular Mechanism of Asymmetric Electron Transfer on the Electron…

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Fig. 12.8.  Hydrogen bond structures of the (a) YZ and (b) YD sites and the changes upon oxidation. The hydrogen-bond acceptor and donor properties of the nitrogen sites of the coupled His are expressed by red and blue backgrounds, respectively. Protons released from YZ and YD upon oxidation are colored red

hydrogen bond partner at the Nπ of D2-H189, which is coupled with YD, is D2-R294 that functions as a hydrogen bond donor (Fig.  12.8b). Hence, Nπ of D2-H189 is always deprotonated and instead Nτ is protonated. Thus, YD is a hydrogen bond acceptor even in the reduced form, and its proton should be hydrogen bonded with a neighboring water molecule. Upon oxidation of YD, its proton has to be released to this water. In this scenario, the proton from YD is transferred away from the YD site through a hydrogen bond network (Fig.  12.8b), whereas the proton from YZ can only be shifted to the neighboring His through a strong hydrogen bond (Fig. 12.8a). This difference in the hydrogen bond pattern and the proton release process between YZ and YD

originates from the difference in the hydrogen bond partner of the coupled His residue, i.e., D1-N298 and D2-R294 for YZ and YD, respectively. B. FTIR Detection of Proton Release from YD to the Bulk

If the above scenario of the proton release processes of YZ and YD is the case, it is highly likely that a proton from YD is released out of the PS II proteins through a hydrogen bond network in the proteins. Indeed, Saito et al. (2013) theoretically predicted that the proton from YD is transferred up to D2-H61, the border of the QM region, though a hydrogen bond network following the movement of a hydronium ion from YD

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to D2-R180 (Fig.  12.6b). In contrast, the proton from YZ is not be released to the bulk if it is trapped at the neighboring His. We attempted to verify this prediction by monitoring protons released to the bulk upon YZ and YD oxidation using the FTIR method of proton detection (Nakamura and Noguchi 2015), which was previously developed to monitor the proton release from the Mn4CaO5 cluster during water oxidation (Suzuki et  al. 2009). In this method, virtually all protons released from PS II were trapped by high-concentration Mes buffer, and this protonation reaction of Mes (Fig.  12.9a) was monitored by isotope-­ edited Mes signals using deuterated Mes (Mes-d12; Fig. 12.9b) in light-induced FTIR difference spectra. From the Mes-minus-­ Mes-d12 double difference spectra, it was shown that 0.84 ± 0.10 protons were released into the bulk upon YD oxidation, whereas virtually no proton was released from PS II upon YZ oxidation (Fig.  12.9c) (Nakamura and Noguchi 2015). It was thus experimentally proved that the proton from YD is transferred to the bulk through a long proton pathway in the PS II proteins, whereas the proton from YZ is trapped in PS II most probably at D1-H190 (Fig. 12.8).

T. Noguchi IV. Mechanism of Asymmetric Electron Transfer from Tyrosines to P680+ The above FTIR data suggest that the asymmetric electron transfer from YZ and YD to P680+ is caused by both the factors of P680+ and YZ/YD (Fig. 12.10). As for P680+, a positive charge mostly resides on PD1, making an electron transfer distance from YZ to P680+ shorter than that from YD (the distance from the Mg center of PD1 to YZ and YD is ~8 and ~ 17 Å, respectively). The charge localization on one side in P680+ also increases the redox potential of P680 (Em  =  +1100– 1200 mV) (Rappaport et al. 2002; Kato et al. 2009) to the level that is capable of oxidizing YZ (Em = +900–1000 mV) (Metz et al. 1989; Vass and Styring 1991). Indeed, previous DFT calculation estimated the Em increase by ~140 mV upon complete charge localization (Takahashi et  al. 2008). Thus, charge localization on PD1 in P680+ promotes electron abstraction from YZ by two factors of the shortened electron transfer distance and the increased Em. In contrast to P680, the mechanism of electron transfer regulation of YZ and YD is mainly related to the proton transfer that

Fig. 12.9.  (a) Protonation and deprotonation reactions of a Mes buffer. (b) Chemical structure of deuterated Mes (Mes-d12). (c) Isotope-edited Mes bands (Mes-minus-Mes-d12) in the (a) 3000–2100 cm−1 and (b) 1350– 1130 cm−1 regions of the YZ•/YZ (blue lines) and YD•/YD (red lines) FTIR difference spectra together with those of the standard spectrum of proton release from the water oxidizing center (WOC) representing one proton per oneelectron transfer. YZ•/YZ and YD•/YD difference spectra were measured with Mn-depleted PS II core complexes from T. elongatus in high-concentration (~400 mM) Mes and Mes-d12 buffers (pH 6.5), and Mes-minus-Mes-d12 double difference spectra were calculated

12  Molecular Mechanism of Asymmetric Electron Transfer on the Electron…

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Fig. 12.10.  Factors to determine the asymmetric electron transfer from YZ and YD to P680+. A positive charge is mainly localized on PD1 in P680+, making an electron transfer distance shorter from YZ than from YD (green dotted lines). A proton is transferred to the neighboring D1-H190 upon YZ oxidation, whereas a proton is released to the bulk through a long hydrogen bond network in the PS II proteins upon YD oxidation, which imposes a high energy barrier

takes place concomitantly with electron transfer. A proton shift from YZ to the neighboring D1-H190 through a strong hydrogen bond is virtually a barrier-less reaction, whereas proton transfer through a long hydrogen-bond network in the PS II proteins from YD to the lumen should provide a high energy barrier in the photoreaction of YD (Fig. 12.10). The proton rocking between YZ and D1-H190 through a low barrier hydrogen bond is also advantageous to rapid rereduction of YZ• by electron transfer from the Mn4CaO5 cluster, which occurs at 30–50μs for the fastest S0→S1 transition (Haumann et al. 2005; Shimizu et al. 2018). In contrast, proton diffusion into the bulk upon YD oxidation contributes to the high stability of YD• by increasing entropy, resulting in the relatively low redox potential of YD (Em  =  +700– 800  mV) (Boussac and Etienne 1984; Vass and Styring 1991). Thus, rereduction of YD• is significantly slow, which also contributes to the high quantum yield of electron transfer from YZ to P680+ under steady-state light conditions.

These asymmetric properties of charge distribution of P680+ and proton release reactions in YZ and YD mainly originate from the asymmetric amino acid residue pairs of the D1 and D2 subunits. Theoretical calculation by Saito et al. (2011) showed that D1/ D2 residue pairs such as D1-N181/D2-R180, D1-N298/D2-R294, and D1-D61/D2-H61 contribute to the lower Em of PD1 than that of PD2, which induces the asymmetric charge distribution in P680+. These residue pairs are also responsible for the difference in proton release reactions in YZ and YD (Saito et al. 2011). The hydrogen-bond acceptor and donor properties of D1-N298 and D2-R294, respectively, determine the hydrogen bonding patterns in the YZ-(D1-H190)-(D1-N298) and YD-(D2-H189)-(D2-R294) triads, which cause a proton shift to D1-H190 from YZ and proton transfer to the bulk from YD upon their oxidation (Fig.  12.8). In addition, D2-R180 and D2-H61 form a hydrogen bond network as a proton pathway from YD (Fig. 12.6b) (Saito et al. 2013). In contrast, the corresponding residues, D1-N181 and

T. Noguchi

336

D1-D61, form a hydrogen bond network around a Cl ion (Fig. 12.6a), which probably functions as a proton pathway during water oxidation at the Mn4CaO5 cluster (Rivalta et al. 2011; Pokhrel et al. 2013; Suzuki et al. 2013). Thus, asymmetric amino acid distribution on the electron donor side of PS II realizes the efficient water oxidation by regulating asymmetric proton-coupled electron transfer between YZ/YD and P680. V.

Conclusions

Asymmetric electron transfer reactions from tyrosines, YZ and YD, to P680+ on the electron donor side of PS II, which is significant for the high quantum yield of water oxidation, is caused by asymmetric charge distribution on P680+ and different proton transfer processes at YZ and YD. A positive charge is localized mainly on PD1 in P680+ (Nagao et al. 2017), making the electron transfer distance from YZ much shorter than that from YD. Also, the proton from YZ is only shifted to the neighing His through a strong hydrogen bond in a barrierless way, whereas the proton from YD is released into the bulk by long-distance proton transfer in the PS II proteins imposing a high energy barrier in the proton-coupled electron transfer (Nakamura and Noguchi 2015). These asymmetric electron and proton transfer reactions on the donor side of PS II are realized by the asymmetric distribution of amino acid residue pairs between the D1 and D2 proteins. In particular, the different proton transfer reactions between YZ and YD originate from the difference in the hydrogen bonding pattern determined by the amino acid residues, D1-N298 and D2-R294, functioning as a hydrogen-bond acceptor and donor in the YZ-His-Asn and YD-His-Arg triads, respectively. Including this D1-N298/D2-R294 pair, several ionized/neutral amino acid residue pairs in the D1 and D2 proteins significantly contribute to the asymmetric charge distribution in P680+ (Saito et al. 2011).

In contrast to the main electron transfer pathway on the D1 side, which is directly involved in water oxidation as a primary function of PS II, electron donation from the D2 side may be related to photoprotection of PS II. It is known that YD• oxidizes the S0 state to the S1 state of the Mn4CaO5 cluster, while YD reduces the S2 and S3 states (Styring and Rutherford 1987; Messinger and Renger 1994). Although the physiological roles of these electron transfer reactions have not been well understood, slow oxidation of S0 to S1 by YD• could play a role in preventing overreduction of the Mn4CaO5 cluster that will lead to its impairment (Rutherford et  al. 2004). It has also been suggested that YD plays a photoprotection role during photoassembly of the Mn4CaO5 cluster (Magnuson et  al. 1999; Ananyev et al. 2002; Rutherford et al. 2004). In addition, the presence of Cytb559 only on the D2 side induces a secondary electron transfer pathway involving β-carotene (CarD2), monomeric Chl (ChlZD2), and Cytb559, which removes a positive charge on P680+ to the acceptor side to protect PS II against oxidative damage when the Mn4CaO5 cluster is inactivated (Tracewell et  al. 2001; Wang et al. 2002; Shinopoulos and Brudvig 2012). Thus, the asymmetric electron transfer on the electron-donor side of PS II is essential for both its productive and photoprotective functions. Acknowledgements The author thanks Drs. Ryo Nagao and Shin Nakamura for acquisition of the data involved in this review article. Quantum chemical calculations were performed at the Research Center for Computational Science, Okazaki, Japan, and Information Technology Center, Nagoya University. This study was supported by JSPS KAKENHI Grant Number JP17H06433, JP17H06435, and JP17H03662.

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Part IV Membrane Dynamics and Regulation of Excitation Energy/Electron Transfer Processes

Chapter 13 Structure-Function Relationships in Chloroplasts: EPR Study of Temperature-­ Dependent Regulation of Photosynthesis, an Overview Alexander N. Tikhonov*

Department of Biophysics, Faculty of Physics, M. V. Lomonosov Moscow State University, Moscow, Russia

Summary��������������������������������������������������������������������������������������������������������������������������������������� 343 I. Introduction�������������������������������������������������������������������������������������������������������������������������� 344 II. Electron and Proton Transport in Chloroplasts��������������������������������������������������������������������� 346 A. Structural and Functional Organization of Photosynthetic Electron Transport Chain������ 346 1. Photosystem I������������������������������������������������������������������������������������������������������������ 346 2. Photosystem II����������������������������������������������������������������������������������������������������������� 348 3. Cytochrome b6f Complex������������������������������������������������������������������������������������������� 348 4. Lateral Heterogeneity of Thylakoids, Linear and Cyclic Electron Transport��������������� 349 B. Rate-Limiting Steps in the Chain of the Intersystem Electron Transport������������������������� 351 C. Proton Pumping Across the Thylakoid Membrane and ATP Synthesis���������������������������� 354 III. Lipid-Soluble Nitroxide Radicals as Molecular Probes for Membrane Fluidity��������������������� 354 IV. Temperature-Dependent Regulation of Electron and Proton Transport and ATP Synthesis in Chloroplasts�������������������������������������������������������������������������������������� 358 A. Regulation of Electron Transport������������������������������������������������������������������������������������� 358 1. Photosystem II����������������������������������������������������������������������������������������������������������� 358 2. The Intersystem Electron Transport��������������������������������������������������������������������������� 360 B. Proton Transport, ATP Synthesis, and Carbon Fixation��������������������������������������������������� 362 1. Trans-thylakoid Transfer of Protons���������������������������������������������������������������������������� 362 2. ATP Synthesis and ATP Hydrolysis���������������������������������������������������������������������������� 363 3. Chloroplasts In Situ: Electron Transport and Carbon Fixation����������������������������������� 365 V. Discussion and Concluding Remarks���������������������������������������������������������������������������������� 366 Acknowledgements������������������������������������������������������������������������������������������������������������������������ 368 References������������������������������������������������������������������������������������������������������������������������������������ 368

Summary This chapter presents the retrospective overview of temperature-dependent regulation of photosynthetic electron transport processes in chloroplasts as studied by the electron paramagnetic resonance (EPR) method. The performance of photosynthetic electron transport *Author for correspondence, e-mail: [email protected] © Springer Nature Switzerland AG 2021 J.-R. Shen et al. (eds.), Photosynthesis: Molecular Approaches to Solar Energy Conversion, Advances in Photosynthesis and Respiration 47, https://doi.org/10.1007/978-3-030-67407-6_13

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chain was evaluated from the redox transients of P700, the primary electron donor in photosystem I. The spin labeling technique was applied to monitor structural changes in the lipid phase of thylakoid membranes. Summarizing the results of these studies, one can conclude that the physical state (fluidity) of the thylakoid membrane is one of the crucial factors that determine the regulation of photosynthetic performance of chloroplasts providing sustainable development of plants upon variations of ambient temperature.

I.

Introduction

Photosynthetic organisms of oxygenic type (cyanobacteria, algae, higher plants) assimilate carbon dioxide (CO2) and produce molecular oxygen (O2), using the solar energy absorbed by the light-harvesting complexes of photosystem I (PS I) and photosystem II (PS II). In photoreaction centers of PS I and PS II, the energy of light quanta is converted into the energy of separated charges (Nelson and Yocum 2006; Shelaev et  al. 2010; Blankenship 2002; Cardona et  al. 2012; Müh et  al. 2012; Ruban 2012; Mamedov et  al. 2015). The light-driven actuation of PS I and PS II reaction centers initiate electron transfer along the photosynthetic electron transport chain (ETC). Two electrons, extracted from the water molecule in the water-oxidizing complex (WOC) of PS II (Chap. 1), are delivered to the terminal electron acceptor of PS I, nicotinamide adenine dinucleotide phosphate (NADP+). PS I sequentially donates two electrons to NADP+ Abbreviations: CBC cycle  – Calvin-Benson cycle; CEF  – Cyclic electron flow around photosystem I; Chl – Chlorophyll; DGDG – Digalactosyldiacylglycerol; EPR – Electron paramagnetic resonance; ETC – Electron transport chain; Fd – Ferredoxin; FNR – Ferredoxin:NADP oxidoreductase; ISP  – Iron-sulfur protein; MGDG  – Monogalactosyldiacylglycerol; NADP+  – Nicotinamide adenine dinucleotide phosphate; P680  – Special chlorophyll pair in PS II; P700 – Special chlorophyll pair in PS I; Pc – Plastocyanin; Pheo – Pheophytin; PQ and PQH2 – Plastoquinone and plastoquinol, respectively; PSA  – Photosynthetic apparatus; PS I and PS II – Photosystem I and photosystem II, respectively; SASL – Spin-labeled derivatives of stearic acid; WOC  – Water-oxidizing complex

through ferredoxin (Fd) and ferredoxinNADP-oxidoreductase (FNR): PS I → 2Fd→FNR→NADP+ (Chap. 7). PS II and PS I are interconnected via the membranebound cytochrome (Cyt) b6f complex and mobile electron carriers, plastoquinone (PQ) and plastocyanin (Pc): PS II→PQ→Cyt b6f→Pc→PS I (Siggel 1976; Sigfridsson 1998). The Cyt b6f complex plays a crucial role in operation of photosynthetic ETC (Kurisu et  al. 2003; Stroebel et  al. 2003; Hasan et al. 2013a; Hasan and Cramer 2014; Kono and Terashima 2014; Tikhonov 2014, 2018). Standing at the crossroad of the linear and cyclic electron transport pathways, the Cyt b6f complex provides the oxidation of plastoquinol (PQH2, the fully reduced form of plastoquinone) and reduction of Pc. PQH2 oxidation by the Cyt b6f complex represents the “bottle-­neck” chain in the chloroplast ETC, which virtually determines the overall rate of the intersystem electron transport. The light-induced electron transfer is accompanied by generation of the trans-­ thylakoid difference in electrochemical Ü potentialsÜ of hydrogen ions,  H . The build up of  H occurs due to the light-­induced uptake of protons from the bulk of stroma (the space between the chloroplast envelope and thylakoids) and the proton released into the lumenÜ (the internal volume of thylakoids).  H serves as the driving force for operation of the ATP synthase complex CF0– CF1 (ADP  +  Pi→ATP  +  H2O) (Mitchell 1966; Boyer 1997; Nickolls and Ferguson 2002; Romanovsky and Tikhonov 2010; 





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The main purpose of this chapter is to Junge and Nelson 2015). ATP and NADPH, the main products of the light-­induced stages present a retrospective overview of function relationships in thylaof photosynthesis, are used mainly in bio- structure-­ synthetic reactions of the Calvin-Benson koids in the context of temperature-­ cycle (CBC) (Edwards and Walker 1983; dependent regulation of photosynthetic processes in chloroplasts. Thylakoid memBlankenship 2002). Photosynthetic apparatus (PSA) is sensi- branes consist of the lipid matrix impregtive to changes in plant environments, nated with plentiful electron transport including fluctuations in intensity and spec- complexes. They also perform the role of maintaining the tral composition of the photosynthetically the hydrophobic barrier Ü active radiation, composition of the atmo- required level of  H . In order to analyze spheric gas, nutrition of plants, and temper- the role of thylakoid membranes in thermal ature. The flexibility of PSA provides its regulation of photosynthesis, we consider optimal functioning under variable environ- here the relationships between the funcmental conditions. This is achieved by coop- tional properties of chloroplasts (photosyneration of several feedbacks that regulate the thetic electron and proton transport and photosynthetic processes. There are the operation of the ATP synthase), on the one short-term (seconds-minutes) and long-term hand, and the physical state of the mem(hours-days) mechanisms of PSA response brane lipids, on the other hand. For illustrato varying environmental conditions tion of the structure-function relationships (Kramer et  al. 2004; Murata et  al. 2007; in the thylakoid membranes, the temperaEberhard et al. 2008; Demmig-Adams et al. ture dependences of photosynthetic pro2012; Horton 2012; Rochaix 2014; Kono cesses and membrane fluidity have been and Terashima 2014; Puthiyaveetil et  al. overviewed. In previous works of our group, 2016; Niu and Xiang 2018). At the level of we used the electron paramagnetic resothylakoid membranes, the short-term regu- nance (EPR) method, which provided some latory mechanisms include: (i) ion-­methodological advantages for scrutinizing dependent control of the electron transport, structure-function relationships in thyla(ii) optimization of the light quanta parti- koids. This method allows an experimenter tioning between PS I and PS II, (iii) redistri- to monitor electron transport and probe the bution of electron fluxes between alternative membrane physical state under close experpathways, and (iv) PSA responses to varia- imental conditions. EPR signals of electron tions of temperature. The significance of carriers give information about operation of temperature-dependent regulation of photo- the chloroplasts ETC (Webber and Lubitz synthetic performance in plants is deter- 2001; Möbius and Savitsky 2009; Tikhonov mined by the fact that plants are 2015). Using lipid-soluble paramagnetic poikilothermic organisms, in which their probes, the EPR spectra of which are sensiown temperature varies with the environ- tive to their surroundings, one can detect mental temperatures. Thermo-­induced thermo-­ induced structural changes in the structural changes in the lipid domains of thylakoid membranes (Berliner 1976). thylakoid membrane are considered as one This chapter begins with a brief descripof the major factors that determine the chlo- tion of structural and functional organization roplast response to fluctuations of tempera- of plant chloroplasts, illustrated by some picture (Barber et  al. 1984; Los and Murata torial examples, followed by the analysis of 2004; Los et  al. 2013; Yamamoto 2016; the nature of the rate-limiting step in operaMaksimov et al. 2017). tion of ETC. The performance of the chloro

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koid lumen) (or cytochrome c6 in the case of cyanobacteria) to Fd molecules located in the stroma (Brettel 1997; Fromme et  al. 2001; Nelson and Yocum 2006, Mamedov et al. 2015). The light-induced excitation of P700 induces charge separation: the excited ∗ center P700 donates an electron to the primary electron acceptor (Chl2A or Chl2B). Electron carriers are arranged as two quasi-­ symmetrical branches. Two chlorophyll (Chl) molecules (Chl2A and Chl3A) and one phylloquinone molecule (A1A) belong to the A-branch; two other Chl molecules (Chl2B and Chl3B) and another phylloquinone molecule (A1B) belong to the B-branch. Branches A and B converge at the acceptor FX (the [FeS]4 cluster). Reduced FX donates an elecII. Electron and Proton Transport tron to ferredoxin (Fd) via the iron-­sulfur in Chloroplasts redox centers FA and FB bound to PS I (FX→FA→FB→Fd). Electron transfer on the A. Structural and Functional Organization of Photosynthetic acceptor side of PS I occurs without the Electron Transport Chain long-range spatial movements of electron carriers (DeVault 1980; Moser et  al. 1992; Figure 13.1a shows cartoons of the cross-­ Page et al. 1999; Möbius and Savitsky 2009). section of a chloroplast, in which two kinds Fd molecules located in stroma donate elecof thylakoids (closed vesicles) are depicted: trons to FNR, providing the reduction of (i) the piles of appressed granal thylakoids NADP+ to NADPH (NADP+  +  2e−  +  H+ and (ii) stroma-exposed inter-granal thyla- → NADPH). + koids. The multisubunit electron-transport Oxidized center of PS I, P700 , accepts an − protein complexes (PS I, PS II, and Cyt b6f) electron from Pc reduced by the Cyt b6f and the ATP synthase (CF0–CF1) complexes complex. Diffusing within the thylakoid are embedded into the lamellar membranes lumen, Pc− connects spatially separated of thylakoids. The arrangement of electron complexes Cyt b6f and PS I.  Under certain transport complexes and mobile carriers in conditions (i.e., in dark-adapted chlorothylakoids is shown schematically in plasts), steric constraints may restrict the Fig. 13.1b, where different routes of electron long-range lateral diffusion of Pc− from the and proton transfer are indicated by blue and granal to stromal domains of the narrow red arrows, respectively. The structural and (~4–5 nm) thylakoid lumen (Kirchhoff et al. functional peculiarities of PS I, PS II, and 2011). A width of the lumen can increase Cyt b6f complexes, which are involved in lin- upon illumination of chloroplasts; therefore, ear and cyclic electron transport pathways, the lateral Pc− diffusion would not restrict the are briefly outlined below. intersystem electron transport. In this case, diffusion of Pc− and its oxidation by PS I 1. Photosystem I occur more rapidly (t1/2